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>

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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 border="0" cellpadding="0" cellspacing="0" width="1505">
	<tbody>
		<tr height="43">
			<td dir="LTR" height="43" style="height:43px;width:335px;">Equipment</td>
			<td dir="LTR" style="width:208px;">Manufacturer</td>
			<td dir="LTR" style="width:311px;">Serial/Catalog #</td>
			<td dir="LTR" style="width:363px;">Notes</td>
		</tr>
		<tr height="56">
			<td dir="LTR" height="94" rowspan="2" style="height:94px;width:335px;">Pressure Transducer</td>
			<td dir="LTR" rowspan="2" style="width:208px;">Becton Dickinson</td>
			<td dir="LTR" rowspan="2" style="width:311px;">P23XL-1</td>
			<td dir="LTR" style="width:291px;">1+ (1 for each artery) 
			&#160;Used with water-filled diaphragm domes</td>
		</tr>
		<tr height="172">
			<td dir="LTR" height="172" style="height:172px;width:335px;">Amplifier and transducer box</td>
			<td dir="LTR" style="width:208px;">Gould</td>
			<td dir="LTR" style="width:311px;">5900 Signal Conditioner Cage</td>
			<td dir="LTR" style="width:291px;">1 
			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 height="121">
			<td dir="LTR" height="121" style="height:121px;width:335px;">T403 Console with TS420 perivascular flowmeter module(x2)&#160;</td>
			<td dir="LTR" style="width:208px;">Transonic Systems</td>
			<td dir="LTR" style="width:311px;">T403 module and TS420 (x2)</td>
			<td dir="LTR" style="width:291px;">1 
			Flow probes measuring flow through each of the carotid arteries will connect to each of the TS420 units.&#160;&#160;</td>
		</tr>
		<tr height="126">
			<td dir="LTR" height="126" style="height:126px;width:335px;">Digital ultrasonic measurement unit</td>
			<td dir="LTR" style="width:208px;">Sonometrics</td>
			<td dir="LTR" style="width:311px;">TR-USB</td>
			<td dir="LTR" style="width:291px;">1</td>
		</tr>
		<tr height="48">
			<td dir="LTR" height="48" style="height:48px;width:335px;">Flow Probe Precision S-Series 4mm</td>
			<td dir="LTR" rowspan="4" style="width:208px;">Transonic Systems Inc.</td>
			<td dir="LTR" rowspan="4" style="width:311px;">MC4PSS-LS-WC100-CM4B-GA</td>
			<td dir="LTR" rowspan="4" style="width:291px;">2</td>
			<td dir="LTR" rowspan="4" style="width:363px;"></td>
		</tr>
		<tr height="24">
			<td dir="LTR" height="49" rowspan="2" style="height:49px;width:335px;">1mm&#160; Sonometrics Crystals</td>
			<td dir="LTR" rowspan="2" style="width:208px;">Sonometrics Systems</td>
			<td dir="LTR" rowspan="2" style="width:311px;">1R-38S-20-NC-SH</td>
			<td dir="LTR" style="width:291px;">&#160;2-4 (2 for each artery)</td>
		</tr>
		<tr height="81">
			<td dir="LTR" height="134" rowspan="2" style="height:134px;width:335px;">Catheter for implantation</td>
			<td dir="LTR" style="width:208px;">BD (Becton Dickinson)</td>
			<td dir="LTR" rowspan="2" style="width:311px;">&#160;381447</td>
			<td dir="LTR" style="width:291px;">&#160;1+ (1 for each artery) 
			Catheter is cut and secured to microbore tubing, stylette is utilized for insertion&#160;</td>
		</tr>
		<tr height="24">
			<td dir="LTR" height="73" rowspan="2" style="height:73px;width:335px;">Tygon Microbore Tubing</td>
			<td dir="LTR" rowspan="2" style="width:208px;">Norton Performance Plastics</td>
			<td dir="LTR" style="width:311px;">(AAQ04127) Formulation S-54-HL</td>
			<td dir="LTR" style="width:291px;">NA (cut to length for an extension set)</td>
		</tr>
		<tr height="24">
			<td dir="LTR" height="49" rowspan="2" style="height:49px;width:335px;">Luer Stub Adapter</td>
			<td dir="LTR" style="width:208px;">BD (Becton Dickinson)</td>
			<td dir="LTR" style="width:311px;">427564 (20 gauge)</td>
			<td dir="LTR" style="width:291px;">1+ (1 for each arterial catheter)</td>
		</tr>
		<tr height="24">
			<td dir="LTR" height="82" rowspan="2" style="height:82px;width:335px;">Surflo Injection Plug</td>
			<td dir="LTR" rowspan="2" style="width:208px;">Terumo</td>
			<td dir="LTR" rowspan="2" style="width:311px;">SR-IP2</td>
			<td dir="LTR" style="width:291px;">1+ (1 for each arterial catheter)</td>
		</tr>
		<tr height="39">
			<td dir="LTR" height="128" rowspan="2" style="height:128px;width:335px;">Meadox</td>
			<td dir="LTR" rowspan="2" style="width:208px;">PTFE (Teflon) Felt</td>
			<td dir="LTR" rowspan="2" style="width:311px;">19306</td>
			<td dir="LTR" style="width:291px;">NA (cut to size) 
			The PTFE felt used in our studies was discontinued. However, comparable companies such as &#8220;Surgical Mesh&#8221; offer products which are equivalent.&#160;</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...

Watch this Scientific Journal Video about Surgical Technique for the Implantation of Tissue Engineered Vascular Grafts and Subsequent In Vivo Monitoring at JoVE.com

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

11.6K vistas.  State University of New York Buffalo School of Medicine.  Un protocolo de paso a paso para la colocación inter-posicional de ingeniería tisular buques (TEV) en la arteria carótida de una oveja mediante anastomosis de extremo a extremo y la evaluación digital en tiempo real en vivo hasta el sacrificio de animales. 

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

Bioluminescence Imaging for Assessment of Immune Responses Following Implantation of Engineered Heart Tissue (EHT)

Various techniques of cardiac tissue engineering have been pursued in the past decades including scaffolding strategies using either native or bioartificial scaffold materials, entrapment of cardiac myocytes in hydrogels such as fibrin or collagen and stacking of myocyte monolayers 1. These concepts aim at restoration of compromised cardiac function (e.g. after myocardial infarction) or as experimental models (e.g. predictive toxicology and substance screening or disease modelling). Precise monitoring of cell survival after implantation of engineered heart tissue (EHT) has now become possible using in-vivo bioluminescence imaging (BLI) techniques 2. Here we describe the generation of fibrin-based EHT from a transgenic rat strain with ubiquitous expression of firefly luciferase (ROSA/luciferase-LEW Tg; 3). Implantation is performed into the greater omentum of different rat strains to assess immune responses of the recipient organism following EHT implantation. Comparison of results generated by BLI and the Enzyme Linked Immuno Spot Technique (ELISPOT) confirm the usability of BLI for the assessment of immune responses.

Bioluminescence Imaging for Assessment of Immune Responses Following Implantation of Engineered Heart Tissue EHT

This video demonstrates the use of in vivo bioluminescence imaging to study immune responses after implantation of Engineered Heart Tissue (EHT) in rats.

Various techniques of cardiac tissue engineering have been pursued in the past decades including scaffolding strategies using either native or ...

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 ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Production, Characterization and Potential Uses of a 3D Tissue-engineered Human Esophageal Mucosal Model

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore esophageal adenocarcinoma generally has a poor prognosis, with little improvement in survival rates in recent years. These are difficult conditions to study and there has been a lack of suitable experimental platforms to investigate disorders of the esophageal mucosa.
A model of the human esophageal mucosa has been developed in the MacNeil laboratory which, unlike conventional 2D cell culture systems, recapitulates the cell-cell and cell-matrix interactions present in vivo and produces a mature, stratified epithelium similar to that of the normal human esophagus. Briefly, the model utilizes non-transformed normal primary human esophageal fibroblasts and epithelial cells grown within a porcine-derived acellular esophageal scaffold. Immunohistochemical characterization of this model by CK4, CK14, Ki67 and involucrin staining demonstrates appropriate recapitulation of the histology of the normal human esophageal mucosa.
This model provides a robust, biologically relevant experimental model of the human esophageal mucosa. It can easily be manipulated to investigate a number of research questions including the effectiveness of pharmacological agents and the impact of exposure to environmental factors such as alcohol, toxins, high temperature or gastro-esophageal refluxate components. The model also facilitates extended culture periods not achievable with conventional 2D cell culture, enabling, inter alia, the study of the impact of repeated exposure of a mature epithelium to the agent of interest for up to 20 days. Furthermore, a variety of cell lines, such as those derived from esophageal tumors or Barrett&#8217;s Metaplasia, can be incorporated into the model to investigate processes such as tumor invasion and drug responsiveness in a more biologically relevant environment.

This manuscript describes the production, characterization and potential uses of a tissue engineered 3D esophageal construct prepared from normal primary human esophageal fibroblast and squamous epithelial cells seeded within a de-cellularized porcine scaffold. The results demonstrate the formation of a mature stratified epithelium similar to the normal human esophagus.

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore ...

Athymic Rat Model for Evaluation of Engineered Anterior Cruciate Ligament Grafts

Anterior cruciate ligament (ACL) rupture is a common ligamentous injury that often requires surgery because the ACL does not heal well without intervention. Current treatment strategies include ligament reconstruction with either autograft or allograft, which each have their associated limitations. Thus, there is interest in designing a tissue-engineered graft for use in ACL reconstruction. We describe the fabrication of an electrospun polymer graft for use in ACL tissue engineering. This polycaprolactone graft is biocompatible, biodegradable, porous, and is comprised of aligned fibers. Because an animal model is necessary to evaluate such a graft, this paper describes an intra-articular athymic rat model of ACL reconstruction that can be used to evaluate engineered grafts, including those seeded with xenogeneic cells. Representative histology and biomechanical testing results at 16 weeks postoperatively are presented, with grafts tested immediately post-implantation and contralateral native ACLs serving as controls. The present study provides a reproducible animal model with which to evaluate tissue engineered ACL grafts, and demonstrates the potential of a regenerative medicine approach to treatment of ACL rupture.

Animal models are important tools for the evaluation of tissue-engineered grafts. This paper presents the protocol for preparing an electrospun biodegradable polymer graft for use in anterior cruciate ligament tissue engineering, as well as a surgical protocol for implantation in a rat model.

Anterior cruciate ligament (ACL) rupture is a common ligamentous injury that often requires surgery because the ACL does not heal well without ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from traumatic injury, disease and various congenital, genetic and acquired conditions are quite common. Tissue engineering and regenerative medicine technologies have enormous potential to provide a therapeutic solution. However, utilization of biologically relevant animal models in combination with longitudinal assessments of pertinent functional measures are critical to the development of improved regenerative therapeutics for treatment of VML-like injuries. In that regard, a commercial muscle lever system can be used to measure length, tension, force and velocity parameters in skeletal muscle. We used this system, in conjunction with a high power, bi-phase stimulator, to measure in vivo force production in response to activation of the anterior crural compartment of the rat hindlimb. We have previously used this equipment to assess the functional impact of VML injury on the tibialis anterior (TA) muscle, as well as the extent of functional recovery following treatment of the injured TA muscle with our tissue engineered muscle repair (TEMR) technology. For such studies, the left foot of an anaesthetized rat is securely anchored to a footplate linked to a servomotor, and the common peroneal nerve is stimulated by two percutaneous needle electrodes to elicit muscle contraction and dorsiflexion of the foot. The peroneal nerve stimulation-induced muscle contraction is measured over a range of stimulation frequencies (1-200 Hz), to ensure an eventual plateau in force production that allows for an accurate determination of peak tetanic force. In addition to evaluation of the extent of VML injury as well as the degree of functional recovery following treatment, this methodology can be easily applied to study diverse aspects of muscle physiology and pathophysiology. Such an approach should assist with the more rational development of improved therapeutics for muscle repair and regeneration.

Applications of In Vivo Functional Testing of the Rat Tibialis Anterior for Evaluating Tissue Engineered Skeletal Muscle Repair

We describe an in vivo protocol to measure dorsiflexion of the foot following stimulation of the peroneal nerve and contraction of the anterior crural compartment of the rat hindlimb. Such measurements are an indispensable translational tool for evaluating skeletal muscle pathology and tissue engineering approaches to muscle repair and regeneration.

Despite the regenerative capacity of skeletal muscle, permanent functional and/or cosmetic deficits (e.g., volumetric muscle loss (VML) resulting from ...

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, ...

A Novel Tenorrhaphy Suture Technique with Tissue Engineered Collagen Graft to Repair Large Tendon Defects

Surgical management of large tendon defects with tendon grafts is challenging, as there are a finite number of sites where donors can be readily identified and used. Currently, this gap is filled with tendon auto-, allo-, xeno-, or artificial grafts, but clinical methods to secure them are not necessarily translatable to animals because of the scale. In order to evaluate new biomaterials or study a tendon graft made up of collagen type 1, we have developed a modified suture technique to help maintain the engineered tendon in alignment with the tendon ends. Mechanical properties of these grafts are inferior to the native tendon. To incorporate engineered tendon into clinically relevant models of loaded repair, a strategy was adopted to offload the tissue engineered tendon graft and allow for the maturation and integration of the engineered tendon in vivo until a mechanically sound neo-tendon was formed. We describe this technique using incorporation of the collagen type 1 tissue engineered tendon construct.

In this paper, we present an in vitro and in situ protocol to repair a tendon gap of up to 1.5 cm by filling it with engineered collagen graft. This was performed by developing a modified suture technique to take the mechanical load until the graft matures into the host tissue.

Surgical management of large tendon defects with tendon grafts is challenging, as there are a finite number of sites where donors can be readily ...

A Novel Surgical Technique As a Foundation for In Vivo Partial Liver Engineering in Rat

Organ engineering is a novel strategy to generate liver organ substitutes that can potentially be used in transplantation. Recently, in vivo liver engineering, including in vivo organ decellularization followed by repopulation, has emerged as a promising approach over ex vivo liver engineering. However, postoperative survival was not achieved. The aim of this study is to develop a novel surgical technique of in vivo selective liver lobe perfusion in rats as a prerequisite for in vivo liver engineering. We generate a circuit bypass only through the left lateral lobe. Then, the left lateral lobe is perfused with heparinized saline. The experiment is performed with 4 groups (n = 3 rats per group) based on different perfusion times of 20 min, 2 h, 3 h, and 4 h. Survival, as well as the macroscopically visible change of color and the histologically determined absence of blood cells in the portal triad and the sinusoids, is taken as an indicator for a successful model establishment. After selective perfusion of the left lateral lobe, we observe that the left lateral lobe, indeed, turned from red to faint yellow. In a histological assessment, no blood cells are visible in the branch of the portal vein, the central vein, and the sinusoids. The left lateral lobe turns red after reopening the blocked vessels. 12/12 rats survived the procedure for more than one week. We are the first to report a surgical model for in vivo single liver lobe perfusion with a long survival period of more than one week. In contrast to the previously published report, the most important advantage of the technique presented here is that perfusion of 70% of the liver is maintained throughout the whole procedure. The establishment of this technique provides a foundation for in vivo partial liver engineering in rats, including decellularization and recellularization.

A Novel Surgical Technique As a Foundation for In Vivo Partial Liver Engineering in Rat

We establish a novel surgical technique for an in vivo single liver lobe perfusion model in rat as a prerequisite for further studying in vivo partial liver engineering in the future.