Name: engineered vascularized muscle flap
Uploaded: 2016-01-11T09:30:00.000+00:00
Duration: 8 min 18 s
Description: Watch this Scientific Journal Video about Engineered Vascularized Muscle Flap at JoVE.com

This research was supported by the FP7 European Research Council Grant 281501, ENGVASC.

All animal studies were approved by the Committee of the Ethics of Animal Experiments of the Technion. For this procedure, athymic nude mice were used to avoid immunological rejection. If using another type of mouse, the mice should be shaved prior to the surgical procedure and administration of cyclosporine (or another anti-rejection substitute) is recommended.
1. Scaffold Preparation and Cell Embedding
<ol>
	<li>Prepare scaffolds composed of 1:1 mixture of poly-L-lactic-acid (PLLA) and polylactic-co-glycolic-acid (PLGA), in the following manner:
	<ol>
		<li>Dissolve 500 mg of PLLA and 500 mg of PLGA in 10 ml chloroform.</li>
		<li>Add 0.24 ml of polymer solution to 0.4 g of sodium chloride particles gathered in a Teflon molds. Use a salt diameter of 212-600 &#181;m. Allow the chloroform to evaporate O/N.</li>
		<li>Leach the salt out using distilled water leading to interconnected porous 3D scaffolds.</li>
		<li>Cut polymer pieces using a scissors, then soak in 70% ethanol (v/v) for 30 min and wash 3 times in PBS for 2 min before usage.</li>
	</ol>
	</li>
	<li>Prepare a tri-culture of the following cells in 1:1 endothelial cell medium (EGM-2 supplemented with the components of its bullet kit and muscle cell medium (Dulbecco&#39;s minimal essential medium (DMEM), supplemented with 10% fetal bovine serum (FBS), 2.5% HEPES buffer, 100 U/ml penicillin, and 0.1 mg/ml streptomycin (Pen-Strep Solution).
	<ol>
		<li>Use 0.5 x 106 C2C12 myoblasts, 0.9 x 106 Human umbilical vein endothelial cells&#160;(HUVECs), and 0.2 x 106 Normal human dermal fibroblasts (NHDF).</li>
	</ol>
	</li>
	<li>Mix the cells together in a microcentrifuge tube and centrifuge at 200 x g for 4 min, aspirate the medium and re-suspend the pellet in a mixture of 4 &#181;l ice cold extracellular matrix solution and 4 &#181;l cell medium.</li>
	<li>Cut a piece of 10 mm x 7 mm x 1 mm (length x width x thickness) PLLA/PLGA scaffold and seed the cell suspension in a 6-well plate, allow the extracellular matrix solution to solidify (30 min, 37 &#176;C, 5% CO2) before adding 4 ml cell medium.</li>
	<li>Incubate the scaffolds for ten days before implantation and replace the medium every other day.</li>
</ol>
2. Before Starting the Surgical Procedure
<ol>
	<li>Prepare and sterilize (autoclave) the following surgical tools: a small, fine, straight scissors, a spring scissors, a straight, fine-tipped forceps, a serrated forceps and a needle holder.</li>
	<li>Prepare a mixture of ketamine:xylazine solution in an microcentrifuge tube by mixing 425 &#181;l ketamine and 75 &#181;l xylazine in a 1 ml syringe.</li>
	<li>Dilute 0.3 mg/ml buprenorphine in sterile saline (1:20).</li>
</ol>
3. Flap Construction (Graft Implantation)
<ol>
	<li>Using an insulin syringe, anesthetize the mouse by an intraperitoneal injection of ketamine:xylazine (35 &#181;l per 20 g of body weight).</li>
	<li>Subcutaneously inject buprenorphine (final concentration of 0.05-0.1 mg/kg) before the opperation.</li>
	<li>Place the mouse on a stage heated to 37 &#176;C, to ensure normal body temperature, and then secure the mouse using an adhesive bandage.</li>
	<li>Using cotton tips, administer lubricating eye ointment to the mouse&#39;s eyes (to avoid dehydration).</li>
	<li>Using cotton tips, clean the incision site with iodine and then with 70% ethanol, to establish an aseptic working field.</li>
	<li>Using small scissors and forceps, make an incision through the skin, from the level of the knee to the inguinal ligament, parallel to the femoral vessels, until the femoral artery and vein (AV) vessels are exposed.</li>
	<li>Hold the skin down using a retractor, to gain better access to the tissue.</li>
	<li>Using spring scissors and forceps, carefully isolate the femoral artery and vein vessels from the surrounding tissue. Start from the level of the inguinal ligament to the bifurcation of the superior epigastric vessels. Leave the profunda untouched, in order to preserve blood flow to the leg and prevent subsequent ischemia.</li>
	<li>Fold the graft (from Section 1) around the exposed femoral AV, below the profunda and above the bifurcation to the tibial and proneal AV, and join its ends using 8-0 silk sutures.</li>
	<li>To ensure implant vascularization by the capillaries sprouting from the femoral AV vessels only, wrap a piece of sterilized latex around the graft, and suture with 6-0 sutures.</li>
	<li>Close the overlying skin using 4-0 silk sutures.</li>
	<li>Monitor the mouse closely until recovery from the anesthesia and every day thereafter, until the graft is transferred as a flap. Subcutaneously inject buprenorphine (final concentration of 0.05-0.1 mg/kg) twice a day for 2-3 days. 
	NOTE: Use saline in any stage, to avoid dehydration of the exposed tissue.</li>
</ol>
4. Flap Transfer
<ol>
	<li>At one or two weeks postimplantation, anesthetize the mouse by an intraperitoneal injection of ketamine:xylazine (35 &#181;l/20 g), using an insulin syringe.</li>
	<li>Subcutaneously inject buprenorphine (final concentration of 0.05-0.1 mg/kg) before the opperation.</li>
	<li>Place the mouse on a stage heated to 37 &#176;C, to ensure normal body temperature, and then secure the mouse using an adhesive bandage.</li>
	<li>Using cotton tips, apply lubricating eye ointment to the mouse&#39;s eyes (to avoid dehydration).</li>
	<li>Using cotton tips, clean the incision site (from the knee area to the abdomen) with iodine and then with 70% ethanol, to establish an aseptic working field.</li>
	<li>Using a scissors and forceps, carefully open the sutures in the skin and, using a scissors, make an incision through the skin parallel to the inguinal ligament, continuing through the ventral skin.</li>
	<li>Carefully remove the latex piece with forceps and expose the vascularized flap.</li>
	<li>Using a scissors and forceps, dissect the tissue flap from the surrounding tissue.</li>
	<li>Using a forceps and a needle holder, ligate the distal end of the femoral AV with 8-0 silk sutures to prevent bleeding from the AV. Then, cauterize the AV at the level of the knee, distally to the folded implanted tissue and the 8-0 sutures.</li>
	<li>Gently transfer the femoral AV enveloped by the graft, towards the ventral abdominal wall, avoiding damage to the artery, to determine at which distance the full thickness defect should be made. 
	CAUTION: Pulling the artery too far will damage the artery.</li>
	<li>Using a fine scissors, make an incision in the ventral abdominal wall.</li>
	<li>Using a spring scissors, make an incision in the rectus abdominus muscle. Remove an approximately 1 cm x 0.8 cm segment of the rectus abdominus muscle with the overlying skin.</li>
	<li>Transfer the femoral AV, enveloped by the vascularized graft, as an axial flap, to the full-thickness defect in the ventral abdominal wall.</li>
	<li>Suture the flap to the surrounding muscle tissue, using 8-0 silk sutures, to prevent hernia.</li>
	<li>To avoid suturing the internal organs, raise the muscle with a forceps when suturing the flap to the abdominal muscle. Leave the ventral skin exposed to mimic the effect of a full abdominal wall defect.</li>
	<li>Cover the wound in the skin with iodinated gauze and a sterile bandage to prevent contamination of the exposed area.</li>
	<li>Suture the skin of the leg using 4-0 silk sutures.</li>
	<li>Monitor the mouse closely until recovery from the anesthesia and every day thereafter, until retrieval of the flap.&#160; Subcutaneously inject buprenorphine (final concentration of 0.05-0.1 mg/kg) twice a day for 2-3 days.</li>
	<li>NOTE: Use saline in any stage to avoid dehydration of the exposed tissue.</li>
</ol>
5. Ultrasound Determination of Vascular Perfusion of the Graft
NOTE: Before transfer, vascular perfusion of the graft is measured at one and two weeks after implantation, by ultrasonography.
<ol>
	<li>Anesthetize the mouse using 2% isoflurane.</li>
	<li>Place the mouse on a movable stage heated to 38 &#176;C, to ensure normal body temperature, and then secure the mouse using an adhesive bandage.</li>
	<li>Find the femoral artery and vein vessels with non-linear transducer set on the B-mode.</li>
	<li>Examine the patency of the femoral vessels of the graft using the transducer set on the color Doppler mode.</li>
	<li>Prepare non-targeted contrast agent according to the manufacturer&#8217;s instructions.</li>
	<li>Connect a tail vein catheter (12 cm tubing with a 27 G needle) to a 1 ml syringe filled with saline. Secure the syringe next to the mouse on the heating stage (to avoid movement of the syringe).</li>
	<li>Connect the catheter to the mouse tail vein and secure the needle.</li>
	<li>Inject some of the saline to both ensure that injections are being made into the vein and to fill the tubing with saline (to avoid air bubbles).</li>
	<li>Fill a 1 ml syringe with 70 &#181;l of contrast agent and replace the saline syringe with the contrast agent syringe.</li>
	<li>Set up the ultrasound to image injection of the contrast agent. In properties of the non-linear mode set up the number of frames to 1,500 and the disruption pulse to 30% of the frames. 
	NOTE: After the disruption&#160;pulse, the microbubbles re-fill the vessels at a rate that depends on the flow rate of the microbubbles in the capillaries and is independent of the injection rate of the microbubbles. The number of frames can be changed according to the filling time 19,20.</li>
	<li>Slowly inject the contrast agent into the tail vein.</li>
	<li>Capture images in the non-linear contrast mode of the high-resolution ultrasound system.</li>
	<li>Analyze the results using the software, as previously described 19,20.
	<ol>
		<li>Draw a region of interest (ROI) on the image obtained immediately after the disruption&#160;pulse in the contrast-mode. The ROI should outline the area filled by the contrast agent only.</li>
		<li>Calculate the peak enhancement (PE), which is the ratio of the mean intensity of the non-linear signal after the injection of the microbubbles versus after the disruption&#160;pulse, and is a measure of the perfusion volume.</li>
		<li>Calculate the flow rate, which is determined from the time (T) that elapses until the peak signal (P).</li>
	</ol>
	</li>
</ol>
6. Determination of the Extent of Graft Vascularization
NOTE: The extent of tissue graft vascularization is determined one or two weeks after implantation.
<ol>
	<li>Using an insulin syringe, anesthetize the mouse by an intraperitoneal injection of ketamine:xylazine (35 &#181;l/20 g).</li>
	<li>Using 1 ml syringe, intravenously inject 200 &#181;l of 10&#160;mg/ml fluorescein isothiocyanate-conjugated dextran (FITC-Dextran, MW = 500,000) into the tail vein.</li>
	<li>Ten sec after completion of the injection, euthanize the mouse by cervical dislocation.</li>
	<li>Open the leg of the mouse and expose the graft.</li>
	<li>Wash the graft area with phosphate buffered saline (PBS) and apply 10% neutral buffered formalin.</li>
	<li>Image the graft in the mouse leg, using a confocal microscope.</li>
	<li>Quantify functional vessel density (FVD).
	<ol>
		<li>Isolate the green channel (excitation = 488 nm) of the confocal microscopic images.</li>
		<li>Pass the resulting images through a high-pass filter, in order to accentuate the high-contrast structures.</li>
		<li>Use a despeckling filter to remove noise that might have been amplified by the high-pass filter.</li>
		<li>Set up the value of threshold on the resulting image; the threshold value is adjusted so that the features and structures in the original image are visible in the binary image.</li>
		<li>Apply a size-threshold so that only groups of connected pixels larger than the defined size threshold remain in the binary image.</li>
		<li>Outline the skeleton of the vessels in the graft using the Zhang-Suen&#39;s algorithm 21.</li>
		<li>Calculate the FVD by summing the lengths of the midline of each vessel and dividing the result by the area of the ROI.</li>
	</ol>
	</li>
</ol>
7. Immunohistological and Histological Staining of the Graft
<ol>
	<li>Immunohistological Staining
	<ol>
		<li>After confocal imaging, retrieve the graft using a small scissors and forceps. Dissect the AV vessels in the two ends of the graft and retrieve the graft.</li>
		<li>Fix the grafts in 10% neutral buffered formalin for 30 min - 2 hr.</li>
		<li>Place the grafts in histology cassettes and store in 70% ethanol until paraffin embedding.</li>
		<li>Embed in paraffin using a standard fixation and embedding protocol 22 and slice the paraffin embedded tissues into 5 &#181;m slices using a microtome. Slice the whole tissue perpendicularly to the AV vessels.</li>
		<li>Deparaffinize the paraffin-embedded sections by immersion in 100% xylene twice, for 10 min each.</li>
		<li>Arrange the slides in a 250 ml plastic Coplin jar.</li>
		<li>Rehydrate by serial immersions in decreasing concentrations of ethanol (100%, 96%, 90%, 80%, 70%, 50%, 30% and double distilled water (DDW)) for 3 min each.</li>
		<li>Prepare antigen unmasking solution by diluting 2.35 ml of antigen unmasking solution with 250 mL of DDW. Heat the antigen unmasking solution for 2 min in a microwave set to 95 &#176;C.</li>
		<li>Insert the slides into the pre-heated antigen unmasking solution and heat again for 3 min in a microwave set to 95 &#176;C. Cool the slides to 50 &#176;C.</li>
		<li>Heat again for 2 min and then cool to RT.</li>
		<li>Incubate the slides in 3.3% H2O2solution in methanol for 10 min, at RT to quench the activity of endogenous peroxidase and then rinse twice in PBS.</li>
		<li>Prepare an incubation chamber: place wet absorbent sheets inside a histology box and dry the slides using delicate task wipers.</li>
		<li>Circle the slice area on the slide using a hydrophobic pen and incubate for 30 min at RT, with approximately 50 &#181;l of goat serum blocking solution (2%, prepared in PBS) per slice.</li>
		<li>Dilute the rabbit anti-CD31 antibody 1:50 in blocking solution, apply approximately 50 &#181;l antibody per slice and incubate O/N, at 4 &#176;C. Rinse 3 times with PBS, 5 min each.</li>
		<li>Dilute the biotinylated goat anti-rabbit secondary antibody 1:400 in PBS, place over slides approximately 50 &#181;l of antibody per slice and incubate for 30 min at RT.</li>
		<li>Rinse 3 times with PBS, 5 min each.</li>
		<li>Dilute streptavidin-peroxidase 1:400 in PBS, apply approximately 50 &#181;l per slice and incubate with slides for 30 min, at RT. Rinse 3 times with PBS, 5 min each.</li>
		<li>Apply approximately 50 &#181;l of aminoethylcarbazole (AEC) substrate per slice, according to the manufacturer&#8217;s instructions. Dip the slides in DDW to stop the reaction.</li>
		<li>Stain the nuclei in blue by immersing the sections in approximately 50 &#181;l of Mayer&#8217;s hematoxylin solution for 2 min. Gently rinse in water and then cover the sections with mounting medium.</li>
		<li>Quantify CD31-positive staining using a double-blind approach. CD31 staining appears as a brown stain. Choose 4-5 different fields in a 40X magnification viewing field under a binocular microscope. Count the number of CD31-positive lumens.
		<ol>
			<li>Calculate the average number of CD31-positive lumens per scaffold by dividing the total number of counted lumens with the number of fields and the fieldarea (according to the binocular data). 
			NOTE: The applied reagent volume must be sufficient to cover the entire outlined slice.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Histological Staining
	<ol>
		<li>Fix grafts in 10% neutral buffered formalin.</li>
		<li>Embed grafts in paraffin using standard fixation and embedding procedures.</li>
		<li>Remove the paraffin from the paraffin-embedded sections by immersion in 100% xylene twice for 10 min.</li>
		<li>Fast wash twice for rehydration in 100% isopropanol.</li>
		<li>Fast wash twice 95% Ethanol.</li>
		<li>Wash three times in tap water.</li>
		<li>Immerse in Hematoxylin for 10 min.</li>
		<li>Wash three times in tap water.</li>
		<li>Immere in Eosin for 2 min.</li>
		<li>Fast wash dehydrate in 95% Ethanol twice.</li>
		<li>Fast wash 100% isopropanol twice.</li>
		<li>Fast wash 100% xylene twice.</li>
		<li>Cover with DPX glue with cover glass 1.5 mm thick.</li>
		<li>Image using microscope in 40X magnification.</li>
	</ol>
	</li>
</ol>
8. Mechanical Properties Assessment
<ol>
	<li>Retrieve the graft using a small scissors and forceps.</li>
	<li>Rinse the graft in PBS.</li>
	<li>Measure the width and thickness of the graft and calculate the cross-section area as width multiplied by thickness.</li>
	<li>Use the test instrument to generate a stress-strain curve of hydrated grafts.
	<ol>
		<li>Mount the graft between the system tensile grips and measure the final length between the two grips (i.e., initial length).</li>
		<li>Apply a strain rate of 0.01 mm/sec, until failure.</li>
		<li>Record the developed force and displacement using the manufacturer&#8217;s protocol.</li>
		<li>Generate a stress-strain curve in Matlab.
		<ol>
			<li>Calculate strain according to displacement divided by the initial length. Stress is calculated as the measured force divided by the cross-sectional area.</li>
			<li>Determine stiffness as the slope of the linear-region in the stress-strain curve and the maximum point of the curve is the ultimate tensile strength.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abdominal+wall+reconstruction+using+biological+tissue+grafts:+present+status+and+future+opportunities.">Abdominal wall reconstruction using biological tissue grafts: present status and future opportunities.</a> Exp rev of med dev. 3 (5), 657-675 (2006).</li><li>Caspi, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+engineering+of+vascularized+cardiac+muscle+from+human+embryonic+stem+cells.">Tissue engineering of vascularized cardiac muscle from human embryonic stem cells.</a> Circ Res. 100 (2), 263-272 (2007).</li><li>Kaufman-Francis, K., Koffler, J., Weinberg, N., Dor, Y., Levenberg, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineered+vascular+beds+provide+key+signals+to+pancreatic+hormone-producing+cells.">Engineered vascular beds provide key signals to pancreatic hormone-producing cells.</a> PloS one. 7 (7), e40741 (2012).</li><li>Kaully, T., Kaufman-Francis, K., Lesman, A., Levenberg, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vascularization--the+conduit+to+viable+engineered+tissues.">Vascularization--the conduit to viable engineered tissues.</a> Tiss eng. Part B, Reviews. 15 (2), 159-169 (2009).</li><li>Koffler, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+vascular+organization+enhances+functional+integration+of+engineered+skeletal+muscle+grafts.">Improved vascular organization enhances functional integration of engineered skeletal muscle grafts.</a> PNAS of the USA. 108 (36), 14789-14794 (2011).</li><li>Lesman, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transplantation+of+a+tissue-engineered+human+vascularized+cardiac+muscle.">Transplantation of a tissue-engineered human vascularized cardiac muscle.</a> Tisseng. Part A. 16 (1), 115-125 (2010).</li><li>Levenberg, 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=Engineering+vascularized+skeletal+muscle+tissue.">Engineering vascularized skeletal muscle tissue.</a> Nat Biotechnol. 23 (7), 879-884 (2005).</li><li>Bearzi, 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=PlGF-MMP9-engineered+iPS+cells+supported+on+a+PEG-fibrinogen+hydrogel+scaffold+possess+an+enhanced+capacity+to+repair+damaged+myocardium.">PlGF-MMP9-engineered iPS cells supported on a PEG-fibrinogen hydrogel scaffold possess an enhanced capacity to repair damaged myocardium.</a> Cell death &amp; disease. 5, e1053 (2014).</li><li>Zhang, 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=SDF-1+expression+by+mesenchymal+stem+cells+results+in+trophic+support+of+cardiac+myocytes+after+myocardial+infarction.">SDF-1 expression by mesenchymal stem cells results in trophic support of cardiac myocytes after myocardial infarction.</a> FASEB J : official publication of the .Fed Am Soc Exp Biol. 21 (12), 3197-3207 (2007).</li><li>Dvir, 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=Prevascularization+of+cardiac+patch+on+the+omentum+improves+its+therapeutic+outcome.">Prevascularization of cardiac patch on the omentum improves its therapeutic outcome.</a> PNAS. 106 (35), 14990-14995 (2009).</li><li>Marsano, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+controlled+expression+of+VEGF+by+transduced+myoblasts+in+a+cardiac+patch+on+vascularization+in+a+mouse+model+of+myocardial+infarction.">The effect of controlled expression of VEGF by transduced myoblasts in a cardiac patch on vascularization in a mouse model of myocardial infarction.</a> Biomaterials. 34 (2), 393-401 (2013).</li><li>Rufaihah, A. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhanced+infarct+stabilization+and+neovascularization+mediated+by+VEGF-loaded+PEGylated+fibrinogen+hydrogel+in+a+rodent+myocardial+infarction+model.">Enhanced infarct stabilization and neovascularization mediated by VEGF-loaded PEGylated fibrinogen hydrogel in a rodent myocardial infarction model.</a> Biomaterials. 34 (33), 8195-8202 (2013).</li><li>Nillesen, S. T. 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=Increased+angiogenesis+in+acellular+scaffolds+by+combined+release+of+FGF2+and+VEGF.">Increased angiogenesis in acellular scaffolds by combined release of FGF2 and VEGF.</a> J of Contr Release. 116 (2), e88-e90 (2006).</li><li>Sekine, 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=In+vitro+fabrication+of+functional+three-dimensional+tissues+with+perfusable+blood+vessels.">In vitro fabrication of functional three-dimensional tissues with perfusable blood vessels.</a> Nat Commun. 4, 1399 (2013).</li><li>Tee, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transplantation+of+engineered+cardiac+muscle+flaps+in+syngeneic+rats.">Transplantation of engineered cardiac muscle flaps in syngeneic rats.</a> Tiss eng. Part A. (19-20), 1992-1999 (2012).</li><li>Morritt, A. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+tissue+engineering+in+an+in+vivo+vascularized+chamber.">Cardiac tissue engineering in an in vivo vascularized chamber.</a> Circulation. 115 (3), 353-360 (2007).</li></ol>

The authors declare that they have no competing financial interests.

The advances in tissue engineering have been met with a growing demand for substitute tissues for reconstruction of various tissue types. A variety of synthetic 1,17,18 and biological 14-16 materials as well as fabrication methods have been assessed for their capacity to address these demands. However, despite the progress in clinical care and in tissue engineering, the restoration of full-thickness abdominal wall defects remains a challenge. A tissue adequate for reconstruction of such massive defe...

Abdominal wall defects often arise following severe trauma, cancer treatment, burns and removal of infected mesh. These defects often involve significant tissue loss, requiring complicated surgical procedures and presenting a major challenge for plastic reconstruction surgeons 1-4. Tissue engineering researchers seeking new sources for artificial tissues have explored different materials, cell sources and growth factors. Successful restorations of various tissues, such as trachea 5,6, bladder 7, cornea 8, bone 9and skin 10, by implantation of engineered tissues were previously reported. However, fabrication of a thick vascularized engineered tissue, particularly for reconstruction of large defects, remains a significant challenge in tissue engineering.
One of the main factors limiting the thickness of a viable tissue construct is the control of oxygen supply to its constituent cells. When relying on diffusion, construct thickness is limited to that of a very thin layer. The maximum distance between oxygen- and nutrient-supplying capillaries in vivo is approximately 200&#160;&#181;m, which correlates with the diffusion limit of oxygen11,12. Insufficient vascularization can result in tissue ischemia and escalate to tissue resorption or necrosis 13.
In addition, the ideal material used for tissue reconstruction must be biocompatible and non-immunogenic. It must also be capable of promoting further integration of host cells with the biomaterial, and maintaining structural integrity. Various biological 14-16 and synthetic 1,17,18 matrices have been previously explored for tissue reconstruction, however their use remain limited due to lack of effective blood supply, infections or insufficient tissue strength.
In this study, a biocompatible, cell-embedded scaffold comprised of Food and Drug Administration (FDA)-approved poly L-lactic acid (PLLA)/poly lactic-co-glycolic acid (PLGA), was implanted around the femoral artery and vein (AV) vessels of a nude mouse and separated from the surrounding tissue, ensuring vascularization from the AV vessels only. One week post-implantation, the graft was viable, thick and well vascularized. This thick vascularized tissue with the AV vessels, was then transferred as a pedicled flap to an abdominal full-thickness defect in the same mouse. One week post-transfer, the flap was viable, vascularized and well integrated with the surrounding tissue, bearing sufficient strength to support abdominal viscera. Thus, the engineered thick, vascularized tissue flap, bearing an autologous pedicle, presents a novel method for repairing full-thickness abdominal wall defects.

<table><tbody><tr><td>small fine straight scissors</td><td>Fine Science Tools (FST)</td><td>14090-09</td><td></td></tr><tr><td>spring scissors</td><td>Fine Science Tools (FST)</td><td>15003-08</td><td></td></tr><tr><td>straight forceps with fine tip</td><td>Fine Science Tools (FST)</td><td>11251-20</td><td></td></tr><tr><td>serrated forceps&amp;nbsp;</td><td>Fine Science Tools (FST)</td><td>11050-10</td><td></td></tr><tr><td>needle holder</td><td>Fine Science Tools (FST)</td><td>12500-12</td><td></td></tr><tr><td>Small vessel cauterizer&amp;nbsp;</td><td>Fine Science Tools (FST)</td><td>18000-00</td><td></td></tr><tr><td>Duratears</td><td>Alcon</td><td>5686</td><td></td></tr><tr><td>Sedaxylan</td><td>Euravet</td><td>DJ03</td><td></td></tr><tr><td>Clorketam 1000</td><td>Vetoquinol</td><td>4A0726B</td><td></td></tr><tr><td>Buprenorphine</td><td>vetmarket</td><td>B15100</td><td></td></tr><tr><td>4-0 silk sutures</td><td>Assut sutures</td><td>647</td><td></td></tr><tr><td>6-0 polypropylene sutures</td><td>Assut sutures</td><td>9351F</td><td></td></tr><tr><td>8-0 silk sutures</td><td>Assut sutures</td><td>684568</td><td></td></tr><tr><td>Insulin syringe (6 mm needle)</td><td>BD</td><td>324911</td><td></td></tr><tr><td>Vevo 2100 high-resolution ultrasound system</td><td>VisualSonics inc.</td><td></td><td></td></tr><tr><td>MS250 non-linear transducer</td><td>VisualSonics inc.</td><td></td><td></td></tr><tr><td>Micromarker non-targeted contrast agent</td><td>VisualSonics inc.</td><td>VS-11694</td><td></td></tr><tr><td>tail vein catheter</td><td>VisualSonics inc.</td><td>VS-11912</td><td></td></tr><tr><td>Vevo 2100 software</td><td>VisualSonics inc.</td><td></td><td></td></tr><tr><td>fluorescein isothiocyanate-conjugated dextran</td><td>Sigma</td><td>FD500S</td><td></td></tr><tr><td>Matlab</td><td>Mathworks, MA, USA</td><td></td><td></td></tr><tr><td>Kimwipes</td><td>Kimtech</td><td>34120</td><td></td></tr><tr><td>antigen unmasking solution</td><td>Vector laboratories</td><td>H-3300</td><td></td></tr><tr><td>anti-CD31 antibody</td><td>Abcam&amp;nbsp;</td><td>ab28364</td><td></td></tr><tr><td>biotinylated goat anti-rabbit (secondary) antibody</td><td>Vector laboratories</td><td>BA-1000</td><td></td></tr><tr><td>streptavidin-peroxidase</td><td>Jackson&amp;nbsp;</td><td>016-030-084</td><td></td></tr><tr><td>Mayer&#039;s hamatoxylin solution</td><td>Sigma-Aldrich</td><td>MHS-16</td><td></td></tr><tr><td>aminoethylcarbazole (AEC) substrate kit</td><td>Life technologies, Invitrogen&amp;nbsp;</td><td>00-2007</td><td></td></tr><tr><td>Vectamount</td><td>Vector laboratories</td><td>H-5501</td><td></td></tr></tbody></table>

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.

Graft vascularization and perfusion in vivo
The grafts were implanted one or two weeks prior to their transfer as axial flaps. At one and two weeks post-implantation, gross observation of the graft area revealed viable and vascularized tissue grafts. These grafts proved to be highly vascularized, as determined by positive CD31 immunostaining (Figure 1A), and highly perfused, as evidenced by FITC-dextran tail vein injection and ultrasound measurements. Many vessels wer...

Watch this Scientific Journal Video about Engineered Vascularized Muscle Flap at JoVE.com

Engineered Vascularized Muscle Flap

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 ...

Research

JoVE Journal

Bioengineering

8.2K Views.  Kaplan Medical Center. 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. 

Video: Engineered Vascularized Muscle Flap

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 3D Silk-collagen-based Model of Polarized Neural Tissue

Despite huge efforts to decipher the anatomy, composition and function of the brain, it remains the least understood organ of the human body. To gain a deeper comprehension of the neural system scientists aim to simplistically reconstruct the tissue by assembling it in vitro from basic building blocks using a tissue engineering approach. Our group developed a tissue-engineered silk and collagen-based 3D brain-like model resembling the white and gray matter of the cortex. The model consists of silk porous sponge, which is pre-seeded with rat brain-derived neurons, immersed in soft collagen matrix. Polarized neuronal outgrowth and network formation is observed with separate axonal and cell body localization. This compartmental architecture allows for the unique development of niches mimicking native neural tissue, thus enabling research on neuronal network assembly, axonal guidance, cell-cell and cell-matrix interactions and electrical functions.

Insight into the complex actions of the brain requires advanced research tools. Here we demonstrate a novel silk-collagen-based 3D engineered model of neural tissue resembling brain-like architecture. The model can be used to study neuronal network assembly, axonal guidance, cell-cell interactions and electrical activity.

Despite huge efforts to decipher the anatomy, composition and function of the brain, it remains the least understood organ of the human body. To gain a ...

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 ...

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 ...

A Model of Free Tissue Transfer: The Rat Epigastric Free Flap

Free tissue transfer has been increasingly used in clinical practice since the 1970s, allowing reconstruction of complex and otherwise untreatable defects resulting from tumor extirpation, trauma, infections, malformations or burns. Free flaps are particularly useful for reconstructing highly complex anatomical regions, like those of the head and neck, the hand, the foot and the perineum. Moreover, basic and translational research in the area of free tissue transfer is of great clinical potential. Notwithstanding, surgical trainees and researchers are frequently deterred from using microsurgical models of tissue transfer, due to lack of information regarding the technical aspects involved in the operative procedures. The aim of this paper is to present the steps required to transfer a fasciocutaneous epigastric free flap to the neck in the rat.
This flap is based on the superficial epigastric artery and vein, which originates from and drain into the femoral artery and vein, respectively. On average the caliber of the superficial epigastric vein is 0.6 to 0.8 mm, contrasting with the 0.3 to 0.5 mm of the superficial epigastric artery. Histologically, the flap is a composite block of tissues, containing skin (epidermis and dermis), a layer of fat tissue (panniculus adiposus), a layer of striated muscle (panniculus carnosus), and a layer of loose areolar tissue.
Succinctly, the epigastric flap is raised on its pedicle vessels that are then anastomosed to the external jugular vein and to the carotid artery on the ventral surface of the rat's neck. According to our experience, this model guarantees the complete survival of approximately 70 to 80% of epigastric flaps transferred to the neck region. The flap can be evaluated whenever needed by visual inspection. Hence, the authors believe this is a good experimental model for microsurgical research and training.

This paper describes the steps required to raise a fasciocutaneous epigastric free flap and transfer it to the neck in the rat.

Free tissue transfer has been increasingly used in clinical practice since the 1970s, allowing reconstruction of complex and otherwise untreatable defects ...

Medicine

A Reliable Porcine Fascio-Cutaneous Flap Model for Vascularized Composite Allografts Bioengineering Studies

Vascularized Composite Allografts (VCA) such as hand, face, or penile transplant represents the cutting-edge treatment for devastating skin defects, failed by the first steps of the reconstructive ladder. Despite promising aesthetic and functional outcomes, the main limiting factor remains the need for a drastically applied lifelong immunosuppression and its well-known medical risks, preventing broader indications. Therefore, lifting the immune barrier in VCA is essential to tip the ethical scale and improve patients&#39; quality of life using the most advanced surgical techniques. De novo creation of a patient-specific graft is the upcoming breakthrough in reconstructive transplantation. Using tissue engineering techniques, VCAs can be freed of donor cells and customized for the recipient through perfusion-decellularization-recellularization. To develop these new technologies, a large-scale animal VCA model is necessary. Hence, swine fascio-cutaneous flaps, composed of skin, fat, fascia, and vessels, represent an ideal model for preliminary studies in VCA. Nevertheless, most VCA models described in the literature include muscle and bone. This work reports&#160;a reliable and reproducible technique for saphenous fascio-cutaneous flap harvest in swine, a practical tool for various research fields, especially vascularized composite tissue engineering.

The present protocol describes the porcine fascio-cutaneous flap model and its potential use in vascularized composite tissue research.

Vascularized Composite Allografts (VCA) such as hand, face, or penile transplant represents the cutting-edge treatment for devastating skin defects, ...

Fabrication of Engineered Vascular Flaps Using 3D Printing Technologies

Engineering implantable, functional, thick tissues requires designing a hierarchical vascular network. 3D bioprinting is a technology used to create tissues by adding layer upon layer of printable biomaterials, termed bioinks, and cells in an orderly and automatic manner, which allows for creating highly intricate structures that traditional tissue engineering techniques cannot achieve. Thus, 3D bioprinting is an appealing in vitro approach to mimic the native vasculature complex structure, ranging from millimetric vessels to microvascular networks. 
Advances in 3D bioprinting in granular hydrogels enabled the high-resolution extrusion of low-viscosity extracellular matrix-based bioinks. This work presents a combined 3D bioprinting and sacrificial mold-based 3D printing approach for fabricating engineered vascularized tissue flaps. 3D bioprinting of endothelial and support cells using recombinant collagen-methacrylate bioink within a gelatin support bath is utilized for the fabrication of a self-assembled capillary network. This printed microvasculature is assembled around a mesoscale vessel-like porous scaffold, fabricated using a sacrificial 3D printed mold, and is seeded with endothelial cells. 
This assembly induces the endothelium of the mesoscale vessel to anastomose with the surrounding capillary network, establishing a hierarchical vascular network within an engineered tissue flap. The engineered flap is then directly implanted by surgical anastomosis to a rat femoral artery using a cuff technique. The described methods can be expanded for the fabrication of various vascularized tissue flaps for use in reconstruction surgery and vascularization studies.

Engineered flaps require an incorporated functional vascular network. In this protocol, we present a method of fabricating a 3D printed tissue flap containing a hierarchical vascular network and its direct microsurgical anastomoses to rat femoral artery.

Engineering implantable, functional, thick tissues requires designing a hierarchical vascular network. 3D bioprinting is a technology used to create ...

Procurement and Perfusion-Decellularization of Porcine Vascularized Flaps in a Customized Perfusion Bioreactor

Large volume soft tissue defects lead to functional deficits and can greatly impact the patient&#39;s quality of life. Although surgical reconstruction can be performed using autologous free flap transfer or vascularized composite allotransplantation (VCA), such methods also have disadvantages. Issues such as donor site morbidity and tissue availability limit autologous free flap transfer, while immunosuppression is a significant limitation of VCA. Engineered tissues in reconstructive surgery using decellularization/recellularization methods represent a possible solution. Decellularized tissues are generated using methods that remove native cellular material while preserving the underlying extracellular matrix (ECM) microarchitecture. These acellular scaffolds can then be subsequently recellularized with recipient-specific cells.
This protocol details the procurement and decellularization methods used to achieve acellular scaffolds in a pig model. In addition, it also provides a description of the perfusion bioreactor design and setup. The flaps include the porcine omentum, tensor fascia lata, and the radial forearm. Decellularization is performed via ex vivo perfusion of low concentration sodium dodecyl sulfate (SDS) detergent followed by DNase enzyme treatment and peracetic acid sterilization in a customized perfusion bioreactor.
Successful tissue decellularization is characterized by a white-opaque appearance of flaps macroscopically. Acellular flaps show the absence of nuclei on histological staining and a significant reduction in DNA content. This protocol can be used efficiently to generate decellularized soft tissue scaffolds with preserved ECM and vascular microarchitecture. Such scaffolds can be used in subsequent recellularization studies and have the potential for clinical translation in reconstructive surgery.

The protocol describes the surgical procurement and subsequent decellularization of vascularized porcine flaps by the perfusion of sodium dodecyl sulfate detergent through the flap vasculature in a customized perfusion bioreactor.

Large volume soft tissue defects lead to functional deficits and can greatly impact the patient&#39;s quality of life. Although surgical reconstruction ...

Porcine As a Training Module for Head and Neck Microvascular Reconstruction

Live models that resemble surgical conditions of humans are needed for training free-flap harvesting and anastomosis. Animal models for training purposes have been available for years in many surgical fields. We used the female (because they are easy to handle for the procedure) Yorkshire pigs for the head and neck reconstruction by harvesting the deep inferior epigastric artery perforator or the superior epigastric artery perforator flap. The anastomosis site (neck skin defect or tracheal wall defect) was prepared via the dissection of the common carotid artery and the internal jugular vein, in which 3.5&#215; loupe magnification was used for anastomosis as we use on human cases in real life. This procedure demonstrates a new training method using a reliable learning model and provides a detailed anatomy in a live scenario. We focused on the ischemia time, harvesting, vessel anastomosis, and designing the flap to fit the defect site. This model improves tissue handling and with the use of proper instruments can be repeated many times so that the surgeon is fully confident before starting the surgery on humans.

Here we present a protocol for the use of the pig superior epigastric artery perforator flap as a learning module for head and neck microvascular reconstruction.

Live models that resemble surgical conditions of humans are needed for training free-flap harvesting and anastomosis. Animal models for training purposes ...

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand and face transplants and, more recently, abdominal wall, uterus, and urogenital transplants. Functional outcomes have exceeded initial expectations, and most recipients enjoy an improved quality of life. However, as clinical experience accumulates, chronic rejection and complications from the immunosuppression must be addressed. In many cases where grafts have failed, the causative pathology has been ischemic vasculopathy. <span style="font-size: 11pt; line-height: 115%; font-family: Arial, sans-serif; color: rgb(41, 43, 49); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">The biological mechanisms of the acute and chronic rejection associated with VCA, especially ischemic vasculopathy, are important areas of research. However, due to the very small number of VCA patients, the evaluation of proposed mechanisms is better addressed in an experimental model. Multiple groups have used animal models to address some of the relevant unsolved questions in VCA rejection and vasculopathy. Several model designs involving a variety of species are described in the literature. Here we present a reproducible model of VCA heterotopic hindlimb osteomyocutaneous flap in the rat that can be utilized for translational VCA research. This model allows for the serial evaluation of the graft, including biopsies and different imaging modalities, while maintaining a low level of morbidity.

Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand ...

Engineered Vascularized Muscle Flap

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Engineered 3D Silk-collagen-based Model of Polarized Neural Tissue

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A Model of Free Tissue Transfer: The Rat Epigastric Free Flap

A Reliable Porcine Fascio-Cutaneous Flap Model for Vascularized Composite Allografts Bioengineering Studies

Fabrication of Engineered Vascular Flaps Using 3D Printing Technologies

Procurement and Perfusion-Decellularization of Porcine Vascularized Flaps in a Customized Perfusion Bioreactor

Porcine As a Training Module for Head and Neck Microvascular Reconstruction

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research