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>

<ol>
	<li>Engelsman, A. F., van der Mei, H. C., Ploeg, R. J., Busscher, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+phenomenon+of+infection+with+abdominal+wall+reconstruction.">The phenomenon of infection with abdominal wall reconstruction.</a> Biomaterials. 28 (14), 2314-2327 (2007).</li><li>De Coppi, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myoblast-acellular+skeletal+muscle+matrix+constructs+guarantee+a+long-term+repair+of+experimental+full-thickness+abdominal+wall+defects.">Myoblast-acellular skeletal muscle matrix constructs guarantee a long-term repair of experimental full-thickness abdominal wall defects.</a> Tissue Eng. 12 (7), 1929-1936 (2006).</li><li>Shi, 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=Regeneration+of+full-thickness+abdominal+wall+defects+in+rats+using+collagen+scaffolds+loaded+with+collagen-binding+basic+fibroblast+growth+factor.">Regeneration of full-thickness abdominal wall defects in rats using collagen scaffolds loaded with collagen-binding basic fibroblast growth factor.</a> Biomaterials. 32 (3), 753-759 (2011).</li><li>Yezhelyev, M. V., Deigni, O., Losken, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Management+of+full-thickness+abdominal+wall+defects+following+tumor+resection.">Management of full-thickness abdominal wall defects following tumor resection.</a> Ann Plast Surg. 69 (2), 186-191 (2012).</li><li>Macchiarini, P., Walles, T., Biancosino, C., Mertsching, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=First+human+transplantation+of+a+bioengineered+airway+tissue.">First human transplantation of a bioengineered airway tissue.</a> J Thorac Cardiovasc Surg. 128 (4), 638-641 (2004).</li><li>Macchiarini, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinical+transplantation+of+a+tissue-engineered+airway.">Clinical transplantation of a tissue-engineered airway.</a> Lancet. 372 (9665), 2023-2030 (2008).</li><li>Atala, A., Bauer, S. B., Soker, S., Yoo, J. J., Retik, A. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-engineered+autologous+bladders+for+patients+needing+cystoplasty.">Tissue-engineered autologous bladders for patients needing cystoplasty.</a> Lancet. 367 (9518), 1241-1246 (2006).</li><li>Nishida, K., 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=Corneal+reconstruction+with+tissue-engineered+cell+sheets+composed+of+autologous+oral+mucosal+epithelium.">Corneal reconstruction with tissue-engineered cell sheets composed of autologous oral mucosal epithelium.</a> N Engl J Med. 351 (12), 1187-1196 (2004).</li><li>Petite, 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=Tissue-engineered+bone+regeneration.">Tissue-engineered bone regeneration.</a> Nat Biotechnol. 18 (9), 959-963 (2000).</li><li>Banta, M. N., Kirsner, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulating+diseased+skin+with+tissue+engineering:+actinic+purpura+treated+with+Apligraf.">Modulating diseased skin with tissue engineering: actinic purpura treated with Apligraf.</a> Dermatol Surg. 28 (12), 1103-1106 (2002).</li><li>Vunjak-Novakovic, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Challenges+in+cardiac+tissue+engineering.">Challenges in cardiac tissue engineering.</a> Tissue engineering. Part B, Reviews. 16 (2), 169-187 (2010).</li><li>Novosel, E. C., Kleinhans, C., Kluger, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vascularization+is+the+key+challenge+in+tissue+engineering.">Vascularization is the key challenge in tissue engineering.</a> Advanced Drug Delivery Reviews. 63 (4-5), 300-311 (2011).</li><li>Lesman, A., Gepstein, L., 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+shaping+the+heart.">Vascularization shaping the heart.</a> Ann N Y Acad Sci. 1188, 46-51 (2010).</li><li>Patton Jr, H., Berry, S., Kralovich, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+human+acellular+dermal+matrix+in+complex+and+contaminated+abdominal+wall+reconstructions.">Use of human acellular dermal matrix in complex and contaminated abdominal wall reconstructions.</a> The Am J of Surg. 193 (3), 360-363 (2007).</li><li>Menon, N. G., 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=Revascularization+of+human+acellular+dermis+in+full-thickness+abdominal+wall+reconstruction+in+the+rabbit+model.">Revascularization of human acellular dermis in full-thickness abdominal wall reconstruction in the rabbit model.</a> Ann Plast Surg. 50 (5), 523-527 (2003).</li><li>Buinewicz, B., Rosen, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acellular+cadaveric+dermis+(AlloDerm):+a+new+alternative+for+abdominal+hernia+repair.">Acellular cadaveric dermis (AlloDerm): a new alternative for abdominal hernia repair.</a> Ann Plast Surg. 52 (2), 188-194 (2004).</li><li>Bringman, 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=Hernia+repair:+the+search+for+ideal+meshes.">Hernia repair: the search for ideal meshes.</a> Hernia. 14 (1), 81-87 (2010).</li><li>Meintjes, J., Yan, S., Zhou, L., Zheng, S., Zheng, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synthetic+biological+and+composite+scaffolds+for+abdominal+wall+reconstruction.">Synthetic biological and composite scaffolds for abdominal wall reconstruction.</a> Exp rev of med dev. 8 (2), 275-288 (2011).</li><li>Cheng, G., 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=Engineered+blood+vessel+networks+connect+to+host+vasculature+via+wrapping-and-tapping+anastomosis.">Engineered blood vessel networks connect to host vasculature via wrapping-and-tapping anastomosis.</a> Blood. 118 (17), 4740-4749 (2011).</li><li>Shandalov, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+engineered+muscle+flap+for+reconstruction+of+large+soft+tissue+defects.">An engineered muscle flap for reconstruction of large soft tissue defects.</a> PNAS of the USA. 111 (16), 6010-6015 (2014).</li><li>Zhang, T. Y., Suen, C. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+fast+parallel+algorithm+for+thinning+digital+patterns.">A fast parallel algorithm for thinning digital patterns.</a> Commun. ACM. 27 (3), 236-239 (1984).</li><li>Luna, L. G., Luna, L. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Manual of Histo Stain Meth ; of the Arm Forcs Inst of Path. , (1968).</li><li>Choi, J. 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=Adipose+tissue+engineering+for+soft+tissue+regeneration.">Adipose tissue engineering for soft tissue regeneration.</a> Tissue engineering. Part B, Reviews. 16 (4), 413-426 (2010).</li><li>Bellows, C. F., Alder, A., Helton, W. 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 border="0" cellpadding="0" cellspacing="0">
	<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&#160;</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&#160;</td>
			<td>Fine Science Tools (FST)</td>
			<td>18000-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Duratears</td>
			<td>Alcon</td>
			<td align="right">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 align="right">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 align="right">684568</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin syringe (6mm needle)</td>
			<td>BD</td>
			<td align="right">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 align="right">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&#160;</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&#160;</td>
			<td>016-030-084</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayer&#39;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&#160;</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

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 ...

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 ...

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic bioprinting to generate a scaffold with multilayer interlacing hydrogel microfibers. To achieve smooth bioprinting, a core-sheath microfluidic printhead containing a composite bioink formulation extruded from the core flow and the crosslinking solution carried by the sheath flow, was designed and fitted onto the bioprinter. By blending gelatin methacryloyl (GelMA) with alginate, a polysaccharide that undergoes instantaneous ionic crosslinking in the presence of select divalent ions, followed by a secondary photocrosslinking of the GelMA component to achieve permanent stabilization, a microfibrous scaffold could be obtained using this bioprinting strategy. Importantly, the endothelial cells encapsulated inside the bioprinted microfibers can form the lumen-like structures resembling the vasculature over the course of culture for 16 days. The endothelialized microfibrous scaffold may be further used as a vascular bed to construct a vascularized tissue through subsequent seeding of the secondary cell type into the interstitial space of the microfibers. Microfluidic bioprinting provides a generalized strategy in convenient engineering of vascularized tissues at high fidelity.

We provide a generalized protocol based on a microfluidic bioprinting strategy for engineering a microfibrous vascular bed, where a secondary cell type could be further seeded into the interstitial space of this microfibrous structure to generate vascularized tissues and organoids.

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic ...

Tissue-Engineered Graft for Circumferential Esophageal Reconstruction in Rats

The use of biocompatible materials for circumferential esophageal reconstruction is a technically challenging task in rats and requires an optimal implant technique with nutritional support. Recently, there have been many attempts at esophageal tissue engineering, but the success rate has been limited due to difficulty in early epithelization in the special environment of peristalsis. Here, we developed an artificial esophagus that can improve the regeneration of the esophageal mucosa and muscle layers through a two-layered tubular scaffold, a mesenchymal stem cell-based bioreactor system, and a bypass feeding technique with modified gastrostomy. The scaffold is made of polyurethane (PU) nanofibers in a cylindrical shape with a three-dimensional (3D) printed polycaprolactone strand wrapped around the outer wall. Prior to transplantation, human-derived mesenchymal stem cells were seeded into the lumen of the scaffold, and bioreactor cultivation was performed to enhance cellular reactivity. We improved the graft survival rate by applying surgical anastomosis and covering the implanted prosthesis with a thyroid gland flap, followed by temporary nonoral gastrostomy feeding. These grafts were able to recapitulate the findings of initial epithelialization and muscle regeneration around the implanted sites, as demonstrated by histological analysis. In addition, increased elastin fibers and neovascularization were observed in the periphery of the graft. Therefore, this model presents a potential new technique for circumferential esophageal reconstruction.

Esophageal reconstruction is a challenging procedure, and development of a tissue-engineered esophagus that enables regeneration of esophageal mucosa and muscle and that can be implanted as an artificial graft is necessary. Here, we present our protocol to generate an artificial esophagus, including scaffold manufacturing, bioreactor cultivation, and various surgical techniques.

The use of biocompatible materials for circumferential esophageal reconstruction is a technically challenging task in rats and requires an optimal implant ...

Fabrication of Zero Mode Waveguides for High Concentration Single Molecule Microscopy

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to nanomolar ranges, several orders of magnitude less than many physiological ligand concentrations. Optical nanostructures called zero mode waveguides (ZMWs), which are 100&#8722;200 nm in diameter apertures fabricated in a thin conducting metal such as aluminum or gold, allow imaging of individual molecules at micromolar concentrations of fluorophores by confining visible light excitation to zeptoliter effective volumes. However, the need for expensive and specialized nanofabrication equipment has precluded the widespread use of ZMWs. Typically, nanostructures such as ZMWs are obtained by direct writing using electron beam lithography, which is sequential and slow. Here, colloidal, or nanosphere, lithography is used as an alternative strategy to create nanometer-scale masks for waveguide fabrication. This report describes the approach in detail, with practical considerations for each phase. The method allows thousands of aluminum or gold ZMWs to be made in parallel, with final waveguide diameters and depths of 100&#8722;200 nm. Only common lab equipment and a thermal evaporator for metal deposition are required. By making ZMWs more accessible to the biochemical community, this method can facilitate the study of molecular processes at cellular concentrations and rates.

Described here is a nanosphere lithography method for parallel fabrication of zero mode waveguides, which are arrays of nanoapertures in a metal-clad glass microscopy coverslip for single molecule imaging at nano- to micromolar concentrations of fluorophores. The method takes advantage of colloidal crystal self-assembly to create a waveguide template.

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to ...

Generation of Self-assembled Vascularized Human Skin Equivalents

Human skin equivalents (HSEs) are tissue engineered constructs that model epidermal and dermal components of human skin. These models have been used to study skin development, wound healing, and grafting techniques. Many HSEs continue to lack vasculature and are additionally analyzed through post-culture histological sectioning which limits volumetric assessment of the structure. Presented here is a straightforward protocol utilizing accessible materials to generate vascularized human skin equivalents (VHSE); further described are volumetric imaging and quantification techniques of these constructs. Briefly, VHSEs are constructed in 12 well culture inserts in which dermal and epidermal cells are seeded into rat tail collagen type I gel. The dermal compartment is made up of fibroblast and endothelial cells dispersed throughout collagen gel. The epidermal compartment is made up of keratinocytes (skin epithelial cells) that differentiate at the air-liquid interface. Importantly, these methods are customizable based on needs of the researcher, with results demonstrating VHSE generation with two different fibroblast cell types: human dermal fibroblasts (hDF) and human lung fibroblasts (IMR90s). VHSEs were developed, imaged through confocal microscopy, and volumetrically analyzed using computational software at 4- and 8-week timepoints. An optimized process to fix, stain, image, and clear VHSEs for volumetric examination is described. This comprehensive model, imaging, and analysis techniques are readily customizable to the specific research needs of individual labs with or without prior HSE experience.

The goal of this protocol is to describe the generation and volumetric analysis of vascularized human skin equivalents using accessible and simple techniques for long term culture. To the extent possible, the rationale for steps is described to allow researchers the ability to customize based on their research needs.

Human skin equivalents (HSEs) are tissue engineered constructs that model epidermal and dermal components of human skin. These models have been used to ...

Assessing Functional Metrics of Skeletal Muscle Health in Human Skeletal Muscle Microtissues

Three-dimensional (3D) in vitro models of skeletal muscle are a valuable advancement in biomedical research as they afford the opportunity to study skeletal muscle reformation and function in a scalable format that is amenable to experimental manipulations. 3D muscle culture systems are desirable as they enable scientists to study skeletal muscle ex vivo in the context of human cells. 3D in vitro&#160;models closely mimic aspects of the native tissue structure of adult skeletal muscle. However, their universal application is limited by the availability of platforms that are simple to fabricate, cost and user-friendly, and yield relatively high quantities of human skeletal muscle tissues. Additionally, since skeletal muscle plays an important functional role that is impaired over time in many disease states, an experimental platform for microtissue studies is most practical when minimally invasive calcium transient and contractile force measurements can be conducted directly within the platform itself. In this protocol, the fabrication of a 96-well platform known as &#39;MyoTACTIC&#39;, and en masse production of 3D human skeletal muscle microtissues (hMMTs) is described. In addition, the methods for a minimally invasive application of electrical stimulation that enables repeated measurements of skeletal muscle force and calcium handling of each microtissue over time are reported.

This manuscript describes a detailed protocol to produce arrays of 3D human skeletal muscle microtissues and minimally invasive downstream in situ assays of function, including contractile force and calcium handling analyses.

Three-dimensional (3D) in vitro models of skeletal muscle are a valuable advancement in biomedical research as they afford the opportunity to study ...

Engineered Lung Tissues Prepared from Decellularized Lung Slices

There is a need for improved 3-dimensional (3D) lung models that recapitulate the architectural and cellular complexity of the native lung alveolus ex vivo. Recently developed organoid models have facilitated the expansion and study of lung epithelial progenitors in vitro, but these platforms typically rely on mouse tumor-derived matrix and/or serum, and incorporate just one or two cellular lineages. Here, we describe a protocol for generating engineered lung tissues (ELTs) based on the multi-lineage recellularization of decellularized precision-cut lung slices (PCLS). ELTs contain alveolar-like structures comprising alveolar epithelium, mesenchyme, and endothelium, within an extracellular matrix (ECM) substrate closely resembling that of native lung. To generate the tissues, rat lungs are inflated with agarose, sliced into 450 &#181;m-thick slices, cut into strips, and decellularized. The resulting acellular ECM scaffolds are then reseeded with primary endothelial cells, fibroblasts, and alveolar epithelial type 2 cells (AEC2s). AEC2s can be maintained in ELT culture for at least 7 days with a serum-free, chemically-defined growth medium. Throughout the tissue preparation and culture process, the slices are clipped into a cassette system that facilitates handling and standardized cell seeding of multiple ELTs in parallel. These ELTs represent an organotypic culture platform that should facilitate investigations of cell-cell and cell-matrix interactions within the alveolus as well as biochemical signals regulating AEC2s and their niche.

This protocol describes a method to generate reproducible, small-scale engineered lung tissues, by repopulating decellularized precision-cut lung slices with alveolar epithelial type 2 cells, fibroblasts, and endothelial cells.

There is a need for improved 3-dimensional (3D) lung models that recapitulate the architectural and cellular complexity of the native lung alveolus ex ...

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, ...

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 ...