Figure 3A was created in BioRender.com.

Cadaveric male Lewis rats (300-430 g) obtained from the Toronto General Hospital Research Institute were used for all experiments. For all surgical procedures, sterile instruments and supplies were used to maintain aseptic technique (see the Table of Materials). All procedures were performed in compliance with guidelines from the Animal Care Committee at Toronto General Hospital Research Institute, University Health Network (Toronto, ON, Canada). A total of four hindlimbs were decellularized.
1. Presurgical preparation
<ol>
	<li>Prepare 50 mL of 5% heparinized saline. From a 50 mL saline bag, take out 2.5 mL of saline solution using a 5 mL syringe and discard. Using a 5 mL syringe, add 2.5 mL of heparin to the saline bag. Invert the saline bag to mix its contents.</li>
	<li>Place a cadaveric rat in a supine position under a blue pad. Shave the hindlimb and groin area circumferentially using an electric shaver and remove hair.</li>
	<li>Bring the rat to the surgical station and apply Povidone iodine scrub solution using a gauze to the hindlimb and groin area. Subsequently, apply 70% isopropyl alcohol to wipe off the scrub solution using gauze.</li>
	<li>Discard the blue pad from step 1.2 and change into new, sterile gloves.</li>
</ol>
2. Procurement of rat hindlimb
<ol>
	<li>Make a skin incision using a #10 surgical blade and a #3 blade runner along the inguinal ligament level, moving from a lateral to a medial direction (Figure 1A). Use Adson forceps to hold the surrounding skin to ensure a smooth incision.</li>
	<li>When the underlying fat is exposed, use blunt dissection to carefully dissect through the fat. Locate the superior epigastric vessels.</li>
	<li>Use microscissors to dissect proximally and expose the underlying femoral nerve, artery, and vein at the inguinal ligament level.</li>
	<li>Under a dissection microscope, identify the femoral vessels and dissect the artery and vein proximally using fine forceps to obtain sufficient length from the bifurcation points of the arterial network (Figure 1B).</li>
	<li>Ligate the femoral artery and vein separately using 6-0 sutures.</li>
	<li>Conduct circumferential dissection around the remainder of the hindlimb without disrupting the ligated femoral vessels.</li>
	<li>Transect the femoral bone mid-length using a bone cutter.</li>
	<li>To fully isolate the hindlimb, transect the ligated femoral vessels below the ligatures using microscissors.</li>
	<li>Cannulate the femoral artery using a 24 G angiocatheter under the dissection microscope. Use fine forceps to insert the cannula carefully. Flush with heparinized saline until clear outflow is observed from the femoral vein.</li>
	<li>Secure the cannula by tying one suture around the cannulated vessel and another suture distally around the cannula itself. Ensure the cannula is placed proximally to prevent blocking the bifurcation points.</li>
	<li>Submerge the procured hindlimbs in phosphate-buffered saline (PBS) until decellularization.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64069/64069fig01.jpg" /> 
Figure 1: Procurement of rat hindlimb. (A) Marking of skin incision at the inguinal ligament level from lateral to medial. (B) View of the femoral vein and the femoral artery, which have been dissected proximally toward the inguinal ligament, indicated by the dotted line. Abbreviations: L = lateral; M = medial; FV = femoral vein; FA = femoral artery. <a href="https://www.jove.com/files/ftp_upload/64069/64069fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Preparation of solutions
<ol>
	<li>In a 6 L glass flask, prepare a 5 L detergent reservoir of 0.25% sodium dodecyl sulfate (SDS) by dissolving 12.5 g of SDS powder in 5 L of ultrapure distilled water. Cover the opening of the flask with parafilm to seal it.</li>
	<li>In a 1 L glass jar, prepare 1 L of 0.25% SDS solution separately by dissolving 2.5 g of SDS in 1 L of ultrapure distilled water. Add a stir bar to mix the solution on a magnetic stirrer until all the SDS is dissolved.</li>
	<li>Prepare 1 L of 1x PBS wash solution with 1% Antibiotic-Antimycotic (AA) solution. In a 1 L flask, add 990 mL of 1x PBS solution and 10 mL of AA.</li>
	<li>Separately, in a 500 mL glass jar, prepare 1x PBS + 1% AA solution again using 495 mL of 1x PBS solution and 5 mL of AA.</li>
	<li>Prepare 200 mL of 1% peracetic acid (PAA)/4% ethanol (EtOH) solution. Prepare this solution under a fume hood.</li>
</ol>
4. Bioreactor and perfusion circuit construction
NOTE: Refer to Figure 2 for the configuration of the bioreactor and perfusion circuit throughout the listed steps.
<ol>
	<li>In a 500 mL chamber (plastic container), drill three holes (1/4 in) at the labeled locations in Figure 2: Port A is the inlet line, Port B is the replenishing line, and Port C is the outlet line. Remove and discard excess plastic. Spray and wipe the chamber with 70% ethanol.</li>
	<li>Cut three 5 cm long silicone tubes and insert one halfway into each port of the chamber. Connect a male Luer connector on the opening of Port A facing the inside of the chamber and a female Luer connector on the other end of the tube outside of the chamber.</li>
	<li>Connect a female Luer connector at Port B and Port C on the ends of the tubes facing out of the chamber.</li>
	<li>In an air-tight lid for the plastic container, drill one hole on the surface of the lid.</li>
	<li>Cut a 3 cm silicone tube and insert it into the hole of the lid. Ensure approximately 2 cm of the tube is located out of the lid, as shown in Figure 2.</li>
	<li>Sterilize both the chamber and the lid with the tubing.</li>
	<li>Cut two 30 cm silicone tubes using scissors. Connect a 1/16 in to a 1/8 in connector on one end of each tube.</li>
	<li>Separately, cut two 12 cm silicone tubes using scissors. Connect a 1/16 in to a 1/8 in connector on one end of both tubes. Connect a male Luer connector on the other end of one tube and a female Luer connector to the other tube.</li>
	<li>Sterilize all the tubing material from step 4.7 and step 4.8. Include two 3-stop pump tubings (1.85 mm) in the sterilization.</li>
	<li>Prepare three 3-way stopcocks, one syringe filter, two 1 mL serological pipettes, and one 10 mL syringe for the decellularization setup. Remove the filter from the 1 mL serological pipettes.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64069/64069fig02.jpg" /> 
Figure 2: Preparation of bioreactor and perfusion circuit construction. Apparatus shown of the perfusion circuit including (A) peristaltic pump and (B) corresponding cassettes for both inlet and outlet lines. (C, D) Silicone tubings of 12 cm and 30 cm are also shown with respective connectors. (E) Tubing for peristaltic pump (1.85 mm). Bioreactor chamber with labeled ports for (F) inflow, (G) replenishing port, and (H) outflow. (I) Bioreactor lid shown with ventilation port. <a href="https://www.jove.com/files/ftp_upload/64069/64069fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Decellularization of rat hindlimbs
<ol>
	<li>Place the sterilized chamber and screw on three single-use 3-way stopcocks at the inlet, outlet, and replenishing lines. Ensure the stopcock for the replenishing port is capped at its remaining two ports to prevent leakage.</li>
	<li>Attach the tubes made in step 4.8 to the stopcocks at the inlet and outlet lines.</li>
	<li>Connect peristaltic tubing to the tubes in the step above. Secure the cassette on the peristaltic tubing and place it on the peristaltic pump. Do not secure the cassettes with tubing in place yet.</li>
	<li>Connect one tube from step 4.7 to the end of the peristaltic tubing of the outlet line from the step above. Connect a 1 mL serological pipette on the other end. Suspend the end attached with a serological pipette in the waste reservoir flask.</li>
	<li>Repeat step 4.7 for the inlet line. Suspend the end of the tube attached to the serological pipette into the detergent reservoir. Seal the opening of the detergent reservoir flask with parafilm immediately. See Figure 3A,B for an overview of the decellularization circuit.</li>
	<li>Add 0.25% SDS from the 1 L glass jar (step 3.2) into the bioreactor chamber at the halfway level.</li>
	<li>Take the procured hindlimb using Adson forceps and suspend it into the bioreactor chamber carefully.</li>
	<li>Use two pairs of Adson forceps to guide the cannulated portion of the hindlimb to the inlet line. While holding the cannula with one pair of forceps, use the other pair of forceps to twist and secure the inlet line to the cannula. Ensure the cannula is not pulled or twisted to prevent decannulation.</li>
	<li>Once secured, add more 0.25% SDS from the 1 L glass jar to fully submerge the limb, as needed. Ensure the outlet port is also submerged in the bioreactor reservoir to ensure that consistent outflow is maintained (Figure 3C).</li>
	<li>Attach a single-use syringe filter to the ventilation port on the lid of the bioreactor chamber, referring to step 4.4 and step 4.5.</li>
	<li>Secure the lid on the bioreactor and ensure the chamber is sealed from all sides (Figure 3B).</li>
	<li>To remove air from tubing and prime the perfusion circuit, use a new, single-use 10 mL syringe to draw detergent from the detergent reservoir using the 3-way stopcock at the inlet line.</li>
	<li>Once drawn, use the same fluid to insert into the 3-way stopcock at the outlet line. Ensure that there is detergent present throughout the tubing at both the inlet and outlet lines.</li>
	<li>Press down and secure the cassettes with the tubing into the peristaltic pump. Turn the peristaltic pump on using the power button.
	<ol>
		<li>On the peristaltic pump screen, proceed to the second tab using the arrow key to set the perfusion rate for the first channel. Input flow rate as the mode of delivery and set the perfusion rate at 1 mL/min. Ensure the direction of flow is correct according to apparatus set-up. Repeat for the second channel.</li>
		<li>Calibrate the peristaltic pump to ensure the amount of fluid delivered through the inlet line and/or taken from the outlet line is flowing at a consistent rate between the two. Ensure that the tubing ID is set to 1.85 mm.</li>
	</ol>
	</li>
	<li>Begin decellularization via machine perfusion at 1 mL/min for both the inlet and outlet lines by pressing the power button on the keypad. Monitor and ensure that the flow is consistent and ongoing at both the inlet and outlet lines.</li>
	<li>Continue decellularization and monitor daily. Use the 1 L of 0.25% SDS (step 3.2) to replenish the bioreactor reservoir through the replenishing port, as needed. Look for a white, translucent appearance of the tissue, which will appear by day 5, indicating decellularization of the rat hindlimbs.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64069/64069fig03.jpg" /> 
Figure 3: Overview of perfusion decellularization bioreactor circuit of rat hindlimb. (A) Schematic representation of bioreactor perfusion circuit. Blue arrows indicate the direction of detergent and waste flow. (B) Overview of the decellularization circuit with bioreactor containing rat hindlimb. The SDS reservoir (left flask) leads into the peristaltic pump and into the inlet tubing of the bioreactor. The outflow is connected to the waste reservoir (right flask) through the peristaltic pump. (C) (I) Bioreactor containing rat hindlimb with inlet tubing connected to the cannulated femoral artery. (II) Replenishing port located in the corner for perfusing detergent. (III) Outflow tubing suspended in suspension reservoir. Abbreviation: SDS = sodium dodecyl sulfate. <a href="https://www.jove.com/files/ftp_upload/64069/64069fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Post-decellularization washing and sterilization
<ol>
	<li>Following the confirmation of decellularization, replace the detergent reservoir with the 1x PBS + 1% AA reservoir. Seal the opening of the flask with parafilm. Begin 1x PBS + 1% AA perfusion at 1 mL/min and continue for 2 days.</li>
	<li>Replace the 1x PBS + 1% AA reservoir with 200 mL of 0.1% PAA/4% EtOH reservoir and begin perfusion at 1 mL/min for 2 h.</li>
	<li>Disconnect the hindlimb from the inlet line using two pairs of sterile Adson forceps, with one pair of forceps to twist the inlet line and the other holding the cannula. Ensure that the cannula is not pulled to prevent decannulation.</li>
	<li>Store the limb in a 500 mL glass jar containing 1x PBS + 1% AA at 4 &#176;C until further use.</li>
</ol>

<ol>
	<li>Kueckelhaus, 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=Vascularized+composite+allotransplantation:+Current+standards+and+novel+approaches+to+prevent+acute+rejection+and+chronic+allograft+deterioration.">Vascularized composite allotransplantation: Current standards and novel approaches to prevent acute rejection and chronic allograft deterioration.</a> Transplant International. 29 (6), 655-662 (2016).</li><li>Duisit, 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=Bioengineering+a+human+face+graft:+The+matrix+of+identity.">Bioengineering a human face graft: The matrix of identity.</a> Annals of Surgery. 266 (5), 754-764 (2017).</li><li>Zhang, Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decellularized+skin/adipose+tissue+flap+matrix+for+engineering+vascularized+composite+soft+tissue+flaps.">Decellularized skin/adipose tissue flap matrix for engineering vascularized composite soft tissue flaps.</a> Acta Biomaterialia. 35, 166-184 (2016).</li><li>Londono, R., Gorantla, V. S., Badylak, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emerging+implications+for+extracellular+matrix-based+technologies+in+vascularized+composite+allotransplantation.">Emerging implications for extracellular matrix-based technologies in vascularized composite allotransplantation.</a> Stem Cells International. 2016 (10), 1-16 (2016).</li><li>Fleissig, Y. Y., Beare, J. E., LeBlanc, A. J., Kaufman, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolution+of+the+rat+hind+limb+transplant+as+an+experimental+model+of+vascularized+composite+allotransplantation:+Approaches+and+advantages.">Evolution of the rat hind limb transplant as an experimental model of vascularized composite allotransplantation: Approaches and advantages.</a> SAGE Open Medicine. 8, 205031212096871 (2020).</li><li>Tao, 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=Sterilization+and+disinfection+methods+for+decellularized+matrix+materials:+Review,+consideration+and+proposal.">Sterilization and disinfection methods for decellularized matrix materials: Review, consideration and proposal.</a> Bioactive Materials. 6 (9), 2927-2945 (2021).</li><li>Chen, Y., Geerts, S., Jaramillo, M., Uygun, B. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+decellularized+liver+scaffolds+and+recellularized+liver+grafts.">Preparation of decellularized liver scaffolds and recellularized liver grafts.</a> Methods in Molecular Biology. 1577, 255-270 (2018).</li><li>Ahmed, S., Chauhan, V. M., Ghaemmaghami, A. M., Aylott, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+generation+of+bioreactors+that+advance+extracellular+matrix+modelling+and+tissue+engineering.">New generation of bioreactors that advance extracellular matrix modelling and tissue engineering.</a> Biotechnology Letters. 41 (1), 1-25 (2019).</li><li>Cohen, 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=Generation+of+vascular+chimerism+within+donor+organs.">Generation of vascular chimerism within donor organs.</a> Scientific Reports. 11 (1), 13437 (2021).</li><li>Lupon, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+vascularized+composite+allografts+using+natural+scaffolds:+A+systematic+review.">Engineering vascularized composite allografts using natural scaffolds: A systematic review.</a> Tissue Engineering Part B: Reviews. , (2021).</li><li>Urciuolo, 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=Decellularised+skeletal+muscles+allow+functional+muscle+regeneration+by+promoting+host+cell+migration.">Decellularised skeletal muscles allow functional muscle regeneration by promoting host cell migration.</a> Scientific Reports. 8 (1), 8398 (2018).</li><li>Jank, B. 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The authors have no conflicts of interest to declare.

Rat hindlimbs are useful as experimental models in VCA5. Tissue engineering of acellular scaffolds represents the first step in addressing the shortcomings of long-term immunosuppression regimens associated with VCA. The use of composite grafts poses an added challenge given the presence of multiple tissues, each having unique functional, immunogenic, and structural properties. The present protocol shows a successful method for obtaining acellular composite rat hindlimbs. These scaffolds can be fu...

VCA is an emerging option for patients requiring complex surgical reconstruction. Traumatic injuries or tumor resections result in volumetric tissue loss that can be difficult to reconstruct. VCA offers the transplantation of multiple tissues such as the skin, bone, muscle, nerves, and vessels as a composite graft from a donor to a recipient1. Despite its promising nature, VCA is limited due to long-term immunosuppressive regimens. Lifelong use of such drugs results in increased risk for opportunistic infections, malignancies, and end-organ toxicity1,2,3. To help reduce and/or eliminate the need for immunosuppression, tissue-engineered scaffolds using decellularization approaches for VCA show great promise.
Tissue decellularization entails retaining the extracellular matrix structure while removing the cellular and nuclear contents. This decellularized scaffold can be repopulated with patient-specific cells4. However, preserving the ECM network of composite tissues is an added challenge. This is due to the presence of multiple tissue types with varying tissue densities, architectures, and anatomic locations within a scaffold. The present protocol offers a surgical technique and a decellularization method for a rat hindlimb. This is a proof-of-concept model for applying this tissue engineering technique to composite tissues. This can also prompt subsequent efforts to regenerate composite tissues through recellularization.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.9% Sodium Chloride Injection USP 50 mL</td>
			<td>Baxter Corporation</td>
			<td>JB1308M</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL Disposable Serological Pipets</td>
			<td>VWR</td>
			<td>75816-102</td>
			<td></td>
		</tr>
		<tr>
			<td>10 cc Disposable Syringes</td>
			<td></td>
			<td></td>
			<td>Obtained from Research Institution</td>
		</tr>
		<tr>
			<td>3-way Stopcock</td>
			<td></td>
			<td></td>
			<td>Obtained from Research Institution</td>
		</tr>
		<tr>
			<td>5cc Disposable Syringes</td>
			<td></td>
			<td></td>
			<td>Obtained from Research Institution</td>
		</tr>
		<tr>
			<td>70% Isopropyl Alcohol</td>
			<td></td>
			<td></td>
			<td>Obtained from Research Institution</td>
		</tr>
		<tr>
			<td>Acrodisc Syringe Filter 0.2 &#181;m</td>
			<td>VWR</td>
			<td>CA28143-310</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson Forceps, Straight</td>
			<td>Fine Science Tools</td>
			<td>11006-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Angiocatheter 24 G 19 mm (&#190;&#8221;)&#160;</td>
			<td>VWR</td>
			<td>38112</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic Solution (100x) 100 mL</td>
			<td>Multicell</td>
			<td>450-115-EL</td>
			<td></td>
		</tr>
		<tr>
			<td>Bone Cutter</td>
			<td>Fine Science Tools</td>
			<td>12029-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Connectors for&#160; 1/16&#34; to 1/8&#34; Tubes</td>
			<td>McMasterCarr</td>
			<td>5117K52</td>
			<td></td>
		</tr>
		<tr>
			<td>Female Luer to barbed adapter (PVDF) - 1/8&#34; ID</td>
			<td>McMasterCarr</td>
			<td>51525K328</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Forceps</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Forceps with Micro-Blunted Tips</td>
			<td>Fine Science Tools</td>
			<td>11253-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin Sodium Injection 10,000 IU/10 mL</td>
			<td>LEO Pharma Inc.</td>
			<td>006174-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Male Luer to barbed adapter (PVDF) - 1/8&#34; ID</td>
			<td>McMasterCarr</td>
			<td>51525K322</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Needle Holder</td>
			<td>WLorenz</td>
			<td>04-4125</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscissors</td>
			<td>WLorenz</td>
			<td>SP-4506</td>
			<td></td>
		</tr>
		<tr>
			<td>Peracetic Acid</td>
			<td>Sigma Aldrich</td>
			<td>269336-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic Pump, 3-Channel</td>
			<td>Cole Parmer</td>
			<td>RK-78001-68</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline 1x 500 mL</td>
			<td>Wisent</td>
			<td>311-425-CL</td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone Surgical Scrub Solution</td>
			<td></td>
			<td></td>
			<td>Obtained from Research Institution</td>
		</tr>
		<tr>
			<td>Pump Tubing, 3-Stop, Tygon E-LFL</td>
			<td>Cole Parmer</td>
			<td>RK-96450-40</td>
			<td></td>
		</tr>
		<tr>
			<td>Pump Tubing, Platinum-Cured Silicone</td>
			<td>Cole Parmer</td>
			<td>RK-96410-16</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Blade - #10</td>
			<td>Fine Science Tools</td>
			<td>10010-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Handle - #3&#160;</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Dodecyl Sulfate Reagent Grade: Purity: &#62;99%, 1 kg</td>
			<td>Bioshop</td>
			<td>SDS003.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Suture #6-0</td>
			<td>Covidien</td>
			<td>VS889</td>
			<td></td>
		</tr>
	</tbody>
</table>

procurement and decellularization of rat hindlimbs using an ex vivo perfusion-based bioreactor for vascularized composite allotransplantation

Patients with severe traumatic injuries and tissue loss require complex surgical reconstruction. Vascularized composite allotransplantation (VCA) is an evolving reconstructive avenue for transferring multiple tissues as a composite subunit. Despite the promising nature of VCA, the long-term immunosuppressive requirements are a significant limitation due to the increased risk of malignancies, end-organ toxicity, and opportunistic infections. Tissue engineering of acellular composite scaffolds is a potential alternative in reducing the need for immunosuppression. Herein, the procurement of a rat hindlimb and its subsequent decellularization using sodium dodecyl sulfate (SDS) is described. The procurement strategy presented is based upon the common femoral artery. A machine perfusion-based bioreactor system was constructed and used for ex vivo decellularization of the hindlimb. Successful perfusion decellularization was performed, resulting in a white translucent-like appearance of the hindlimb. An intact, perfusable, vascular network throughout the hindlimb was observed. Histological analyses showed the removal of nuclear contents and the preservation of tissue architecture across all tissue compartments.

The procurement protocol was successful in isolating and cannulating the common femoral arteries for subsequent perfusion steps. The representative dissection images in Figure 1A,B show the incision location and exposure of the femoral vessels with sufficient distance from the bifurcation points. Figure 2 shows the apparatus required for preparing the bioreactor and perfusion circuit.&#160;The endpoint of decellularization was determined by obse...

Watch this Scientific Journal Video about Procurement and Decellularization of Rat Hindlimbs Using an Ex Vivo Perfusion-Based Bioreactor for Vascularized Composite Allotransplantation at JoVE.com

Procurement and Decellularization of Rat Hindlimbs Using an Ex Vivo Perfusion-Based Bioreactor for Vascularized Composite Allotransplantation

We describe the surgical technique and decellularization process for composite rat hindlimbs. Decellularization is conducted using low-concentration sodium dodecyl sulfate through an ex vivo machine perfusion system.

Procurement and Decellularization of Rat Hindlimbs Using an Ex Vivo Perfusion-Based Bioreactor for Vascularized Composite Allotransplantation

Patients with severe traumatic injuries and tissue loss require complex surgical reconstruction. Vascularized composite allotransplantation (VCA) is an ...

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JoVE Journal

Bioengineering

2.1K Views.  University Health Network, Toronto General Hospital. We describe the surgical technique and decellularization process for composite rat hindlimbs. Decellularization is conducted using low-concentration sodium dodecyl sulfate through an ex vivo machine perfusion system. 

Video: Procurement and Decellularization of Rat Hindlimbs Using an Ex Vivo Perfusion-Based Bioreactor for Vascularized Composite Allotransplantation

Decellularization and Recellularization of Whole Livers

The liver is a complex organ which requires constant perfusion for delivery of nutrients and oxygen and removal of waste in order to survive1. Efforts to recreate or mimic the liver microstructure with grounds up approach using tissue engineering and microfabrication techniques have not been successful so far due to this design challenge. In addition, synthetic biomaterials used to create scaffolds for liver tissue engineering applications have been limited in inducing tissue regeneration and repair in large part due to the lack of specific cell binding motifs that would induce the proper cell functions2. Decellularized native tissues such blood vessels3and skin4on the other hand have found many applications in tissue engineering, and have provided a practical solution to some of the challenges. The advantage of decellularized native matrix is that it retains, to an extent, the original composition, and the microstructure, hence enhancing cell attachment and reorganization5.
In this work we describe the methods to perform perfusion-decellularization of the liver, such that an intact liver bioscaffold that retains the structure of major blood vessels is obtained. Further, we describe methods to recellularize these bioscaffolds with adult primary hepatocytes, creating a liver graft that is functional in vitro, and has the vessel access necessary for transplantation in vivo.

Perfusion decellularization is a novel technique to produce whole liver scaffolds that retains the organ's extracellular matrix composition and microarchitecture. Herein, the method of preparing whole organ scaffolds using perfusion decellularization and subsequent repopulation with hepatocytes is described. Functional and transplantable liver grafts can be generated using this technique.

The liver is a complex organ which requires constant perfusion for delivery of nutrients and oxygen and removal of waste in order to survive1. Efforts to ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Determination of the Transport Rate of Xenobiotics and Nanomaterials Across the Placenta using the ex vivo Human Placental Perfusion Model

Decades ago the human placenta was thought to be an impenetrable barrier between mother and unborn child. However, the discovery of thalidomide-induced birth defects and many later studies afterwards proved the opposite. Today several harmful xenobiotics like nicotine, heroin, methadone or drugs as well as environmental pollutants were described to overcome this barrier. With the growing use of nanotechnology, the placenta is likely to come into contact with novel nanoparticles either accidentally through exposure or intentionally in the case of potential nanomedical applications. Data from animal experiments cannot be extrapolated to humans because the placenta is the most species-specific mammalian organ 1. Therefore, the ex vivo dual recirculating human placental perfusion, developed by Panigel et al. in 1967 2 and continuously modified by Schneider et al. in 1972 3, can serve as an excellent model to study the transfer of xenobiotics or particles.
Here, we focus on the ex vivo dual recirculating human placental perfusion protocol and its further development to acquire reproducible results.
The placentae were obtained after informed consent of the mothers from uncomplicated term pregnancies undergoing caesarean delivery. The fetal and maternal vessels of an intact cotyledon were cannulated and perfused at least for five hours. As a model particle fluorescently labelled polystyrene particles with sizes of 80 and 500 nm in diameter were added to the maternal circuit. The 80 nm particles were able to cross the placental barrier and provide a perfect example for a substance which is transferred across the placenta to the fetus while the 500 nm particles were retained in the placental tissue or maternal circuit. The ex vivo human placental perfusion model is one of few models providing reliable information about the transport behavior of xenobiotics at an important tissue barrier which delivers predictive and clinical relevant data.

Determination of the Transport Rate of Xenobiotics and Nanomaterials Across the Placenta using the ex vivo Human Placental Perfusion Model

The ex vivo dual recirculating human placental perfusion model can be used to investigate the transfer of xenobiotics and nanoparticles across the human placenta. In this video protocol we describe the equipment and techniques required for a successful execution of a placenta perfusion.

Decades ago the human placenta was thought to be an impenetrable barrier between mother and unborn child. However, the discovery of thalidomide-induced ...

Operation of a Benchtop Bioreactor

Fermentation systems are used to provide an optimal growth environment for many different types of cell cultures. The ability afforded by fermentors to carefully control temperature, pH, and dissolved oxygen concentrations in particular makes them essential to efficient large scale growth and expression of fermentation products. This video will briefly describe the advantages of the fermentor over the shake flask. It will also identify key components of a typical benchtop fermentation system and give basic instruction on setup of the vessel and calibration of its probes. The viewer will be familiarized with the sterilization process and shown how to inoculate the growth medium in the vessel with culture. Basic concepts of operation, sampling, and harvesting will also be demonstrated. Simple data analysis and system cleanup will also be discussed.

Fermentors are used to increase culture yield and productivity of bioengineered cells. After screening multiple microbial or animal cell culture candidates in shake flasks, the next logical step is to increase the selected culture&#x2019;s biomass with the fermentor. This video demonstrates the setup and operation of a typical benchtop bioreactor system.

Fermentation systems are used to provide an optimal growth environment for many different types of cell cultures. The ability afforded by fermentors to ...

Nonhuman Primate Lung Decellularization and Recellularization Using a Specialized Large-organ Bioreactor

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an attractive alternative to classic organ transplantation. This technology utilizes an organ&#39;s natural biological extracellular matrix (ECM) as a scaffold onto which autologous or stem/progenitor cells may be seeded and cultured in such a way that facilitates regeneration of the original tissue. The natural ECM is isolated by a process called decellularization. Decellularization is accomplished by treating tissues with a series of detergents, salts, and enzymes to achieve effective removal of cellular material while leaving the ECM intact. Studies conducted utilizing decellularization and subsequent recellularization of rodent lungs demonstrated marginal success in generating pulmonary-like tissue which is capable of gas exchange in vivo. While offering essential proof-of-concept, rodent models are not directly translatable to human use. Nonhuman primates (NHP) offer a more suitable model in which to investigate the use of bioartificial organ production for eventual clinical use.
The protocols for achieving complete decellularization of lungs acquired from the NHP rhesus macaque are presented. The resulting acellular lungs can be seeded with a variety of cells including mesenchymal stem cells and endothelial cells. The manuscript also describes the development of a bioreactor system in which cell-seeded macaque lungs can be cultured under conditions of mechanical stretch and strain provided by negative pressure ventilation as well as pulsatile perfusion through the vasculature; these forces are known to direct differentiation along pulmonary and endothelial lineages, respectively. Representative results of decellularization and cell seeding are provided.

Whole-organ decellularization produces natural biological scaffolds that may be used for regenerative medicine. The description of a nonhuman primate model of lung regeneration in which whole lungs are decellularized and then seeded with adult stem cells and endothelial cells in a bioreactor that facilitates vascular circulation and liquid media ventilation is presented.

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an ...

Procedure for Decellularization of Rat Livers in an Oscillating-pressure Perfusion Device

Decellularization and recellularization of parenchymal organs may enable the generation of functional organs in vitro, and several protocols for rodent liver decellularization have already been published. We aimed to improve the decellularization process by construction of a proprietary perfusion device enabling selective perfusion via the portal vein and/or the hepatic artery. Furthermore, we sought to perform perfusion under oscillating surrounding pressure conditions to improve the homogeneity of decellularization. The homogeneity of perfusion decellularization has been an underestimated factor to date. During decellularization, areas within the organ that are poorly perfused may still contain cells, whereas the extracellular matrix (ECM) in well-perfused areas may already be affected by alkaline detergents. Oscillating pressure changes can mimic the intraabdominal pressure changes that occur during respiration to optimize microperfusion inside the liver. In the study presented here, decellularized rat liver matrices were analyzed by histological staining, DNA content analysis and corrosion casting. Perfusion via the hepatic artery showed more homogenous results than portal venous perfusion did. The application of oscillating pressure conditions improved the effectiveness of perfusion decellularization. Livers perfused via the hepatic artery and under oscillating pressure conditions showed the best results. The presented techniques for liver harvesting, cannulation and perfusion using our proprietary device enable sophisticated perfusion set-ups to improve decellularization and recellularization experiments in rat livers.

The presented techniques for liver harvesting, cannulation and perfusion using our proprietary device enable sophisticated perfusion set-ups to improve decellularization and recellularization experiments in rat livers.

Decellularization and recellularization of parenchymal organs may enable the generation of functional organs in vitro, and several protocols for rodent ...

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

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

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

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

Organ engineering is a novel strategy to generate liver organ substitutes that can potentially be used in transplantation. Recently, in vivo liver ...

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 ...

A Novel 3D Printed Bioreactor for Ex Vivo Culture of Blood Vessels in Perfusion

Ex Vivo Perfusion Culture of Large Blood Vessels in a 3D Printed Bioreactor

Vascular disease forms the basis of most cardiovascular diseases (CVDs), which remain the primary cause of mortality and morbidity worldwide. Efficacious surgical and pharmacological interventions to prevent and treat vascular disease are urgently needed. In part, the shortage of translational models limits the understanding of the cellular and molecular processes involved in vascular disease. Ex vivo perfusion culture bioreactors provide an ideal platform for the study of large animal vessels (including humans) in a controlled dynamic environment, combining the ease of in vitro culture and the complexity of the live tissue. Most bioreactors are, however, custom manufactured and therefore difficult to adopt, limiting the reproducibility of the results. This paper presents a 3D printed system that can be easily produced and applied in any biological lab, and provides a detailed protocol for its setup, enabling users&#39; operation. This innovative and reproducible ex vivo perfusion culture system enables the culture of blood vessels for up to 7 days in physiological conditions. We expect that adopting a standardized perfusion bioreactor will support a better understanding of physiological and pathological processes in large animal blood vessels and accelerate the discovery of new therapeutics.

Author Spotlight: EasyFlow - An Economical and Adaptable Perfusion Bioreactor for Large Blood Vessel Culture 

This protocol presents the setup and operation of a newly developed, 3D printed bioreactor for the ex vivo culture of blood vessels in perfusion. The system is designed to be easily adopted by other users, practical, affordable, and adaptable to different experimental applications, such as basic biology and pharmacological studies.

Vascular disease forms the basis of most cardiovascular diseases (CVDs), which remain the primary cause of mortality and morbidity worldwide. Efficacious ...

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Subtitles

Procurement and Decellularization of Rat Hindlimbs Using an Ex Vivo Perfusion-Based Bioreactor for Vascularized Composite Allotransplantation