Name: procurement and perfusion-decellularization of porcine vascularized flaps in a customized perfusion bioreactor
Uploaded: 2022-08-01T13:00:00
Description: Watch this Scientific Journal Video about Procurement and Perfusion-Decellularization of Porcine Vascularized Flaps in a Customized Perfusion Bioreactor at JoVE.com

All procedures involving animal subjects have been approved by the University Health Network Institutional Animal Care and Use Committee (IACUC) and are performed in accordance with University Health Network Animal Resource Centre protocol and procedures and Canadian Council on Animal Care Guidelines.&#160;Five Yorkshire pigs (35-50 kg; age approximately 12 weeks old) were used for all experiments.
1. Perfusion bioreactor fabrication
<ol>
	<li>See Figure 1 for all t...

<ol>
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Global Open. 6 (2), 1651 (2018).</li><li>Issa, F. <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+allograft-specific+characteristics+of+immune+responses.">Vascularized composite allograft-specific characteristics of immune responses.</a> Transplant International. 29 (6), 672-681 (2016).</li><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>Iske, 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=Composite+tissue+allotransplantation:+Opportunities+and+challenges.">Composite tissue allotransplantation: Opportunities and challenges.</a> Cellular and Molecular Immunology. 16 (4), 343-349 (2019).</li><li>Londono, R., Gorantla, V. 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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vascularization+in+tissue+engineering.">Vascularization in tissue engineering.</a> Trends in Biotechnology. 26 (8), 434-441 (2008).</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>Jank, B. 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=Creation+of+a+bioengineered+skin+flap+scaffold+with+a+perfusable+vascular+pedicle.">Creation of a bioengineered skin flap scaffold with a perfusable vascular pedicle.</a> Tissue Engineering - Part A. 23 (13-14), 696-707 (2017).</li><li>Giatsidis, G., Guyette, J. P., Ott, H. C., Orgill, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+large-volume+human-derived+adipose+acellular+allogenic+flap+by+perfusion+decellularization.">Development of a large-volume human-derived adipose acellular allogenic flap by perfusion decellularization.</a> Wound Repair and Regeneration. 26 (2), 245-250 (2018).</li><li>Zhang, 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=Perfusion-decellularized+skeletal+muscle+as+a+three-dimensional+scaffold+with+a+vascular+network+template.">Perfusion-decellularized skeletal muscle as a three-dimensional scaffold with a vascular network template.</a> Biomaterials. 89, 114-126 (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=Decellularization+of+the+porcine+ear+generates+a+biocompatible,+nonimmunogenic+extracellular+matrix+platform+for+face+subunit+bioengineering.">Decellularization of the porcine ear generates a biocompatible, nonimmunogenic extracellular matrix platform for face subunit bioengineering.</a> Annals of Surgery. 267 (6), 1191-1201 (2018).</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=Perfusion-decellularization+of+human+ear+grafts+enables+ECM-based+scaffolds+for+auricular+vascularized+composite+tissue+engineering.">Perfusion-decellularization of human ear grafts enables ECM-based scaffolds for auricular vascularized composite tissue engineering.</a> Acta Biomaterialia. 73, 339-354 (2018).</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>Haughey, B. 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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+of+tissue+decellularization+used+for+preparation+of+biologic+scaffolds+and+in+vivo+relevance.">Methods of tissue decellularization used for preparation of biologic scaffolds and in vivo relevance.</a> Methods. 84, 25-34 (2015).</li><li>Mendibil, U., 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-specific+decellularization+methods:+Rationale+and+strategies+to+achieve+regenerative+compounds.">Tissue-specific decellularization methods: Rationale and strategies to achieve regenerative compounds.</a> International Journal of Molecular Sciences. 21 (15), 5447 (2020).</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. 28 (3), 677-693 (2022).</li><li>Duisit, J., Maistriaux, L., Bertheuil, N., Lellouch, A. G. <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+tissues+by+perfusion+decellularization/recellularization:+Review.">Engineering vascularized composite tissues by perfusion decellularization/recellularization: Review.</a> Current Transplantation Reports. 8, 44-56 (2021).</li><li>Adil, A., Xu, M., Haykal, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recellularization+of+bioengineered+scaffolds+for+vascular+composite+allotransplantation.">Recellularization of bioengineered scaffolds for vascular composite allotransplantation.</a> Frontiers in Surgery. 9, 843677 (2022).</li><li>Phelps, E. A., Garc&#237;a, A. 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The proposed protocol uses the perfusion of low concentration SDS to decellularize a range of porcine-derived flaps. With this procedure, acellular omentum, tensor fascia lata, and radial forearm flaps can be successfully decellularized using a protocol that favors low concentration SDS. Preliminary optimization experiments have determined that SDS at a low concentration (0.05%) between 2 days to 5 days is capable of removing cellular material for the omentum, tensor fascia lata, and radial forearm flap when analyzed wit...

Traumatic injury and tumor removal can lead to large and complex soft tissue defects. These defects can impair patient quality of life, cause loss of function, and result in permanent disability. While techniques such as autologous tissue flap transfer have been commonly practiced, issues with flap availability and donor site morbidity are major limitations1,2,3. Vascularized composite allotransplantation (VCA) is a promising alternative that transfers composite tissues, e.g., muscle, skin, vasculature, as a single unit to recipients. However, VCA requires long-term immunosuppression, which leads to drug toxicity, opportunistic infections, and malignancies4,5,6.
Tissue-engineered acellular scaffolds are a potential solution to these limitations7. Acellular tissue scaffolds can be obtained using decellularization methods, which remove cellular material from native tissues while preserving the underlying extracellular matrix (ECM) microarchitecture. In contrast to the use of synthetic materials in tissue engineering, the use of biologically derived scaffolds offers a biomimetic ECM substrate that allows biocompatibility and the potential for clinical translation8. Following decellularization, the subsequent recellularization of scaffolds with recipient-specific cells can then generate functional, vascularized tissues with little to no immunogenicity9,10,11. By developing an effective protocol to obtain acellular tissues using perfusion decellularization techniques, a broad range of tissue types can be engineered. In turn, building on this technique allows the application to more complex tissues. To date, perfusion decellularization of vascularized soft tissues has been investigated using simple vascularized tissues such as a full thickness fasciocutaneous flap in rodent12, porcine13, and human models14, as well as porcine rectus abdominis skeletal muscle15. Additionally, complex vascularized tissues have also been perfusion decellularized as demonstrated in porcine and human ear16,17 models and human full-face graft models18.
Here, the protocol describes the decellularization of vascularized free flaps using biologically derived ECM scaffolds. We present the decellularization of three clinically relevant flaps: 1) the omentum, 2) the tensor fascia lata, and 3) the radial forearm, all of which are representative of workhorse flaps used routinely in reconstructive surgery and have not been previously examined in animal studies within the context of tissue decellularization. These bioengineered flaps offer a versatile and readily available platform that has the potential for clinical applications for use in the field of large soft tissue defect repair and reconstruction.

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		<tr >
			<td >0.2 &#181;m pore Acrodisk Filter</td>
			<td >VWR</td>
			<td >CA28143-310</td>
			<td ></td>
		</tr>
		<tr >
			<td >0.9 % Sodium Chloride Solution (Normal Saline)</td>
			<td>Baxter</td>
			<td>JF7123</td>
			<td></td>
		</tr>
		<tr >
			<td >20 L Polypropylene Carboy</td>
			<td>Cole-Parmer</td>
			<td>RK-62507-20</td>
			<td></td>
		</tr>
		<tr >
			<td >3-0 Sofsilk Nonabsorbable Surgical Tie</td>
			<td>Covidien</td>
			<td>&#160;LS639</td>
			<td></td>
		</tr>
		<tr >
			<td >3-way Stopcock</td>
			<td>Cole-Parmer</td>
			<td>UZ-30600-04</td>
			<td></td>
		</tr>
		<tr >
			<td >Adson Forceps</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
		</tr>
		<tr >
			<td >Antibiotic-Antimycotic Solution, 100X</td>
			<td>Wisent</td>
			<td>450-115-EL</td>
			<td></td>
		</tr>
		<tr >
			<td >Atropine Sulphate 15 mg/30ml</td>
			<td>Rafter 8 Products</td>
			<td>238481</td>
			<td></td>
		</tr>
		<tr >
			<td >BD Angiocath 20-Gauge</td>
			<td>VWR</td>
			<td>BD381134</td>
			<td></td>
		</tr>
		<tr >
			<td >BD Angiocath 22-Gauge</td>
			<td>VWR</td>
			<td>BD381123</td>
			<td></td>
		</tr>
		<tr >
			<td >BD Angiocath 24-Gauge</td>
			<td>VWR</td>
			<td>BD381112</td>
			<td></td>
		</tr>
		<tr >
			<td >Calcium Chloride</td>
			<td>Sigma-Aldrich</td>
			<td>C4901</td>
			<td>DNAse Co-factor</td>
		</tr>
		<tr >
			<td >DNase I from bovine pancreas</td>
			<td>Sigma-Aldrich</td>
			<td>DN25</td>
			<td></td>
		</tr>
		<tr >
			<td >DNA assay (Quant-iT PicoGreen dsDNA Assay Kit)</td>
			<td>Invitrogen</td>
			<td>P7589</td>
			<td></td>
		</tr>
		<tr >
			<td >DPBS, 10X</td>
			<td>Wisent</td>
			<td>311-415-CL</td>
			<td>&#160;without Ca++/Mg++</td>
		</tr>
		<tr >
			<td >Halsted-Mosquito Hemostat</td>
			<td>Fine Science Tools</td>
			<td>13008-12</td>
			<td></td>
		</tr>
		<tr >
			<td >Heparin, 1000 I.U./mL</td>
			<td>Leo Pharma A/S</td>
			<td>453811</td>
			<td></td>
		</tr>
		<tr >
			<td >Ketamine Hydrochloride&#160; 5000 mg/50 ml</td>
			<td>Bimeda-MTC Animal Health Inc.</td>
			<td>612316</td>
			<td></td>
		</tr>
		<tr >
			<td >Ismatec Pump Tygon 3-Stop Tubing</td>
			<td>Cole-Parmer</td>
			<td>RK-96450-40</td>
			<td>Internal Diameter:&#160; 1.85 mm</td>
		</tr>
		<tr >
			<td >Ismatec REGLO 4-Channel Pump</td>
			<td>Cole-Parmer</td>
			<td>78001-78</td>
			<td></td>
		</tr>
		<tr >
			<td >Ismatec Tubing Cassettes</td>
			<td>Cole-Parmer</td>
			<td>RK-78016-98</td>
			<td></td>
		</tr>
		<tr >
			<td >Isoflurane 99.9%, 250 ml</td>
			<td>Pharmaceutical Partners of Canada Inc.</td>
			<td>2231929</td>
			<td></td>
		</tr>
		<tr >
			<td >LB Agar Lennox</td>
			<td>Bioshop Canada</td>
			<td>LBL406.500</td>
			<td>Sterility testing agar plates</td>
		</tr>
		<tr >
			<td >Magnesium Sulfate</td>
			<td>Sigma-Aldrich</td>
			<td>M7506</td>
			<td>DNAse Co-factor</td>
		</tr>
		<tr >
			<td >Masterflex L/S 16 Tubing</td>
			<td>Cole-Parmer</td>
			<td>RK-96410-16</td>
			<td></td>
		</tr>
		<tr >
			<td >Midazolam 50 mg/10 ml</td>
			<td>Pharmaceutical Partners of Canada Inc.</td>
			<td>2242905</td>
			<td></td>
		</tr>
		<tr >
			<td >Monopolar Cautery Pencil</td>
			<td>Valleylab</td>
			<td>E2100</td>
			<td></td>
		</tr>
		<tr >
			<td >Normal Buffered Formalin, 10%</td>
			<td>Sigma-Aldrich</td>
			<td>HT501128</td>
			<td></td>
		</tr>
		<tr >
			<td >N&#176;11 scalpel blade</td>
			<td>Swann Morton</td>
			<td>303</td>
			<td></td>
		</tr>
		<tr >
			<td >Papain from papaya latex</td>
			<td>Sigma-Aldrich</td>
			<td>P3125</td>
			<td></td>
		</tr>
		<tr >
			<td >Peracetic Acid</td>
			<td>Sigma-Aldrich</td>
			<td>269336</td>
			<td></td>
		</tr>
		<tr >
			<td >Plastic Barbed Connector for 1/4&#34; to 1/8&#34; Tube ID</td>
			<td>McMaster-Carr</td>
			<td>5117K61</td>
			<td></td>
		</tr>
		<tr >
			<td >Plastic Barbed Tube 90&#176; Elbow Connectors</td>
			<td>McMaster-Carr</td>
			<td>5117K76</td>
			<td></td>
		</tr>
		<tr >
			<td >Plastic Quick-Turn Tube Plugs</td>
			<td>McMaster-Carr</td>
			<td>51525K143</td>
			<td>Male Luer</td>
		</tr>
		<tr >
			<td >Plastic Quick-Turn Tube Sockets</td>
			<td>McMaster-Carr</td>
			<td>51525K293</td>
			<td>Female Luer</td>
		</tr>
		<tr >
			<td >Punch Biopsy Tool</td>
			<td>Integra Miltex</td>
			<td>3332</td>
			<td></td>
		</tr>
		<tr >
			<td >Potassium Chloride 40 mEq/20 ml</td>
			<td>Hospira Healthcare Corporation</td>
			<td>37869</td>
			<td></td>
		</tr>
		<tr >
			<td >Povidone-Iodine, 10%</td>
			<td>Rougier</td>
			<td>833133</td>
			<td></td>
		</tr>
		<tr >
			<td >Serological Pipet, 2mL</td>
			<td>Fisher Science</td>
			<td>13-678-27D</td>
			<td></td>
		</tr>
		<tr >
			<td >Snap Lid Airtight Containers</td>
			<td>SnapLock</td>
			<td>142-3941-4</td>
			<td></td>
		</tr>
		<tr >
			<td >Sodium Dodecyl Sulfate Powder</td>
			<td>Sigma-Aldrich</td>
			<td>L4509</td>
			<td></td>
		</tr>
		<tr >
			<td >Surgical Metal Ligation Clips, Small</td>
			<td >Teleflex</td>
			<td >001200</td>
			<td></td>
		</tr>
		<tr >
			<td >Stevens Tenotomy Scissors, 115 mm, straight</td>
			<td >B. Braun</td>
			<td >BC004R</td>
			<td></td>
		</tr>
		<tr >
			<td >TruWave Pressure Monitoring Set</td>
			<td >Edwards Lifesciences</td>
			<td >PX260</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

This protocol to decellularize vascularized porcine flaps relies on the perfusion of an ionic-based detergent, SDS, through the flap vasculature in a customized perfusion bioreactor. Prior to decellularization, three vascularized flaps in a porcine model were procured and cannulated according to their main supplying vessels. The flaps were immediately flushed after procurement in order to maintain a patent, perfusable vasculature to allow for successful decellularization. Using airtight snap-lid containers, a customized ...

Watch this Scientific Journal Video about Procurement and Perfusion-Decellularization of Porcine Vascularized Flaps in a Customized Perfusion Bioreactor at JoVE.com

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

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 ...

This method describes the surgical procurement of three vascularized porcine flaps and their subsequent perfusion decellularization. This method can aid efforts to regenerate porcine flaps using a tissue decellularization and recellularization approach. The main advantage of this technique is its versatility to achieve profusion decellularization across a variety of vascularized porcine flaps in a simple to assemble bioreactor setup.The technique described here can be broadly applied to other vascularized flaps as a potential tissue engineering solution using porcine flaps in reconstructive surgery. To begin, make an inflow and outflow tubing by measuring approximately 50 centimeters of LS16 platinum-coated silicone tubing. Connect one end to a yellow 3-stop pump tubing and the other to a sterile 2 milliliter serological pipette to carry perfusate from the detergent reservoirs into the bioreactor chamber.Autoclave the chamber and tubings in individually wrapped sterile packaging. For omental flap procurement, place a 12-week old anesthetized Yorkshire pig in the supine position, and then prepare the abdomen in the usual sterile fashion. Perform a xipho-umbilical laparotomy to enter the abdomen and mobilize the stomach cephalad, and place the omentum into the operative field.Starting at the left aspect of the greater curvature, where the left gastroepiploic artery joins the splenic artery, skeletonize the left gastroepiploic artery and vein using straight Stevens tenotomy scissors. Then ligate and divide both separately using surgical ties or clips. Proceed along the greater curvature of the stomach from left to right to ligate and divide the short vascular branches arising from the omentum, supplying the greater curvature of the stomach.Continue the mobilization of the greater omentum until the junction of the right gastroepiploic vessels and gastroduodenal artery is encountered. Then ligate and divide the right gastroepiploic artery and vein to free the omental flap and allow for flap cannulation later. For tensor fascia lata flap procurement, place the pig in the lateral decubitus position.Mark the interior flap boundary from the anterior superior iliac spine, or ASIS, towards the lateral patella, and a posterior boundary approximately 8 to 10 centimeters caudally. Use a scalpel to make a sharp incision at the distal margin at the patella. Deepen the incision through the subcutaneous tissues to reach the fascia overlying the rectus femoris, and continue the incisions towards the ASIS along the marked boundaries.To isolate the fascial flap alone, remove the overlying skin component from the underlying fascia layer. Then ligate or cauterize small perforator vessels to the skin. Return to the distal border of the flap, and incise the deep fascia.Mobilize the fascia off the underlying vastus lateralis muscle towards the ASIS. After tracing the pedicle toward the deep femoral vessels, skeletonize the lateral circumflex femoral artery and vein supplying the fascial flap. Then ligate and divide flap vessels proximally, where they join the deep femoral vessels and allow for later cannulation.For radial forearm flap procurement, mark a 3 by 3 centimeter square on the radial aspect of an extended pig forearm. Draw a line between the midpoint of the proximal flap margin and the antecubital fossa to denote the approximate course of the radial artery pedicle. Confirm the arterial course with palpation.Next, incise the skin using a scalpel at the distal aspect with a depth up to the antebrachial fascia. Blunt dissect with tenotomy scissors until the distal radial artery and its vena comitantes are encountered. Then ligate and divide the artery and veins with surgical ties or clips.Continue the skin incisions on the radial and ulnar margins of the marked skin square. Then mobilize the flap off the underlying radial bone with a surgical blade or cautery, proceeding from an ulnar to radial direction while raising the skin flap proximally toward the antecubital fossa. Using tenotomy scissors, skeletonize the radial artery and vena comitantes proximally at the antecubital fossa, where they join the brachial artery and veins, respectively.Dissect the vessels from the surrounding fibro-fatty tissues to isolate the vascular pedicle. Then ligate and divide the artery and vein separately to free the radial forearm flap. Next, cannulate the pedicle vessels with 20 to 24-gauge angiocatheter secured with 3-0 silk ties.Flush heparinized saline into the flap arterial cannula until a clear venous outflow is observed. While working in a laminar flow Biosafety cabinet, place flaps in the bioreactor tissue chamber. Prime inflow tubing with sterile heparinized saline to remove residual air and arrange flaps to allow connection between the flap arterial cannula and the male luer connector.Then use sterile hemostats to screw the arterial cannula onto the connector. Then flush the heparinized saline through the stopcock to check that the luer connection is without leaks, and that the flops can be perfused with evidence of venous outflow. After closing the tissue chamber lid, connect the inflow tubing with a yellow 3-stop pump tubing to the inflow stopcock of the tissue chamber.Also, attach an inline pressure sensor transducer to the inflow 3-way stopcock for perfusion pressure monitoring. Next, turn on the inflow peristaltic pump and on the initial screen, move to the third tab using the arrow to set the tubing ID to 1.85 millimeters. Then set the perfusion rate from the second tab.The input rate as the mode of delivery is set to 2 milliliters per minute. Ensure the flow direction is correct as displayed on the screen. Load the 3-stop pump tubing to the peristaltic pump with a compatible cassette.Set the outflow pump setting to a rate at 4 milliliters per minute. Connect the outflow tubing with a yellow 3-stop pump tubing to the outflow stopcock of the tissue chamber. Then load the 3-stop pump tubing to the peristaltic pump, and start the flow for both pumps by pressing the power button on each pump.Begin the perfusion decellularization of flaps with heparinized saline in the tissue chamber for 15 minutes to remove any retained blood clots. Upon completion, replace the residual saline with 500 milliliters of sodium dodecyl sulfate, or SDS, decellularization solution to submerge the flap. Next, transfer the inflow tubing to the SDS solution and profuse the tissue.After SDS decellularization, aspirate the remaining SDS before perfusing the flap with PBS for one day. After SDS perfusion decellularization, sequentially perfuse the flaps in DNA solution for two hours, and then in PBS for one day, followed by transferring the inflow tubing into peracetic acid and ethanol in distilled water to sterilize the flaps by perfusing for three hours. Once done, remove the flaps aseptically and immerse them into two washes of PBS containing 1%antibiotics antimycotics for 15 minutes each.Store flaps at 4 degrees Celsius until ready for recellularization. Use a 3 to 10 millimeter punch biopsy tool to obtain tissue samples for evaluation of decellularized flaps. The isolated decellularized flaps flushed under manual control demonstrated evidence of outflow from the freely draining venous cannula of omentum, tensor fascia lata, and radial forearm flap.The gross morphology of the native omentum, tensor fascia lata, and radial forearm flaps appeared pink colored immediately after the procurement. In comparison, decellularized tissues were characteristically white or opaque in appearance. Histological examination of native tissues with hematoxylin and eosin showed the presence of blue nuclei.In decellularized flaps, histological staining revealed loss of cellular material without blue nuclear staining, indicating an acellular tissue scaffold. Additional quantification of DNA content showed a significant decrease in DNA in acellular scaffolds compared to native tissues. The most important consideration is a surgical technique used during procurement, taking care to avoid damage to the flap pedicle in order to permit subsequent perfusion decellularization.Following perfusion decellularization, the resulting flaps may be recellularized with tissue specific cell populations in order to regenerate the native tissue compartments.

procurement-perfusion-decellularization-porcine-vascularized-flaps

Research

JoVE Journal

Bioengineering

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 ...

Engineered Vascularized Muscle Flap

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

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

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

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

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

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

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

Fabrication of Engineered Vascular Flaps Using 3D Printing Technologies

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

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

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

Multi-Stream Perfusion Bioreactor Integrated with Outlet Fractionation for Dynamic Cell Culture

Certain cell and tissue functions operate within the dynamic time scale of minutes to hours that are poorly resolved by conventional culture systems. This work has developed a low-cost perfusion bioreactor system that allows culture medium to be continuously perfused into a cell culture module and fractionated in a downstream module to measure dynamics on this scale. The system is constructed almost entirely from commercially available parts and can be parallelized to conduct independent experiments in conventional multi-well cell culture plates simultaneously. This video article demonstrates how to assemble the base setup, which requires only a single multichannel syringe pump and a modified fraction collector to perfuse up to six cultures in parallel. Useful variants on the modular design are also presented that allow for controlled stimulation dynamics, such as solute pulses or pharmacokinetic-like profiles. Importantly, as solute signals travel through the system, they are distorted due to solute dispersion. Furthermore, a method for measuring the residence time distributions (RTDs) of the components of the perfusion setup with a tracer using MATLAB is described. RTDs are useful to calculate how solute signals are distorted by the flow in the multi-compartment system. This system is highly robust and reproducible, so basic researchers can easily adopt it without the need for specialized fabrication facilities.

This paper presents a method to construct and operate a low-cost, multichannel perfusion cell culture system for measuring the dynamics of secretion and absorption rates of solutes in cellular processes. The system can also expose cells to dynamic stimulus profiles.

Certain cell and tissue functions operate within the dynamic time scale of minutes to hours that are poorly resolved by conventional culture systems. This ...

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.

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.

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

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 ...

Assembly and Priming of the 3D-Printed Bioreactor for Perfusion Culture

Begin assembling the bioreactor under laminar flow by first assembling the two 3D printed easy flow inserts. Then using one insert as the reservoir, and the other as the reaction chamber, proceed to assemble the perfusion system. Attach two connected three-way valves to the reservoir R1 port using a system tube equipped with a one-way valve.Connect the resulting outlet with the pump head. Then connect the pump head to the reaction chamber's C4 port using similar tubing with a one-way valve. Next, connect the reaction chamber port C1 via soft wall tubing to the resistance channel with a smaller inner diameter.Create a perfusion return channel by extending the resistance channel with the system tubing, and closing the luminal circulation loop by connecting to the reservoir at R5.Create an overflow channel by connecting the reaction chamber port C3 to the reservoir port R3 using system tubing, Attach a ventilator filter through R2.Construct a pressure dampener by connecting a syringe filled with air and media to the reaction chamber C6.Using a perfusion media, prime the system through the media exchange ports and the reservoir.

Perfusion Culture of Blood Vessels in the 3D-Printed Bioreactor

To prepare the sample for perfusion culture using the 3D-printed bioreactor. In a laminar flow cabinet, place the isolated porcine carotid artery samples in cold transport media. Using a scalpel blade, remove excess connective tissue and trim the ends of the tissue.Then wash the tissue twice in cold transport media. Next, place the tissue in transport media on an orbital shaker for a minimum of 30 minutes for a thorough final washing. After that, connect a non-branching segment of the washed artery to the fabricated bioreactor system using two barbed lure connections.And secure it using a vessel bond made of vascular silicone ties. Using a small syringe attached to the first lure connection, gently flow media through the artery to ensure its patency. After securing the artery to the fabricated insert, fill the reaction space with perfusion media.Next, carefully fill the luminal circulation loop with media, removing any remaining air from the system. Lastly, connect the reaction space to the assembled perfusion system, completing the circulation. To perform the perfusion culture, place the perfusion system in an incubator.After connecting it to a peristaltic pump, connect any additional acquisition systems, such as pressure sensors. Allow the system to equilibrate overnight with a low media flow rate of approximately 10 to 15 milliliters per minute. The following day, incrementally increase the flow until a final flow rate of 35 milliliters per minute is achieved.Every three days, replace 50%of the media by connecting one syringe with fresh media to the media exchange port closer to the pump and an empty syringe to the port closer to the reservoir to collect the spent medium. After the experiment, using sterile surgical scissors, harvest the tissue from the reaction chamber by trimming the tissue ends connected to the bioreactor. Confocal imaging using endothelial and smooth muscle-specific markers revealed that the normal structure of an artery was well preserved and maintained throughout, starting from the time of harvest, to cleaning and processing, and even after perfusion culture.However, incorrect handling during processing resulted in the loss of the endothelium ahead of the culture, and the application of non-physiological culture conditions like abrupt initiation of high flow showed luminal damage. Histological evaluation of the tissue using hematoxylin and eosin staining and immunofluorescence staining at the time of harvest and after seven days of perfusion culture revealed that the morphology and overall distribution of the cells in the vessel wall were maintained throughout.

Enhancing CVD Research with Modified Langendorff Perfusion

Modified Langendorff Perfusion for Extended Perfusion Times of Rodent Cardiac Grafts

Despite important advancements in the diagnosis and treatment of cardiovascular diseases (CVDs), the field is in urgent need of increased research and scientific advancement. As a result, innovation, improvement and/or repurposing of the available research toolset can provide improved testbeds for research advancement. Langendorff perfusion is an extremely valuable research technique for the field of CVD research that can be modified to accommodate a wide array of experimental needs. This tailoring can be achieved by personalizing a large number of perfusion parameters, including perfusion pressure, flow, perfusate, temperature, etc. This protocol demonstrates the versatility of Langendorff perfusion and the feasibility of achieving longer perfusion times (4 h) without graft function loss by utilizing lower perfusion pressures (30-35 mmHg). Achieving extended perfusion times without graft damage and/or function loss caused by the technique itself has the potential to eliminate confounding elements from experimental results. In effect, in scientific circumstances where longer perfusion times are relevant to the experimental needs (i.e., drug treatments, immunological response analysis, gene editing, graft preservation, etc.), lower perfusion pressures can be key for scientific success.

Author Spotlight: Advancing Cardiovascular Research &#8212; Tailored Langendorff Perfusion Techniques for Improved Experimental Outcomes

This article demonstrates the feasibility of achieving longer perfusion times (4 h) of murine cardiac grafts without function loss by employing lower (30-35 mmHg) than physiological (60-80 mmHg) perfusion pressures during Langendorff.

Despite important advancements in the diagnosis and treatment of cardiovascular diseases (CVDs), the field is in urgent need of increased research and ...