This work was supported by The British Heart Foundation and The Oak Foundation.

Perform all animal experiments according to protocols approved by the Institutional Committee for Use and Care of Laboratory Animals.
1. Preparation of Culture Media
<ol>
	<li>Make culture media for human fibroblast cell line CCL-153: F-12K Medium, 10% fetal bovine serum (FBS) and 100 U/ml penicillin and streptomycin.</li>
	<li>Make Reprogramming Media for PiPSC generation: Knockout Dulbecco&#8217;s modified eagle&#8217;s medium (DMEM) containing 20% Knockout Serum Replacement, 0.1 mM &#946;-mercaptoethanol, 0.1 mM Minimum Essential Medium (MEM) Non-Essential Amino Acids, 10 ng/ml basic Fibroblast Growth Factor 2 (bFGF-2) and 100 U/ml penicillin and streptomycin. Add bFGF-2 freshly each time before using the media.</li>
	<li>Make Differentiation Media (DM) to induce smooth muscle cell (SMC) differentiation: &#945;-MEM media containing 10% FBS, 0.1 mM &#946;-mercaptoethanol, 100 U/ml penicillin and streptomycin and 25 ng/ml Platelet Derived Growth Factor (PDGF-&#946;&#946;).</li>
	<li>Make Endothelial Differentiation Media (EC-DM) to induce endothelial cell (EC) differentiation: Endothelial growth media-2 (EGM-2) media containing 50 ng/ml Vascular Endothelial Growth Factor (VEGF) and 100 U/ml penicillin and streptomycin.</li>
</ol>
2. Reprogramming Human Fibroblasts into Partially-induced Pluripotent Stem Cells (PiPSC)
<ol>
	<li>Culture human fibroblast cell line CCL-153 on 0.04% gelatin solution coated flasks in culture medium.</li>
	<li>Passage cells every 3 days at a ratio of 1:6. Use cells before passage 9 for optimal efficiency of PiPSC generation. Cells are ready for transfection upon reaching 80 - 90% confluency.</li>
	<li>Linearize polycistronic plasmid pCAG2LMKOSimO containing 4 reprogramming factors octamer-binding transcription factor 4 (OCT4), SOX2, Kruppel-like factor 4 (KLF4) and C-MYC (pCAG-OSKM). Digest 5 &#181;g of plasmid with 5 units of PvuI restriction enzyme for 3 hr at 37 &#176;C. Purify the plasmid with a commercial kit according to manufacturer&#8217;s protocol.</li>
	<li>Transfect 2 x 106 human fibroblasts with 4 &#181;g of linearized pCAG-OSKM plasmid by electroporation with human dermal fibroblast (NHDF) nucleofection kit according to manufacturer&#8217;s protocol. Gently seed transfected cells onto pre- 0.04% gelatin coated T25 flask containing 5 ml of pre-warmed Reprogramming Media.</li>
	<li>After 24 hr, change medium with supplements of 25 &#181;g/ml neomycin to select a pure population of transfected cells.</li>
	<li>Change medium every other day until day 4. 
	Note: Human fibroblasts become PiPSC after 4 days of reprogramming.</li>
</ol>
3. Differentiation of PiPSC into Endothelial and Smooth Muscle Cells
<ol>
	<li>Seed PiPSC on collagen IV pre-coated dishes containing DM for 4 days to induce differentiation into SMCs. Change medium every other day until day 4.</li>
	<li>Seed PiPSC on collagen IV pre-coated dishes containing endothelial-DM for 6 days to induce differentiation into endothelial cells. Change medium every other day until day 6.</li>
</ol>
4. Decellularized Aorta Graft Preparation
Note: All solutions and equipment should be sterile.
<ol>
	<li>Sacrifice the mouse by cervical dislocation and fix the mouse in a supine position under dissection microscope.</li>
	<li>Cut through the sternum and around the rib cage to open the thoracic cavity. Remove the heart, lungs and esophagus to expose the aorta. Gently remove the peri-aorta fat with forceps.</li>
	<li>Cut the aorta from the anterior end with scissors and gently hold the end with blunt forceps. Detach the aorta from the spine column behind by blunt dissection. Carefully close all the branching arteries from the aorta by ligation with a bipolar electrocoagulator and cutting from the far end of the ligation with scissors.</li>
	<li>Use a 5 ml syringe to flush the aorta lumen with 3 ml of saline solution containing 100 U of heparin to prevent the formation of blood clots.</li>
	<li>Cut the posterior end of the aorta before it branches into the renal. Keeping the integrity of the aorta is very important during the whole procedure.</li>
	<li>Flush the aorta lumen with 5 ml of 0.075% sodium dodecyl sulphate (SDS) solution diluted in phosphate buffered saline (PBS).</li>
	<li>Soak the thoracic aorta in 0.075% SDS solution in a 10 cm Petri dish. Place the Petri dish on an orbital shaker for 2 hr at 150 rpm at RT.</li>
	<li>Flush the aorta lumen with 5 ml of PBS.</li>
	<li>Soak the aorta in PBS in a 10 cm Petri dish. Place the Petri dish on the orbital shaker for 2 hr at 150 rpm at RT. Refresh PBS every 20 min.</li>
	<li>Flush the aorta lumen with 5 ml of PBS.</li>
	<li>Keep decellularized aorta graft in PBS at 4 &#176;C for up to one week.</li>
</ol>
5. Dual Seeding of PiPSC and Bioreactor Conditioning
Note: All solutions and equipment should be sterile. Perform all operations in a tissue culture hood.
<ol>
	<li>Assemble the bioreactor flow circuit as illustrated in Figure 1. Connect incubation chamber, media reservoir, peristaltic pump and compliance chamber with tubing. Minimal volume of medium needed for circulation is 40 ml.</li>
	<li>Pre-condition the decellularized vessel with culture medium by soaking the aorta in DM medium in a 10 cm Petri dish. Place the Petri dish on the orbital shaker for 1 hr at 50 rpm at RT.</li>
	<li>Insert 1 cm length Nylon tubes (OD 0.9 mm, ID 0.75 mm) into both ends of the decellularized vessel under microscope. Tie the vessel and the tubes with 8-0 silk sutures.</li>
	<li>Assemble the decellularized aorta graft in the incubation chamber by connecting the Nylon tubes with the inlet and outlet ports (1/32&#8221; tubing) from the chamber wall. Maintain incubation chamber with 5mls of medium each time.</li>
	<li>Trypsinize PiPSC by adding pre-warmed trypsin to cover the culture dish and gently rock the dish 10 times. Discard the trypsin and leave the dish in incubator for 2 - 3 min. Add pre-warmed culture medium in the dish and mix the medium with the cells.</li>
	<li>Count cell number using a hemocytometer. Centrifuge two aliquots of cell suspensions containing 5 x 105 cells each for 5 min at 300 x g. Aspirate the supernatant completely.</li>
	<li>After aspirating the supernatant completely, resuspend 1 pellet of the PiPS cell pellets in 50 &#181;l of DM. Inject this cell suspension into the decellularized vessel lumen.</li>
	<li>Then, resuspend the other PiPS cell pellet in 100 &#181;l of Matrigel. Carefully pipette the mixture onto the decellularized aorta graft.</li>
	<li>Wait for 10 - 15 min until the mixture turns into a gel-like state and evenly wraps around the vessel external surface. Fill the incubation chamber with DM.</li>
	<li>Place the whole bioreactor setting in a 5% CO2 incubator at 37 &#176;C. Manually rotate the decellularized graft 90&#176; around the longitudinal axis every half an hour in the first 2 hr.</li>
	<li>Keep static culture for 12 hr to enable cell adhesion.</li>
	<li>Deliver DM through the lumen by the peristaltic pump to induce smooth muscle cell differentiation. Start initial flow rate at 5 ml/min and stepwise increase to 20 ml/min over 24 hr.</li>
	<li>Keep the circulating medium flow rate at 20 ml/min for 24 hr.</li>
	<li>Stop the medium flow and move the bioreactor setting out of the incubator.</li>
	<li>Re-seed the lumen of the graft with PiPSC. Trypsinize, count and centrifuge 1 x 106 PiPSC as in steps 5.5 - 5.6. Resuspend the cells in 50 &#181;l of EGM-2 medium containing 50 ng/ml VEGF. Inject the cell mixture into the lumen of the graft.</li>
	<li>Change the medium in the incubation chamber and whole circulation to EGM-2 medium containing 50 ng/ml VEGF to induce endothelial differentiation.</li>
	<li>Move the bioreactor setting back to the incubator. Manually rotate the graft 90&#176; every 30 min in the first 2 hr and then keep a static culture for 12 hr.</li>
	<li>Start circulating flow from 5 ml/min and stepwise increase to 35 ml/min. Keep the flow rate at 35 ml/min for 5 days. Change circulating and chamber medium every other day.</li>
	<li>Harvest the engineered graft for further grafting to mouse.</li>
</ol>
6. Grafting of Double-seeded PiPSC Graft into Mice
<ol>
	<li>Use NOD.CB17-Prkdcscid/NcrCrl male mice as vessel graft recipients. Always ensure that the mice are about 10 weeks old, weigh approximately 25 - 35 g, and are in good health conditions.</li>
	<li>Anesthetize recipient mouse by administering a combination of Hypnorm (25 mg/kg) and Hypnovel (25 mg/kg) intraperitoneally. Apply petrolatum ophthalmic ointment on eyes to prevent dryness while under anesthesia. Confirm efficient anesthetization by ensuring that mice have relaxed muscles and are breathing steadily.
	<ol>
		<li>Fix the mouse in a supine position with its neck shaved and extended. Administer atropine sulfate (1.7 mg/kg) in combination with the anaesthetics to ensure that the mouse respiratory tract remains clear.</li>
	</ol>
	</li>
	<li>Prepare a midline incision from the mandible to sternum of the mouse. Under a dissecting microscope with 5- to 10-fold amplification, lift the right salivary glands laterally and remove the right cleidomastoid muscle to expose the right common carotid artery.</li>
	<li>Gently remove attached tissues to mobilize the right common carotid artery from the distal end toward the proximal end. Ligate the middle portion of the common carotid artery twice with an 8-0 silk suture and dissect between the two ties.</li>
	<li>Pass through a cuff made of autoclavable nylon tube in each vessel end and fix each end with microhemostat clamps.</li>
	<li>Remove suture tie&#160;on one end and apply a drop of heparin (100 U/ml). Immediately after, invert the distal end of the artery to cover the whole cuff body and fix the inverted vessel with an 8-0 silk suture to the cuff.
	<ol>
		<li>Repeat the same procedure to the other portion of the artery. Flush artery with saline solution to remove blood clots.</li>
	</ol>
	</li>
	<li>Implant a decellularized vessel graft between two ends of the carotid artery by sliding the decellularized vessel ends of the over the artery cuffs and fixing it with 8-0 silk sutures. Remove vascular clamps from either ends before evaluating the pulsation of the graft.</li>
	<li>Place the right salivary gland back to its original position. Close the skin on the surgical location with an interrupted suture using a 6-0 polyglactin suture.</li>
	<li>Consistently observe vital signs of mouse after the procedure is completed, until it resumes consciousness. Sacrifice recipient mice by cervical dislocation and harvest vessel grafts either 24 hr or 3 weeks later for further analysis.</li>
</ol>

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The authors have no conflicting financial interests.

The current protocol indicates a sound, fast, simple, efficient and reproducible strategy in which functional tissue engineered vessels can be generated using PiPSC from human fibroblasts. This technique represents a valuable tool for regenerative medicine, tissue engineering and potentially patient-specific cell therapy in the near future. Critical steps to ensure the efficacy of the protocol include the preparation of PiPSC, preparation of sterile and fully decellularized aortic grafts, successful seeding and differentiation of PiPSC in the scaffolds and maintaining sterility of aortic grafts in the vascular bioreactor.
To initiate the successful generation of tissue-engineered grafts, it is of utmost importance to ensure that the preparation of PiPSC is carried out effectively. The protocol has successfully &#39;reprogrammed&#39; a human fibroblast cell line into PiPSC via nucleofection with a pCAG2LMKOSimO OSKM plasmid. The survival of fibroblasts after nucleofection relies on several key points such as the use of high quality OSKM plasmids, plasmids of concentrations ranging between 500 - 1,000 ng/&#181;l, avoid delays in seeding of fibroblasts immediately after nucleofection as cells are particularly sensitive following the introduction of relatively large plasmids and finally the addition of FGF-2 which is crucial and should be added into freshly prepared media just before use. Additionally, we found that it was crucial to use cells of up to passage 9 to avoid replicative senescence, to ensure efficient &#39;reprogramming&#39; into PiPSC and subsequent differentiation into endothelial and smooth muscle cells. The purity of PiPSC can be improved by selection of the nucleofected cells with neomycin. The pCAG2LMKOSimO plasmid has a neomycin resistant gene and thus the addition of neomycin in a range of 25 &#181;g/ml one day after the nucleofection for 3 days will result in the selection of a pure population of PiPSC. To engineer an efficient PiPSC vessel for grafting into mice, we considered the integrity and sterility of the decellularized aortas to be of highest priority. To maintain integrity of the aortas before and after decellularization, it was necessary to constantly apply manual dexterity whilst handling the vessels, as they were relatively fragile and considerably diminutive in size. The efficient removal of perivascular tissue and sealing of branching arteries using a bipolar electro-coagulator was critical to ensure subsequent competent cell seeding and to avoid leakage of media. At any point, we ensured that all materials and apparatus used to generate the bioreactor (i.e., forceps, tubes, bottles, chambers, scissors, adaptors and connectors) were autoclavable, thus reducing risks of bacterial contamination and failure of vascular grafts prior to surgery.
Whilst the current protocol holds promising clinical applications in cardiovascular diseases, there are a few limitations which should be improved in the near future to further enhance the methodology. Thus far, the OSKM transcription factors were introduced into human fibroblasts via nucleofection. While this technique is relatively simple and straightforward, it also resulted in variable transfection efficiencies at different experimental times. Notably, this inconsistency may be overcome by introducing an additional step of the aforementioned neomycin selection. Similarly, one could consider the introduction of the four factors into human fibroblasts using lentivirus integration, which may potentially increase the efficiency of &#39;reprogramming&#39; and enhance the survival of cells following infection. In terms of the bioreactor, we found that complete decellularization of aortas can be achieved using 0.075% SDS. Although this technique was considered to be optimal for cell removal as compared to other detergents (e.g., Triton X-100), it may however cause potential alterations within the ultra-structures of the extracellular matrix proteins23. This phenomenon therefore warrants more efficient decellularization strategies to minimize disruption of the associated extracellular matrix proteins. Additionally, we also considered that the diameter and wall thickness of decellularized aortas vary between mice, depending on their age, strain and background, and thus cause potential differences in the speed of flow and shear stress within bioreactors and the distribution of PiPSC following PiPSC seeding. All this remains to be clarified.
This protocol herein described the generation of functional tissue engineered grafts using PiPSC that hold a promising potential for future clinical applications in stem cell therapy for cardiovascular disease. This process which utilizes the direct reprogramming from one somatic lineage (fibroblasts) to another (either endothelial or smooth muscle cells) by skipping pluripotency may be a way to obtain safe and suitable cells of interest. Indeed, several protocols have been published to describe the generation of stem cell lines from human embryos, iPS cells and adult stem cells24-26, all of which however still raise concerning issues with regards to their use in clinic. It is now known that PiPSC do not develop teratomas in SCID mice, thus eliminating the concern of tumor formation. Furthermore, the cells take a significantly shorter time to generate, culture and differentiate10,11, and have been shown to generate successful and functional tissue engineered vessels grafts that are comparable to native aortas. Therefore, PiPSC present as a promising cell source for treatment of patients using personalized cell therapy because fibroblasts (e.g., from skin) can be derived from a specific individual. The decellularised vessel system in this protocol provides a more biomimetic model that provides seeded cells with an extracellular matrix protein that is more representative to that of a native aorta11. These key points are critical for ensuring the survival, differentiation and functionality of the seeded cells and tissue engineered vessels thereafter. Recent study by Sumitran-Holgersson and colleagues demonstrated a successful repopulation of a previously decellularized illiac vein graft from a deceased person prior to engraftment27, thus postulating a translational value of the current protocol into humans in the near future. Furthermore, the decellularized vessel grafts may represent a good model in which the role and behavior of other vascular cell types that also contribute to tissue-engineered grafts can be assessed. This model system in mice could provide a powerful tool for future drug screening and also in elucidating the cellular and molecular mechanisms involved in vascular stem cell biology. Taken together, the current protocol holds promising value for its application in vascular tissue engineering and a breakthrough in personalized medicine.

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. To date, graft materials used in surgery include biocompatible synthetic polymers (polytetrafluoroethylene [Teflon], expanded polytetrafluoroethylene [ePTFE; Gore-Tex] or polyethylene terephthalate [Dacron]), allografts, autologous tissue (pericardium or saphenous vein) and xenografts1. Whilst artificial grafts (e.g., Gore-Tex and Dacron) are most commonly used, these materials likely cause numerous short- and long-term complications that include stenosis, calcium deposition, thrombo-embolization and infections. Although patients with biological grafts present with decreased thrombo-embolic events, they still encounter limitations such as secondary graft failure and shortened durability owing to calcification degradation2. Therefore, despite significant improvements in surgical techniques over the years, researchers and clinicians are still burdened with the need for identifying the ideal conduit for vascular diseases. More recently, the research field of vascular tissue engineering has generated a concept in which cells are incorporated into biodegradable scaffolds, with the aim of creating a biomimetic environment that epitomizes a functional vessel for successful grafting1. Fundamentally, the success of the vascular constructs depend on three essential components; cells that comprise the scaffold, i.e., an endothelial cell inner layer and a smooth muscle cell layer, a scaffold containing the appropriate extracellular matrix to provide mechanical properties comparable to the native vasculature, and the molecular/cellular signaling that is required for initiating/regulating repair.
Long term graft patency and sustained development of the neo-tissues are highly dependent on effective cell seeding of scaffolds, thereby rendering the decision of cell type of critical importance. Several reports demonstrate the use of mature endothelial and smooth muscle cells from various sources to develop small diameter conduits3-6. Although promising, the lack of sufficient autologous vessels to obtain mature endothelial and smooth muscle cells remain a considerable burden. More recently, stem cells from various sources have been exploited for vascular tissue engineering applications. Indeed, a variety of stem cell types including embryonic stem cells7, induced pluripotent stem cells (iPSCs)8,9, PiPSC10,11, bone marrow-derived mononuclear cells12, mesenchymal stem cells13, endothelial progenitor cells and adult vessel wall-derived stem cell antigen-1 (Sca-1)+ stem/progenitor cells14,15 have all been demonstrated to be capable of differentiation into either functional endothelial or smooth muscle cells in response to defined media and culture conditions. Furthermore, the unlimited self renewal capacity of the stem cells make them better candidates unlike mature endothelial and smooth muscle cells which can only divide for a finite number of times before undergoing growth arrest and senescence.
The selection of scaffold material to generate successful tissue engineered vessel for grafting depends on several factors such as biocompatibility, biomechanical properties, and rate of biodegradation. Fundamentally, materials used to create scaffolds for the grafts should be biodegradable and will not mount unnecessary recipient immune responses. Additionally, it must encompass a suitable porosity and microstructure for cell attachment and subsequent survival. To date, the most common materials used for scaffolds in vascular tissue engineering include polymers of polyglycolic acid, polylactic acid, and poly &#949;-caprolactone16. More recently, decellularized biological materials have also been applied with some success. Several laboratories have shown that seeding decellularized human, canine or porcine vessels with autologous cells provided a biological graft that resisted clotting and intimal hyperplasia17-19. Other strategies in vascular tissue engineering include extracellular matrix proteins-based vascular grafts e.g., seeding cells in fibrin gel13 and generating cell sheets without scaffold support20, 21.
The current protocol demonstrates the differentiation of human PiPSC into functional endothelial and smooth muscle cells, the generation of a bioreactor consisting of a decellularized vessel scaffold to harbor functional PiPSC-derived vascular cells, and grafting of the tissue engineered vessels into severe combined immunodeficiency (SCID) mice. PiPSC are an optimum cell type to use for tissue engineering of vessel grafts because these cells do not form tumors in mice or raise ethical and allo-immune responses. Furthermore, we have shown that the strategy for generating PiPS-endothelial cells and PiPS-smooth muscle cells is efficient and reproducible10,11. Thereafter, we designed a decellularized vessel for seeding of PiPSC-derived vascular cells to closely mimic the matrix proteins that exists within a native vessel, thus enhancing grafting and survival efficacy. Furthermore, the decellularization of the vessels prior to PiPSC seeding prevents the occurrence of inflammatory responses mounted by immune cell types such as macrophages. More importantly, this protocol does not only represent a methodology to generate promising vascular conduits for translation into humans, but also provides valuable means of studying and understanding the molecular mechanisms that govern vascular tissue regeneration through mouse models.

<table border="0" cellpadding="0" cellspacing="0" >
	<tbody>
		<tr >
			<td >Human Fibroblasts CCL-153</td>
			<td >ATCC</td>
			<td >CCL-153</td>
			<td>Prenatal human embryonic fibroblasts</td>
		</tr>
		<tr >
			<td >ATCC F-12K Medium (Kaighn&#39;s Modification of Ham&#39;s F-12 Medium)</td>
			<td >ATCC</td>
			<td >30-2004</td>
			<td></td>
		</tr>
		<tr >
			<td >Fetal Bovine Serum</td>
			<td >ATCC</td>
			<td >30-2020</td>
			<td></td>
		</tr>
		<tr >
			<td >Knockout DMEM medium optimized for embryonic stem cells</td>
			<td >Life technologies (Gibco)</td>
			<td >12660-012</td>
			<td></td>
		</tr>
		<tr >
			<td >Knockout Serum Replacement</td>
			<td >Life technologies (Invitrogen)</td>
			<td >10828-028</td>
			<td></td>
		</tr>
		<tr >
			<td >Human Basic FGF-2&#160;</td>
			<td >Miltenyi Biotech</td>
			<td >130-093-837</td>
			<td></td>
		</tr>
		<tr >
			<td >alpha-MEM medium</td>
			<td >Life technologies (Invitrogen)</td>
			<td >32571093</td>
			<td></td>
		</tr>
		<tr >
			<td >Human PDGF</td>
			<td >R&#38;D System</td>
			<td >120-HD-001</td>
			<td></td>
		</tr>
		<tr >
			<td >Gelatin Solution 2%</td>
			<td >Sigma</td>
			<td >G1393</td>
			<td></td>
		</tr>
		<tr >
			<td >Plasmid 20866: pCAG2LMKOSimO (SOX2, OCT4, KLF4, C-MYC)</td>
			<td >Addgene</td>
			<td >20866</td>
			<td></td>
		</tr>
		<tr >
			<td >&#160;PvuI Restriction Enzyme</td>
			<td >New England Biolabs</td>
			<td >RO150S</td>
			<td></td>
		</tr>
		<tr >
			<td >SureClean Plus</td>
			<td >Bioline</td>
			<td >BIO-37047</td>
			<td></td>
		</tr>
		<tr >
			<td >Nucelofection Kit (NHDF Kit)</td>
			<td >LONZA</td>
			<td >VPD-1001</td>
			<td></td>
		</tr>
		<tr >
			<td >Neomycin</td>
			<td >SIGMA</td>
			<td >G418</td>
			<td>Selection of&#160;</td>
		</tr>
		<tr >
			<td >KL 1500 LCD, Illumination for Stereo Microscopy</td>
			<td >SCHOTT</td>
			<td >KL 1500 LCD</td>
			<td>Cold light illumination for stereo microscopy</td>
		</tr>
		<tr >
			<td >Nikon Zoom Steromicroscope SMZ800</td>
			<td >Nikon</td>
			<td >SMZ800</td>
			<td></td>
		</tr>
		<tr >
			<td >Heparin sodium salt</td>
			<td >Sigma</td>
			<td >H3393</td>
			<td></td>
		</tr>
		<tr >
			<td >10% SDS Stock Solution Molecular Biology Reagent</td>
			<td >Severn Biotech</td>
			<td >CAS 151-21-3</td>
			<td></td>
		</tr>
		<tr >
			<td >Dulbecco&#39;s Phosphate Buffered Saline</td>
			<td >Sigma</td>
			<td >D8537</td>
			<td></td>
		</tr>
		<tr >
			<td >Matrigel (10mg/ml)</td>
			<td >BD</td>
			<td >A6661</td>
			<td></td>
		</tr>
		<tr >
			<td >Shaker IKA Vibrax with Shaking platform VX 7&#160;</td>
			<td >Jepson Bolton&#39;s, Janke&#38;Kunkel</td>
			<td >S32-102</td>
			<td></td>
		</tr>
		<tr >
			<td >Masterflex L/S Digital Pump Drive</td>
			<td >Cole-Parmer</td>
			<td >WZ-07523-80</td>
			<td></td>
		</tr>
		<tr >
			<td >Masterflex L/S 6-channel, 6-roller cartridge pump head</td>
			<td >Cole-Parmer</td>
			<td >EW-07519-15</td>
			<td></td>
		</tr>
		<tr >
			<td >Masterflex L/S large cartridges for pump head</td>
			<td >Cole-Parmer</td>
			<td >EW-07519-75</td>
			<td></td>
		</tr>
		<tr >
			<td >Masterflex platinum-cured silicone pump tubing, L/S 14, 25 ft</td>
			<td >Cole-Parmer&#160;</td>
			<td >WZ-96410-14</td>
			<td >Tubing goes through the peristaltic pump</td>
		</tr>
		<tr >
			<td >0.5mm ID, 0.8 mm OD Silicone Tubing</td>
			<td >SILEX</td>
			<td >N/A</td>
			<td>Tubings connect incubation chamber, media reservoir and compliance chamber&#160;</td>
		</tr>
		<tr >
			<td >Fitting Reducer 0.5 to 1.6, natural Polypropyline</td>
			<td >Ibidi</td>
			<td >10829</td>
			<td>Adapter connect above two types of tubings</td>
		</tr>
		<tr >
			<td >1/32&#34; Tubing, ID 0.01&#34; (250&#181;m) Material: PEEK</td>
			<td >LabSmith</td>
			<td >T-132-010P</td>
			<td>Tubing through the incubation chamber wall which connects the graft with outside tubing</td>
		</tr>
		<tr >
			<td >One-Piece Fittings&#160;</td>
			<td >LabSmith</td>
			<td >T-132-100</td>
			<td>Fix the above tubings through the incubation chamber wall</td>
		</tr>
		<tr >
			<td >Nylon tubes (OD 0.9mm, ID 0.75mm)&#160;</td>
			<td >Smiths Medical</td>
			<td >N/A</td>
			<td>Tubings insert into two ends of the aorta graft</td>
		</tr>
		<tr >
			<td >NOD.CB17-Prkdcscid/NcrCrl mouse</td>
			<td >Charles River</td>
			<td ></td>
			<td ></td>
		</tr>
		<tr >
			<td >Surgical sutures, 8-0&#160; silk</td>
			<td >ETHICON</td>
			<td >W819</td>
			<td ></td>
		</tr>
		<tr >
			<td >Hypnorm</td>
			<td >Vetapharm</td>
			<td >Vm21757/4000</td>
			<td >Neuroleptanalgesic for use in mice</td>
		</tr>
		<tr >
			<td >Hypnovel (Midazolam)</td>
			<td >Roche</td>
			<td >59467-70-8</td>
			<td >Induction of anaesthesia</td>
		</tr>
		<tr >
			<td >Dissecting microscope</td>
			<td >Carl Zeiss</td>
			<td >Stemi 2000</td>
			<td ></td>
		</tr>
		<tr >
			<td >Nylon Tubing</td>
			<td >Portex LTD</td>
			<td >800/200/100/200</td>
			<td >0.65 mm in diameter and 1 mm in length; to make artery cuff</td>
		</tr>
		<tr >
			<td >Electrocoagulator</td>
			<td >Martin&#160;</td>
			<td >SN 54.131</td>
			<td >Ligation of artery branches on aorta</td>
		</tr>
		<tr >
			<td >Bipolar micro hemostat forceps</td>
			<td >Martin</td>
			<td >80-91-12-04</td>
			<td >Fixation of vessel ends</td>
		</tr>
		<tr >
			<td >Vessel Dilator</td>
			<td >S&#38;T</td>
			<td >JFX-7</td>
			<td ></td>
		</tr>
		<tr >
			<td >Vessel Dilator</td>
			<td >S&#38;T</td>
			<td >JFL-3dZ</td>
			<td ></td>
		</tr>
		<tr >
			<td >Vessel Dilator</td>
			<td >S&#38;T</td>
			<td >D-5aZ</td>
			<td ></td>
		</tr>
		<tr >
			<td >Mini applier&#160;</td>
			<td >AESCULAP</td>
			<td >FE572K</td>
			<td ></td>
		</tr>
		<tr >
			<td >Micro hemostats clips</td>
			<td >AESCULAP</td>
			<td >FE720K</td>
			<td ></td>
		</tr>
		<tr >
			<td >Surgical sutures, 6-0 VICRYL</td>
			<td >ETHICON</td>
			<td >V489</td>
			<td ></td>
		</tr>
	</tbody>
</table>

generation and grafting of tissue-engineered vessels in a mouse model

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. The current protocol presents an efficient and reproducible strategy in which functional tissue engineered vessel grafts can be generated using partially induced pluripotent stem cell (PiPSC) from human fibroblasts. We designed a decellularized vessel scaffold bioreactor, which closely mimics the matrix protein structure and blood flow that exists within a native vessel, for seeding of PiPSC-endothelial cells or smooth muscle cells prior to grafting into mice. This approach was demonstrated to be advantageous because immune-deficient mice engrafted with the PiPSC-derived grafts presented with markedly increased survival rate 3 weeks after surgery. This protocol represents a valuable tool for regenerative medicine, tissue engineering and potentially patient-specific cell-therapy in the near future.

The successful generation of PiPSC was confirmed 4 days after nucleofecting human fibroblasts with a linearized pCAG2LMKOSimO plasmid carrying 4 transcription factors, OCT4, SOX2, KLF4 and c-MYC (OSKM). PiPSC displayed a markedly distinct phenotype when compared to fibroblasts (Figure 2A) and expressed the 4 reprogramming factors at mRNA (Figure 2B) and protein (Figure 2C) levels10. The efficacy of a PiPSC-based vascular graft is highly dependent on the ability of the PiPSC to differentiate into both endothelial and smooth muscle lineages. It is therefore critical to confirm these properties of PiPSC in vitro by differentiating the cells in defined media before utilizing them to generate vessel grafts. Figure 3 shows the capacity of PiPSC to differentiate into functional endothelial cells at both gene (Figure 3A) and protein (Figure 3B-D) levels, based on endothelial-specific markers10. Similarly, PiPSC are also able to differentiate into the smooth muscle lineage in response to specific media (Figure 3E-H)11.
Whole aorta decellularization was achieved by treatment with SDS, an anionic detergent that lyses cells and solubilizes cytoplasmic components. After 2 hr of treatment, the decellularized aorta appeared as a translucent acellular scaffold when compared to a freshly harvested aorta (Figure 4A-B)22. Histological evaluation in Figure 4C-F illustrates the absence of nuclei or cytoplasmic staining in decellularized vessels, but not in the normal mouse aorta. Following double seeding of PiPSC-derived endothelial and smooth muscle cells within decellularized aortas, the engineered vascular grafts display endothelial and smooth muscle cell properties respectively. Hematoxylin and eosin (HE) staining of the double-seeded grafts revealed an architecture similar to that of native vessels (Figure 4G) with multiple layers of smooth muscle cells and a monolayer of endothelial cells as confirmed by positive staining for smooth muscle-22&#945; (SM22), smooth muscle-&#945; actin (SMA), CD31 and CD144 markers respectively (Figure 4H-M)11.
To confirm the patency of the tissue engineered vessels in vivo, the double seeded PiPSC-derived vessels were grafted into mice - both normal mouse and decellularized arteries were used as controls. Subsequent analysis showed that while mice grafted with decellularized vessels presented with markedly higher mortality rates as early as day 1, PiPSC vessel grafts conferred a survival rate of 60% 3 weeks after transplantation (Figure 5A)11. Additionally, 3 week old PiPSC-derived grafts were fully characterized for the presence of human PiPSC-derived endothelial and smooth muscle cells and host macrophage infiltration (Figure 5B and Table 1)11. Taken together, results indicate that PiPSC have the capacity to differentiate into both vascular lineages and are potent for generating functional tissue-engineered vessels that can substitute native vessels in vivo.
<img alt="Figure 1" src="/files/ftp_upload/52565/52565fig1.jpg" /> 
Figure 1. Schematic representation of the decellularized graft bioreactor flow circuit. The decellularized vessel graft is assembled in the incubation chamber. A peristaltic pump is at the upstream of the incubation chamber to provide stable medium perfusion flow. The media reservoir is at the downstream of the incubation chamber. The compliance chamber is to improve the flow regime. The flow direction is indicated by arrows. <a href="https://www.jove.com/files/ftp_upload/52565/52565fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52565/52565fig2.jpg" /> 
Figure 2. Generation of PiPSC from human fibroblasts. Human fibroblasts were nucleofected with either four reprogramming genes (OCT4, SOX2, KLF4 and C-MYC) or an empty vector (control). (A) Phase contrast pictures show the morphology of PiPS after 4 days. PiPSC expressed all four transcription factors, both at mRNA (B) and protein (C) levels. Scale bar for images shown are 50 &#181;m and data are means &#177; SEM (n = 3); *P &#60; 0.05, ***P &#60; 0.001. This figure has been modified from Margariti, A. et al.10 <a href="https://www.jove.com/files/ftp_upload/52565/52565fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52565/52565fig3.jpg" /> 
Figure 3. PiPSC can differentiate into both endothelial and smooth muscle cells. PiPSC or control cells were differentiated into endothelial cells . (A) In comparison to control cells, PiPSC-derived endothelial cells expressed endothelial-specific markers such as CD31, CD144, kinase insert domain receptor (KDR), endothelial nitric oxide synthase (eNOS) and von willebrand factor (vWF) at the mRNA level. (B) This was confirmed at the protein level using western blot analysis. (C) Immunofluorescence staining indicated positive staining of CD144 (VE-cadherin) in PiPSC-endothelial, but not control cells. (D) Flow cytometric analysis confirmed PiPSC-endothelial cell expression of CD31 and CD144. Collagen IV-seeded PiPSC (or control cells) were also differentiated into smooth muscle cells. (E,F) Contrary to control cells, PiPSC-derived smooth muscle cells expressed smooth muscle-specific markers at the gene level, including SMA, calponin, SM22, smooth muscle myosin heavy chain II (SMMHC), serum response factor (SRF), myocardin, smoothelin, elastin and collagen 1A1. (G) The expression levels of SMA, calponin and SM22 in PiPSC-derived cells were confirmed using western blot analysis. (H) Immunofluorescence staining using confocal microscopy showed a typical smooth muscle marker staining for calponin and SM22. Data represent means &#177; SEM (n = 3); *P &#60; 0.05, **P &#60; 0.01, ***P &#60; 0.001. Scale bar for images shown are 25 &#181;m or 50 &#181;m . These figures have been modified from Margariti, A. et al.10 and Karamariti, E. et al.11 <a href="https://www.jove.com/files/ftp_upload/52565/52565fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52565/52565fig4.jpg" /> 
Figure 4. Generation of double-seeded PiPSC-derived tissue engineered vascular grafts. (A,B) Decellularization of mouse thoracic aorta preserved its structure and still resembled that of a normal vessel, original magnification 2X. (C-F) HE staining of decellularized aorta confirmed successful decellularization and complete removal of cells when compared to controls, original magnification: 50X or 200X. (G) HE staining of PiPSC-derived endothelial and smooth muscle cells double-seeded vessels revealed an architecture that was similar to a native vessel. (J,M) Double-seeded PiPSC tissue engineered vessels stained positive for endothelial (CD31 and CD144) and smooth muscle cells (SM22 and SMA) markers unlike decellularized vessel grafts. (H,K) The concomitant expression of vascular markers as well as the characteristic morphology and localization of PiPSC-derived endothelial and smooth muscle cells within medial and intimal areas was comparable to native vessels. Scale bar for images shown are 50 &#181;m. These figures have been modified from Tsai, T. et al. 22 and Karamariti, E. et al.11 <a href="https://www.jove.com/files/ftp_upload/52565/52565fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52565/52565fig5.jpg" /> 
Figure 5. Double-seeded PiPSC-derived tissue engineered vascular grafts are patent for substituting native vessels in vivo. PiPSC-derived vessels were grafted into the carotid artery of severe combined immunodeficiency (SCID) mice, normal mouse and decellularized arteries were used as controls. (A) Analysis of mice grafted with PiPSC-derived vessels showed 60% survival rate 3 weeks post surgery, whereas mice grafted with decellularized vessels presented with markedly higher mortality rates and lower survival (20%). Data represent means (n = 10). (B) The tissue-engineered vessels were also harvested 3 weeks after grafting and subjected to HE (left panels) or immunofluorescence staining (right panels) for the presence of human CD144 and calponin or mouse CD11b/CD18 (Mac-1); quantification of staining is summarized in Table 1. Original magnification for images shown are 400X unless stated. This figure has been modified from Karamariti, E. et al.11 <a href="https://www.jove.com/files/ftp_upload/52565/52565fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Positive cells Field (40X)</td>
			<td>Engineered-vessel before grafting</td>
			<td>Engineered-vessel graft (3 weeks)</td>
		</tr>
		<tr>
			<td>calponin</td>
			<td>72 &#177; 9</td>
			<td>22 &#177; 11*</td>
		</tr>
		<tr>
			<td>CD144</td>
			<td>20 &#177; 8</td>
			<td>5 &#177; 6*</td>
		</tr>
		<tr>
			<td>Mac-1</td>
			<td>0</td>
			<td>71 &#177; 11*</td>
		</tr>
	</tbody>
</table>
Table 1. Characterization of tissue-engineered vessels 3 weeks after grafting. The number of PiPSC-derived endothelial and smooth muscle cells and host macrophages was quantified following immunofluorescence staining with human calponin, human CD144 and mouse Mac-1 respectively. Data represents mean &#177; SEM (n = 6); *P &#60; 0.05, significant difference from tissue-engineered vessels before grafting. This table has been modified from Karamariti, E. et al.11

Watch this Scientific Journal Video about Generation and Grafting of Tissue-engineered Vessels in a Mouse Model at JoVE.com

Generation and Grafting of Tissue-engineered Vessels in a Mouse Model

Here, we present a protocol to generate tissue engineered vessel grafts that are functional for grafting into mice by double seeding partially induced pluripotent stem cell (PiPSC) - derived smooth muscle cells and PiPSC - derived endothelial cells on a decellularized vessel scaffold bioreactor.

The construction of vascular conduits is a fundamental strategy for surgical repair of damaged and injured vessels resulting from cardiovascular diseases. ...

generation-and-grafting-of-tissue-engineered-vessels-in-a-mouse-model

Nanyang Technological University

Research

JoVE Journal

Bioengineering

11.9K Views.  King's College London BHF Centre. Here, we present a protocol to generate tissue engineered vessel grafts that are functional for grafting into mice by double seeding partially induced pluripotent stem cell (PiPSC) - derived smooth muscle cells and PiPSC - derived endothelial cells on a decellularized vessel scaffold bioreactor.

Video: Generation and Grafting of Tissue-engineered Vessels in a Mouse Model

Protocol for Relative Hydrodynamic Assessment of Tri-leaflet Polymer Valves

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of tri-leaflet polymer valve prostheses. However, identification of a protocol for initial assessment of polymer valve hydrodynamic functionality is paramount during the early stages of the design process. Traditional in vitro pulse duplicator systems are not configured to accommodate flexible tri-leaflet materials; in addition, assessment of polymer valve functionality needs to be made in a relative context to native and prosthetic heart valves under identical test conditions so that variability in measurements from different instruments can be avoided. Accordingly, we conducted hydrodynamic assessment of i) native (n = 4, mean diameter, D = 20 mm), ii) bi-leaflet mechanical (n= 2, D = 23 mm) and iii) polymer valves (n = 5, D = 22 mm) via the use of a commercially available pulse duplicator system (ViVitro Labs Inc, Victoria, BC) that was modified to accommodate tri-leaflet valve geometries. Tri-leaflet silicone valves developed at the University of Florida comprised the polymer valve group. A mixture in the ratio of 35:65 glycerin to water was used to mimic blood physical properties. Instantaneous flow rate was measured at the interface of the left ventricle and aortic units while pressure was recorded at the ventricular and aortic positions. Bi-leaflet and native valve data from the literature was used to validate flow and pressure readings. The following hydrodynamic metrics were reported: forward flow pressure drop, aortic root mean square forward flow rate, aortic closing, leakage&#160;and regurgitant volume, transaortic closing, leakage, and total energy losses. Representative results indicated that hydrodynamic metrics from the three valve groups could be successfully obtained by incorporating a custom-built assembly into a commercially available pulse duplicator system and subsequently, objectively compared to provide insights on functional aspects of polymer valve design.

There has been renewed interest in developing polymer valves. Here, the objectives are to demonstrate the feasibility of modifying a commercial pulse duplicator to accommodate tri-leaflet geometries and to define a protocol to present polymer valve hydrodynamic data in comparison to native and prosthetic valve data collected under near-identical conditions.

Limitations of currently available prosthetic valves, xenografts, and homografts have prompted a recent resurgence of developments in the area of ...

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

The development of Tissue Engineered Vessels (TEVs) is advanced by the ability to routinely and effectively implant TEVs (4-5 mm in diameter) into a large animal model. A step by-step protocol for inter-positional placement of the TEV and real-time digital assessment of the TEV and native carotid arteries is described here. In vivo monitoring is made possible by the implantation of flow probes, catheters and ultrasonic crystals (capable of recording dynamic diameter changes of implanted TEVs and native carotid arteries) at the time of surgery. Once implanted, researchers can calculate arterial blood flow patterns, invasive blood pressure and artery diameter yielding parameters such as pulse wave velocity, augmentation index, pulse pressures and compliance. Data acquisition is accomplished using a single computer program for analysis throughout the duration of the experiment. Such invaluable data provides insight into TEV matrix remodeling, its resemblance to native/sham controls and overall TEV performance in vivo.

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

A step-by-step protocol for the inter-positional placement of Tissue Engineered Vessels (TEVs) into the carotid artery of a sheep using end-to-end anastomosis and real-time digital assessment in vivo until animal sacrifice.

The development of Tissue Engineered Vessels (TEVs) is advanced by the ability to routinely and effectively implant TEVs (4-5 mm in diameter) into a large ...

Generation of a Three-dimensional Full Thickness Skin Equivalent and Automated Wounding

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models reflect the human wound situation better than animal models. Although two-dimensional models are widely used to investigate processes such as cellular migration and proliferation, models that are more complex are required to gain a deeper knowledge about wound healing. Besides a suitable model system, the generation of precise and reproducible wounds is crucial to ensure comparable results between different test runs. In this study, the generation of a three-dimensional full thickness skin equivalent to study wound healing is shown. The dermal part of the models is comprised of human dermal fibroblast embedded in a rat-tail collagen type I hydrogel. Following the inoculation with human epidermal keratinocytes and consequent culture at the air-liquid interface, a multilayered epidermis is formed on top of the models. To study the wound healing process, we additionally developed an automated wounding device, which generates standardized wounds in a sterile atmosphere.

The goal of this protocol is to build up a three-dimensional full thickness skin equivalent, which resembles natural skin. With a specifically constructed automated wounding device, precise and reproducible wounds can be generated under maintenance of sterility.

In vitro models are a cost effective and ethical alternative to study cutaneous wound healing processes. Moreover, by using human cells, these models ...

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

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

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

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

A Combined 3D Tissue Engineered In Vitro/In Silico Lung Tumor Model for Predicting Drug Effectiveness in Specific Mutational Backgrounds

In the present study, we combined an in vitro 3D lung tumor model with an in silico model to optimize predictions of drug response based on a specific mutational background. The model is generated on a decellularized porcine scaffold that reproduces tissue-specific characteristics regarding extracellular matrix composition and architecture including the basement membrane. We standardized a protocol that allows artificial tumor tissue generation within 14 days including three days of drug treatment. Our article provides several detailed descriptions of 3D read-out screening techniques like the determination of the proliferation index Ki67 staining&#39;s, apoptosis from supernatants by M30-ELISA and assessment of epithelial to mesenchymal transition (EMT), which are helpful tools for evaluating the effectiveness of therapeutic compounds. We could show compared to 2D culture a reduction of proliferation in our 3D tumor model that is related to the clinical situation. Despite of this lower proliferation, the model predicted EGFR-targeted drug responses correctly according to the biomarker status as shown by comparison of the lung carcinoma cell lines HCC827 (EGFR -mutated, KRAS wild-type) and A549 (EGFR wild-type, KRAS-mutated) treated with the tyrosine-kinase inhibitor (TKI) gefitinib. To investigate drug responses of more advanced tumor cells, we induced EMT by long-term treatment with TGF-beta-1 as assessed by vimentin/pan-cytokeratin immunofluorescence staining. A flow-bioreactor was employed to adjust culture to physiological conditions, which improved tissue generation. Furthermore, we show the integration of drug responses upon gefitinib treatment or TGF-beta-1 stimulation - apoptosis, proliferation index and EMT - into a Boolean in silico model. Additionally, we explain how drug responses of tumor cells with a specific mutational background and counterstrategies against resistance can be predicted. We are confident that our 3D in vitro approach especially with its in silico expansion provides an additional value for preclinical drug testing in more realistic conditions than in 2D cell culture.

A Combined 3D Tissue Engineered In Vitro/In Silico Lung Tumor Model for Predicting Drug Effectiveness in Specific Mutational Backgrounds

We present a three-dimensional (3D) lung cancer model based on a biological collagen scaffold to study sensitivity towards non-small-cell-lung-cancer-(NSCLC)-targeted therapies. We demonstrate different read-out techniques to determine the proliferation index, apoptosis and epithelial-mesenchymal transition (EMT) status. Collected data are integrated into an in silico model for prediction of drug sensitivity.

In the present study, we combined an in vitro 3D lung tumor model with an in silico model to optimize predictions of drug response based on a specific ...

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

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

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

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

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

Three-dimensional Tissue Engineered Aligned Astrocyte Networks to Recapitulate Developmental Mechanisms and Facilitate Nervous System Regeneration

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to replace lost neurons and regenerate axonal pathways. However, during nervous system development, neuronal migration and axonal extension often occur along pathways formed by other cells, referred to as &#34;living scaffolds&#34;. Seeking to emulate these mechanisms and to design a strategy that circumvents the inhibitory environment of the CNS, this manuscript presents a protocol to fabricate tissue engineered astrocyte-based &#34;living scaffolds&#34;. To create these constructs, we employed a novel biomaterial encasement scheme to induce astrocytes to self-assemble into dense three-dimensional bundles of bipolar longitudinally-aligned somata and processes. First, hollow hydrogel micro-columns were assembled, and the inner lumen was coated with collagen extracellular-matrix. Dissociated cerebral cortical astrocytes were then delivered into the lumen of the cylindrical micro-column and, at a critical inner diameter of &#60;350 &#181;m, spontaneously self-aligned and contracted to produce long fiber-like cables consisting of dense bundles of astrocyte processes and collagen fibrils measuring &#60;150 &#181;m in diameter yet extending several cm in length. These engineered living scaffolds exhibited &#62;97% cell viability and were virtually exclusively comprised of astrocytes expressing a combination of the intermediate filament proteins glial-fibrillary acidic protein (GFAP), vimentin, and nestin. These aligned astrocyte networks were found to provide a permissive substrate for neuronal attachment and aligned neurite extension. Moreover, these constructs maintain integrity and alignment when extracted from the hydrogel encasement, making them suitable for CNS implantation. These preformed constructs structurally emulate key cytoarchitectural elements of naturally occurring glial-based &#34;living scaffolds&#34; in vivo. As such, these engineered living scaffolds may serve as test-beds to study neurodevelopmental mechanisms in vitro or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding following CNS degeneration in vivo.

We showcase the development of self-assembled, three-dimensional bundles of longitudinally aligned astrocytic somata and processes within a novel biomaterial encasement. These engineered &#34;living scaffolds&#34;, exhibiting micron-scale diameter yet extending centimeters in length, may serve as test-beds to study neurodevelopmental mechanisms or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding.

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to ...

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

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

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

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

Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional tissue. Tissue engineering holds great promise as a potential solution with its ability to integrate cells and signaling molecules into a 3-dimensional scaffold. Recent work with tissue engineered tracheal grafts (TETGs) has seen some success but their translation has been limited by graft stenosis, graft collapse, and delayed epithelialization. In order to investigate the mechanisms driving these issues, we have developed a mouse model for tissue engineered tracheal graft implantation. TETGs were constructed using electrospun polymers polyethylene terephthalate (PET) and polyurethane (PU) in a mixture of PET and PU (20:80 percent weight). Scaffolds were then seeded using bone marrow mononuclear cells isolated from 6-8 week-old C57BL/6 mice by gradient centrifugation. Ten million cells per graft were seeded onto the lumen of the scaffold and allowed to incubate overnight before implantation between the third and seventh tracheal rings. These grafts were able to recapitulate the findings of stenosis and delayed epithelialization as demonstrated by histological analysis and lack of Keratin 5 and Keratin 14 basal epithelial cells on immunofluorescence. This model will serve as a tool for investigating cellular and molecular mechanisms involved in host remodeling.

Graft stenosis poses a critical obstacle in tissue engineered airway replacement. To investigate cellular mechanisms underlying stenosis, we utilize a murine model of tissue engineered tracheal replacement with seeded bone marrow mononuclear cells (BM-MNC). Here, we detail our protocol, including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation.

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional ...

Tissue-Engineered Graft for Circumferential Esophageal Reconstruction in Rats

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

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

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