This research was supported in part by the NIH Director's New Innovator Award Program, part of the NIH Roadmap for Medical Research, through grant number 1-DP2-OD004309-01. S.B.S-N. would like to thank the NSF for a Graduate Research Fellowship.

1. Pre-processing Tissue Preparation
<ol class="lalpha">
<li>	Before using this protocol, one must have already decellularized the tissue of choice. For this example, fresh porcine and human pericardium samples are decellularized using hypotonic and hypertonic rinses in deionized (DI) water and sodium dodecyl sulfate (SDS).</li>
<li>	Specifically, first wash the porcine pericardium in DI water for 30 minutes, then stir continuously in 1% SDS in phosphate buffered saline (PBS) for 24 hours, followed by a 5 hour DI water rinse. For human pericardia, first rinse in DI water for 30 minutes, then stir continuously in 1% SDS in PBS for 60-65 hours, followed by an overnight DI rinse. Remove all specimens from their final solution and rinse again under running DI water 1. </li>
<li>	To verify decellularization, remove a small piece, fresh freeze it in O.C.T. freezing medium, and take 10 &mu;m tissue sections every 100 &mu;m throughout the sample for examination via histological analysis, as previously reported 34-36.</li>
<li>	Use hematoxylin and eosin (H&amp;E) stains to examine the tissue for the absence of nuclei. One could also use a fluorescent Hoescht 33342 stain (1.0 &mu;g/mL) for DNA to verify the H&amp;E results; briefly, fix the sections, rehydrate and rinse in water, stain for 10 minutes, and then rinse well and store in the dark. Alternatively, one can use a kit such as the Qiagen DNeasy Blood and Tissue kit, which is designed to quantify the total DNA content of a sample.</li>
</ol>
2. Preparation of Injectable ECM
<ol class="lalpha">
<li>	Lyophilization
<ol class="lroman">
<li>	After appropriate preparation, freeze the decellularized samples with liquid nitrogen or by storing at -80 &deg;C. Lyophilize the samples until completely dry. Depending on your system and the water content of your samples, this may take anywhere from 12 to 72 hours.</li>
</ol>
</li>
<li>	Milling
<ol class="lroman">
<li>	Once completely dry, a Wiley Mini Mill is used to grind the dry ECM into a fine powder. Choose the appropriate sieve size for your purposes; here, the 40 gauge mesh is used. Once the majority of the sample has come through the filter, remove the collection jar with the milled ECM powder. If there is only a small sample, the rest of the sample can be extracted from the mill in a variety of ways; here, a Q-tip is used to remove the ECM stuck behind the stationary blades of the mill.</li>
<li>	Always make sure that the mill is clean and free of debris. It is also important to make sure that both the mill and your sample are totally dry; any remaining moisture will cause the ECM to aggregate and clump together, preventing successful milling. The ECM may pick up moisture from the air and so should go directly from the lyophilizer to the mill.</li>
</ol>
</li>
<li>	Digestion
<ol class="lroman">
<li>	To form the injectable ECM, the powder is digested pepsin. The following protocol was modified from Freyetes, et al. (2).</li>
<li>	It is important to maintain as sterile a product as possible, as it will be injected in vivo and contamination may cause complications. Thus, use sterile vials and weigh spoons and filter all solutions before use. A lyophilization step after milling and before digestion will also help maintain sterility.</li>
<li>	Weigh out the desired amount of ECM powder into an appropriate scintillation vial. For total volumes smaller than 1 mL, it is recommended you use a 2 mL vial and for larger volumes, a 20 mL vial. This ensures that there is enough liquid in the bottom of the vial to stir effectively.</li>
<li>	Weigh out the desired amount of pepsin into a scintillation vial. Add 0.1 M HCl such that the pepsin is at a concentration of 1 mg/mL. Make sure the pepsin is fully dissolved (no particulates) before use this can be hastened by vortexing the pepsin solution.</li>
<li>	Add the pepsin/HCl solution to the scintillation vial with the ECM powder so that the ECM is at a concentration of 10 mg/mL.</li>
<li>	Stir solution continuously for 60-65 hours, periodically scraping down the sides of the vial with a weigh spoon or spatula.</li>
</ol>
</li>
<li>	pH Adjustment
<ol class="lroman">
<li>	This step is performed to bring the liquid matrix material to physiological pH and to inactivate the pepsin, keeping it from cleaving the ECM further. Again, be sure to use filtered solutions made with Millipore water.</li>
<li>	1 M NaOH is added to be 1/10 the original volume. Test the pH at this point and add small amounts (2-10 &mu;l) of NaOH or HCl such that the desired pH (7.4) is reached. Keep track of all small amounts added for neutralization or removed for pH testing. Once the solution has been neutralized, add 10x PBS to be 1/10 the final volume (1/9 the current volume). Determine the concentration of the injectable ECM and then add 1x PBS to reach desired final concentration, here 6 mg/mL.</li>
<li>	Characterization of the injectable ECM can be done via SDS-PAGE, Blyscan Colorimetric GAG Assay, and mass spectroscopy.</li>
<li>	At this point, the matrix material can be injected in vivo to form a scaffold for myocardial tissue engineering.</li>
</ol>
</li> 
<li>	Biotin-Labeling
<ol class="lroman">
<li>	The ECM can be tagged with biotin before injection for easy visualization of the injected ECM.</li>
<li>	Prepare a 10mM solution of Biotin and add to the injectable ECM at a ratio of 0.3mg/mg. The ECM should be at the desired concentration for injection. If injecting at a concentration of 6 mg/mL, add 40 &mu;L Biotin per 1 mL of ECM. Prior to injection, keep the biotin-ECM solution on ice for at least 1 hour.</li>
</ol>
</li>
<li>	All processing steps can be performed at room temperature. To keep the injectable matrix from gelling, the pH adjustment step may be done on ice. </li>
</ol>
3. Myocardial Injections
<ol class="lalpha">
<li>Injection preparation
<ol class="lroman">
<li>For the female rats (225-250 g) used here, 75 &mu;L injections of pH 7.4 injectable ECM were prepared in 0.1 mL syringes tipped with 30 gauge needles. If male rats (375-400 g) were used, 90 &mu;L could be injected.</li>
<li>All surgical supplies should be sterilized prior to surgery; here we use an autoclaved surgical pack that contains all the necessary tools.</li>
</ol>
</li>
<li>On the day of surgery, a Harlan Sprague-Dawley rat is anesthetized using isoflurane at 5%, intubated using an otoscope, and then maintained at 2.5% isoflurane throughout the procedure.</li>
<li>Add artificial tear ointment to the animal's eyes to protect against dryness and hair. Administer 3 mL of Lactated Ringers solution for hydration during surgery. This should be done via subcutaneous injection in the lower abdominal area.</li>
<li>Place the animal in supine position on the surgical table and gently tape down limbs. Use clippers to remove the hair on the abdomen and vacuum free hair prior to scrubbing.</li>
<li>Make a series of small injections (approximately 50 &mu;L each) of 2% lidocaine along a diagonal from the zyphoid process to the lower right-hand side of the abdomen. Then scrub three times with betadine, starting in the middle and moving outward. Repeat with 70% ethanol. Cover the animal using a surgical drape with a pre-made circular window; secure with towel clamps, if necessary.</li>
<li>Using a No. 10 scalpel make a 3-4 cm incision from the xyphoid process to the lower right-hand side of the abdomen. Locate xyphoid process and dissect vertically down through the muscle to the right, being careful to avoid the large vessel on the right. Once you have dissected through the muscle, use scissors to cut laterally through the muscle, exposing the diaphragm. Be careful to avoid the liver.</li>
<li>Using a 36 inch length of 3-0 Vicrile suture and a pair of hemostats, drive one needle through the zyphoid process and pull the suture halfway through. Tape the ends of the suture together and fix that to a point above and behind the animal essentially lifting the zyphoid up and exposing the diaphragm. With another 36 inch length of 3-0 Vicrile suture, drive a needle through the muscle closest to the xyphoid process and, again, pull through and fix the ends to another point to the right of the surgical table, fully exposing the surgical cavity.</li>
<li>At this point, you should be able to see the heart through the diaphragm. Using a small pair of rat tooth microforceps, grab the center of diaphragm, pull it out toward you and make a very small incision with blunt scissors. It is extremely important to use very blunt scissors in order to avoid poking the lungs. Once this hole has been made, the lungs will retract, allowing you to make a 2-3 cm incision vertically through the diaphragm to visualize the heart.</li>
<li>Insert a 3-inch Q-tip to the left of the cavity to push the lungs out of the way. Use a towel clamp to secure the Q-tip in place. Use another Q-tip to move the right lung out of the way in order to locate the pericardial sac. Using a pair of microforceps and a pair of large serrated forceps, tear through the pericardial sac, exposing the apex of the heart. If the animal has previously undergone an infarction procedure, the pericardium will already be absent.</li>
<li>Grab the apex of the heart with the rat tooth microforceps and inject the liquid matrix material a third of the way up from the apex. Insert the needle parallel to the epicardial surface and inject with the beveled edge of the needle to the left. Be sure to avoid the great cardiac vein. Wait a couple seconds before removing the needle. You should see a whitening of the tissue at the site of injection.</li>
<li>Clean up blood in the cavity and remove the towel clamp and Q-tip that was holding the right lung. Use curved microhemostats and the rat tooth microforceps to close the diaphragm. Make an initial knot and then use the serrated forceps and microhemostats to close the diaphragm with a continuous suture and a tapered needle. Before closing entirely, insert PE160 suction tubing and use a 10 mL syringe to evacuate the chest cavity as the suture is tightened.</li>
<li>When properly evacuated and closed, the diaphragm should be concave. A convex diaphragm indicates that there is still air in the cavity.</li>
<li>At this point the isoflurane can be turned down to 1%.</li>
<li>Take out both 3-0 Vicrile sutures holding the cavity open. Use sterile water to hydrate the muscle, wiping excess away with a Q-tip or sterile gauze. Use a new length of 3-0 suture to close the muscle layer with intermittent sutures.</li>
<li>At this point, the isoflurane can be turned off.</li>
<li>Clean area with sterile water and use staples to close skin. If staples will interfere with the study, 5-0 proline suture can also be used. In that case, use a reverse cutting (RC) needle to close the skin with intermittent sutures. In both cases, spot surgical glue over the closed incision.</li>
<li>Use a Q-tip to anoint the incision site with a triple-antibiotic ointment.</li>
<li>Allow the animal to recover under 100% oxygen.</li>
<li>When the animals starts to breathe around the ventilator, remove the trachea tube.</li>
<li>Once the animals are sternal, administer a subcutaneous injection of 0.05 mg/kg of buprenorphine hydrochloride and return them to their home cage on a sterile towel.</li>
<li>The animals should receive post-operative observation and care.</li>
<li>At the time points of interest, the animals are euthanized and the hearts removed for analysis. Short axis cross sections are taken and can be stained with hematoxylin and eosin (H&amp;E) for gross tissue analysis. At short time points, the injected region will appear as a pink, fibrous network that has spread interstitially. At later time points, one will observe an influx of cells and an eventual degradation of the injected matrix after 2 to 3 weeks. If the ECM was labeled with biotin before injecting, it can be more directly visualized by staining the biotin with HRP-conjugated streptavidin. Immunohistochemical staining can be used to identify specific cells or structures in the injection region, here the slides have been co-stained with a FITC-conjugated isolectin that binds to endothelial cells and fluoresces green and an anti-smooth muscle actin antibody with an Alexafluor secondary antibody that reports the SMCs in red.</li>
</ol>
4. Representative Results
Previous characterization of matrix materials prepared in this way demonstrated the retention of a variety of macromolecules. Specifically, multiple fibrous proteins and glycoproteins were identified via mass spectrometry (Table 1). If the matrix materials processed according to this protocol are found to no longer contain a complex array of macromolecules, it could be that the decellularization protocol employed is too harsh. Once fully lyophilized, the dried pericardial ECM looks like very stiff, crumpled paper. Other tissue types will resemble packing peanuts. If milled well, the ECM should fall through the sieve and be collected in the jar at the bottom. If there is moisture left in the sample, the ECM will clump together and get stuck in the milling chamber. After digestion, the ECM solution should be a milky color and completely free of visible particulates. It will also be slightly more viscous than the HCl alone. If the ECM does not digest well, there will still be large particulates in the vial; alternatively, the ECM may crash out of solution. After pH adjustment, there is no visible change in the solution, and the ECM is still a cloudy, homogeneous liquid. If it starts to gel, the viscosity will increase and it will be difficult or impossible to draw the material up into a syringe. If this happens, perform the pH adjustment step on ice, with pre-chilled solutions. Once the injections have been prepared, keep them on ice until use. If the injection is successful, the region around the needle tip whitens and a small bolus is visible. If this is not observed, the injection may have gone into the chamber instead of into the wall. When suturing the animal up after injection, careful closing of the diaphragm is the most important step. If done correctly, the diaphragm will be tight against the lungs and will appear concave. If there is any air left inside the lungs, the diaphragm will remain convex, or have an area where it balloons out. When examining the histology of the injection region at an early time point (30 minutes to 4 hours post-injection), one should see a pink, fibrous area that is devoid of cells this is the reassembled matrix material post-injection (Figure 2). The network often spreads interstitially and may be present throughout many sections. A possible explanation for seeing only a very small area of matrix gel is that a portion of the injection missed the intramural target and was either injected into the LV chamber or leaked out of the injection location. Additionally, depending on the gelation time, the interstitial spread may vary.
<img src="/files/ftp_upload/2109/2109fig1.jpg" alt="Figure 1" /> 
Figure 1. Polyacrylamide gel electrophoresis (PAGE) results. (A) Molecular weight standard. (B) Rat tail collagen Type I (2.5 mg/mL) compared to the solubilized human (C) and porcine (D) pericardial ECM (7 mg/mL). Note the presence of collagen as well as several other proteins and peptides in the pericardial matrix samples.
<img src="/files/ftp_upload/2109/2109table1.jpg" alt="Table 1" /> 
Table 1. ECM components identified with mass spectroscopy.
<img src="/files/ftp_upload/2109/2109fig2.jpg" alt="Figure 2" /> 
Figure 2. Myocardial injections: in vivo gelation. H&amp;E stain of human (A) and porcine (B) pericardial matrix injections that have gelled in vivo after 45 minutes. Arrows denote matrix location, stained lighter pink than the myocardium. Scale bar is 500 &mu;m.
<img src="/files/ftp_upload/2109/2109fig3.jpg" alt="Figure 3" /> 
Figure 3.Vascular cell infiltration. Fluorescent stains for vessels in the injected human (A-C) and porcine (D-F) matrix gels at two weeks. Endothelial cells are labeled green (A, D), while smooth muscle cells are labeled red (B, E). Merged images are shown in C and F. Scale bar is 100 &mu;m. The white dotted lines indicate the area of matrix injection, as determined by H&amp;E analysis of a nearby section.
<img src="/files/ftp_upload/2109/2109fig4.jpg" alt="Figure 4" /> 
Figure 4. Stem cells within matrix injection region. A Hoescht stain for nuclei (blue) and c-kit (green) identifies stem cells in the human (A) and porcine (B) matrix injection regions. Scale bar is 50 &mu;m.

<ol>
	<li>Seif-Naraghi, S. B., Salvatore, M. A., Magoffin-Schup, P. J., Hu, D. P., Christman, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+and+characterization+of+an+injectable+pericardial+matrix+gel:+A+potentially+autologous+scaffold+for+cardiac+tissue+engineering.">Design and characterization of an injectable pericardial matrix gel: A potentially autologous scaffold for cardiac tissue engineering.</a> Tissue Engineering. , (2009).</li><li>Freytes, D. O., Martin, J., Velankar, S. S., Lee, A. S., Badylak, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+and+rheological+characterization+of+a+gel+form+of+the+porcine+urinary+bladder+matrix.">Preparation and rheological characterization of a gel form of the porcine urinary bladder matrix.</a> Biomaterials. 29, 1630-1630 (2008).</li><li>Gilbert, T. W., Sellaro, T. L., Badylak, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decellularization+of+tissues+and+organs.">Decellularization of tissues and organs.</a> Biomaterials. 27, 3675-3675 (2006).</li><li>Liao, J., Joyce, E. M., Sacks, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+decellularization+on+the+mechanical+and+structural+properties+of+the+porcine+aortic+valve+leaflet.">Effects of decellularization on the mechanical and structural properties of the porcine aortic valve leaflet.</a> Biomaterials. 29, 1065-1065 (2008).</li><li>Singelyn, J. M., DeQuach, J. A., Seif-Naraghi, S. B., Littlefield, R. B., Schup-Magoffin, P. J., Christman, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Naturally+derived+myocardial+matrix+as+an+injectable+scaffold+for+cardiac+tissue+engineering.">Naturally derived myocardial matrix as an injectable scaffold for cardiac tissue engineering.</a> Biomaterials. 30, 5409-5409 (2009).</li><li>Christman, K. L., Vardanian, A. J., Fang, Q., Sievers, R. E., Fok, H. H., Lee, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Injectable+fibrin+scaffold+improves+cell+transplant+survival,+reduces+infarct+expansion,+and+induces+neovasculature+formation+in+ischemic+myocardium.">Injectable fibrin scaffold improves cell transplant survival, reduces infarct expansion, and induces neovasculature formation in ischemic myocardium.</a> J Am Coll Cardiol. 44, 654-654 (2004).</li><li>Christman, K. L., Fok, H. H., Sievers, R. E., Fang, Q., Lee, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibrin+glue+alone+and+skeletal+myoblasts+in+a+fibrin+scaffold+preserve+cardiac+function+after+myocardial+infarction.">Fibrin glue alone and skeletal myoblasts in a fibrin scaffold preserve cardiac function after myocardial infarction.</a> Tissue Eng. 10, 403-410 (2004).</li><li>Huang, N. F., Sievers, R. E., Park, J. S., Fang, Q., Li, S., Lee, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rodent+model+of+myocardial+infarction+for+testing+the+efficacy+of+cells+and+polymers+for+myocardial+reconstruction.">A rodent model of myocardial infarction for testing the efficacy of cells and polymers for myocardial reconstruction.</a> Nat Protoc. (1), 1596-1609 (2006).</li><li>Ott, H. C., Matthiesen, T. S., Goh, S. K., Black, L. D., Kren, S. M., Netoff, T. I. <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+matrix:+using+nature's+platform+to+engineer+a+bioartificial+heart.">Perfusion-decellularized matrix: using nature's platform to engineer a bioartificial heart.</a> Nat Med. 14, 213-221 (2008).</li><li>Badylak, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+extracellular+matrix+as+a+biologic+scaffold+material.">The extracellular matrix as a biologic scaffold material.</a> Biomaterials. 28, 3587-3593 (2007).</li></ol>

Animal Use: All experiments in this study were performed in accordance with the guidelines published by the Institutional Animal Care and Use Committee at the University of California, San Diego and the American Association for Accreditation of Laboratory Animal Care.

This method allows for the generation of biologically derived, injectable scaffolds for myocardial tissue engineering. Although these methods were initially developed for the fabrication and in vivo testing of a myocardial matrix gel and presented here with a pericardial matrix gel, this protocol can be adapted for use with any tissue, provided the tissue can be appropriately decellularized. Decellularization should be performed and verified prior to the use of these methods, as the presence of DNA in the matr...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Reagents:</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Pepsin</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>p6887-1G</td>
<td>Lyophilized </td>
</tr>
<tr>
<td>Biotin</td>
<td>&nbsp;</td>
<td>Thermo Scientific</td>
<td>21217</td>
<td>&nbsp;</td>
<tr>
<td>Neutravidin-HRP</td>
<td>&nbsp;</td>
<td>Thomas Scientific</td>
<td>21130</td>
<td>&nbsp;</td>
<tr>
<td>Equipment: </td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Wiley Mini Mill</td>
<td>&nbsp;</td>
<td>Thomas Scientific</td>
<td>3383L10</td>
<td>&nbsp;</td>
<tr>
<td>Labconco Lyophilizer</td>
<td>&nbsp;</td>
<td>Labconco, Inc</td>
<td>7670520</td>
<td>&nbsp;</td>
<tr>
<td>Surgical supplies:</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Betadine</td>
<td>&nbsp;</td>
<td>Purdue Products, L.P.</td>
<td>67618-154-16</td>
<td>&nbsp;</td>
<tr>
<td>Lactated Ringers Solution</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>003966</td>
<td>&nbsp;</td>
<tr>
<td>KY Jelly</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>28658</td>
<td>&nbsp;</td>
<tr>
<td>Lidocaine, 2%</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>17767</td>
<td>&nbsp;</td>
<tr>
<td>Buprenorphine hydrochloride</td>
<td>&nbsp;</td>
<td>Reckitt Benckiser Healthcare (UK) Ltd.</td>
<td>12496-0757-1</td>
<td>&nbsp;</td>
<tr>
<td>Artificial tear ointment</td>
<td>&nbsp;</td>
<td>Fisher </td>
<td>NC9860843</td>
<td>&nbsp;</td>
<tr>
<td>Triple antibiotic ointment</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>19082795</td>
<td>&nbsp;</td>
<tr>
<td>Isoflurane</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>60307-120-25</td>
<td>&nbsp;</td>
<tr>
<td>Otoscope </td>
<td>&nbsp;</td>
<td>MWI</td>
<td>008699</td>
<td>&nbsp;</td>
<tr>
<td>Stop cock</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>006245</td>
<td>&nbsp;</td>
<tr>
<td>3-0 Vicrile suture</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>J327H</td>
<td>&nbsp;</td>
<tr>
<td>5-0 Proline suture</td>
<td>&nbsp;</td>
<td>MWI</td>
<td>s-1173</td>
<td>&nbsp;</td>
<tr>
<td>Reverse cutting (RC) needle</td>
<td>&nbsp;</td>
<td>Ethicon</td>
<td>8684G</td>
<td>&nbsp;</td>
<tr>
<td>Microhemostats </td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>13013-14</td>
<td>&nbsp;</td>
<tr>
<td>Rat tooth microforceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11084-07</td>
<td>&nbsp;</td>
<tr>
<td>No. 10 scalpel</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>10110-01</td>
<td>&nbsp;</td>
<tr>
<td>Blunt scissors</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>14108-09</td>
<td>&nbsp;</td>
<tr>
<td>Sharp, curved scissors</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>14085-08</td>
<td>&nbsp;</td>
<tr>
<td>Large, serrated forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>1106-12</td>
<td>&nbsp;</td>
<tr>
<td>PE160 suction tubing</td>
<td>&nbsp;</td>
<td>BD</td>
<td>427430 </td>
<td>&nbsp;</td>
<tr>
<td>Clippers </td>
<td>&nbsp;</td>
<td>MWI</td>
<td>21608</td>
<td>&nbsp;</td>
<tr>
<td>Skin staples/stapler </td>
<td>&nbsp;</td>
<td>Ethicon</td>
<td>PRR35</td>
<td>&nbsp;</td>
<tr>
<td>General supplies:</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Stir plates</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>0.1 M HCl</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>1 M NaOH</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>10x PBS</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>1x PBS</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>70% Ethanol</td>
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<td>0.1 mL syringes</td>
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<td>10 mL syringe</td>
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<td>Surgical glue</td>
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<td>Surgical drape</td>
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<td>Towel clamps</td>
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<td>Small hand-held vacuum </td>
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fabrication of biologically derived injectable materials for myocardial tissue engineering

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications. Briefly, decellularized tissue is lyophilized, milled, enzymatically digested, and then brought to physiological pH. The lyophilization removes all water content from the tissue, resulting in dry ECM that can be ground into a fine powder with a small mill. After milling, the ECM powder is digested with pepsin to form an injectable matrix. After adjustment to pH 7.4, the liquid matrix material can be injected into the myocardium. Results of previous characterization assays have shown that matrix gels produced from decellularized pericardial and myocardial tissue retain native ECM components, including diverse proteins, peptides and glycosaminoglycans. Given the use of this material for tissue engineering, in vivo characterization is especially useful; here, a method for performing an intramural injection into the left ventricular (LV) free wall is presented as a means of analyzing the host response to the matrix gel in a small animal model. Access to the chest cavity is gained through the diaphragm and the injection is made slightly above the apex in the LV free wall. The biologically derived scaffold can be visualized by biotin-labeling before injection and then staining tissue sections with a horse radish peroxidase-conjugated neutravidin and visualizing via diaminobenzidine (DAB) staining. Analysis of the injection region can also be done with histological and immunohistochemical staining. In this way, the previously examined pericardial and myocardial matrix gels were shown to form fibrous, porous networks and promote vessel formation within the injection region.

Watch this Scientific Journal Video about Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering at JoVE.com

Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering

Methods for preparing an injectable matrix gel from decellularized tissue and injecting it into rat myocardium in vivo are described.

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications.  ...

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Research

JoVE Journal

Bioengineering

14.9K Views.  University of California, San Diego. Methods for preparing an injectable matrix gel from decellularized tissue and injecting it into rat myocardium in vivo are described.

Video: Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

Postproduction Processing of Electrospun Fibres for Tissue Engineering

Electrospinning is a commonly used and versatile method to produce scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo undergo biaxial distension to varying extents such as skin, bladder, pelvic floor and even the hard palate as children grow. In producing scaffolds for these purposes there is a need to develop scaffolds of appropriate biomechanical properties (whether achieved without or with cells) and which are sterile for clinical use. The focus of this paper is not how to establish basic electrospinning parameters (as there is extensive literature on electrospinning) but on how to modify spun scaffolds post production to make them fit for tissue engineering purposes - here thickness, mechanical properties and sterilisation (required for clinical use) are considered and we also describe how cells can be cultured on scaffolds and subjected to biaxial strain to condition them for specific applications.
Electrospinning tends to produce thin sheets; as the electrospinning collector becomes coated with insulating fibres it becomes a poor conductor such that fibres no longer deposit on it. Hence we describe approaches to produce thicker structures by heat or vapour annealing increasing the strength of scaffolds but not necessarily the elasticity. Sequential spinning of scaffolds of different polymers to achieve complex scaffolds is also described. Sterilisation methodologies can adversely affect strength and elasticity of scaffolds. We compare three methods for their effects on the biomechanical properties on electrospun scaffolds of poly lactic-co-glycolic acid (PLGA).
Imaging of cells on scaffolds and assessment of production of extracellular matrix (ECM) proteins by cells on scaffolds is described. Culturing cells on scaffolds in vitro can improve scaffold strength and elasticity but the tissue engineering literature shows that cells often fail to produce appropriate ECM when cultured under static conditions. There are few commercial systems available that allow one to culture cells on scaffolds under dynamic conditioning regimes - one example is the Bose Electroforce 3100 which can be used to exert a conditioning programme on cells in scaffolds held using mechanical grips within a media filled chamber.4 An approach to a budget cell culture bioreactor for controlled distortion in 2 dimensions is described. We show that cells can be induced to produce elastin under these conditions. Finally assessment of the biomechanical properties of processed scaffolds cultured with or without cells is described.

Electrospun scaffolds can be processed post production for tissue engineering applications. Here we describe methods for spinning complex scaffolds (by consecutive spinning), for making thicker scaffolds (by multi-layering using heat or vapour annealing), for achieving sterility (aseptic production or sterilisation post production) and for achieving appropriate biomechanical properties.

Electrospinning is a commonly used and versatile method to produce  scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Design and Construction of Artificial Extracellular Matrix (aECM) Proteins from Escherichia coli for Skin Tissue Engineering

Recombinant technology is a versatile platform to create novel artificial proteins with tunable properties. For the last decade, many artificial proteins that have incorporated functional domains derived from nature (or created de novo) have been reported. In particular, artificial extracellular matrix (aECM) proteins have been developed; these aECM proteins consist of biological domains taken from fibronectin, laminins and collagens and are combined with structural domains including elastin-like repeats, silk and collagen repeats. To date, aECM proteins have been widely investigated for applications in tissue engineering and wound repair. Recently, Tjin and coworkers developed integrin-specific aECM proteins designed for promoting human skin keratinocyte attachment and propagation. In their work, the aECM proteins incorporate cell binding domains taken from fibronectin, laminin-5 and collagen IV, as well as flanking elastin-like repeats. They demonstrated that the aECM proteins developed in their work were promising candidates for use as substrates in artificial skin. Here, we outline the design and construction of such aECM proteins as well as their purification process using the thermo-responsive characteristics of elastin.

Design and Construction of Artificial Extracellular Matrix aECM Proteins from Escherichia coli for Skin Tissue Engineering

Recombinant technologies have enabled material designers to create novel artificial proteins with customized functionalities for tissue engineering applications. For example, artificial extracellular matrix proteins can be designed to incorporate structural and biological domains derived from native ECMs. Here, we describe the construction and purification of aECM proteins containing elastin-like repeats.

Recombinant technology is a versatile platform to create novel artificial proteins with tunable properties. For the last decade, many artificial proteins ...

Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer (SALB) Method

In order to mimic cell membranes, the supported lipid bilayer (SLB) is an attractive platform which enables in vitro investigation of membrane-related processes while conferring biocompatibility and biofunctionality to solid substrates. The spontaneous adsorption and rupture of phospholipid vesicles is the most commonly used method to form SLBs. However, under physiological conditions, vesicle fusion (VF) is limited to only a subset of lipid compositions and solid supports. Here, we describe a one-step general procedure called the solvent-assisted lipid bilayer (SALB) formation method in order to form SLBs which does not require vesicles. The SALB method involves the deposition of lipid molecules onto a solid surface in the presence of water-miscible organic solvents (e.g., isopropanol) and subsequent solvent-exchange with aqueous buffer solution in order to trigger SLB formation. The continuous solvent exchange step enables application of the method in a flow-through configuration suitable for monitoring bilayer formation and subsequent alterations using a wide range of surface-sensitive biosensors. The SALB method can be used to fabricate SLBs on a wide range of hydrophilic solid surfaces, including those which are intractable to vesicle fusion. In addition, it enables fabrication of SLBs composed of lipid compositions which cannot be prepared using the vesicle fusion method. Herein, we compare results obtained with the SALB and conventional vesicle fusion methods on two illustrative hydrophilic surfaces, silicon dioxide and gold. To optimize the experimental conditions for preparation of high quality bilayers prepared via the SALB method, the effect of various parameters, including the type of organic solvent in the deposition step, the rate of solvent exchange, and the lipid concentration is discussed along with troubleshooting tips. Formation of supported membranes containing high fractions of cholesterol is also demonstrated with the SALB method, highlighting the technical capabilities of the SALB technique for a wide range of membrane configurations.

Biomembrane Fabrication by the Solvent-assisted Lipid Bilayer SALB Method

We present an experimental protocol to form a supported lipid bilayer on solid substrates without using lipid vesicles. We demonstrate a one-step method to form a lipid bilayer on silicon dioxide and gold as well as supported membranes with cholesterol-enriched domain for various biological applications.

In order to mimic cell membranes, the supported lipid bilayer (SLB) is an attractive platform which enables in vitro investigation of membrane-related ...

Fabrication of Mechanically Tunable and Bioactive Metal Scaffolds for Biomedical Applications

Biometal systems have been widely used for biomedical applications, in particular, as load-bearing materials. However, major challenges are high stiffness and low bioactivity of metals. In this study, we have developed a new method towards fabricating a new type of bioactive and mechanically reliable porous metal scaffolds-densified porous Ti scaffolds. The method consists of two fabrication processes, 1) the fabrication of porous Ti scaffolds by dynamic freeze casting, and 2) coating and densification of the porous scaffolds. The dynamic freeze casting method to fabricate porous Ti scaffolds allowed the densification of porous scaffolds by minimizing the chemical contamination and structural defects. The densification process is distinctive for three reasons. First, the densification process is simple, because it requires a control of only one parameter (degree of densification). Second, it is effective, as it achieves mechanical enhancement and sustainable release of biomolecules from porous scaffolds. Third, it has broad applications, as it is also applicable to the fabrication of functionally graded porous scaffolds by spatially varied strain during densification.

Bioactive and mechanically reliable metal scaffolds have been fabricated through a method which consists of two processes, dynamic freeze casting for the fabrication of porous Ti, and coating and densification of the Ti scaffolds. The densification process is simple, effective and applicable to the fabrication of functionally graded scaffolds.

Biometal systems have been widely used for biomedical applications, in particular, as load-bearing materials. However, major challenges are high stiffness ...

Fabrication of Inverted Colloidal Crystal Poly(ethylene glycol) Scaffold: A Three-dimensional Cell Culture Platform for Liver Tissue Engineering

The ability to maintain hepatocyte function in vitro, for the purpose of testing xenobiotics&#39; cytotoxicity, studying virus infection and developing drugs targeted at the liver, requires a platform in which cells receive proper biochemical and mechanical cues. Recent liver tissue engineering systems have employed three-dimensional (3D) scaffolds composed of synthetic or natural hydrogels, given their high water retention and their ability to provide the mechanical stimuli needed by the cells. There has been growing interest in the inverted colloidal crystal (ICC) scaffold, a recent development, which allows high spatial organization, homotypic and heterotypic cell interaction, as well as cell-extracellular matrix (ECM) interaction. Herein, we describe a protocol to fabricate the ICC scaffold using poly (ethylene glycol) diacrylate (PEGDA) and the particle leaching method. Briefly, a lattice is made from microsphere particles, after which a pre-polymer solution is added, properly polymerized, and the particles are then removed, or leached, using an organic solvent (e.g., tetrahydrofuran). The dissolution of the lattice results in a highly porous scaffold with controlled pore sizes and interconnectivities that allow media to reach cells more easily. This unique structure allows high surface area for the cells to adhere to as well as easy communication between pores, and the ability to coat the PEGDA ICC scaffold with proteins also shows a marked effect on cell performance. We analyze the morphology of the scaffold as well as the hepatocarcinoma cell (Huh-7.5) behavior in terms of viability and function to explore the effect of ICC structure and ECM coatings. Overall, this paper provides a detailed protocol of an emerging scaffold that has wide applications in tissue engineering, especially liver tissue engineering.

Fabrication of Inverted Colloidal Crystal Polyethylene glycol Scaffold: A Three-dimensional Cell Culture Platform for Liver Tissue Engineering

This manuscript presents a detailed protocol for the fabrication of an emerging three-dimensional hepatocyte culture platform, the inverted colloidal crystal scaffold, and the concomitant techniques to assess hepatocyte behavior. The size-controllable pores, interconnectivity and ability to conjugate extracellular matrix proteins to the poly(ethylene glycol) (PEG) scaffold enhance Huh-7.5 cell performance.

The ability to maintain hepatocyte function in vitro, for the purpose of testing xenobiotics&#39; cytotoxicity, studying virus infection and developing ...

Fabrication of Extracellular Matrix-derived Foams and Microcarriers as Tissue-specific Cell Culture and Delivery Platforms

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus of this article is on the methods for fabricating ECM-derived porous foams and microcarriers for use as biologically-relevant substrates in advanced 3D in vitro cell culture models or as pro-regenerative scaffolds and cell delivery systems for tissue engineering and regenerative medicine. Using decellularized tissues or purified insoluble collagen as a starting material, the techniques can be applied to synthesize a broad array of tissue-specific bioscaffolds with customizable geometries. The approach involves mechanical processing and mild enzymatic digestion to yield an ECM suspension that is used to fabricate the three-dimensional foams or microcarriers through controlled freezing and lyophilization procedures. These pure ECM-derived scaffolds are highly porous, yet stable without the need for chemical crosslinking agents or other additives that may negatively impact cell function. The scaffold properties can be tuned to some extent by varying factors such as the ECM suspension concentration, mechanical processing methods, or synthesis conditions. In general, the scaffolds are robust and easy to handle, and can be processed as tissues for most standard biological assays, providing a versatile and user-friendly 3D cell culture platform that mimics the native ECM composition. Overall, these straightforward methods for fabricating customized ECM-derived foams and microcarriers may be of interest to both biologists and biomedical engineers as tissue-specific cell-instructive platforms for in vitro and in vivo applications.

The tissue-specific extracellular matrix (ECM) is a key mediator of cell function. This article describes methods for synthesizing pure ECM-derived foams and microcarriers that are stable in culture without the need for chemical crosslinking for applications in advanced 3D in vitro cell culture models or as pro-regenerative bioscaffolds.

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus ...

Isolation of Murine Adipose Tissue-derived Microvascular Fragments as Vascularization Units for Tissue Engineering

A functional microvascular network is of pivotal importance for the survival and integration of engineered tissue constructs. For this purpose, several angiogenic and prevascularization strategies have been established. However, most cell-based approaches include time-consuming in vitro steps for the formation of a microvascular network. Hence, they are not suitable for intraoperative one-step procedures. Adipose tissue-derived microvascular fragments (ad-MVF) represent promising vascularization units. They can be easily isolated from fat tissue and exhibit a functional microvessel morphology. Moreover, they rapidly reassemble into new microvascular networks after in vivo implantation. In addition, ad-MVF have been shown to induce lymphangiogenesis. Finally, they are a rich source of mesenchymal stem cells, which may further contribute to their high vascularization potential. In previous studies we have demonstrated the remarkable vascularization capacity of ad-MVF in engineered bone and skin substitutes. In the present study, we report on a standardized protocol for the enzymatic isolation of ad-MVF from murine fat tissue.

We present a protocol to isolate adipose tissue-derived microvascular fragments that represent promising vascularization units. They can be rapidly isolated, do not require in vitro processing and, thus, may be used for one-step prevascularization in different fields of tissue engineering.

A functional microvascular network is of pivotal importance for the survival and integration of engineered tissue constructs. For this purpose, several ...

Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human patients. Human Organ-on-a-Chip (Organ Chip) microfluidic cell culture devices, which provide an experimental in vitro platform to assess efficacy, toxicity, and pharmacokinetic (PK) profiles in humans, may be better predictors of therapeutic efficacy and safety in the clinic compared to animal studies. These devices may be used to model the function of virtually any organ type and can be fluidically linked through common endothelium-lined microchannels to perform in vitro studies on human organ-level and whole body-level physiology without having to conduct experiments on people. These Organ Chips consist of two perfused microfluidic channels separated by a permeable elastomeric membrane with organ-specific parenchymal cells on one side and microvascular endothelium on the other, which can be cyclically stretched to provide organ-specific mechanical cues (e.g., breathing motions in lung). This protocol details the fabrication of flexible, dual channel, Organ Chips through casting of parts using 3D printed molds, enabling combination of multiple casting and post-processing steps. Porous poly (dimethyl siloxane) (PDMS) membranes are cast with micrometer sized through-holes using silicon pillar arrays under compression. Fabrication and assembly of Organ Chips involves equipment and steps that can be implemented outside of a traditional cleanroom. This protocol provides researchers with access to Organ Chip technology for in vitro organ- and body-level studies in drug discovery, safety and efficacy testing, as well as mechanistic studies of fundamental biological processes.

Here, we present a protocol that describes the fabrication of stretchable, dual channel, organ chip microfluidic cell culture devices for recapitulating organ-level functionality in vitro.

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human ...