The authors would like to acknowledge Brogan Guest, Michelle Weaver, and Kristen Lippert. Funding for this study was provided by NIH Grant R03EB009237, as well as NIH Training Grants T32EB001026-06 from the National Institute of Biomedical Imaging And Bioengineering and T32HL076124-05.

1. Tissue Preparation and Experiment Setup
<ol>
	<li>Harvest porcine organ immediately after euthanasia from an abattoir or research facility and rinse off excess blood. Trim the heart of excess fat and tissue, keeping the atria and aorta intact. Trim away fat to separate the pulmonary artery from the aorta. If there are any cuts in the tissue, discard appropriately.</li>
	<li>Wrap each heart individually in freezer paper and store all tissue in a -80 &deg;C freezer for at least 24 hr to ensure complete freezing.</li>
	<li>When ready for use (usually less than 3 months), thaw one intact frozen porcine heart in Type 1 water overnight submerged in a 4 L beaker at 4 &deg;C.</li>
	<li>After the heart is completely thawed, pat the heart dry, weigh the heart, and record the weight. The heart of a market weight pig should weigh approximately 375-450 g.</li>
	<li>Connect size 18 Masterflex tubing to the &frac14;" end of a barbed reducer. Insert the barbed reducer and tubing inside the aorta. Place 2 hose clamps or secure zip ties around the aorta, just below the brachiocephalic trunk. The reducer and tubing must remain above the aortic valve, so the coronary arteries can be perfused (Figure 1).</li>
	<li>Use a 30 or 60 ml syringe to fill the tubing with Type I water. Insert the tubing within the cartridge of a Masterflex roller pump at its approximate midpoint. Submerge the inflow end of the tubing in the bottom of a 4 L beaker filled with 2.5 L of water and secure the tubing.</li>
	<li>Place the heart in the beaker filled with water, and prime the pump to remove air bubbles. If bubbles are observed coming from the aorta where the tubing is inserted, the aorta may need to be repositioned or secured with additional ties. An airtight seal is important to maintain adequate pressure during the decellularization process (Figure 2).</li>
	<li>Place the 4 L beaker containing 3 L of a 0.2% Trypsin/0.05% EDTA/0.05% NaN3 solution on stir plate and warm it to 37 &deg;C in preparation of the decellularization process.</li>
</ol>
2. Tissue Rinses
<ol>
	<li>Set the pump to a flow rate of 400 ml/min, ensuring that the correct tubing size is selected. Flush the heart with Type I water for 15-25 min. As the pump is started, the heart should swell and effuse blood from the ventricles. Fresh solution should be substituted every 5-10 min, or as needed based on the amount of blood removed from the heart. If blood is not effused from the heart, adjust the tubing and clamps as necessary.</li>
	<li>Stop the pump and transfer the heart to a separate beaker filled with 2X Phosphate Buffered Saline (PBS). After the tubing is submerged in solution, start the pump and increase the flow rate to 700 ml/min. The heart should remain in solution for 15 min, changing the solution every 5 min. Each solution change requires the pump to be stopped temporarily while the tissue and tubing is moved to the new beaker.</li>
	<li>Transfer the heart to Type I water for 10 min and increase the flow rate to 750 ml/min.</li>
</ol>
3. Decellularization and Solution Perfusion
<ol>
	<li>Transfer the heart to the beaker containing 0.2% Trypsin/0.05% EDTA/0.05% NaN3 at 37 &deg;C. Increase the pump speed to 1,200 ml/min and start the pump. Use a stir bar placed at the bottom of the beaker to circulate solution in the beaker. The heart should remain in the 0.2% Trypsin/0.05% EDTA/0.05% NaN3 solution at 37 &deg;C for a total of three hours. After 1 hr, increase the pump speed to 1,500 ml/min. After an additional hour, increase the pump speed to 1,800 ml/min. The tissue is slowly subjected to increased perfusion speeds to condition the tissue and prevent rupture of the vessels. The heart will swell and nearly double in size during this step of the protocol. The tissue will lose its natural color, progressing from the atria to the apex throughout the protocol (Figure 3).</li>
	<li>After each solution perfusion, a two step rinse is performed to remove cellular debris, chemical residue, and aid cell lysis. Each rinse consists of a 10 min rinse in Type I water followed by a 10 min rinse with 2X PBS solution at room temperature. Each wash consists of removal of solution from the original beaker, adding rinse solutions, and circulating the perfusate within the beaker containing the submerged heart. After the 0.2% Trypsin/0.05% EDTA/0.05% NaN3 solution, perfuse water at 1,900 ml/min and then perfuse 2X PBS at 1950 ml/min.</li>
	<li>Transfer the heart to a solution of 3% Triton X-100/0.05% EDTA/0.05% NaN3 at room temperature. Increase the pump speed to 2,000 ml/min and perfuse solution for one hour. Remove the solution from the beaker and replace with fresh solution, increase the pump speed to 2100 ml/min, and perfuse the fresh solution for an additional hour and a half, bringing the total time in 3% Triton X-100/0.05% EDTA/0.05% NaN3 to 2.5 hr.</li>
	<li>Rinse the tissue in Type I water at 2150 ml/min and 2X PBS at 2180 ml/min for 10 min each.</li>
	<li>Transfer the heart to a 4% Sodium Deoxycholate solution at room temperature. Increase the pump speed to 2,200 ml/min and perfuse solution for 3 hr.</li>
	<li>Rinse the tissue in Type I water at and 2X PBS at 2,200 ml/min for 15 min each, changing the solutions after 5-10 min for each solution. The described perfusion steps may be split over multiple days by performing the rinse step twice and storing the heart with attached tubing overnight at 4 &deg;C and submerged in Type I water.</li>
	<li>The following day, perform a 5 min rinse with Type I water at 750 ml/min, followed by a 5 min rinse in 1X PBS at 1,500 ml/min. The protocol may then be continued at the described flow rate in the proper solution.</li>
</ol>
4. Disinfection and Final Processing
<ol>
	<li>Transfer the heart to a 0.1% peracetic acid/4% ethanol solution and perfuse solution for 1.5 hr at 2,200 ml/min.</li>
	<li>The final rinses for the tissue are all performed at 2,200 ml/min. Perfuse the tissue with 1X PBS for 15 min, followed by two 5 min washes in Type I water. This series of rinses is repeated once more in order to complete the solution perfusion procedure.</li>
	<li>Turn the pump off and remove the heart from solution to drain the heart. Cut the ties from the aorta, remove all tubing, and place the heart in an empty beaker to drain for 1 hr. Excess liquid will need to be drained periodically. Lay the heart on an absorbent pad to fully drain the heart (Figure 4).</li>
	<li>After most of the water is removed, record the weight of the cardiac extracellular matrix (C-ECM). The heart can be expected to lose approximately 20-25% of its initial weight during the decellularization process.</li>
	<li>Dissect the right and left ventricles, as well as the ventricular septum for DNA quantification and histological processing in order to confirm complete decellularization of the tissue (Figure 5).</li>
	<li>Freeze the C-ECM at -80 &deg;C for at least 2 hr before lyophilization.</li>
</ol>

<ol>
	<li>Ott, H. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regeneration+and+orthotopic+transplantation+of+a+bioartificial+lung.">Regeneration and orthotopic transplantation of a bioartificial lung.</a> Nat. Med. 16, 927-933 (2010).</li><li>Ott, H. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=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. , (2008).</li><li>Petersen, T. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-engineered+lungs+for+in+vivo+implantation.">Tissue-engineered lungs for in vivo implantation.</a> Science. 329, 538-541 (2010).</li><li>Uygun, B. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organ+reengineering+through+development+of+a+transplantable+recellularized+liver+graft+using+decellularized+liver+matrix.">Organ reengineering through development of a transplantable recellularized liver graft using decellularized liver matrix.</a> Nat. Med. , (2010).</li><li>Wainwright, J. 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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+for+tissue+and+organ+decellularization.">Strategies for tissue and organ decellularization.</a> Journal of cellular biochemistry. , (2012).</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, 3675-3683 (2006).</li><li>Akhyari, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+quest+for+an+optimized+protocol+for+whole-heart+decellularization:+a+comparison+of+three+popular+and+a+novel+decellularization+technique+and+their+diverse+effects+on+crucial+extracellular+matrix+qualities.">The quest for an optimized protocol for whole-heart decellularization: a comparison of three popular and a novel decellularization technique and their diverse effects on crucial extracellular matrix qualities.</a> Tissue Eng. 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Part A. 16, 2565-2580 (1089).</li><li>Remlinger, N. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hydrated+xenogeneic+decellularized+tracheal+matrix+as+a+scaffold+for+tracheal+reconstruction.">Hydrated xenogeneic decellularized tracheal matrix as a scaffold for tracheal reconstruction.</a> Biomaterials. 31, 3520-3526 (2010).</li><li>Sellaro, T. L., Ravindra, A. K., Stolz, D. B., 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=Maintenance+of+hepatic+sinusoidal+endothelial+cell+phenotype+in+vitro+using+organ-specific+extracellular+matrix+scaffolds.">Maintenance of hepatic sinusoidal endothelial cell phenotype in vitro using organ-specific extracellular matrix scaffolds.</a> Tissue Eng. 13, 2301-2310 (2007).</li><li>Wainwright, J. 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Dr. Gilbert was on the Scientific Advisory Board at ACell, Inc. while the study was being done, and recently became the VP of Research and Development. ACell, Inc. sells urinary bladder matrix and has no commercial interest in the present study.

The current study described methodology for consistent and efficient decellularization of a porcine heart. The protocol was a modification to a previously published report 1, and included longer exposure to flow and increased pressure, which provided more repeatable results. The resulting decellularized tissue met all of the published criteria for successful decellularization of tissue 2. Frequent solution changes were performed to limit the reintroduction of cellular material to the tissue,...

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Trypsin</td>
 <td>Gibco</td>
 <td>15090</td>
 <td></td>
 </tr>
 <tr>
 <td>EDTA</td>
 <td>Fisher</td>
 <td>BP120-500</td>
 <td></td>
 </tr>
 <tr>
 <td>NaN3</td>
 <td>Sigma</td>
 <td>S2002-500G</td>
 <td></td>
 </tr>
 <tr>
 <td>Triton X-100</td>
 <td>Sigma</td>
 <td>X100-1L</td>
 <td></td>
 </tr>
 <tr>
 <td>10X PBS</td>
 <td>Fisher</td>
 <td>BP399-20</td>
 <td></td>
 </tr>
 <tr>
 <td>Sodium Deoxycholate</td>
 <td>Sigma</td>
 <td>D6750-500G</td>
 <td></td>
 </tr>
 <tr>
 <td>Peracetic Acid</td>
 <td>Pfaltz and Bauer</td>
 <td>P05020</td>
 <td>35% CAS# 79-21-0</td>
 </tr>
 <tr>
 <td>Ethanol</td>
 <td>Pharmco</td>
 <td>111000200</td>
 <td></td>
 </tr>
 <tr>
 <td>Masterflex Pump Drive</td>
 <td>Cole Parmer</td>
 <td>SI-07524-50</td>
 <td></td>
 </tr>
 <tr>
 <td>Masterflex Tubing</td>
 <td>Cole Parmer</td>
 <td>96400-18</td>
 <td>Size 18</td>
 </tr>
 <tr>
 <td>Barbed Reducer</td>
 <td>Cole Parmer</td>
 <td>EW-30612-20</td>
 <td></td>
 </tr>
 <tr>
 <td>4L Beaker</td>
 <td>Fisher Scientific</td>
 <td>02-540T</td>
 <td></td>
 </tr>
 </tbody>
</table>

procedure for decellularization of porcine heart by retrograde coronary perfusion

Perfusion-based whole organ decellularization has recently gained interest in the field of tissue engineering as a means to create site-specific extracellular matrix scaffolds, while largely preserving the native architecture of the scaffold. To date, this approach has been utilized in a variety of organ systems, including the heart, lung, and liver 1-5. Previous decellularization methods for tissues without an easily accessible vascular network have relied upon prolonged exposure of tissue to solutions of detergents, acids, or enzymatic treatments as a means to remove the cellular and nuclear components from the surrounding extracellular environment6-8. However, the effectiveness of these methods hinged upon the ability of the solutions to permeate the tissue via diffusion. In contrast, perfusion of organs through the natural vascular system effectively reduced the diffusion distance and facilitated transport of decellularization agents into the tissue and cellular components out of the tissue. Herein, we describe a method to fully decellularize an intact porcine heart through coronary retrograde perfusion. The protocol yielded a fully decellularized cardiac extracellular matrix (c-ECM) scaffold with the three-dimensional structure of the heart intact. Our method used a series of enzymes, detergents, and acids coupled with hypertonic and hypotonic rinses to aid in the lysis and removal of cells. The protocol used a Trypsin solution to detach cells from the matrix followed by Triton X-100 and sodium deoxycholate solutions to aid in removal of cellular material. The described protocol also uses perfusion speeds of greater than 2 L/min for extended periods of time. The high flow rate, coupled with solution changes allowed transport of agents to the tissue without contamination of cellular debris and ensured effective rinsing of the tissue. The described method removed all nuclear material from native porcine cardiac tissue, creating a site-specific cardiac ECM scaffold that can be used for a variety of applications.

The effect of decellularization on whole porcine hearts naturally varies due to differences in size, pressures, and vessel arrangement. Therefore, the exact composition of the derived extracellular matrix scaffolds will not be the same from heart to heart. The completion of the described protocol will yield a heart that appears white or translucent, indicating the loss of cellular material. However, it is widely accepted that a tissue can be considered "decellularized" based on the combination of a few more quantit...

Watch this Scientific Journal Video about Procedure for Decellularization of Porcine Heart by Retrograde Coronary Perfusion at JoVE.com

Procedure for Decellularization of Porcine Heart by Retrograde Coronary Perfusion

A method to rapidly and completely remove cellular components from an intact porcine heart through retrograde perfusion is described. This method yields a site specific cardiac extracellular matrix scaffold which has the potential for use in multiple clinical applications.

Perfusion-based  whole organ decellularization has recently gained interest in the field of  tissue engineering as a means to create site-specific ...

procedure-for-decellularization-porcine-heart-retrograde-coronary

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23.7K Views.  McGowan Institute for Regenerative Medicine. A method to rapidly and completely remove cellular components from an intact porcine heart through retrograde perfusion is described. This method yields a site specific cardiac extracellular matrix scaffold which has the potential for use in multiple clinical applications.

Video: Procedure for Decellularization of Porcine Heart by Retrograde Coronary Perfusion

Decellularization and Recellularization of Whole Livers

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

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

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

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

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

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

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

Decellularization of Whole Human Heart Inside a Pressurized Pouch in an Inverted Orientation

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ultimately rejection can occur. Creating a functional, autologous bio-artificial heart could solve these challenges. Biofabrication of a heart comprised of scaffold and cells is one option. A natural scaffold with tissue-specific composition as well as micro- and macro-architecture can be obtained by decellularizing hearts from humans or large animals such as pigs. Decellularization involves washing out cellular debris while preserving 3D extracellular matrix and vasculature and allowing &#34;cellularization&#34; at a later timepoint. Capitalizing on our novel finding that perfusion decellularization of complex organs is possible, we developed a more &#34;physiological&#34; method to decellularize non-transplantable human hearts by placing them inside a pressurized pouch, in an inverted orientation, under controlled pressure. The purpose of using a pressurized pouch is to create pressure gradients across the aortic valve to keep it closed and improve myocardial perfusion. Simultaneous assessment of flow dynamics and cellular debris removal during decellularization allowed us to monitor both fluid inflow and debris outflow, thereby generating a scaffold that can be used either for simple cardiac repair (e.g. as a patch or valve scaffold) or as a whole-organ scaffold.

This method enables decellularization of a complex solid organ using a simple protocol based on osmotic shock and perfusion of ionic detergent with minimal organ matrix disruption. It comprises a novel decellularization technique for human hearts inside a pressurized pouch with real-time monitoring of flow dynamics and cellular debris outflow.

The ultimate solution for patients with end-stage heart failure is organ transplant. But donor hearts are limited, immunosuppression is required, and ...

Isolation and Decellularization of a Whole Porcine Pancreas

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an allogeneic or xenogeneic source with human cells. A demonstration of methods for the efficient dissection, decellularization, and recellularization of porcine pancreas might benefit the field. Akin to human pancreases, porcine pancreases have a special anatomical arrangement with three lobes (splenic, duodenal, and connection) rounded by the duodenum and small intestine. The duodenal lobe of the pancreas connects to the duodenum by several small blood vessels. Tissue engineering of the pancreas is complicated because of its exocrine and endocrine nature. In this paper, we show a detailed protocol to dissect the whole porcine pancreas and decellularize it with detergents while saving its structure and some extracellular matrix components. To achieve complete perfusion, the aorta is chosen as inlet and the portal vein as outlet. The other blood vessels (hepatic artery, splenic vein, splenic artery,&#160;mesenteric artery and vein tree) and bile duct are ligated. To prevent the formation of thrombus, the pig is heparinized and, immediately after dissection, the organ is flushed with cold heparin. To inhibit the action of exocrine enzymes, the pancreas decellularization is set at 4 &#176;C. The decellularization is performed by perfusion of Triton X-100, sodium deoxycholate, and deoxyribonuclease, with an intermittent and final extensive washing. With a successful decellularization, the pancreas appears white, and a histological evaluation with hematoxylin and eosin shows an absence of nuclei with a preserved extracellular matrix structure. Thus, the proposed method can be used to successfully dissect and decellularize whole porcine pancreas.

Tissue engineering of the whole pancreas is a challenge because of its exocrine and endocrine functions. We show a method for the dissection of an intact porcine pancreas and the process of successful decellularization by perfusion of detergents Triton X-100, sodium deoxycholate, and deoxyribonuclease.

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an ...

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate (LAL) Assays

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause inflammation, fever, hypo- or hypertension, and, in extreme cases, can lead to tissue and organ damage that may become fatal. The amounts of endotoxin in pharmaceutical products, therefore, are strictly regulated. Among the methods available for endotoxin detection and quantification, the Limulus Amoebocyte Lysate (LAL) assay is commonly used worldwide. While any pharmaceutical product can interfere with the LAL assay, nano-formulations represent a particular challenge due to their complexity. The purpose of this paper is to provide a practical guide to researchers inexperienced in estimating endotoxins in engineered nanomaterials and nanoparticle-formulated drugs. Herein, practical recommendations for performing three LAL formats including turbidity, chromogenic and gel-clot assays are discussed. These assays can be used to determine endotoxin contamination in nanotechnology-based drug products, vaccines, and adjuvants.

Detection of Endotoxin in Nano-formulations Using Limulus Amoebocyte Lysate LAL Assays

Detection of endotoxins in engineered nanomaterials represents one of the grand challenges in the field of nanomedicine. Here, we present a case study that describes the framework composed of three different LAL formats to estimate potential endotoxin contamination in nanoparticles.

When present in pharmaceutical products, a Gram-negative bacterial cell wall component endotoxin (often also called lipopolysaccharide) can cause ...

Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart

The ability to measure biomarkers in vivo relevant to the assessment of disease progression is of great interest to the scientific and medical communities. The resolution of results obtained from current methods of measuring certain biomarkers can take several days or weeks to obtain, as they can be limited in resolution both spatially and temporally (e.g., fluid compartment microdialysis of interstitial fluid analyzed by enzyme-linked immunosorbent assay [ELISA], high-performance liquid chromatography [HPLC], or mass spectrometry); thus, their guidance of timely diagnosis and treatment is disrupted. In the present study, a unique technique for detecting and measuring peptide transmitters in vivo through the use of a capacitive immunoprobe biosensor (CI probe) is reported. The fabrication protocol and in vitro characterization of these probes are described. Measurements of sympathetic stimulation-evoked neuropeptide Y (NPY) release in vivo are provided. NPY release is correlated to the sympathetic release of norepinephrine for reference. The data demonstrate an approach for the fast and localized measurement of neuropeptides in vivo. Future applications include intraoperative real-time assessment of disease progression and minimally invasive catheter-based deployment of these probes.

Time-Resolved In Vivo Measurement of Neuropeptide Dynamics by Capacitive Immunoprobe in Porcine Heart

Established immunochemical methods to measure peptide transmitters in vivo rely on microdialysis or bulk fluid draw to obtain the sample for offline analysis. However, these suffer from spatiotemporal limitations. The present protocol describes the fabrication and application of a capacitive immunoprobe biosensor that overcomes the limitations of the existing techniques.

The ability to measure biomarkers in vivo relevant to the assessment of disease progression is of great interest to the scientific and medical ...

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

Large volume soft tissue defects lead to functional deficits and can greatly impact the patient&#39;s quality of life. Although surgical reconstruction can be performed using autologous free flap transfer or vascularized composite allotransplantation (VCA), such methods also have disadvantages. Issues such as donor site morbidity and tissue availability limit autologous free flap transfer, while immunosuppression is a significant limitation of VCA. Engineered tissues in reconstructive surgery using decellularization/recellularization methods represent a possible solution. Decellularized tissues are generated using methods that remove native cellular material while preserving the underlying extracellular matrix (ECM) microarchitecture. These acellular scaffolds can then be subsequently recellularized with recipient-specific cells.
This protocol details the procurement and decellularization methods used to achieve acellular scaffolds in a pig model. In addition, it also provides a description of the perfusion bioreactor design and setup. The flaps include the porcine omentum, tensor fascia lata, and the radial forearm. Decellularization is performed via ex vivo perfusion of low concentration sodium dodecyl sulfate (SDS) detergent followed by DNase enzyme treatment and peracetic acid sterilization in a customized perfusion bioreactor.
Successful tissue decellularization is characterized by a white-opaque appearance of flaps macroscopically. Acellular flaps show the absence of nuclei on histological staining and a significant reduction in DNA content. This protocol can be used efficiently to generate decellularized soft tissue scaffolds with preserved ECM and vascular microarchitecture. Such scaffolds can be used in subsequent recellularization studies and have the potential for clinical translation in reconstructive surgery.

The protocol describes the surgical procurement and subsequent decellularization of vascularized porcine flaps by the perfusion of sodium dodecyl sulfate detergent through the flap vasculature in a customized perfusion bioreactor.

Large volume soft tissue defects lead to functional deficits and can greatly impact the patient&#39;s quality of life. Although surgical reconstruction ...

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

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

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

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

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

Evaluation of Hydrodynamic Properties in Mounted Porcine Aortic Valves

An Ex Vivo Porcine Model for Hydrodynamic Testing of Experimental Aortic Valve Procedures and Novel Medical Devices

The options for testing new cardiac procedures and investigative medical devices prior to use in an animal model are limited. In this study, we present a method for mounting a porcine aortic valve in a pulse duplicator to evaluate its hydrodynamic properties. These properties can then be evaluated before and after the procedure under investigation is performed and/or the investigative medical device is applied. Securing the inflow segment presents some difficulty owing to the lack of circumferential myocardium in the left ventricular outflow tract. This method addresses that issue by securing the inflow segment using the anterior leaflet of the mitral valve and then suturing the left ventricular free wall around the inflow fixture. The outflow segment is secured simply by inserting the fixture into an incision in the superior aspect of the aortic arch. We found that specimens had significantly different hydrodynamic properties before and after tissue fixation. This finding induced us to use fresh specimens in our testing and should be considered when using this method. In our work, we used this method to test novel intracardiac patch materials for use in the valvular position by performing an aortic valve neocuspidization procedure (Ozaki procedure) on the mounted porcine aortic valves. These valves were tested before and after the procedure to assess the change in hydrodynamic properties in comparison to the native valve. Herein, we report a platform for hydrodynamic testing of experimental aortic valve procedures that enables comparison with the native valve and between different devices and techniques used for the procedure under investigation.

Author Spotlight: Advancing Cardiac Procedure Testing Prior to Embarking on Large Animal Studies 

We present a method for mounting a porcine aortic valve on a pulse duplicator to test its hydrodynamic properties. This method can be used to determine the change in hydrodynamics after the application of an experimental procedure or novel medical device prior to use in a large animal model.

The options for testing new cardiac procedures and investigative medical devices prior to use in an animal model are limited. In this study, we present a ...

Preparation of Left Ventricular Outflow Tract (LVOT) from Porcine Heart for Cannulation 

To begin, collect a fresh porcine cardiac specimen after cardiectomy. Dissect the pulmonary artery off the aorta with Metzenbaum scissors until ventricular tissue is visible. Then, dissect the right and the left coronary arteries, and ligate them using silk ties at their origin from the aortic sinuses without narrowing the sinuses.Transect the coronary arteries distal to the silk ties. Using Metzenbaum scissors, incise the right ventricle between the aorta and the pulmonary artery at the base of the pulmonary valve. Beginning anteriorly, continue the incision circumferentially along the intraventricular septum to remove the right ventricle free wall.Continue the incision posteriorly through the tricuspid valve annulus along the interatrial septum to remove all right atrial tissue. Next, using Metzenbaum scissors, incise the left atrium through the right pulmonary vein ostium parallel to the aorta. Continue this incision towards the anterolateral commissure of the mitral valve.Trim excess left atrium tissue while maintaining the three millimeter cuff of atrial tissue on the aorta and the mitral valve annulus circumferentially. Now, extend the incision onto the left ventricle through the anterolateral commissure of the mitral valve without damaging the anterolateral papillary muscle. Divide the chordae tendineae from the anterolateral papillary muscle to the posterior mitral valve leaflet, preserving attachments to the anterior mitral valve leaflet.Extend the incision to the apex of the heart. Trim excess left ventricle tissue below the papillary muscles, preserving both papillary muscles.