The authors would like to gratefully thank Steffen Lippert, Khalid Aliyev, Korinna J&#246;hrens and Katharina Struecker for their help during this project.

Animals were kept at the Facility for Experimental Medicine (FEM, Charit&#233;, Berlin, Germany), and all experimental protocols were reviewed and approved by the State Office of Health and Local Affairs (LAGeSo, Berlin, Germany; Reg. No. O 0365/11).
1. Liver Harvesting
<ol>
	<li>Pre-surgical Preparation
	<ol>
		<li>Use a cork plate for fixation. Put a medical drape on the plate. Using four needles, fix an inhalation mask on the cork plate for intraoperative inhalation narcosis.</li>
	</ol>
	</li>
	<li>Prearrangement of the Portal Venous Cannula
	<ol>
		<li>Connect a peripheral venous catheter (G 20) to a three-way stopcock. The venous catheter should be cut obliquely, with a length of approx. 1.5 cm.</li>
		<li>De-air the system using a 10 ml syringe filled with Ringer solution. It is very important to carefully de-air because gas embolism may negatively affect the perfusion decellularization.</li>
	</ol>
	</li>
	<li>Prearrangement of the Arterial Cannula
	<ol>
		<li>Use a butterfly cannula and cut the needle to a length of 5 mm. Fit a 5 cm tube (ID 0.58 mm; OD 0.96 mm) to the 5 mm needle and fix it with superglue. Insert a 2 cm-long tube (ID 0.28 mm; OD 0.61 mm) into the bigger tube and fix it with superglue.</li>
		<li>Slip another tube over this construction as an abutment and fix it with superglue. De-air the arterial cannula with a 5 ml syringe filled with Ringer solution.</li>
	</ol>
	</li>
	<li>Anesthesia and Analgesia
	<ol>
		<li>Induce narcosis of the rat in the anesthesia induction chamber by 3.5% isoflurane in oxygen at a flow rate of 1.5 L/min.</li>
		<li>To ensure safe analgesia during the operation, inject metamizol (100 mg/kg body mass) and low-dose ketamine/medetomidine (10 mg/0.1 mg per kg body mass) intraperitoneally.</li>
		<li>Supply 1.5% isoflurane in 1 L/min oxygen via the inhalation mask for subsequent maintenance of surgical anesthesia.</li>
	</ol>
	</li>
	<li>Preoperative Arrangements
	<ol>
		<li>Shave the abdomen liberally with an electric shearing machine. Moisten the shaved incision site on the abdomen with 70% ethanol for disinfection.</li>
		<li>Place the animal in the supine position on the prepared plate, insert the head into the inhalation mask and fix the limbs with medical tape and pins. Remove excess ethanol and fur&#160;with a gauze compress.</li>
	</ol>
	</li>
	<li>Anesthesia Assessment
	<ol>
		<li>Assess the anesthesia depth by watching the respiratory rate. When the anesthesia seems deep enough (40-60 breaths/min), check whether analgesia is sufficient by trying to trigger the toe pinch reflex using surgical forceps to pinch the skin between the toes of both hind limbs.</li>
	</ol>
	</li>
	<li>Abdominal Approach
	<ol>
		<li>Make a median incision into the caudal abdominal wall above the bladder. Cut from this point to both lateral costal arches in a V-shaped manner. Take care not to cut into the diaphragm. Pull the cut abdominal wall over the chest.</li>
		<li>Fix the xiphoid with an overholt clamp, pull it cranially and fix it with a rubber band. Use wet cotton swabs to evacuate the intestine. Wrap the intestine in a wet gauze bandage. Furthermore, cover all abdominal margins with wet bandages.</li>
	</ol>
	</li>
	<li>Liver Preparation
	<ol>
		<li>Dissect the falciform ligament. Use a wet gauze bandage to hold the medial and left liver lobes cranially under the dome of the diaphragm. Expose the hepatoduodenal ligament and the upper omental liver lobe.&#160;</li>
		<li>Use two microforceps and spring scissors to carefully dissect the ligament attached to the upper omental lobe until the lobe is mobile. Move the lobe using wet cotton swabs under the gauze compress,holding the medial and left lobes in place.</li>
		<li>After mobilization of the upper omental liver lobe, move the stomach to the left. Fix the stomach with a clamp and dissect the minor omentum with microforceps and the spring scissors.</li>
		<li>Mobilize the lower omental liver lobe until it is mobile and subsequently move the lobe back into its cavity. Use a Backhaus clamp to retain the duodenum. Move the duodenum to the left to stretch and expose the hepatoduodenal ligament. Circumscribe the common bile duct in the stretched hepatoduodenal ligament.</li>
		<li>Dissect the bile duct from the adipose and pancreatic tissue. Cut the bile duct 1.5 cm from the bile duct bifurcation. Dissect the portal vein from the surrounding adipose tissue. Expose the gastroduodenal vein.</li>
		<li>Ligate the gastroduodenal vein twice with silk 6-0 and divide the vein between the 2 ligatures. Dissect the celiac trunk and the common hepatic artery from the surrounding tissue until all arterial rami are revealed.</li>
		<li>Liberate the gastroduodenal artery from the surrounding tissue. Tie the gastroduodenal artery twice with silk 6-0, with the ties distant from each other. Dissect the gastroduodenal artery between the 2 ligatures.</li>
		<li>Liberate the splenic artery from the surrounding tissue. Tie the splenic artery twice with silk 6-0, with the ties distant from each other. Dissect the splenic artery between the 2 ligatures.</li>
		<li>Liberate the left gastric artery from the surrounding tissue. Tie the left gastric artery twice with silk 6-0, with the ties distant from each other. Dissect the artery between the 2 ligatures.</li>
		<li>Circumscribe the inferior caval vein between the right kidney and the right lateral liver lobe. Dissect the vein from the retroperitoneal space. Expose the aorta by following the celiac trunk.</li>
		<li>Dissect the retroperitoneal adipose tissue. Inject 1,000 I.E. heparin in 1 ml saline into the inferior vena cava. Retract the cannula and close the incision with a cotton pad. Wait 1-2 min&#160;for the systemic effect of heparin.</li>
		<li>Circumscribe the aorta&#160;with forceps. Dissect the aorta and exsanguinate the rat. Furthermore, dissect the inferior vena cava distally from the kidney.</li>
	</ol>
	</li>
	<li>Portal Venous Cannulation (Use the cannula prepared in section 1.2)
	<ol>
		<li>Make a fish-mouth incision in the distal portal vein. Open the incision with forceps and cannulate the portal vein. Flush the liver with 20 ml Ringer solution until the liver decolorizes and fix the cannula with ligatures (silk 6-0).</li>
	</ol>
	</li>
	<li>Arterial Cannulation (Use the cannula prepared in section 1.3)
	<ol>
		<li>Dissect the aorta above the celiac trunk. Liberate the aorta segment from the retroperitoneal space. Cut the aorta segment longitudinally to generate an aortic patch. Insert the self-built cannula into a static holder.</li>
		<li>Use 2 forceps to slip the aortic patch over the cannula. Ligate the hepatic artery with silk 6-0 over the cannula and fix the patch at the abutment.</li>
	</ol>
	</li>
	<li>Liver Explantation
	<ol>
		<li>Open the diaphragm, make a circular incision and dissect the liver from the surrounding structures.</li>
	</ol>
	</li>
	<li>Storage
	<ol>
		<li>Store the liver in a sterile 500 ml tank filled with 350 ml Ringer solution until use.</li>
	</ol>
	</li>
</ol>
2. Detergent Solutions
<ol>
	<li>10% Triton X-100 stock solution
	<ol>
		<li>Fill a 1,000 ml bottle with 900 ml aqua dest. and add 100 ml Triton X-100 (99,9%). Use a stirrer to dissolve the Triton X-100. Fill the bottle with aqua dest. until a total volume of 1,000 ml is reached.</li>
	</ol>
	</li>
	<li>1% Triton X-100
	<ol>
		<li>Add 100 ml of the stock solution (10%) to a 1,000 ml bottle. Add 900 ml aqua dest. and shake the bottle twice. Filter the 1% Triton X-100 solution through a sterile filter.</li>
	</ol>
	</li>
	<li>10% SDS Stock Solution
	<ol>
		<li>Fill a 1,000 ml bottle with 900 ml aqua dest. and add 66.99 g SDS (99.9%, density 0.67). Use a stirrer to dissolve the SDS. Fill the bottle with aqua dest. until a total volume of 1,000 ml is reached.</li>
	</ol>
	</li>
	<li>1% SDS
	<ol>
		<li>Add 100 ml of the stock solution (10%) to a 1,000 ml bottle. Add 900 ml aqua dest. and shake the bottle twice. Filter the 1% SDS solution through a sterile filter.</li>
	</ol>
	</li>
</ol>
3. Decellularization
<ol>
	<li>Set-up of Perfusion Device<img alt="Figure 1" src="/files/ftp_upload/53029/53029fig1.jpg" /></li>
</ol>
Figure 1: Scheme of the Perfusion Device for Selective Arterial and Portal Venous Perfusion of Four Rat Livers under Oscillating Pressure Conditions.&#160;(modified from Struecker et al. 14)
<ol>
	<li>Link the drain to a three-way stopcock and a Heidelberger extension to regulate the fill level and fill the reactor with 500 ml 0,9% NaCl. 
	<img alt="Figure 2" src="/files/ftp_upload/53029/53029fig2.jpg" /></li>
</ol>
Figure 2: Scheme of the perfusion set-up.&#160;Link the perfusion chamber (1,400 cm3)(1) to the distal end (4) of the bubble trap (5) via the portal venous access (2) or via the arterial access (3). The bubble trap (5) should be equipped with an obturation (4) at the distal end and a vent (6). Connect the bubble trap (5) to a Heidelberger extension (7). Link the Heidelberger extension to the pump segment (8). Furthermore, connect the pump segment (8) to another Heidelberger extension (9). Insert the last Heidelberger extension (9) into the bottle of detergent solution (10). Connect the respirator (11) to the perfusion chamber (or to the pressure distributor in the case of multiple perfusions). To drain the reactor fluids, connect the outflow to the waste bottle (12).
<ol>
	<li>Insert the pump segment into the pump. Start the pump to fill the cycle with PBS, close the distal end of the bubble trap and open the vent. When the bottom of the bubble is securely covered with PBS (2-3 cm), close the vent and open the distal end of the bubble trap.</li>
	<li>Liver Installation
	<ol>
		<li>Link the three-way stopcock of the portal vein cannula to the related reactor access. Connect the arterial cannula to the arterial access. Be careful to ensure that no air enters the systems.</li>
	</ol>
	</li>
	<li>Application of Oscillating Pressure Conditions
	<ol>
		<li>Use a respirator with a frequency of 15 bpm and an amplitude of 26 mbar (min. 4 mbar, max. 30 mbar). Connect the respirator to the pressure distribution chamber and connect the chamber to the reactor. Start pressure application.</li>
	</ol>
	</li>
	<li>Decellularization Protocol
	<ol>
		<li>Perform perfusion steps with a flow rate of 5 ml/min. Start the decellularization with perfusion of the liver with 1% Triton X-100 for 90 min. Allow waste effluence via the outflow of the perfusion device to keep the fluid level inside the perfusion chamber constant and above the perfused liver. After 90 min, change the detergent to 1% SDS. 
		<img alt="Figure 3" src="/files/ftp_upload/53029/53029fig3.jpg" /> 
		<img alt="Table 1" src="/files/ftp_upload/53029/53029table1.jpg" /></li>
	</ol>
	</li>
</ol>
&#160;Figure 3: Perfusion Protocol for All Experimental Groups.

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The authors have nothing to disclose.

Although the presented technique for rat liver harvesting and decellularization is easily reproducible, there are certain critical steps to consider:
During preparation for liver harvesting, it is important to avoid severe bleeding because it will activate blood coagulation and may lead to blood clot formation within the liver. In our opinion, it is advantageous to incise the abdominal aorta directly before cannulation of the portal vein to avoid blood inflow via the hepatic artery during perf...

Decellularization and recellularization may enable the generation of functional, transplantable organs in vitro 1. By removing cells and antigenic material (e.g., DNA, alpha-Gal epitopes) from an organ, the non- or less-immunogenic extracellular matrix (ECM) can be obtained. This matrix conserves the three-dimensional microanatomy of an organ and can serve as the ideal biomatrix for repopulation with cells of a different, possibly xenogeneic origin 2. Thus, a decellularized rat liver matrix could be repopulated with human liver cells. This humanized micro-liver could serve as an ex vivo model for research on diseases (e.g., inborn metabolic diseases, viral diseases or malignancies) or for preclinical pharmaceutical testing 3.
Several different protocols for rat liver perfusion decellularization have already been published 4-13. In all protocols, decellularization was achieved by perfusion of alkaline ionic or non-ionic detergents via the cannulated portal vein. To the best of our knowledge, we were the first group to report rat liver decellularization by selective perfusion via the portal vein and/or the rat hepatic artery 14. Enabling the selective perfusion of the different vascular systems in the liver may enable better decellularization results and, furthermore, may play an important role in cellular repopulation.
In the study detailed here, livers were perfused in a custom-made proprietary perfusion device, enabling perfusion under oscillating pressure conditions. These pressure conditions mimic the physiologic respiratory-dependent perfusion of the liver: in situ, the liver hangs under the copula of the diaphragm, whose movement during respiration has a direct impact on liver perfusion. Inspiration specifically leads to lowering of the diaphragm and squeezing of the liver, optimizing hepato-venous outflow, whereas expiration leads to elevation of the liver and lowering of the intraabdominal pressure to optimize portal-venous inflow 15.
Our aim was to evaluate whether oscillating pressure conditions have an impact on the homogeneity of rat liver perfusion decellularization by mimicking intraabdominal conditions ex vivo. The homogeneity of the decellularization process may be an underestimated factor in perfusion decellularization. All known agents used for liver decellularization cause alterations to the ECM. Cells in poorly perfused areas remain within the ECM, whereas other areas are already completely decellularized. To dissolve the remaining cells, the perfusion duration or pressure must be elevated, causing more alterations to the well-perfused areas. Thus, detergents for decellularization should be distributed homogenously within the organ.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Self built arterial cannula</td>
		</tr>
		<tr>
			<td>&#160;Portex Non Sterile Polyethene Tubing</td>
			<td>SIMS Portex</td>
			<td>REF 800/110/100</td>
			<td>0,28mm ID 0,61mm OD</td>
		</tr>
		<tr>
			<td>&#160;Portex Non Sterile Polyethene Tubing</td>
			<td>SIMS Portex</td>
			<td>REF 800/110/200</td>
			<td>0,58mm ID 0,96m OD</td>
		</tr>
		<tr>
			<td>Venodrop Safe butterfly catheter</td>
			<td>Fresenius Kabi</td>
			<td>3275851</td>
			<td>21 G</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>portal vein cannula</td>
		</tr>
		<tr>
			<td>Periphereal Venous Catheter</td>
			<td>BD</td>
			<td>393224</td>
			<td>BD Venflon Pro 20G</td>
		</tr>
		<tr>
			<td>three-way stopcock</td>
			<td>smiths medical</td>
			<td>888-101RE</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>surgery</td>
		</tr>
		<tr>
			<td>Cotton Sticks</td>
			<td>Hecht-Assistent</td>
			<td>4302</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton Pads&#160;</td>
			<td>Shaoxing Zhengde Surgical dressing</td>
			<td>13H118-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze Bandage</td>
			<td>Hubei Haige&#160; Medical Instruments</td>
			<td>14388</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Ringer Solution</td>
			<td>Fresenius Kabi</td>
			<td>13 HKP022</td>
			<td>1000ml</td>
		</tr>
		<tr>
			<td>10ml Syringe</td>
			<td>Braun</td>
			<td>4606108V</td>
			<td>10ml/ Luer Solo</td>
		</tr>
		<tr>
			<td>5ml Syringe</td>
			<td>Braun</td>
			<td>4606061V</td>
			<td>5ml /Luer Solo</td>
		</tr>
		<tr>
			<td>Suture (Silk 6/0)</td>
			<td>Resorba</td>
			<td>H1F</td>
			<td>LOT 105001.81</td>
		</tr>
		<tr>
			<td>medical drape</td>
			<td>Shaoxing Zhengde Surgical dressing</td>
			<td>D0613011</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>surgical instruments</td>
		</tr>
		<tr>
			<td>needle holder</td>
			<td>Geuder</td>
			<td>17570</td>
			<td></td>
		</tr>
		<tr>
			<td>micro-forceps</td>
			<td>Inox-Electronic</td>
			<td>91150-20</td>
			<td></td>
		</tr>
		<tr>
			<td>micro-scissors</td>
			<td>Martin</td>
			<td>11-740-11</td>
			<td></td>
		</tr>
		<tr>
			<td>micro-forceps</td>
			<td>S&#38;T&#160;</td>
			<td>112314</td>
			<td></td>
		</tr>
		<tr>
			<td>Clamp</td>
			<td>Aesculap</td>
			<td>BH111R</td>
			<td></td>
		</tr>
		<tr>
			<td>scissors</td>
			<td>F S T&#160;</td>
			<td>14501-14</td>
			<td></td>
		</tr>
		<tr>
			<td>surgical forceps</td>
			<td>Aesculap</td>
			<td>BD 557</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Decellularisation</td>
		</tr>
		<tr>
			<td>Respirator</td>
			<td>Resmed</td>
			<td>14.24.11.0004</td>
			<td>SmartAIR ST</td>
		</tr>
		<tr>
			<td>Perfusion Device</td>
			<td>Charite, medical engineering laboratory</td>
			<td>custome-made device</td>
			<td>decellularisation device</td>
		</tr>
		<tr>
			<td>peristaltic pump ismatec reglo ICC</td>
			<td>IDEX</td>
			<td>ISM4408&#160;</td>
			<td>4-channel</td>
		</tr>
		<tr>
			<td>heidelberger extension 75 cm&#160;</td>
			<td>Fresenius Kabi</td>
			<td>2873</td>
			<td>75 cm</td>
		</tr>
		<tr>
			<td>MS/CA pump-segment</td>
			<td>IDEX</td>
			<td>IS 3510&#160;</td>
			<td>MS/CA/click&#39;n&#39;go/POM-C</td>
		</tr>
		<tr>
			<td>CA 2-stopper tube</td>
			<td>Pharmed</td>
			<td>BPT NSF-51</td>
			<td></td>
		</tr>
		<tr>
			<td>bubble trap&#160;</td>
			<td></td>
			<td>custome-made item</td>
			<td></td>
		</tr>
		<tr>
			<td>Luer Lock hose connector</td>
			<td>Neolab</td>
			<td>No. 02-1887</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Detergents</td>
		</tr>
		<tr>
			<td>SDS pellets</td>
			<td>Carl Roth&#160;</td>
			<td>CN30.4&#160;</td>
			<td>2,5 kg</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Carl Roth&#160;</td>
			<td>3051.1&#160;</td>
			<td>10l</td>
		</tr>
		<tr>
			<td>PBS&#160;</td>
			<td>Gibco</td>
			<td>14190-094</td>
			<td>DPBS</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>staining</td>
		</tr>
		<tr>
			<td>Eosin 1%</td>
			<td>Morphisto</td>
			<td>10177</td>
			<td></td>
		</tr>
		<tr>
			<td>Mayer hematoxylin</td>
			<td>AppliChem</td>
			<td>A4840</td>
			<td></td>
		</tr>
		<tr>
			<td>gomori staining</td>
			<td>Morphisto</td>
			<td>11104</td>
			<td></td>
		</tr>
		<tr>
			<td>AlcainBlue-PAS staining</td>
			<td>Morphisto</td>
			<td>11388</td>
			<td></td>
		</tr>
		<tr>
			<td>Direct Red 80&#160;</td>
			<td>Sigma Aldrich</td>
			<td>365548</td>
			<td></td>
		</tr>
	</tbody>
</table>

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 homogeneity and thus the effectiveness of different decellularization protocols were evaluated by macroscopic observation, histological analysis, and analysis of the remaining DNA content within decellularized liver matrices. Furthermore, corrosion casting was performed to visualize the intact microanatomy of livers after decellularization.
Macroscopy
During decellularization, livers become lucent, indicating the removal of cellular content. Livers perfused via ...

Watch this Scientific Journal Video about Procedure for Decellularization of Rat Livers in an Oscillating-pressure Perfusion Device at JoVE.com

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

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 ...

procedure-for-decellularization-rat-livers-an-oscillating-pressure

Research

JoVE Journal

Bioengineering

10.2K Views.  Charité – Universitätsmedizin Berlin. 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.

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

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 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.

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 ...

An Ultrasonic Tool for Nerve Conduction Block in Diabetic Rat Models

Nerve conduction block with a high intensity-focused ultrasound (HIFU) transducer has been performed in normal and diabetic animal models recently. HIFU can reversibly block the conduction of peripheral nerves without damaging the nerves while using an appropriate ultrasonic parameter. Temporary and partial block of the action potentials of nerves shows that HIFU has the potential to be a useful clinical treatment for pain relief. This work demonstrates the procedures for suppressing the action potentials of neuropathic nerves in diabetic rats in vivo using an HIFU transducer. The first step is to generate adult male diabetic neuropathic rats by streptozotocin (STZ) injection. The second step is to evaluate the peripheral diabetic neuropathy in STZ-induced diabetic rats by an electronic von Frey probe and a hot plate. The final step is to record in vivo extracellular action potentials of the nerve exposed to HIFU sonication. The method showed here may benefit the study of ultrasound analgesic applications.

This work presents the methodology of applying high intensity-focused ultrasound to block the action potentials of diabetic neuropathic nerves.

Nerve conduction block with a high intensity-focused ultrasound (HIFU) transducer has been performed in normal and diabetic animal models recently. HIFU ...

Development of an Electrochemical DNA Biosensor to Detect a Foodborne Pathogen

Vibrio parahaemolyticus (V. parahaemolyticus) is a common foodborne pathogen that contributes to a large proportion of public health problems globally, significantly affecting the rate of human mortality and morbidity. Conventional methods for the detection of V. parahaemolyticus such as culture-based methods, immunological assays, and molecular-based methods require complicated sample handling and are time-consuming, tedious, and costly. Recently, biosensors have proven to be a promising and comprehensive detection method with the advantages of fast detection, cost-effectiveness, and practicality. This research focuses on developing a rapid method of detecting V. parahaemolyticus with high selectivity and sensitivity using the principles of DNA hybridization. In the work, characterization of synthesized polylactic acid-stabilized gold nanoparticles (PLA-AuNPs) was achieved using X-ray Diffraction (XRD), Ultraviolet-visible Spectroscopy (UV-Vis), Transmission Electron Microscopy (TEM), Field-emission Scanning Electron Microscopy (FESEM), and Cyclic Voltammetry (CV). We also carried out further testing of stability, sensitivity, and reproducibility of the PLA-AuNPs. We found that the PLA-AuNPs formed a sound structure of stabilized nanoparticles in aqueous solution. We also observed that the sensitivity improved as a result of the smaller charge transfer resistance (Rct) value and an increase of active surface area (0.41 cm2). The development of our DNA biosensor was based on modification of a screen-printed carbon electrode (SPCE) with PLA-AuNPs and using methylene blue (MB) as the redox indicator. We assessed the immobilization and hybridization events by differential pulse voltammetry (DPV). We found that complementary, non-complementary, and mismatched oligonucleotides were specifically distinguished by the fabricated biosensor. It also showed reliably sensitive detection in cross-reactivity studies against various food-borne pathogens and in the identification of V. parahaemolyticus in fresh cockles.

A protocol for the development of an electrochemical DNA biosensor comprising a polylactic acid-stabilized, gold nanoparticles-modified, screen-printed carbon electrode to detect Vibrio parahaemolyticus is presented.

Vibrio parahaemolyticus (V. parahaemolyticus) is a common foodborne pathogen that contributes to a large proportion of public health problems globally, ...

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

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

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

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

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

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 ...

Generation of Dynamical Environmental Conditions using a High-Throughput Microfluidic Device

Mimicking in vivo environmental conditions is crucial for in vitro studies on complex life machinery. However, current techniques targeting live cells and organs are either highly expensive, like robotics, or lack nanoliter volume and millisecond time accuracy in liquid manipulation. We herein present the design and fabrication of a microfluidic system, which consists of 1,500 culture units, an array of enhanced peristaltic pumps and an on-site mixing modulus. To demonstrate the capacities of the microfluidic device, neural stem cell (NSC) spheres are maintained in the proposed system. We observed that when the NSC sphere is exposed to CXCL in day 1 and EGF in day 2, the round-shaped conformation is well maintained. Variation in the input order of 6 drugs causes morphological changes to the NSC sphere and the expression level representative marker for NSC stemness (i.e., Hes5 and Dcx). These results indicate that dynamic and complex environmental conditions have great effects on NSC differentiation and self-renewal, and the proposed microfluidic device is a suitable platform for high throughput studies on the complex life machinery.

We present a microfluidic system for high throughput studies on complex life machinery, which consists of 1500 culture units, an array of enhanced peristaltic pumps and an on-site mixing modulus. The microfluidic chip allows for the analysis of the highly complex and dynamic micro-environmental conditions in vivo.

Mimicking in vivo environmental conditions is crucial for in vitro studies on complex life machinery. However, current techniques targeting live cells and ...

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 ...

A Combined Micro-Device System for Promoting and Monitoring Coral Growth

An Integrated Micro-Device System for Coral Growth and Monitoring

Corals are fundamental organisms in marine and coastal ecosystems. With the advancement of coral protection research in recent years, precise control of the coral culture environment is highly in demand for coral conservation and study. Here, we developed a semi-closed coral culture micro-device system as a multi-functional platform, which can provide accurate and programmable temperature control, a sterile initial environment, long-term stable water quality, an adjustable dissolved oxygen concentration, and a customized light spectrum for corals. Owing to the modular design, the coral culture system can be upgraded or modified by installing desirable new modules or removing existing ones. Currently, under appropriate conditions and with proper system maintenance, the sample corals can survive for at least 30 days in a healthy state. Furthermore, due to the controllable and sterile initial environment, this coral culture system can support research into the symbiotic relationship between corals and associated microorganisms. Therefore, this micro-device system can be applied to monitor and investigate sea corals in a relatively quantitative manner.

Author Spotlight: Advancing Coral Culture - Creating a Semi-Quantitatively Controlled Microenvironment System to Counter Current Limitations 

This protocol describes the development of a modular controllable micro-device system that can be applied for the long-term culturing and monitoring of sea corals.

Corals are fundamental organisms in marine and coastal ecosystems. With the advancement of coral protection research in recent years, precise control of ...