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 perfusion via the portal vein. Once the liver is cannulated and clearly perfused, blood clot formation is no longer an issue.
After liver harvesting and transportation to the perfusion device, the connection of livers to the perfusion device is a particularly critical step because the entry of gas bubbles into the perfusion circle must be avoided to prevent gas embolism. It is useful to prefill the three-way valve, which is connected to the liver cannula, with PBS using a syringe and a thin cannula directly before connecting the valve to the perfusion system.
After perfusion is established, whether all tube connectors, three-way valves and tubes are perfectly connected should be re-checked. If livers are perfused for longer durations (e.g., O/N), even a small error in connection can lead to air siphoning into the perfusion system, thus leading to fatal perfusion outcomes.
The presented data are limited by the fact that the optimal pressure and frequency of oscillating pressure changes for the best decellularization outcomes remain unclear. Furthermore, pressure-controlled perfusion may lead to better and more stable results than flow-controlled perfusion does. Pressure-controlled perfusion enables the application of one perfusion protocol to livers of different sizes and weights, with the same outcome, because the flow will automatically be adapted.
The hepatic artery and portal vein are significantly different in terms of size and physiologic flow rates. Thus, a constant perfusion rate (5 ml/min) will certainly result in different perfusion pressures within the two vascular systems. Similarly, a constant perfusion pressure will result in different perfusion rates. One method to compare the decellularization efficacy via the two perfusion routes would be to perform perfusion at physiologic perfusion pressures or perfusion rates via both routes. However, to make the results comparable in the current study, we chose a constant perfusion rate for both routes. Unfortunately, using our device, we were not able to measure the perfusion pressure within perfused livers.
Our next-generation perfusion devices will be smaller to minimize the amount of perfusion media necessary. Furthermore, pump devices will enable pressure-controlled perfusion.
The presented technique appears very useful for reproducible, homogenous rat liver decellularization. Whether the presented technique is appropriate for recellularization and organ maturation in vitro has to be further addressed. Whether oscillating pressure conditions have an impact on repopulated cells will be a subject of further experiments.
The presented technique is more sophisticated than other rat liver perfusion devices and shows several clear advantages: 1) The homogenous decellularization is reproducible, with good results when oscillating pressure conditions are applied. 2) The perfusion device is sealed and enables sterile decellularization, which is a prerequisite for matrix repopulation and long-term perfusion. 3) The presented protocol is shorter than other published protocols but is still very effective. 4) Selective perfusion of the portal vein and the hepatic artery renders selective cell repopulation via different systems possible, which may be an important tool.
The perfusion device will be applied in further decellularization experiments to refine and optimize rat liver decellularization. The effect of pressure-controlled perfusion will be experimentally evaluated and compared with flow-controlled perfusion. The addition of further elements (e.g., a &#8220;dialysis unit&#8221;, a heat exchanger, an oxygenator) for repopulation, culture and maturation experiments appears possible and reasonable to further develop the presented device.

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><tbody><tr><td colspan="4">Self built arterial cannula</td></tr><tr><td>&amp;nbsp;Portex Non Sterile Polyethene Tubing</td><td>SIMS Portex</td><td>REF 800/110/100</td><td>0.28 mm ID 0.61 mm OD</td></tr><tr><td>&amp;nbsp;Portex Non Sterile Polyethene Tubing</td><td>SIMS Portex</td><td>REF 800/110/200</td><td>0.58 mm ID 0.96 mm OD</td></tr><tr><td>Venodrop Safe butterfly catheter</td><td>Fresenius Kabi</td><td>3275851</td><td>21 G</td></tr><tr><td colspan="4">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 colspan="4">surgery</td></tr><tr><td>Cotton Sticks</td><td>Hecht-Assistent</td><td>4302</td><td></td></tr><tr><td>Cotton Pads&amp;nbsp;</td><td>Shaoxing Zhengde Surgical dressing</td><td>13H118-03</td><td></td></tr><tr><td>Gauze Bandage</td><td>Hubei Haige&amp;nbsp; Medical Instruments</td><td>14388</td><td></td></tr><tr><td>&amp;nbsp;Ringer Solution</td><td>Fresenius Kabi</td><td>13 HKP022</td><td>1000 ml</td></tr><tr><td>10 ml Syringe</td><td>Braun</td><td>4606108V</td><td>10 ml/ Luer Solo</td></tr><tr><td>5 ml Syringe</td><td>Braun</td><td>4606061V</td><td>5 ml /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 colspan="4">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&amp;amp;T&amp;nbsp;</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&amp;nbsp;</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 colspan="4">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&amp;nbsp;</td><td>4-channel</td></tr><tr><td>heidelberger extension 75 cm&amp;nbsp;</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&amp;nbsp;</td><td>MS/CA/click&#039;n&#039;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&amp;nbsp;</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 colspan="4">Detergents</td></tr><tr><td>SDS pellets</td><td>Carl Roth&amp;nbsp;</td><td>CN30.4&amp;nbsp;</td><td>2.5 kg</td></tr><tr><td>Triton X-100</td><td>Carl Roth&amp;nbsp;</td><td>3051.1&amp;nbsp;</td><td>10 L</td></tr><tr><td>PBS&amp;nbsp;</td><td>Gibco</td><td>14190-094</td><td>DPBS</td></tr><tr><td colspan="4">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&amp;nbsp;</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 the portal vein showed inhomogeneous elucidation (and thus decellularization), with macroscopically visible remaining cells during and after decellularization (white arrows). Livers perfused via the hepatic artery showed a more homogenous decellularization course and no visible remaining cells. If livers were perfused with the application of oscillating pressure conditions, no remaining cells were visible, irrespective of the perfusion route.
<img alt="Figure 4" src="/files/ftp_upload/53029/53029fig4.jpg" /> 
Figure 4: Macroscopic Results of Rat Liver Decellularization without Oscillating Pressure Conditions.&#160;Left: Rat liver decellularized via the hepatic artery. The liver appears lucent, without macroscopically visible remaining cells. Right: Liver perfused via the portal vein. The liver appears inhomogeneously decellularized, with macroscopically visible cells
<img alt="Figure 5" src="/files/ftp_upload/53029/53029fig5.jpg" /> 
Figure 5: Macroscopic results of rat liver decellularization under oscillating pressure conditions. Left: Rat liver decellularized via the hepatic artery. Right: Liver perfused via the portal vein. Both livers appear lucent, without visible remaining cells.
Histological Evaluation
Histological evaluation of decellularized liver matrices supported the macroscopic findings. Livers perfused via the hepatic artery showed no remaining cells, irrespective of whether they were perfused under oscillating pressure conditions. Livers perfused via the portal vein showed zones of remaining cells, even if they were perfused with the application of oscillating pressure conditions. However, the application of oscillating pressure conditions reduced the remaining cell mass in livers perfused via the portal vein. The ECM of the livers was conserved, without visible differences across all applied protocols.
<img alt="Figure 6" src="/files/ftp_upload/53029/53029fig6.jpg" />
Figure 6: H/E Staining of Decellularized Liver Matrices.&#160;A-P: Decellularized liver matrix perfused via the hepatic artery without oscillating pressure conditions. A+P: Decellularized liver matrix perfused via the hepatic artery under oscillating pressure conditions. PA-P: Decellularized liver matrix perfused via the portal vein without oscillating pressure conditions. PA+P: Decellularized liver matrix perfused via the portal vein under oscillating pressure conditions. Arrows: Remaining cell clusters.
DNA content
The DNA content per dry weight of liver matrix declined in all experimental groups compared with that in native livers, although differences were only statistically significant in the groups perfused under oscillating pressure conditions (PV+P; A+P). The lowest amount of DNA per dry weight was found in livers perfused via the hepatic artery under oscillating pressure conditions.
<img alt="Figure 7" src="/files/ftp_upload/53029/53029fig7.jpg" />
Figure 7: DNA Content Per Dry Weight of Liver Matrix (modified from Struecker et al. 14).&#160;Livers decellularized via the hepatic artery show less remaining DNA than those perfused via the portal vein do. Livers perfused under oscillating pressure conditions show less remaining DNA than those decellularized without these conditions do. Thus, livers perfused via the hepatic artery under oscillating pressure conditions show the least remaining DNA content.
Corrosion Casting
Corrosion casting of decellularized liver matrices confirmed that the microanatomy of livers (including the protein framework of the portal venous system, the arterial system and the biliary system) is conserved during decellularization.
<img alt="Figure 8" src="/files/ftp_upload/53029/53029fig8.jpg" /> 
Figure 8: Photograph of a Corrosion Casting of a Decellularized Rat Liver Matrix (modified from Struecker et al. 14).&#160;Despite removal of all cells, the microanatomy of the organ, including the portal venous (blue) and arterial (red) systems, are conserved. The remaining main branches of the biliary system are casted in yellow.
<img alt="Table 1" src="/files/ftp_upload/53029/53029table1.jpg" /> 
Table 1: Experimental Groups for Assessing the Effect of Oscillating Pressure Conditions and the Perfusion Route on Rat Liver Decellularization.

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

Nanyang Technological University

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 ...

Purification of Hepatocytes and Sinusoidal Endothelial Cells from Mouse Liver Perfusion

This protocol demonstrates a method for obtaining high yield and viability for mouse hepatocytes and sinusoidal endothelial cells (SECs) suitable for culturing or for obtaining cell lysates. In this protocol, the portal vein is used as the site for catheterization, rather than the vena cava, as this limits contamination of other possible cell types in the final liver preparation. No special instrumentation is required throughout the procedure. A water bath is used as a source of heat to maintain the temperature of all the buffers and solutions. A standard peristaltic pump is used to drive the fluid, and a refrigerated table-top centrifuge is required for the centrifugation procedures. The only limitation of this technique is the placement of the catheter within the portal vein, which is challenging on some of the mice in the 18 - 25 g size range. An advantage of this technique is that only one vein is utilized for the perfusion and the access to the vein is quick, which minimizes ischemia and reperfusion of the liver that reduces hepatic cell viability. Another advantage to this protocol is that it is easy to distinguish live from dead hepatocytes by eyesight due to the difference in cellular density during the centrifugation steps. Cells from this protocol may be used in cell culture for any downstream application as well as processed for any biochemical assessment.

The goal of this protocol is to obtain high viability and high yield of hepatocytes and sinusoidal endothelial cells from liver. This is accomplished by perfusing the liver with a type IV collagenase solution via the portal vein, followed by differential centrifugation to obtain hepatocytes and sinusoidal endothelial cells.

This protocol demonstrates a method for obtaining high yield and viability for mouse hepatocytes and sinusoidal endothelial cells (SECs) suitable for ...

Biochemistry

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing modified cannulas and techniques adapted from conventional commercial ex vivo perfusion equipment. The system was utilized to evaluate the preservation outcomes following 12 h of perfusion. C57BL/6J mice served as liver donors, and the livers were explanted by cannulating the portal vein (PV) and bile duct (BD), and subsequently flushing the organ with warm (37 &#176;C) heparinized saline. Then, the explanted livers were transferred to the perfusion chamber and subjected to normothermic oxygenated machine perfusion (NEVLP). Inlet and outlet perfusate samples were collected at 3 h intervals for perfusate analysis. Upon completion of the perfusion, liver samples were obtained for histological analysis, with morphological integrity assessed using modified Suzuki-Score through Hematoxylin-Eosin (HE) staining. The optimization experiments yielded the following findings: (1) mice weighing over 30 g were deemed more suitable for the experiment due to the larger size of their bile duct (BD). (2) a 2 Fr (outer diameter = 0.66 mm) polyurethane cannula was better suited for cannulating the portal vein (PV) when compared to a polypropylene cannula. This was attributed to the polyurethane material&#39;s enhanced grip, resulting in reduced catheter slippage during the transfer from the body to the organ chamber. (3) for cannulation of the bile duct (BD), a 1 Fr (outer diameter = 0.33 mm) polyurethane cannula was found to be more effective compared to the polypropylene UT - 03 (outer diameter = 0.30 mm) cannula. With this optimized protocol, mouse livers were successfully preserved for a duration of 12 h without significant impact on the histological structure. Hematoxylin-Eosin (HE) staining revealed a well-preserved morphological architecture of the liver, characterized by predominantly viable hepatocytes with clearly visible nuclei and mild dilation of hepatic sinusoids.

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

A normothermic ex vivo liver perfusion (NEVLP) system was created for mouse livers. This system requires experience in microsurgery but allows for reproducible perfusion results. The ability to utilize mouse livers facilitates the investigation of molecular pathways to identify novel perfusate additives and enables the execution of experiments focused on organ repair.

This protocol presents an optimized erythrocytes-free NEVLP system using mouse livers. Ex vivo preservation of mouse livers was achieved by employing ...

Engineering

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 ...

Isolation of Rat Portal Fibroblasts by In situ Liver Perfusion

Liver fibrosis is defined by the excessive deposition of extracellular matrix by activated myofibroblasts. There are multiple precursors of hepatic myofibroblasts, including hepatic stellate cells, portal fibroblasts and bone marrow derived fibroblasts 1. Hepatic stellate cells have been the best studied, but portal fibroblasts are increasingly recognized as important contributors to the myofibroblast pool, particularly in biliary fibrosis 2. Portal fibroblasts undergo proliferation in response to biliary epithelial injury, potentially playing a key role in the early stages of biliary scarring 3-5. A method of isolating portal fibroblasts would allow in vitro study of this cell population and lead to greater understanding of the role portal fibroblasts play in biliary fibrosis.
Portal fibroblasts have been isolated using various techniques including outgrowth 6, 7 and liver perfusion with enzymatic digestion followed by size selection 8. The advantage of the digestion and size selection technique compared to the outgrowth technique is that cells can be studied without the necessity of passage in culture. Here, we describe a modified version of the original technique described by Kruglov and Dranoff 8 for isolation of portal fibroblasts from rat liver that results in a relatively pure population of primary cells.

Isolation of Rat Portal Fibroblasts by In situ Liver Perfusion

A technique for isolating portal fibroblasts from rat liver is described. Livers are perfused and digested in situ with collagenase, followed by ex vivo digestion of the liver slurry and size selection of cells. This method provides a pure population of portal fibroblasts without the need for passage in culture.

Liver fibrosis is defined  by the excessive deposition of extracellular matrix by activated  myofibroblasts. There are multiple precursors of hepatic ...

Biology

Isolation and Primary Culture of Rat Hepatic Cells

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology and other related subjects. The method based on two-step collagenase perfusion for isolation of intact hepatocytes was first introduced by Berry and Friend in 1969 1 and, since then, has undergone many modifications. The most commonly used technique was described by Seglenin 1976 2. Essentially, hepatocytes are dissociated from anesthetized adult rats by a non-recirculating collagenase perfusion through the portal vein. The isolated cells are then filtered through a 100 &mu;m pore size mesh nylon filter, and cultured onto plates. After 4-hour culture, the medium is replaced with serum-containing or serum-free medium, e.g. HepatoZYME-SFM, for additional time to culture. These procedures require surgical and sterile culture steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol, which allow consistently in the generation of viable hepatocytes in large numbers.

Primary hepatocytes provide a valuable tool to evaluate biochemical, molecular, and metabolic functions in a physiologically relevant experimental system. We describe a reliable protocol for rat in situ liver perfusion, which consistently generates viable hepatocytes up to 1.0 &times; 108 cells per preparation with cell viability between 88 ~ 96%.

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology ...

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 Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

Medicine

Isolation of Primary Rat Hepatocytes with Multiparameter Perfusion Control

Primary hepatocytes are widely used in basic research on liver diseases and for toxicity testing in vitro. The two-step collagenase perfusion procedure for primary hepatocyte isolation is technically challenging, especially in portal vein cannulation. The procedure is also prone to occasional contamination and variations in perfusion conditions due to difficulties in the assembly, optimization, or maintenance of the perfusion setup. Here, a detailed protocol for an improved two-step collagenase perfusion procedure with multiparameter perfusion control is presented. Primary rat hepatocytes were successfully and reliably isolated by taking the necessary technical precautions at critical steps of the procedure, and by reducing the operational difficulty and mitigating the variability of perfusion parameters through the adoption of a special intravenous catheter, standardized sterile disposable tubing, temperature control, and real-time monitoring and alarm system. The isolated primary rat hepatocytes consistently exhibit high cell viability (85%-95%), yield (2-5 x 108 cells per 200-300 g rat) and functionality (albumin, urea and CYP activity). The procedure was complemented by an integrated perfusion system, which is compact enough to be set up in the laminar flow hood to ensure aseptic operation.

This protocol details the use of a special intravenous catheter, standardized sterile disposable tubing, temperature control complemented by real-time monitoring, and an alarm system for two-step collagenase perfusion procedure to improve the consistency in the viability, yield, and functionality of isolated primary rat hepatocytes.

Primary hepatocytes are widely used in basic research on liver diseases and for toxicity testing in vitro. The two-step collagenase perfusion procedure ...

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

Summary

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Chapters in this video

Decellularization and Recellularization of Whole Livers

Purification of Hepatocytes and Sinusoidal Endothelial Cells from Mouse Liver Perfusion

Normothermic Ex Vivo Liver Machine Perfusion in Mouse

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

Isolation of Rat Portal Fibroblasts by In situ Liver Perfusion

Isolation and Primary Culture of Rat Hepatic Cells

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

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

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

Isolation of Primary Rat Hepatocytes with Multiparameter Perfusion Control