This work was supported by grants from the Ministry of Education Innovation Team Development Program of China (No. IRT16R57), National Natural Science Foundation of China (No. 81470896), and a research fund for Young Talent Recruiting Plans of Xi'an Jiaotong University (RW).

The protocol was carried out in accordance with the Guidelines for the Care and Use of Laboratory Animals and was approved by the Committee on the Ethics of Animal Experiments of Xi&#39;an Jiaotong University, Xi&#39;an, Shaanxi Province, China.
NOTE: None of the procedures in this protocol was performed under a surgical microscope.
1. Preoperative Preparation
<ol>
	<li>The Design and Shape of the Magnetic Rings (Figure 1)
	<ol>
		<li>Assemble the magnetic anastomosis device by matching a pair of neodymium-iron-boron magnetic rings.</li>
		<li>Design the rings according to the parameters of rat SHVC with a major axis diameter, minor axis diameter, thickness and weight of 8 mm, 5.5 mm, 1 mm, 0.5 g, respectively.</li>
		<li>Curve the rings to the desired shape by a wire-electrode cutting technique, and then coat the obtained rings with a titanium oxide airbrush and sterilize with 500 mg/L ethylene oxide at 55-60 &#176;C for 6 h.</li>
	</ol>
	</li>
	<li>The cuffs (Figure 2)
	<ol>
		<li>Prepare the cuff for anastomosis of the IHVC and PV by cutting the polyethylene tube. Ensure that the cuff body is 5 mm long with a 2 mm cuff extension. 
		NOTE: The inner and outer diameters of the PV cuff are 1.8 and 2.1 mm, while those of the IHVC cuff are 2.6 and 2.8 mm, respectively.</li>
	</ol>
	</li>
	<li>The Stent (Figure 2)
	<ol>
		<li>Prepare the stent for anastomosis of the BD by cutting 24-G intravenous cannula with a length of 5 mm to produce slant at both ends.</li>
	</ol>
	</li>
	<li>Experimental Animals
	<ol>
		<li>Use male Sprague Dawley rats weighing between 250 and 280 g as donors and recipients. Ensure that the recipient is the same weight or slightly heavier (&#60; 10 g) than the donor.</li>
		<li>Keep the rats in climate-controlled rooms with free access to food and water before surgery.</li>
	</ol>
	</li>
</ol>
2. Donor Operation
<ol>
	<li>Inject buprenorphine (0.05 mg/kg) subcutaneously as an analgesic 1 hour before operation.</li>
	<li>Anesthetize the donor rat by isoflurane inhalation. Use 5 vol% isoflurane at a flow rate of 1.5 L/min in a Plexiglas box for the induction of anesthesia and 2 vol% isoflurane at a flow rate 0.6-0.8 L/min on a cone mask for the maintenance of anesthesia. Confirm depth of anesthesia by performing a toe pinch and a skin pinch.</li>
	<li>Shave the entire abdominal skin of the rat using an electric shaver to allow for cleaner exposure. Disinfect the corresponding skin with a povidone-iodine solution.</li>
	<li>Open the abdominal cavity via a cruciform incision using surgical scissors from the root of the penis extending to 1 cm above the xiphisternum along linea alba, extending from the midpoint of the longitudinal incision to the midaxillary line. Exteriorize the gastrointestinal tract to the left and cover it with wet gauze.</li>
	<li>Clamp and pull the xiphisternum toward the head using a hemostatic forceps. Dissect the falciform ligament and the connective tissues around the liver. Isolate and ligate the left inferior phrenic vein with a 6-0 silk suture.</li>
	<li>Insert the stent into the bile duct.
	<ol>
		<li>Clamp the BD just above the point where the gastroduodenal vein joins the portal vein and pull it to maintain a little tension using a microforceps. Then perform a &#34;V&#34; incision of 1 mm length on the anterior wall about 5 mm proximal to the biliary confluence by a micro scissors.</li>
		<li>Insert the stent into the lumen of the bile duct using a curved microforceps. Make sure at least a half of the stent lies outside the bile duct.</li>
		<li>Secure the stent with a 6-0 silk suture. Cut one end of the suture. Keep the other one to hold during later anastomosis. Transect the bile duct below the stent and make sure the bile can come out of the stent.</li>
	</ol>
	</li>
	<li>Isolate the IHVC down to the left renal vein level. Free the IHVC from anterior and lateral surrounding tissues by cotton bud dissection until the IHVC is skeletonized. Separate and ligate the right suprarenal vein and the right renal vein with a 6-0 silk suture.</li>
	<li>Heparinize the rat by injecting 50 U of heparin diluted in 2 mL of normal saline solution through the dorsal vein of the penis.</li>
	<li>Perfuse the liver.
	<ol>
		<li>Isolate the abdominal aorta below the left renal vein. Insert a 22-G catheter into the aorta.</li>
		<li>Perform a thoracotomy, clamp the thoracic aorta using a microforceps.</li>
		<li>Perfuse the donor liver through the catheter in the aorta with 20 mL of normal saline solution containing 25 U of heparin (4 &#176;C) at the rate of 3 mL/min with an infusion pump. In the meantime, cut the vena cava below the right renal vein&#160;to allow the perfusion solution to flow out of the liver.</li>
		<li>Transect the SHVC along with part of the diaphragm and IHVC at the level of the left renal vein when the donor liver became pallid.</li>
	</ol>
	</li>
	<li>Dissect the connective tissue surrounding the PV by two micro forceps. Ligate and divide the pyloric veins with an 8-0 prolene suture. Transect the PV at the level of the splenic vein. Excise the donor liver and rinse it with cold normal saline solution. Place it in a cold saline bath 4 &#176;C.</li>
</ol>
3. Graft Preparation
NOTE: All procedures for the liver graft preparation are performed in a cold saline bath at 4 &#176;C.
<ol>
	<li>Attach a cuff to the donor&#39;s IHVC (Figure 3).
	<ol>
		<li>Insert the IHVC into its cuff with the cuff extension pointing towards the liver. Position the cuff extension on the posterior wall of IHVC. Fix the cuff extension and the IHVC with a curved microforceps itself attached using a file clip to the bath container.</li>
		<li>Evert the distal end of the IHVC over the cuff body using two micro forceps.</li>
		<li>Secure the edge with a 6-0 silk suture. Make sure the IHVC is not twisted during the process.</li>
	</ol>
	</li>
	<li>Repeat step 3.1 on the PV to attach a cuff to the donor&#39;s PV.</li>
	<li>Attach a magnetic ring to the donor&#39;s SHVC (Figure 4).
	<ol>
		<li>Pull the diaphragm through the four forceps and trim the SHVC.</li>
		<li>Pass the SHVC through the magnetic ring using the titanium-alloy micro forceps.</li>
		<li>Evert the distal end of the SHVC over the ring.</li>
		<li>Secure the ring with a 6-0 silk suture. Make sure the SHVC is not twisted during the process.</li>
		<li>Remove the excess diaphragm around the liver.</li>
	</ol>
	</li>
	<li>Immerse the donor liver in a cold normal saline bath and store at 4 &#176;C.</li>
</ol>
4. Recipient Operation
NOTE: The schema of the graft implantation in the recipient rat is shown in Figure 5.
<ol>
	<li>Inject buprenorphine (0.05 mg/kg) subcutaneously as an analgesic 1 hour before operation. Anesthetize the donor rat by isoflurane inhalation. Use 5 vol% isoflurane at a flow rate of 1.5 L/min in a Plexiglas box for the induction of anesthesia and 2 vol% isoflurane at a flow rate 0.6-0.8 L/min on a cone mask for the maintenance of anesthesia.</li>
	<li>Shave the entire abdominal skin of the rat. Disinfect the corresponding skin with a povidone-iodine solution.</li>
	<li>Open the abdominal cavity via a 4 cm midline incision. Place the abdominal retractors. Pull the ribs as far apart as possible from the midline.</li>
	<li>Clamp and pull the xiphisternum toward the head using a hemostatic forceps. Dissect the connective tissues and ligaments around the liver. Ligate and divide the hepato-esophageal ligament. Ligate and divide the hepatic artery with an 8-0 prolene suture.</li>
	<li>Isolate the common bile duct from the first hepatic portal. Place a 6-0 silk suture around the common bile duct just below its division and ligate the common bile duct.</li>
	<li>Dissect the IHVC down to the right renal vein. Ligate and divide the right adrenal vein. Place a strap in the SHVC of the liver.</li>
	<li>Clamp the IHVC just above the right renal vein with microvessel clips. Clamp the PV at the level of the pyloric vein with microvessel clips.
	<ol>
		<li>To flush the blood out of the liver, slowly inject 2 mL of normal saline through the PV. Drag down the liver by pulling the previously placed strap. Clamp the SHVC and part of the diaphragm with a Satinsky clamp.</li>
		<li>Transect the IHVC close to the parenchyma and the PV at the porta hepatis. Excise the liver by transecting the SHVC just above the liver. Remove the recipient liver quickly.</li>
	</ol>
	</li>
	<li>Attach a magnetic ring to the recipient&#39;s SHVC.
	<ol>
		<li>Insert the recipient&#39;s remaining SHVC into a magnetic ring.</li>
		<li>Fix the magnetic ring on a Satinsky clamp by magnetic attraction.</li>
		<li>Evert the SHVC over the magnetic ring until the vessel covers the edge of the ring.</li>
	</ol>
	</li>
	<li>Place the donor liver graft orthotopically and cover it with a cold wet gauze.</li>
	<li>Fill the lumen of the donor&#39;s and the recipient&#39;s SHVCs with normal saline solution. Remove air bubbles thoroughly to prevent air embolism. Finish the SHVC reconstruction by coupling the magnetic rings embedded in the donor&#39;s and recipient&#39;s SHVC together through magnetic force.</li>
	<li>Reconstruct the PV.
	<ol>
		<li>Tract the right and left branch of the recipient&#39;s PV by pulling an 8-0 prolene suture to maintain the tension of the PV.</li>
		<li>Move the clamp of recipient&#39;s PV down from the pylorus vein to the splenic vein in order to expose the vein fully.</li>
		<li>Incise the anterior wall of the recipient. Fill the lumen of the PV with normal saline solution.</li>
		<li>Hold the cuff extension of the donor&#39;s PV with a curved forceps.</li>
		<li>Insert the cuff body of the donor&#39;s PV into the recipient&#39;s PV while keep flushing the PV lumen with normal saline.</li>
		<li>Secure it with a circumferential 6-0 silk ligature. Re-establish the blood flow to the graft liver by releasing the clamps on the PV and SHVC.</li>
	</ol>
	</li>
	<li>Repeat step 4.11 on the IHVC to reconstruct the IHVC.</li>
	<li>Reconstruct the bile duct by a stent technique.
	<ol>
		<li>Incise a &#34;V&#34; on the anterior wall of the recipient common bile duct.</li>
		<li>Insert the donor stent into the lumen of the recipient BD and fix it with a 6-0 silk circumferential suture. Pull the sutures on the two bile ducts together and tie the sutures to bring the BD close to each other.</li>
		<li>Wrap omentum tissue around the anastomotic site to prevent bile leak.</li>
	</ol>
	</li>
	<li>Irrigate the abdominal cavity with warm normal saline solution. Close the abdomen incision in two layers with a continuous 3-0 non-absorbable surgical suture.</li>
	<li>Administer 16 mg/kg cefuroxime (an antibiotic) and 0.05 mg/kg buprenorphine (an analgesic) by subcutaneous injection after the operation immediately.</li>
	<li>Place the rat in a clean cage and allow food and water ad libitum.</li>
	<li>Inject cefuroxime (16 mg/kg) and buprenorphine (0.05 mg/kg) into the abdominal cavity every 12 hours for 3 days.</li>
</ol>

<ol>
	<li>Aller, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+half+century+(1961-2011)+of+applying+microsurgery+to+experimental+liver+research.">A half century (1961-2011) of applying microsurgery to experimental liver research.</a> World journal of hepatology. 4, 199-208 (2012).</li><li>Aller, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+value+of+microsurgery+in+liver+research.">The value of microsurgery in liver research.</a> Liver international : official journal of the International Association for the Study of the Liver. 29, 1132-1140 (2009).</li><li>Lee, S., Charters, A. C., Chandler, J. G., Orloff, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technique+for+orthotopic+liver+transplantation+in+the+rat.">A technique for orthotopic liver transplantation in the rat.</a> Transplantation. 16, 664-669 (1973).</li><li>Kamada, N., Calne, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthotopic+liver+transplantation+in+the+rat.+Technique+using+cuff+for+portal+vein+anastomosis+and+biliary+drainage.">Orthotopic liver transplantation in the rat. Technique using cuff for portal vein anastomosis and biliary drainage.</a> Transplantation. 28, 47-50 (1979).</li><li>Oldani, G., Lacotte, S., Morel, P., Mentha, G., Toso, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthotopic+liver+transplantation+in+rats.">Orthotopic liver transplantation in rats.</a> Journal of visualized experiments : JoVE. , (2012).</li><li>Nagai, K., Yagi, S., Uemoto, S., Tolba, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Surgical+procedures+for+a+rat+model+of+partial+orthotopic+liver+transplantation+with+hepatic+arterial+reconstruction.">Surgical procedures for a rat model of partial orthotopic liver transplantation with hepatic arterial reconstruction.</a> Journal of visualized experiments : JoVE. , e4376 (2013).</li><li>Liu, X., He, C., Huang, T., Gu, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+New+Technique+for+Reconstruction+of+Hepatic+Artery+during+Liver+Transplantation+in+Sprague-Dawley+Rat.">Development of a New Technique for Reconstruction of Hepatic Artery during Liver Transplantation in Sprague-Dawley Rat.</a> PloS one. 10, e0145662 (2015).</li><li>Jamshidi, R., Stephenson, J. T., Clay, J. G., Pichakron, K. O., Harrison, M. 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The authors have nothing to disclose.

Many clinical progresses in liver transplantation can be attributed to animal studies. Rat OLT is a widely used and well accepted model in the research of organ preservation, transplant immunology, physiology and pathology. However, it is also a very complicated procedure and requires advanced microsurgical skill. Despite many improvements in the procedure of rat OLT, the SHVC anastomosis remained a challenging step. In this article, we describe a novel magnetic anastomosis technique for SHVC reconstruction in rat OLT. The application of this technique reduced the time for SHVC reconstruction to less than 2 min and the anhepatic phase of rat OLT to approximately 10 min. We believe this technique has great implications in transplant research. 
The critical step in using this magnetic anastomosis technique for the reconstruction of the SHVC is to attach the magnetic rings to the donor's and recipient's SHVC. Every caution should be used to avoid twisting the SHVC. Moreover, when connecting the two magnetic rings, air bubbles in the SHVC should be removed thoroughly. 
There are some limitations in using this technique. Some strains of rats such as Fischer-344 rats and Buffalo rats have an extremely short SHVC. It is impossible to attach the magnetic ring to their SHVC. Thus, they are not suitable for this technique. However, this technique can be easily applied for OLT in most strains of rats including Sprague Dawley rats, Lewis rats and brown Norway rats. Magnetic rings can interfere with the magnetic resonance imaging (MRI) scan. Therefore, this magnetic anastomosis technique cannot be used if an MRI exam is required for the experiment.
In summary, the magnetic anastomosis technique makes easy and fast SHVC reconstruction in rats possible. After a reasonable learning curve, even researchers without advanced microsurgical skills can produce reliable and reproducible results using this rat model of OLT.

The rat model of orthotopic liver transplantation (OLT) is essential for transplant research1,2. The first rat OLT was described by Lee et al. in 19733. In that model, all the vessels were reconstructed by the hand-suture technique. The hand-suture technique requires advanced microsurgical skills, which significantly limits its utilization. Since then, various modifications to the original protocol of rat OLT have been reported. Among them, the cuff technique for anastomosis of the portal vein (PV) and infrahepatic vena cava (IHVC) reported by Kamada et al. in 1979 is considered a major improvement to this model, as it significantly simplified the reconstruction procedures4. However, due to the short anterior wall of the recipients&#39; suprahepatic vena cava (SHVC), the cuff technique is very difficult to use for the reconstruction of the SHVC. Most researchers in this field still use the hand-suture technique for SHVC reconstruction, which makes it the bottleneck step in rat OLT5,6,7.
The magnetic anastomosis technique (i.e., magnamosis) is a method of connecting two vessels or other tubular structures using the attractive force between two magnets8,9,10,11. The magnetic force gradually compresses and remodels the tissue into a strong, sutureless anastomosis12,13. This compression anastomosis has been proven to be effective in humans14,15. We have designed a pair of magnetic rings specifically for the anastomosis of the SHVC in rats. Our recent study has shown that the magnetic anastomosis technique is superior to the hand-suture technique for SHVC reconstruction in rat OLT16. The purpose of this article is to provide a detailed, step-by-step protocol for SHVC reconstruction in rats using the novel magnetic anastomosis technique.

<table border="0" cellpadding="0" cellspacing="0" width="1148">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Anesthesia Machine</td>
			<td style="width:377px;">Harvard</td>
			<td style="width:119px;">&#160;tabletop</td>
			<td style="width:353px;">Animal anaesthesia</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:299px;">PLX7000B HF Mobile Digital C-arm System</td>
			<td style="width:377px;">Perlong Medical</td>
			<td style="width:119px;">PLX7000B</td>
			<td>It is mainly used for the angiography and photography of various operations</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:299px;">Syringe Pump</td>
			<td style="width:377px;">Mindray</td>
			<td style="width:119px;">BeneFusion SP5</td>
			<td>intravenous infusion</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Isoflurane</td>
			<td style="width:377px;">RWD life Science Co.</td>
			<td style="width:119px;"></td>
			<td>anesthetic:for the induction and maintenanceof anesthesia</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">iohexol</td>
			<td style="width:377px;">Shanghai General Pharmaceutical Co</td>
			<td style="width:119px;"></td>
			<td>&#160;intravascular contrast media&#160;&#160;</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:299px;">heparin sodium injection</td>
			<td style="width:377px;">SPH No.1 Biochemical &#38; Pharmaceutical Co., LTD&#160;</td>
			<td style="width:119px;"></td>
			<td>&#160;prevent the formation of thrombosis&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">cefuroxime&#160;</td>
			<td style="width:377px;">Glaxo Operations UK Limited&#160;</td>
			<td style="width:119px;"></td>
			<td>an antibiotic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">buprenorphine&#160;</td>
			<td style="width:377px;">TIPR Pharmaceutical Responsible Co.,Ltd</td>
			<td style="width:119px;"></td>
			<td>an analgesic</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">curved microforceps</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">W40350</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">hemostatic forceps(straight)</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">J31010</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">hemostatic forceps(curved)</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">J31020</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Satinsky clamp</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">XEC050</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">needle holder</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">J32010</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">microneedle holder</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">WBA040</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">notched forceps</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">J42010</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">tissue forceps(with hook)</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">J41010</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">tissue scissor</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">Y00040</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">surgical scissors</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">Y00030</td>
			<td>surgical tool</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:299px;">micro scissors</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">MR-S121T</td>
			<td>surgical tool</td>
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			<td height="21" style="height:21px;width:299px;">microvessel clips</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">XEC240</td>
			<td>surgical tool</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:299px;">straight microforceps(titanium alloy)</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">WCC010</td>
			<td>surgical tool</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">curved microforceps&#160; (titanium alloy)</td>
			<td style="width:377px;">Shanghai Medical Instruments (Group) Ltd., Corp.</td>
			<td style="width:119px;">WCC020</td>
			<td>surgical tool</td>
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a rat model of orthotopic liver transplantation using a novel magnetic anastomosis technique for suprahepatic vena cava reconstruction

The rat model of orthotopic liver transplantation (OLT) is essential for transplant research. It is a very sophisticated animal model and requires a steep learning curve. The introduction of the cuff technique for anastomosis of the portal vein (PV) and infrahepatic vena cava (IHVC) has significantly simplified the transplant procedure in rats. However, due to the short anterior wall of the recipients' suprahepatic vena cava (SHVC), the cuff technique is very difficult to use for the reconstruction of the SHVC. Most researchers in this field still use the hand-suture technique for SHVC reconstruction, which makes it the bottleneck step in rat orthotopic liver transplantation. The magnetic anastomosis technique (i.e., magnamosis) is a method of connecting two vessels using the attractive force between two magnets. Our recent study has shown that the magnetic anastomosis technique is superior to the hand-suture technique for SHVC reconstruction in rats. In this article, we show a step-by-step protocol for SHVC reconstruction in rats using the novel magnetic anastomosis technique. In this model, the reconstruction of the PV and IHVC was performed by the standard cuff technique, while the reconstruction of the bile duct (BD) was performed by a stent technique. The hepatic re-arterialization was not performed. The magnetic anastomosis technique made SHVC reconstruction much easier and significantly shortened the anphepatic phase. After a reasonable learning curve, even researchers without advanced microsurgical skills can produce reliable and reproducible results using this rat model of OLT.

After approximately 10 attempts, the magnetic anastomosis technique for SHVC reconstruction was successfully mastered by a researcher who had no prior microsurgical training. The reconstruction of the SHVC took less than 2 min. The anhepatic phase for the recipient rats was approximately 10 min. No thrombosis, bleeding or angiostegnosis was observed at the SHVC anastomotic site at 1, 5 and 30 days after reperfusion. The inferior vena cavography was performed at 2 weeks after transplantation. As shown in Figure 6, the magnetic rings were intact, and the blood flow was patent through the SHVC anastomosis. The post-transplant survival rate was 95% at day 1, 90% at day 3, and 85% at days 7 to 30 (Figure 7).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56933/56933fig1.jpg" /> 
Figure 1: The design (A) and photo (B) of the magnetic ring for anastomosis of the SHVC. 
SHVC: suprahepatic vena cava.&#160; <a href="/files/ftp_upload/56933/56933fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56933/56933fig2.jpg" /> 
Figure 2: The cuffs and the stent for anastomosis of the IHVC (A), PV (B) and BD (C). 
IHVC: infrahepatic vena cava; PV: portal vein; BD: bile duct. <a href="/files/ftp_upload/56933/56933fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56933/56933fig3.jpg" /> 
Figure 3: Schema of the cuff anastomosis of the IHVC and PV. Insert the IHVC or PV into a cuff (A); Evert the distal end of the IHVC or PV over the cuff body using two microforceps (B); Secure the edge with a 6-0 silk suture (C); Insert the cuff body of the donor&#39;s IHVC or PV into the recipient&#39;s IHVC or PV (D). 
IHVC: infrahepatic vena cava; PV: portal vein. <a href="/files/ftp_upload/56933/56933fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56933/56933fig4.jpg" /> 
Figure 4: Schema of the magnetic anastomosis of the SHVC. Pass the SHVC through a magnetic ring using the titanium-alloy microforceps (A); Evert the distal end of the SHVC over the ring (B); Secure the ring with a 6-0 silk suture (C). Couple the magnetic rings embedded in the donor&#39;s and recipient&#39;s SHVC together through magnetic force (D). 
SHVC: suprahepatic vena cava. <a href="/files/ftp_upload/56933/56933fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56933/56933fig5.jpg" /> 
Figure 5: Schema of the graft implantation in the recipient rat. Reconstruction of the SHVC: magnetic anastomosis; Reconstruction of the IHVC and PV: cuff anastomosis; Reconstruction of the BD: stent anastomosis. Reconstruction of the HA: not performed. 
SHVC: suprahepatic vena cava; IHVC: infrahepatic vena cava; PV: portal vein; BD: bile duct; HA: hepatic artery. <a href="/files/ftp_upload/56933/56933fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56933/56933fig6.jpg" /> 
Figure 6: Inferior vena cavography of the rat at 2 weeks after transplantation. <a href="/files/ftp_upload/56933/56933fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/56933/56933fig7.jpg" /> 
Figure 7: Representative survival cure after transplantation. <a href="/files/ftp_upload/56933/56933fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Rat Model of Orthotopic Liver Transplantation Using a Novel Magnetic Anastomosis Technique for Suprahepatic Vena Cava Reconstruction at JoVE.com

A Rat Model of Orthotopic Liver Transplantation Using a Novel Magnetic Anastomosis Technique for Suprahepatic Vena Cava Reconstruction

The reconstruction of the suprahepatic vena cava (SHVC) remains a difficult step in rat orthotopic liver transplantation. In this article, we show a step-by-step protocol for SHVC reconstruction in rats using a novel magnetic anastomosis technique.

The rat model of orthotopic liver transplantation (OLT) is essential for transplant research. It is a very sophisticated animal model and requires a steep ...

a-rat-model-orthotopic-liver-transplantation-using-novel-magnetic

Nanyang Technological University

Research

JoVE Journal

Medicine

9.7K Views.  First Affiliated Hospital, Xi'an Jiaotong University. The reconstruction of the suprahepatic vena cava (SHVC) remains a difficult step in rat orthotopic liver transplantation. In this article, we show a step-by-step protocol for SHVC reconstruction in rats using a novel magnetic anastomosis technique. 

Video: A Rat Model of Orthotopic Liver Transplantation Using a Novel Magnetic Anastomosis Technique for Suprahepatic Vena Cava Reconstruction

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

Electrolytic Inferior Vena Cava Model (EIM) of Venous Thrombosis

Animal models serve a vital role in deep venous thrombosis (DVT) research in order to study thrombus formation, thrombus resolution and to test potential therapeutic compounds (1). New compounds to be utilized in the treatment and prevention of DVT are currently being developed. The delivery of potential therapeutic antagonist compounds to an affected thrombosed vein has been problematic. In the context of therapeutic applications, a model that uses partial stasis and consistently generates thrombi within a major vein has been recently established. The Electrolytic Inferior vena cava Model (EIM) is mouse model of DVT that permits thrombus formation in the presence of continuous blood flow. This model allows therapeutic agents to be in contact with the thrombus in a dynamic fashion, and is more sensitive than other models of DVT (1). In addition, this thrombosis model closely simulates clinical situations of thrombus formation and is ideal to study venous endothelial cell activation, leukocyte migration, venous thrombogenesis, and to test therapeutic applications (1). The EIM model is technically simple, easily reproducible, creates consistent thrombi sizes and allows for a large sample (i.e. thrombus and vein wall) which is required for analytical purposes.

Electrolytic Inferior Vena Cava Model EIM of Venous Thrombosis

The electrolytic induction of endothelial activation to the internal surface of the Inferior Vena Cava results in venous type thrombus formation due to endothelial activation and partial blood stasis, two components of Virchow's triad.

Animal models serve a vital role in deep venous thrombosis (DVT) research in order to study thrombus formation, thrombus resolution and to test potential ...

Mouse Complete Stasis Model of Inferior Vena Cava Thrombosis

Venous thromboembolism (VTE) includes both deep vein thrombosis (DVT) and pulmonary embolism (PE). In the United States (U.S.), the high morbidity and mortality rates make VTE a serious health concern 1-2. After heart disease and stroke, VTE is the third most common vascular disease 3. In the U.S. alone, there is an estimated 900,000 people affected each year, with 300,000 deaths occurring annually 3. A reliable in vivo animal model to study the mechanisms of this disease is necessary.
The advantages of using the mouse complete stasis model of inferior vena cava thrombosis are several. The mouse model allows for the administration of very small volumes of limited availability test agents, reducing costs dramatically. Most promising is the potential for mice with gene knockouts that allow specific inflammatory and coagulation factor functions to be delineated. Current molecular assays allow for the quantitation of vein wall, thrombus, whole blood, and plasma for assays. However, a major concern involving this model is the operative size constraints and the friability of the vessels. Also, due to the small IVC sample weight (mean 0.005 grams) it is necessary to increase animal numbers for accurate statistical analysis for tissue, thrombus, and blood assays such as real-time polymerase chain reaction (RT-PCR), western blot, enzyme-linked immunosorbent (ELISA), zymography, vein wall and thrombus cellular analysis, and whole blood and plasma assays 4-8.
The major disadvantage with the stasis model is that the lack of blood flow inhibits the maximal effect of administered systemic therapeutic agents on the thrombus and vein wall.

The mouse complete stasis model of inferior vena cava thrombosis yields quantifiable amounts of vein wall tissue and thrombus. It has proven useful for evaluating interactions between the vein wall and the occlusive thrombus and in assessing the progression from acute to chronic inflammation.

Venous thromboembolism (VTE) includes both deep vein thrombosis (DVT) and pulmonary embolism (PE).  In the United States (U.S.), the high morbidity and ...

Orthotopic Liver Transplantation in Rats

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat liver transplantation in 1979 by Kamada et al.3, this model has remained the gold standard despite various proposed alternative techniques4. Nevertheless, its broader use is limited by its steep learning curve5.
In this video paper, we show a simple and easy-to-establish revision of Kamada's two-cuff technique. The suprahepatic vena cava anastomosis is performed manually with a running suture, and the vena porta and infrahepatic vena cava anastomoses are performed utilizing a quick-linker cuff system6. Manufacturing the quick-linker kit is shown in a separate video paper.

We present an easy-to-establish revision of the classical two-cuff technique for orthotopic liver transplantation in rat.

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat ...

Surgical Procedures for a Rat Model of Partial Orthotopic Liver Transplantation with Hepatic Arterial Reconstruction

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as studies on graft preservation and ischemia-reperfusion injury 1,2, immunological responses 3,4, hemodynamics 5,6, and small-for-size syndrome 7. The rat OLT is among the most difficult animal models in experimental surgery and demands advanced microsurgical skills that take a long time to learn. Consequently, the use of this model has been limited. Since the reliability and reproducibility of results are key components of the experiments in which such complex animal models are used, it is essential for surgeons who are involved in rat OLT to be trained in well-standardized and sophisticated procedures for this model.
While various techniques and modifications of OLT in rats have been reported 8 since the first model was described by Lee et al. 9 in 1973, the elimination of the hepatic arterial reconstruction 10 and the introduction of the cuff anastomosis technique by Kamada et al. 11 were a major advancement in this model, because they simplified the reconstruction procedures to a great degree. In the model by Kamada et al., the hepatic rearterialization was also eliminated. Since rats could survive without hepatic arterial flow after liver transplantation, there was considerable controversy over the value of hepatic arterialization. However, the physiological superiority of the arterialized model has been increasingly acknowledged, especially in terms of preserving the bile duct system 8,12 and the liver integrity 8,13,14.
In this article, we present detailed surgical procedures for a rat model of OLT with hepatic arterial reconstruction using a 50% partial graft after ex vivo liver resection. The reconstruction procedures for each vessel and the bile duct are performed by the following methods: a 7-0 polypropylene continuous suture for the supra- and infrahepatic vena cava; a cuff technique for the portal vein; and a stent technique for the hepatic artery and the bile duct.

Orthotopic liver transplantation in rats is an indispensable experimental model for biomedical research. Here we present our surgical procedures for orthotopic rat liver transplantation with hepatic arterial reconstruction using a 50% partial graft.

Orthotopic liver transplantation (OLT) in rats using a whole or partial graft is an indispensable experimental model for transplantation research, such as ...

Implantation of Inferior Vena Cava Interposition Graft in Mouse Model

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. The long-term clinical results showed excellent patency rates, however, with significant incidence of stenosis. To investigate the cellular and molecular mechanisms of vascular neotissue formation and prevent stenosis development in tissue engineered vascular grafts (TEVGs), we developed a mouse model of the graft with approximately 1 mm internal diameter. First, the TEVGs were assembled from biodegradable tubular scaffolds fabricated from a polyglycolic acid nonwoven felt mesh coated with &#949;-caprolactone and L-lactide copolymer. The scaffolds were then placed in a lyophilizer, vacuumed for 24 hr, and stored in a desiccator until cell seeding. Second, bone marrow was collected from donor mice and mononuclear cells were isolated by density gradient centrifugation. Third, approximately one million cells were seeded on a scaffold and incubated O/N. Finally, the seeded scaffolds were then implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. The implanted grafts demonstrated excellent patency (&#62;90%) without evidence of thromboembolic complications or aneurysmal formation. This murine model will aid us in understanding and quantifying the cellular and molecular mechanisms of neotissue formation in the TEVG.

To improve our knowledge of cellular and molecular neotissue formation, a murine model of the TEVG was recently developed. The grafts were implanted as infrarenal vena cava interposition grafts in C57BL/6 mice. This model achieves similar results to those achieved in our clinical investigation, but over a far shortened time-course.

Biodegradable scaffolds seeded with bone marrow mononuclear cells (BMCs) are often used for reconstructive surgery to treat congenital cardiac anomalies. ...

Technique of Porcine Liver Procurement and Orthotopic Transplantation using an Active Porto-Caval Shunt

The success of liver transplantation has resulted in a dramatic organ shortage. Each year, a considerable number of patients on the liver transplantation waiting list die without receiving an organ transplant or are delisted due to disease progression. Even after a successful transplantation, rejection and side effects of immunosuppression remain major concerns for graft survival and patient morbidity. 
Experimental animal research has been essential to the success of liver transplantation and still plays a pivotal role in the development of clinical transplantation practice. In particular, the porcine orthotopic liver transplantation model (OLTx) is optimal for clinically oriented research for its close resemblance to human size, anatomy, and physiology. 
Decompression of intestinal congestion during the anhepatic phase of porcine OLTx is important to guarantee reliable animal survival. The use of an active porto-caval-jugular shunt achieves excellent intestinal decompression. The system can be used for short-term as well as long-term survival experiments. The following protocol contains all technical information for a stable and reproducible liver transplantation model in pigs including post-operative animal care.

Experimental animal research plays a pivotal role in the development of clinical transplantation practice. The porcine orthotopic liver transplantation model (OLTx) closely resembles human conditions and is frequently used in clinically oriented research. The following protocol contains all information for a reliable porcine OLTx model using an active porto-caval-jugular shunt.

The success of liver transplantation has resulted in a dramatic organ shortage. Each year, a considerable number of patients on the liver transplantation ...

Mouse Model for Pancreas Transplantation Using a Modified Cuff Technique

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the availability of the widest range of molecular probes and reagents to perform in vivo as well as in vitro studies. Based on our experience with various murine transplantation models, we developed a heterotopic pancreas transplantation model in mice with the intent to analyze mechanisms underlying severe ischemia reperfusion injury-associated early graft damage. In contrast to previously described techniques using suture techniques, herein we describe a new procedure using a non-suture cuff technique. 
In recent years, we have performed more than 300 pancreas transplantations in mice with an overall success rate of &gt;90%, a success rate never described before in mouse pancreas transplantation. The backbone of this non-suture cuff technique for graft revascularization consists of two major steps: (I) pulling the recipient vessel over a polyethylene/polyamide cuff and fixing it with a circumferential ligature, and (II) placing the donor vessel over the everted recipient vessel and fixing it with a second circumferential ligature. The resultant continuity of the endothelial layer results in less thrombogenic lesions with high patency rates and, finally, high success rates.
In this model, arterial anastomosis is achieved by pulling the abdominal aorta of the donor graft over the everted common carotid artery of the recipient animal. Venous drainage of the graft is achieved by pulling the portal vein of the graft over the everted external jugular vein of the recipient. This manuscript provides details and crucial steps of the organ recovery and organ implantation procedures, which will allow researchers with microsurgical skills to perform the transplantation successfully in their laboratories.

Among abdominal solid organ transplantation, pancreatic grafts are prone to develop severe ischemia reperfusion injury-associated graft damage, leading eventually to early graft loss. This protocol describes a model of murine pancreas transplantation using a non-suture cuff technique, ideally suited for analyzing these early, deleterious damages.

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the ...

Patch Angioplasty in the Rat Aorta or Inferior Vena Cava

Pericardial patches are commonly used in vascular surgery to close vessels. To facilitate studies of the neointimal hyperplasia that forms on the patch, we developed a rat model of patch angioplasty that can be used in either a vein or an artery, creating a patch venoplasty or a patch arterioplasty, respectively. Technical aspects of this model are discussed. The infra-renal IVC or aorta are dissected and then clamped proximally and distally. A 3 mm venotomy or arteriotomy is performed in the infrarenal inferior vena cava or aorta of 6 to 8 week-old Wistar rats. A bovine pericardial patch (3 mm x 1.5 mm x 0.6 mm) is then used to close the site using a 10-0 nylon suture. Compared to arterial patches, venous patches show increased neointimal thickness on postoperative day 7. This novel model of pericardial patch angioplasty can be used to examine neointimal hyperplasia on vascular biomaterials, as well as to compare the differences between the arterial and venous environments.

We have established a model of pericardial patch angioplasty that can be used in either small-diameter veins or arteries. This model can be used to compare venous and arterial neointimal hyperplasia formation.

Pericardial patches are commonly used in vascular surgery to close vessels. To facilitate studies of the neointimal hyperplasia that forms on the patch, ...

Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

Kidney transplantation offers increased survival rates and improved quality of life for patients with end-stage renal disease, as compared to any type of renal replacement therapy. Over the past few decades, the rat kidney transplantation model has been used to study the immunological phenomena of rejection and tolerance. This model has become an indispensable tool to test new immunomodulatory pharmaceuticals and regimens prior to proceeding with expensive preclinical large animal studies.
This protocol provides a detailed overview of how to reliably perform orthotopic kidney transplantation in rats. This protocol includes three distinctive steps that increase the probability of success: perfusion of the donor kidney by flushing through the portal vein and the use of a cuff system to anastomose the renal veins and ureters, thereby decreasing cold and warm ischemia times. Using this technique, we have achieved survival rates beyond 6 months with normal serum creatinine in animals with syngeneic or tolerant kidney transplants. Depending on the aim of the study, this model can be modified by pre- or posttransplant treatments to study the acute, chronic, cellular, or antibody-mediated rejection. It is a reproducible, reliable, and cost-effective animal model to study different aspects of kidney transplantation.

The purpose of this manuscript and protocol is to explain and demonstrate in detail the surgical procedure of orthotopic kidney transplantation in rats. This method is simplified to achieve the correct perfusion of the donor kidney and shorten the reperfusion time by using the venous and ureteral cuff anastomosis technique.

Kidney transplantation offers increased survival rates and improved quality of life for patients with end-stage renal disease, as compared to any type of ...