This work was funded by a generous gift from the Bombeck Family Estate.

Lewis (RT11) and Dark Agouti (DA) (RT1Aa) rats were purchased from commercial vendors (see the Table of Materials). These fully MHC-mismatched strains are often used to study acute renal allograft rejection. All animals were housed and maintained according to the National Institutes of Health&#8217;s (NIH) guidelines in a specific pathogen-free facility at the Johns Hopkins University. All procedures were approved by the institutional animal care and use committee.
1. Donor procedure
<ol>
	<li>Prepare and autoclave all surgical instruments to be used in this procedure as a means of sterilization and use disposable sterile gloves to prevent infectious complications.</li>
	<li>Anesthetize the donor rat by isoflurane inhalation (induction at 3%&#8211;4% and maintenance at 1%&#8211;2%) for the rest of the procedure. Give all donor and recipient animals preemptive buprenorphine subcutaneously at 0.1 mg/kg body weight for analgesia.</li>
	<li>Now, place the rat in a supine position and immobilize the limbs with sterile masking tape.</li>
	<li>Use a mechanical clipper to remove hair from the abdominal area.
	<ol>
		<li>Apply an eye lubricant and use sterile gauze soaked in povidone-iodine, followed by gauze soaked in isopropyl alcohol, to sterilize the surgical field.</li>
		<li>Prior to the first incision, make sure that the rat is adequately anesthetized by checking the absence of the toe pinch withdrawal reflex.</li>
	</ol>
	</li>
	<li>Using scissors, start off by making a large longitudinal midline skin and muscle incision from the symphysis pubis to the xiphoid, and enter the peritoneal cavity.</li>
	<li>Insert two retractors on either side of the abdominal wall in order to expose the intra-abdominal cavity.</li>
	<li>Cover the intestine with a moist sterile gauze and shift it to the right lateral side of the abdomen, exposing the aorta, vena cava, and left kidney. Apply 1 mL of preheated saline with a 1 cc syringe to keep the intestines and the abdominal organs moist and at a normal temperature.
	<ol>
		<li>Apply a second moist gauze to cover and mobilize the stomach and spleen cranial to the kidney and a small moist gauze to cover the exposed kidney (Figure 1A).</li>
	</ol>
	</li>
	<li>Use microsurgical dissecting forceps to isolate and mobilize the left renal artery and vein from the connective tissue and each other. Isolate the left renal vein by cauterizing the left gonadal vein and isolate the left renal artery by cauterizing the adrenal artery. After that, mobilize the aorta and vena cava superior and inferior of the left renal pedicle by dissecting the connective tissue with dissecting forceps (Figure 1B).</li>
	<li>Divide and mobilize the ureter from the connective tissue using dissecting forceps, and make a diagonal incision at a length of 2 cm measured from the renal pelvis, using microscissors. Insert a polyamide cuff (see Table of Materials) halfway into the ureter and secure the cuff by placing a knot with 8-0 silk suture (Figure 1C). 
	&#8203;NOTE: It is important not to remove all fat and connective tissue from the ureter, as they provide protection against obstruction caused by adhesions, and their removal may cause ureteral necrosis. Pay extra attention to preserving the vessel supplying oxygen to the ureter.</li>
	<li>Mobilize the left kidney by separating it from the perinephric fat using dissecting forceps or microscissors. Leave the adipose capsule of the kidney attached and use that site for handling the kidney.
	<ol>
		<li>Expose the inferior vena cava.</li>
	</ol>
	</li>
	<li>Administer 200 units of heparin using a syringe with a 27 G needle through the penile vein. Pressure the site of injection with a cotton swab for at least 1 min to prevent bleeding.</li>
	<li>Identify the portal vein (pv) and inferior vena cava (ivc) (Figure 1D). Flush the kidney by injecting 50 mL of cold saline mixed with 500 units of heparin into the portal vein using a 16 G needle (Figure 1E). Before flushing, cut the inferior vena cava at the infrahepatic level and the portal vein caudal at the needle insertion site to allow the blood to exit the circulation. Start flushing the kidney by gradually infusing the saline solution. Observe a change of color of the kidney from dark red to a uniform grey and pale color (Figure 1F).</li>
	<li>After flushing, ligate the renal artery and vein proximal to the aorta and vena cava and place the flushed kidney in a Petri dish in cold saline on ice. Figure 2A represents the schematic overview of the donor procedure.</li>
	<li>Once the kidney is in cold saline, fix and immobilize the handle of the vein cuff (see the Table of Materials) and gently pull the renal vein through the cuff. Then, fix the renal vein over the cuff by placing three knots using eight-zero silk suture (Figure 2B).&#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; &#160; 
	NOTE: Pay special attention to the orientation of the vein while securing it in place. Rotated veins cause an obstruction of the blood flow and lead to thrombosis.</li>
</ol>
2. Recipient procedure
<ol>
	<li>Repeat steps 1.1&#8211;1.11 from the donor procedure.</li>
	<li>Place two atraumatic micro-vessel clamps on the left renal artery and vein proximal to the aorta and vena cava (Figure 3A).</li>
	<li>Ligate the recipient renal vein proximal to the inlet of the kidney. Flush the renal vein with heparinized saline to remove all the remaining blood out of the vessel.</li>
	<li>Slide the ligated renal vein over the cuffed renal vein previously positioned in the donor kidney and secure it with an 8-0 silk suture (Figure 3B). Maintain the same positional orientation when securing the renal vein over the cuff.</li>
	<li>Ligate the ureter at the level of the lower pole of the left kidney. Mobilize the kidney from the perinephric fat.</li>
	<li>Ligate the renal artery proximal to the inlet of the recipient kidney. Flush it with heparinized saline to remove any excess blood in the vessel. Perform an end-to-end anastomosis of the renal artery with 8 to 10 interrupted sutures using a 10-0 nylon suture (Figure 3C). Maneuver the artery by using the adventitial layer.</li>
	<li>Remove the vessel clamps to reinitiate the reperfusion of the kidney. Start by removing the clamp on the vein followed by the clamp on the artery (Figure 3D). Use a sterile cotton swab to lightly pressure any oozing areas around the anastomosis region. A few minutes should be sufficient to achieve a patent anastomosis.</li>
	<li>Briefly observe the kidney to assess for adequate perfusion. Immediately after reperfusion, the kidney should change color and gradually regain its natural dark red color after a few minutes (Figure 3E). Visible peristalsis of the ureter and on-site urine production are sometimes observed.</li>
	<li>Finish by inserting the exposed tip of the ureteral cuff into the recipient ureter and secure the recipient ureter with an 8-0 silk suture&#160;(Figure 2C and Figure 3F).</li>
	<li>In order to keep the donor and recipient ureters in position, tie off the ends of each ureter side to each other.</li>
	<li>Optionally, the right kidney can be nephrectomized by tying off the right renal artery and vein with a 4-0 silk suture and removing the kidney.</li>
	<li>Remove all gauzes from the abdominal cavity, return all the organs to their natural position, squirt 1 mL of saline over the intestines to keep them moist, and close the abdomen by using a 4-0 absorbable suture on the rectus muscle and a 4-0 silk suture to close the skin layer in an interrupted fashion.</li>
</ol>
3. Postoperative care
<ol>
	<li>Place the animal in a clean cage with access to ad libitum water and food and allow for recovery on a 37 &#176;C heating pad.</li>
	<li>Inject 0.1 mg/kg buprenorphine subcutaneously for analgesia and monitor the animal for recovery.&#160;Additional analgesics administration may be needed in the next few days, depending on signs of discomfort or pain.&#160;Antibiotics are not routinely administered, as infectious complications are rare.</li>
	<li>Observe the recovery for 1&#8211;2 h before returning the animal back to the animal facility. Inspect the animal 2x&#8211;3x a day for the first 24 h, followed by a daily inspection. Pay attention to signs of pain and distress, oral intake, and urinary output.</li>
	<li>Remove the stitches 7&#8211;10 days after the operation.</li>
</ol>

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The authors have nothing to disclose.

In this manuscript, we describe the surgical method for orthotopic KT in rats in detail, including all the necessary equipment needed to perform this procedure (Figure 5). In 1965, Fisher and Lee published the first report on KT in rats, which became the start of an exciting investigative field18. Since then, many modifications have been introduced to improve the reproducibility of this model. It has served as an effective animal model for studying ischemia-reperfusio...

Historically, the first transplant rejection studies were performed by Brent and Medawar using skin transplants in rodents1. It soon became clear that skin has distinct immunological features, making it a highly immunogenic organ that is different in rejection from other vascularized solid organs2. Rat studies of solid organ transplant rejection are habitually limited to heart, liver, and kidney transplants. Although each of these organs is suitable to study rejection, there are advantages and disadvantages to each of them. Heart transplants are often transplanted into the abdomen and anastomosed to the aorta and vena cava, with the recipient&#8217;s native heart in place3. This does not recreate human clinical, anatomical, and physiological conditions. Additionally, hearts are very sensitive to cold ischemia and have to be reperfused preferentially within 1 h in order to be able to recover their function4. Liver transplants are generally considered to be surgically more challenging and time-sensitive to perform. After removing the native liver, the donor liver has to be implanted and reperfused within 30 min as the recipients cannot last longer without a functioning liver5. The hepatic artery, portal vein, and especially the bile duct reconstruction requires refined surgical skills. Besides the surgical challenges, the liver is known to possess tolerogenic properties and rodents and humans can become operationally tolerant6,7,8. The kidney, unlike the aforementioned organs, can be transplanted in an orthotopic fashion, is known to be an immunogenic organ with consistent, reproducible rejection episodes (if not immunosuppressed), and allows for prolonged cold ischemia times of several hours. This makes the rat kidney transplant an ideal model for studying allograft rejection and tolerance.
Kidney transplantation (KT) is the preferred choice of treatment for patients with end-stage renal disease. Over the last few decades, short-term survival outcomes after KT have improved dramatically, but long-term survival outcomes are stagnant9. Conventional immunosuppressive regimens remain the standard anti-rejection therapy. However, the chronic use of immunosuppressive therapies causes significant morbidity and mortality, such as nephrotoxicity, diabetes, and secondary malignancies10,11,12. In the long-term, chronic antibody- and cellular-mediated rejection threaten graft survival, with limited therapeutic options available.
A major goal in transplantation is the induction of transplant tolerance in order to obviate the need for chronic immunosuppression. The rat KT model is a robust tool to investigate the immunological rejection process and to evaluate new approaches to immunomodulation and transplant tolerance. The rat also serves as a suitable model to study acute and chronic cell- and antibody-mediated rejection13,14,15,16,17. This surgical model has proven to be a reliable, reproducible, and cost-effective tool to study various aspects of allograft rejection and tolerance. It is often used to test novel tolerance-inducing protocols prior to undertaking expensive and cumbersome large-animal studies. Performing KT in rats requires extensive surgical training and expertise to reach survival rates of &#62;90%. In this manuscript and in the accompanying instructional video, we provide a step-by-step outline for orthotopic KT in the rat, as successfully performed for many years at our institution.
Prior to starting any procedure, donor and recipient selection is critical and depends on the nature of the experiment. Ideally, donors and recipients should weigh between 220&#8211;260 g and be between 8&#8211;12 weeks of age. Animals under 220 g have small-diameter arteries, veins, and ureters, making the anastomosis in the recipient particularly challenging. Minor blood loss can cause hypovolemia and lead to death in smaller animals. Animals heavier than 260 g display more fat around their vessels, and vessel isolation will require more operative time and increase the cold ischemia time.

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			<td height="42" style="height:42px;width:216px;">Buprenorphine HCL</td>
			<td style="width:167px;">Reckitt Benckiser Healthcare UK</td>
			<td style="width:167px;">NDC12496-0757-5</td>
			<td style="width:463px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Dissecting forceps, curved</td>
			<td style="width:167px;">Zhenbang, China</td>
			<td style="width:167px;"></td>
			<td style="width:463px;">11cm Flat handle&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Heparin sodium injection USP</td>
			<td style="width:167px;">Sagent Pharmaceuticals&#160;</td>
			<td style="width:167px;">NDC25021-400-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micro-forceps, straight, smooth</td>
			<td style="width:167px;">Jingzhong, China</td>
			<td style="width:167px;">WA3010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micro needle holder</td>
			<td style="width:167px;">Jingzhong, China</td>
			<td style="width:167px;">WA2010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micro vessel clamps</td>
			<td style="width:167px;">Jingzhong, China</td>
			<td style="width:167px;">WA40120</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micro spring sciccor 1</td>
			<td style="width:167px;">ROBOZ</td>
			<td style="width:167px;">RS-5620</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micro spring sciccor 2</td>
			<td style="width:167px;">F.S.T.</td>
			<td style="width:167px;">91501-09</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micro spring sciccor 3</td>
			<td style="width:167px;">Zhenbang, China</td>
			<td style="width:167px;"></td>
			<td style="width:463px;">8.5cm Vannas,curved</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Prograf (Tacrolimus/FK506)</td>
			<td style="width:167px;">Astellas</td>
			<td style="width:167px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Rats</td>
			<td style="width:167px;">Charles River &#38; Taconic Biosciences&#160;</td>
			<td style="width:167px;">LEW/Crl &#38; DA-M&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Shaver</td>
			<td style="width:167px;">Wahl</td>
			<td>79600-2101</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Suture 4-0</td>
			<td style="width:167px;">Ethicon</td>
			<td style="width:167px;">J304H</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Suture, 4-0&#160;</td>
			<td style="width:167px;">Ethicon</td>
			<td style="width:167px;">683G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Suture, 10-0&#160;</td>
			<td style="width:167px;">Ethicon</td>
			<td style="width:167px;">2820G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Syringes &#38; Needles</td>
			<td style="width:167px;">BD</td>
			<td style="width:167px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Thread, 8-0</td>
			<td style="width:167px;">Ashaway</td>
			<td style="width:167px;">75290</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ureteral cuff</td>
			<td style="width:167px;">Microlumen</td>
			<td style="width:167px;">160-1</td>
			<td>Polymide Tubing, Diameter 0.41 mm&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Venous cuff</td>
			<td style="width:167px;">Intramedic BD</td>
			<td style="width:167px;">7441</td>
			<td>PE-200 Non-radiopaque polyethylene tubing ID: 1.4 mm, OD: 1.9 mm</td>
		</tr>
	</tbody>
</table>

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.

We performed syngeneic (N = 5) and allogeneic kidney transplants (N = 5). Animals with a syngeneic transplant achieved long-term survival without any immunosuppressive treatment. Animals that received an allogeneic transplant without immunosuppression rejected their graft and succumbed to renal failure with a median survival of 8 days (Figure 4A). Mean serum creatinine increased modestly in the syngeneic group while it increased by 14-fold in the allogeneic...

Watch this Scientific Journal Video about Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach at JoVE.com

Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

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 ...

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Research

JoVE Journal

Medicine

15.3K Views.  Johns Hopkins University School of Medicine. 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.

Video: Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

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 ...

A Novel Surgical Approach for Intratracheal Administration of Bioactive Agents in a Fetal Mouse Model

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and acquired diseases. Apart from congenital or inherited abnormalities with the requirement for long-term expression of the delivered gene, several non-inherited perinatal conditions, where short-term gene expression or pharmacological intervention is sufficient to achieve therapeutic effects, are considered as potential future indications for this kind of approach. Candidate diseases for the application of short-term prenatal therapy could be the transient neonatal deficiency of surfactant protein B causing neonatal respiratory distress syndrome1,2 or hyperoxic injuries of the neonatal lung3. Candidate diseases for permanent therapeutic correction are Cystic Fibrosis (CF)4, genetic variants of surfactant deficiencies5 and &alpha;1-antitrypsin deficiency6.
Generally, an important advantage of prenatal gene therapy is the ability to start therapeutic intervention early in development, at or even prior to clinical manifestations in the patient, thus preventing irreparable damage to the individual. In addition, fetal organs have an increased cell proliferation rate as compared to adult organs, which could allow a more efficient gene or stem cell transfer into the fetus. Furthermore, in utero gene delivery is performed when the individual's immune system is not completely mature. Therefore, transplantation of heterologous cells or supplementation of a non-functional or absent protein with a correct version should not cause immune sensitization to the cell, vector or transgene product, which has recently been proven to be the case with both cellular and genetic therapies7.
In the present study, we investigated the potential to directly target the fetal trachea in a mouse model. This procedure is in use in larger animal models such as rabbits and sheep8, and even in a clinical setting9, but has to date not been performed before in a mouse model. When studying the potential of fetal gene therapy for genetic diseases such as CF, the mouse model is very useful as a first proof-of-concept because of the wide availability of different transgenic mouse strains, the well documented embryogenesis and fetal development, less stringent ethical regulations, short gestation and the large litter size.
Different access routes have been described to target the fetal rodent lung, including intra-amniotic injection10-12, (ultrasound-guided) intrapulmonary injection13,14 and intravenous administration into the yolk sac vessels15,16 or umbilical vein17. Our novel surgical procedure enables researchers to inject the agent of choice directly into the fetal mouse trachea which allows for a more efficient delivery to the airways than existing techniques18.

We developed a novel surgical approach for intratracheal administration of bioactive agents into the mouse fetus. The delivery route is more efficient in targeting the fetal mouse lungs than the commonly used intra-amniotic injection. This procedure has to date not been described in a mouse model.

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and ...

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 ...

Cardiopulmonary Bypass in a Mouse Model: A Novel Approach

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization and for minimizing organ damage resulting from prolonged extracorporal circulation. The goal of this paper was to demonstrate a fully functional and clinically relevant model of cardiopulmonary bypass in a mouse. We report on the device design, perfusion circuit optimization, and microsurgical techniques. This model is an acute model, which is not compatible with survival due to the need for multiple blood drawings. Because of the range of tools available for mice (e.g., markers, knockouts, etc.), this model will facilitate investigation into the molecular mechanisms of organ damage and the effect of cardiopulmonary bypass in relation to other comorbidities.

This paper describes how to perform cardiopulmonary bypass in mice. This novel model will facilitate the investigation of the molecular mechanisms involved in organ damage.

As prolonged cardiopulmonary bypass becomes more essential during cardiac interventions, an increasing clinical demand arises for procedure optimization ...

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.

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 ...

Reduced Complications after Arterial Reconnection in a Rat Model of Orthotopic Liver Transplantation

The rat orthotopic liver transplantation (OLT) model is a powerful tool to study acute and chronic rejection. However, it is not a complete representation of human liver transplantation due to the absence of arterial reconnection. Described here is a modified transplantation procedure that includes the incorporation of hepatic artery (HA) reconnection, leading to a marked improvement in transplant outcomes. With a mean anhepatic time of 12 min and 14 s, HA reconnection results in improved perfusion of the transplanted liver and an increase in long-term recipient survival from 37.5% to 88.2%. This protocol includes the use of 3D-printed cuffs and holders to connect the portal vein and infrahepatic inferior vena cava. It can be implemented for studying multiple aspects of liver transplantation, from immune response and infection to technical aspects of the procedure. By incorporating a simple and practical method for arterial reconnection using a microvascular technique, this modified rat OLT protocol closely mimics aspects of human liver transplantation and will serve as a valuable and clinically relevant research model.

The goal of this study is to modify the rat orthotopic liver transplant model to better represent human liver transplantation and improve recipient survival. The presented method reestablishes hepatic arterial inflow by connecting the donor liver&#39;s common hepatic artery to the recipient liver&#39;s proper hepatic artery.

The rat orthotopic liver transplantation (OLT) model is a powerful tool to study acute and chronic rejection. However, it is not a complete representation ...

Orthotopic Kidney Auto-Transplantation in a Porcine Model Using 24 Hours Organ Preservation And Continuous Telemetry

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for kidney transplantation (KT). Even though, the use of allografts from extended criteria donors (ECD) could partially ease the shortage of organ donors, ECD organs carry a potentially higher risk for inferior outcomes and postoperative complications. Dynamic organ preservation techniques, modulation of ischemia-reperfusion and preservation injury, and allograft therapies are in the spotlight of scientific interest in an effort to improve allograft utilization and patient outcomes in KT.
Preclinical animal experiments are playing an essential role in translational research, especially in the medical device and drug development. The major advantage of the porcine orthotopic auto-transplantation model over ex vivo or small animal studies lies within the surgical-anatomical and physiological similarities to the clinical setting. This allows the investigation of new therapeutic methods and techniques and ensures a facilitated clinical translation of the findings. This protocol provides a comprehensive and problem-oriented description of the porcine orthotopic kidney auto-transplantation model, using a preservation time of 24 hours and telemetry monitoring. The combination of sophisticated surgical techniques with highly standardized and state-of-the-art methods of anesthesia, animal housing, perioperative follow up, and monitoring ensure the reproducibility and success of this model.

Large animal models play an essential role in preclinical transplantation research. Due to its similarities to the clinical setup, the porcine model of orthotopic kidney auto-transplantation described in this article provides an excellent in vivo setting for the testing of organ preservation techniques and therapeutic interventions.

In the present era of organ transplantation with critical organ shortage, various strategies are employed to expand the pool of available allografts for ...

Robot-Assisted Kidney Transplantation

This paper describes robot-assisted kidney transplantation (RAKT) from a living donor. The robot is docked between the parted legs of the patient, placed in the supine Trendelenburg position. Kidney allografts are provided by a living donor. Before vascular anastomosis, the kidney allograft is prepared by inserting a double-J stent in the ureter, and the temperature for the anastomosis is lowered by wrapping it in an ice-packed gauze. A 12 mm or 8 mm port for the robotic camera and three 8 mm ports for robotic arms are placed. A peritoneal pouch is created for the kidney allograft by raising the peritoneal flaps on both sides over the psoas muscle before dissecting the iliac vessels and bladder. A 6 cm Pfannenstiel incision is made to insert the kidney into the peritoneal pouch, lateral to the right iliac vessels. 
After clamping the external iliac vein with Bulldogs clamps, a venotomy is performed, and the graft renal vein is anastomosed to the external iliac vein in an end-to-side continuous manner with a 6/0 polytetrafluoroethylene suture. After clamping the graft renal vein, the iliac vein is declamped. This is followed by clamping of the external iliac artery, arteriotomy, arterial anastomosis with a 6/0 polytetrafluoroethylene suture, clamping of the graft renal artery, and declamping of the external iliac artery. Reperfusion is then carried out, and ureteroneocystostomy is performed using the Lich-Gregoir technique. The peritoneum is closed at a few locations with polymer locking clips, and a closed-suction drain is placed through one of the working ports. After deflating the pneumoperitoneum, all incisions are closed.

This paper provides technical details for robot-assisted kidney transplantation from a living donor.

This paper describes robot-assisted kidney transplantation (RAKT) from a living donor. The robot is docked between the parted legs of the patient, placed ...

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

Left Lung Orthotopic Transplantation in a Juvenile Porcine Model for ESLP

Lung transplantation is the gold-standard treatment for end-stage lung disease, with over 4,600 lung transplantations performed worldwide annually. However, lung transplantation is limited by a shortage of available donor organs. As such, there is high waitlist mortality. Ex situ lung perfusion (ESLP) has increased donor lung utilization rates in some centers by 15%-20%. ESLP has been applied as a method to assess and recondition marginal donor lungs and has demonstrated acceptable short- and long-term outcomes following transplantation of extended criteria donor (ECD) lungs. Large animal (in vivo) transplantation models are required to validate ongoing in vitro research findings. Anatomic and physiologic differences between humans and pigs pose significant technical and anesthetic challenges. An easily reproducible transplant model would permit the&#160;in vivo&#160;validation of current ESLP strategies and the preclinical evaluation of various interventions designed to improve donor lung function. This protocol describes a porcine model of orthotopic left lung allotransplantation. This includes anesthetic and surgical techniques, a customized surgical checklist, troubleshooting, modifications, and the benefits and limitations of the approach.

This protocol describes a juvenile porcine model of orthotopic left lung allotransplantation designed for use with ESLP research. Focus is made on anesthetic and surgical techniques, as well as critical steps and troubleshooting.

Lung transplantation is the gold-standard treatment for end-stage lung disease, with over 4,600 lung transplantations performed worldwide annually. ...