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-reperfusion injury and renal transplant rejection and tolerance, thanks to the availability of several inbred and outbred strains with partial and full MHC mismatch combinations19. The rat KT model can serve as a tool to test hypotheses prior to extending investigations to swine and nonhuman primate models of KT. Options to study kidney transplant rejection or tolerance in rodents are limited. The kidney transplant model in mice is technically very challenging and requires a long training period to achieve survival rates of &#62;80%20. Another limitation of the mouse model is the spontaneous renal allograft acceptance without the need for immunosuppression in about 30% of the recipients. However, other organ transplants in mice, such as skin and heart, are rejected within 10 days, suggesting that the rejection of kidney allografts in fully MHC-mismatched mice is weak and not representative of the clinical situation21. However, if the technical challenge can be overcome, mice models are preferred for mechanism studies of allograft rejection because of the availability of genetically modified knock-in or knock-out mice.
KT in rats can be performed in a number of ways. We will discuss a few advantages and disadvantages of these various methods. Irrespective of the preferred technique, it is always critical to reduce warm ischemia time and to avoid irreversible injury to the graft and recipient.
Right versus left kidney 
The abdominal anatomy of rats is very similar to that of humans. The left kidney is located superior compared to the right kidney because of the anatomical position of the liver. One of the advantages of using the left kidney is the length of the vessels. Generally, the left renal artery and vein are twice the length of the right renal vessels. This is especially beneficial when performing anastomosis where the length of vessels is not a limiting factor. However, reports exist of right-sided donor kidney retrieval and transplantation22,23. Approaches using both kidneys for transplantation have also been described24.
Flushing donor kidney via the portal vein 
One of the key steps of this procedure is the donor kidney perfusion. Perfusion is necessary to remove all donor blood from the vessels and kidney and to cool the organ down to slow biological deterioration. There are various methods described for perfusing the kidney. We have experimented with flushing the kidney in different ways and concluded that flushing the kidney through the portal vein offers advantages and consistently leads to complete perfusion of the kidney and vessels. The conventional approaches described in the literature entail flushing the donor kidney after ligating the renal artery and vein or retrograde through the infrarenal aorta24,25,26,27,28. These approaches may lead to endothelial damage and renal vasoconstriction because of increased local pressures or to incomplete perfusion due to low perfusion pressure29,30.
By flushing the kidney through the portal vein, the pressure is managed by the heart. During the perfusion, the heart is still active and pumps the perfusion fluid in a normal fashion to the aorta and kidney with pulsatile flow, preventing damage to capillaries and glomeruli due to shear pressure flow. When transplanting kidneys en bloc or using the right kidney for transplantation, this method is suitable to achieve uniform perfusion and to harvest both kidneys at the same time.
Arterial and venous anastomosis 
One of the most critical steps in the rat KT model is performing a reliable microvascular anastomosis in a time-efficient manner. The donor renal artery can be anastomosed to the recipient&#8217;s renal artery or the aorta. Anastomosing the donor vessels to the aorta and inferior vena cava causes ischemic injury to the recipient&#8217;s organs. In this protocol, we demonstrate the end-to-end anastomosis of the renal arteries, as it avoids ischemic injury to other organs. During the arterial anastomosis, it is important not to damage the endothelial surface of the lumen when handling the vessel. For the vein anastomosis, we use a cuff technique to reduce the warm ischemia time and shorten the operative procedure. This has proven to be a very reliable and durable method to ensure adequate venous flow. To ensure adequate venous flow, it is imperative for the veins not to be kinked or twisted when these are secured together. Alternatively, an end-to-end or an end-to-side vein anastomosis is possible, depending on the surgeon&#8217;s preference. Ideally, the arterial and venous vessel anastomosis should take between 20&#8211;30 min.
Pahlavan et al. summarized the complications of each type of technique based on a literature screening31. One of the main complications that may occur after any microsurgical vessel anastomosis is thrombosis. Ligation and adequate flushing of the recipient vessels significantly reduce thrombosis formation, and it is certainly not a complication frequently observed. Other complications are leakage or rupture of the anastomosis after reperfusion. This is related to inadequate microsurgical technique or inadequate handling of the vessels.
Ureteral anastomosis 
The ureter has to be handled with utmost care, especially during isolation of the ureter in the donor. Injury to the periureteric structures can cause ureteral ischemia leading to strictures and obstruction and, in the worst-case scenario, ureteral necrosis. The literature reports different methods for ureteral anastomosis. End-to-end, cuff-assisted end-to-end, bladder patch, and bladder insertion are the most commonly used19,32,33. In previous studies, we have used a cuff with oblique edges on both ends to facilitate the entry into the ureter on both ends. We did not observe any urine leakage or blood clot formations. However, long-term complications (&#62;30 days) of this technique include hydronephrosis and occasionally nephrolithiasis, which can be explained by stricture formation, dislocation, or obstruction of the cuff due to ureteral stones. This finding is consistent with other reports and our own findings of performing ureteral anastomosis with a cuff. Ureteral complications are often noticed postoperatively after significant injury to the kidney, are unsalvageable, and require the animal to be euthanized.
Postoperative care and survival 
The postoperative care of transplanted animals requires adequate pain management and detailed observations of the animals&#8217; overall activity, weight observations, and urine production. Common early postoperative complications include bleeding from the arterial or venous anastomosis, urine leakage, ureteral obstruction, or delayed graft function because of prolonged ischemia time. Animals with these complications show modest activity and usually remain in a hunched-back position with no urinary output and nutrient intake. Generally, it is favorable to administer up to 1&#8211;5 mL of saline to the animals postoperatively to speed up their recovery and prevent dehydration. Animals that receive no immunosuppression can survive between 7 to 10 days, which allows for a sufficient therapeutic window to test novel drugs or other methods. If animals are adequately immunosuppressed (1.0 mg/kg/day FK506 subcutaneously) or tolerant, they can be monitored long-term past 6 months as previously reported13. The rat kidney transplant model allowed the definition of the mechanisms of tolerance induced by using a unique approach of stem cell mobilization prior to confirming this phenomenon in large animals34. Rat KT has provided crucial information to investigators for decades, and it will continue to do so in the future.

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.

<table><tbody><tr><td>Buprenorphine HCL</td><td>Reckitt Benckiser Healthcare UK</td><td>NDC12496-0757-5</td><td></td></tr><tr><td>Dissecting forceps, curved</td><td>Zhenbang, China</td><td></td><td>11cm Flat handle&amp;nbsp;</td></tr><tr><td>Heparin sodium injection USP</td><td>Sagent Pharmaceuticals&amp;nbsp;</td><td>NDC25021-400-10</td><td></td></tr><tr><td>Micro-forceps, straight, smooth</td><td>Jingzhong, China</td><td>WA3010</td><td></td></tr><tr><td>Micro needle holder</td><td>Jingzhong, China</td><td>WA2010</td><td></td></tr><tr><td>Micro vessel clamps</td><td>Jingzhong, China</td><td>WA40120</td><td></td></tr><tr><td>Micro spring sciccor 1</td><td>ROBOZ</td><td>RS-5620</td><td></td></tr><tr><td>Micro spring sciccor 2</td><td>F.S.T.</td><td>91501-09</td><td></td></tr><tr><td>Micro spring sciccor 3</td><td>Zhenbang, China</td><td></td><td>8.5cm Vannas,curved</td></tr><tr><td>Prograf (Tacrolimus/FK506)</td><td>Astellas</td><td></td><td></td></tr><tr><td>Rats</td><td>Charles River &amp;amp; Taconic Biosciences&amp;nbsp;</td><td>LEW/Crl &amp;amp; DA-M&amp;nbsp;</td><td></td></tr><tr><td>Shaver</td><td>Wahl</td><td>79600-2101</td><td></td></tr><tr><td>Suture 4-0</td><td>Ethicon</td><td>J304H</td><td></td></tr><tr><td>Suture, 4-0&amp;nbsp;</td><td>Ethicon</td><td>683G</td><td></td></tr><tr><td>Suture, 10-0&amp;nbsp;</td><td>Ethicon</td><td>2820G</td><td></td></tr><tr><td>Syringes &amp;amp; Needles</td><td>BD</td><td></td><td></td></tr><tr><td>Thread, 8-0</td><td>Ashaway</td><td>75290</td><td></td></tr><tr><td>Ureteral cuff</td><td>Microlumen</td><td>160-1</td><td>Polymide Tubing, Diameter 0.41 mm&amp;nbsp;</td></tr><tr><td>Venous cuff</td><td>Intramedic BD</td><td>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 group (0.5 mg/dL versus 7.0 mg/dL, p &lt;0.01) (Figure 4B). Upon explantation, the macroscopic view of the syngeneic kidney allograft did not show any abnormalities. The kidney color and internal structures remained intact. In contrast, kidney allografts of rejected animals presented red hemorrhagic patches with the destruction of the internal structures (Figure 4C). Hematoxylin and eosin stains of syngeneic grafts showed thin glomerular capillary loops with normal numbers of endothelial and mesangial cells. Rejected allografts displayed destroyed glomerular structures with signs of inflammation and tubulitis (Figure 4D). To confirm T-cell-mediated rejection, we performed CD8+ staining. While syngeneic allografts showed very few positive CD8+ T cells, the rejected allografts showed a significantly higher number of CD8+ cells in and around the glomeruli and tubuli (Figure 4E), confirming T-cell-mediated rejection.
<img alt="Histological liver biopsy process; sequence of liver tissue extraction; microscopic analysis setup." class="xfigimg" src="/files/ftp_upload/59403/59403fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Donor nephrectomy. (A) Upon opening the abdomen, the left kidney is isolated with moist gauzes. (B) The left renal artery and vein are isolated and mobilized from the surrounding fat. (C) The ureter is ligated, cuffed, and secured with a single silk suture. (D) The portal vein (pv) and inferior vena cava (ivc) are identified and the kidney is perfused through the portal vein. (E) Perfusion is successfully executed as the right kidney and liver are becoming pale by flushing the animal’s portal vein. (F) Successful perfusion demonstrates a pale kidney and vessels ready for transplantation. <a href="https://www.jove.com/files/ftp_upload/59403/59403fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Kidney biopsy method diagram; procedural steps A-C showing needle insertion and positioning." class="xfigimg" src="/files/ftp_upload/59403/59403fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Schematic overview of the kidney transplant procedure. (A) Schematic overview of the donor procedure. (B) Schematic overview of a cuffed donor vein. (C) Schematic overview of anastomosis of the recipient and the cuffed donor vein and anastomosis of the ureters. <a href="https://www.jove.com/files/ftp_upload/59403/59403fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Liver transplant procedure steps; surgical clamps in microvascular anastomosis; medical study." class="xfigimg" src="/files/ftp_upload/59403/59403fig3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Kidney transplantation in the recipient. (A) The recipient’s artery and vein are mobilized from the surrounding fat and clamped following separation. (B) The donor kidney is introduced, and the veins are connected via the cuff technique and secured with an 8-0 suture. (C) The arteries are sutured in an end-to-end fashion. (D) The clamps are removed. (E) The kidney is reperfused and recovers its natural color without any bleeding. (F) Finally, the ureters are anastomosed by using the previously placed cuff and secured with an 8-0 suture. <a href="https://www.jove.com/files/ftp_upload/59403/59403fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Kidney transplant outcomes; survival graphs, creatinine levels, kidney photos, histology analysis." class="xfigimg" src="/files/ftp_upload/59403/59403fig4.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Kidney transplant survival. (A) The Kaplan-Meyer figure demonstrates the survival of rats with syngeneic or allogeneic kidney transplants over time. (B) Measurement and comparison of serum creatinine in rats with syngeneic or allogeneic kidney transplants compared to nontransplanted animals. (C) Macroscopic overview of explanted kidneys of syngeneic (top) and allogeneic (bottom) kidney transplant at day 8. Animals were perfused with saline prior to explanting. The last two panels show a microscopic overview of (D) hematoxylin and eosin staining and (E) CD8+ of syngeneic (top) and allogeneic (bottom) kidney explants. The images are taken under 200x magnification. *Results were considered statistically significant if p &lt; 0.05. <a href="https://www.jove.com/files/ftp_upload/59403/59403fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Surgical instruments and supplies setup for medical procedures, including syringes and forceps." class="xfigimg" src="/files/ftp_upload/59403/59403fig5.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Required surgical instruments. (1) Straight scissors. (2) Fine scissors. (3) Micro-spring scissors 1 (ureter ligation). (4) Micro-spring scissors 2. (5) Micro-spring scissors 3. (6) Small-animal surgical retractors. (7) Forceps. (8) Microforceps, straight, smooth. (9) Dissecting forceps, curved. (10) Micro-needle holder. (11) Needle holder. (12) 8-0 braided silk suture without needle. (13) 4-0 silk suture. (14) Micro-vessel clamps (one pair). (15) Micro-vessel clamp applier. (16) Fine-tip clamp. (17) Heparin. (18) Vessel clamp (medium size). (19) Vessel clamp (large). (20) Sterile cotton swabs. (21) 10-0 micro-suture with needle. (22) Sterile gauze. (23) Heparinized saline flush syringe. (24) 60 cc syringe with needle. (25) 10 cc syringe. (26) 1 cc syringe. (27) 25 G 5/8 inch needles. (28) 19 G needles. (29) Trimmer. (30) Bipolar cautery system. (31) Tape. (32) Petri dish with 0.9% normal saline. (33) 60 cc syringe with 50 cc heparinized saline for perfusion. (34) 10 cc syringe with 5 cc heparinized saline flush. (35) Ureter cuff. (36) Vein cuff. <a href="https://www.jove.com/files/ftp_upload/59403/59403fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

orthotopic-rat-kidney-transplantation-novel-simplified-surgical

Nanyang Technological University

Research

JoVE Journal

Medicine

15.6K 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 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 ...

Non-invasive Imaging of Acute Allograft Rejection after Rat Renal Transplantation Using 18F-FDG PET

The number of patients with end-stage renal disease, and the number of kidney allograft recipients continuously increases. Episodes of acute cellular allograft rejection (AR) are a negative prognostic factor for long-term allograft survival, and its timely diagnosis is crucial for allograft function 1. At present, AR can only be definitely diagnosed by core-needle biopsy, which, as an invasive method, bares significant risk of graft injury or even loss. Moreover, biopsies are not feasible in patients taking anticoagulant drugs and the limited sampling site of this technique may result in false negative results if the AR is focal or patchy. As a consequence, this gave rise to an ongoing search for new AR detection methods, which often has to be done in animals including the use of various transplantation models.
Since the early 60s rat renal transplantation is a well-established experimental method for the examination and analysis of AR 2. We herein present in addition small animal positron emission tomography (PET) using 18F-fluorodeoxyglucose (FDG) to assess AR in an allogeneic uninephrectomized rat renal transplantation model and propose graft FDG-PET imaging as a new option for a non-invasive, specific and early diagnosis of AR also for the human situation 3. Further, this method can be applied for follow-up to improve monitoring of transplant rejection 4.

Non-invasive Imaging of Acute Allograft Rejection after Rat Renal Transplantation Using 18F-FDG PET

We herein present a rat renal transplantation model to non-invasively assess acute allograft rejection using positron emission tomography with 18F-fluorodeoxyglucose.

The number of patients with end-stage renal  disease, and the number of kidney allograft recipients continuously increases.  Episodes of acute cellular ...

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 ...

Murine Renal Transplantation Procedure

Renal orthotopic transplantation in mice is a technically challenging procedure. Although the first kidney transplants in mice were performed by Russell et al over 30 years ago (1) and refined by Zhang et al years later (2), few people in the world have mastered this procedure. In our laboratory we have successfully performed 1200 orthotopic kidney transplantations with &gt; 90% survival rate. The key points for success include stringent control of reperfusion injury, bleeding and thrombosis, both during the procedure and post-transplantation, and use of 10-0 instead of 11-0 suture for anastomoses.
Post-operative care and treatment of the recipient is extremely important to transplant success and evaluation. All renal graft recipients receive antibiotics in the form of an injection of penicillin immediately post-transplant and sulfatrim in the drinking water continually. Overall animal health is evaluated daily and whole blood creatinine analyses are performed routinely with a portable I-STAT machine to assess graft function.

Renal transplantation in mice is a technically challenging procedure that requires careful post-operative care and treatment for success.

Renal orthotopic transplantation in mice is a technically challenging procedure. Although the first kidney transplants in mice were performed by Russell ...

Biology

Mouse Kidney Transplantation: Models of Allograft Rejection

Rejection of the transplanted kidney in humans is still a major cause of morbidity and mortality. The mouse model of renal transplantation closely replicates both the technical and pathological processes that occur in human renal transplantation. Although mouse models of allogeneic rejection in organs other than the kidney exist, and are more technically feasible, there is evidence that different organs elicit disparate rejection modes and dynamics, for instance the time course of rejection in cardiac and renal allograft differs significantly in certain strain combinations. This model is an attractive tool for many reasons despite its technical challenges. As inbred mouse strain haplotypes are well characterized it is possible to choose donor and recipient combinations to model acute allograft rejection by transplanting across MHC class I and II loci. Conversely by transplanting between strains with similar haplotypes a chronic process can be elicited were the allograft kidney develops interstitial fibrosis and tubular atrophy. We have modified the surgical technique to reduce operating time and improve ease of surgery, however a learning curve still needs to be overcome in order to faithfully replicate the model. This study will provide key points in the surgical procedure and aid the process of establishing this technique.

Here, we present a protocol to study the immunology of rejection. The surgical model presented reports a short operating time and a concise technique. Depending on the donor-recipient strain combination, the transplanted kidney may develop acute cellular rejection or chronic allograft damage, defined by interstitial fibrosis and tubular atrophy.

Rejection of the transplanted kidney in humans is still a major cause of morbidity and mortality. The mouse model of renal transplantation closely ...

A Modified Surgical Technique for Kidney Transplantation in Mice

Kidney transplantation in mice is a complicated and challenging surgery procedure. There are very few publications demonstrating the key steps of this operation. Therefore, this article introduces the technique and points out the surgical caveats associated with this operation. In addition, important modifications in comparison to the conventional procedure are demonstrated. Firstly, a patch of the abdominal aorta is cut and prepared so that the proximal bifurcations of the renal artery, including the ureteral artery are transected together with the donor kidney en bloc. This reduces the risk of a ureter necrosis and avoids the development of a urinary tract occlusion. Secondly, a new method of the vascular anastomosis is demonstrated that allows the operator to flexibly increase or decrease the size of the anastomosis after renal transplant reperfusion has already been initiated. This avoids the development of vessel strictures and intraabdominal bleeding. Thirdly, a technique that enables the anastomosis of the delicate donor ureter and the recipient bladder that does not cause a trauma is shown. Adopting this protocol can shorten the operation time and reduces the damage to the recipient's bladder, thereby significantly increasing the operation success rate for the recipient mice.

This protocol presents a new surgical technique of mouse kidney transplantation focusing on a modified arterial anastomosis strategy. A vascular suture technique including a simple and safer ureter-bladder anastomosis method is also presented. These modifications shorten the operation time and improve the success rate of the mouse kidney transplantation procedure.

Kidney transplantation in mice is a complicated and challenging surgery procedure. There are very few publications demonstrating the key steps of this ...

Immunology and Infection

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 ...

A Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential model to investigate the mechanism of renal allograft rejection in pre-clinical studies and could aid in the development of novel approaches to improve the long-term survival of renal allografts. Donor kidney implantation in rat orthotopic renal transplantation is commonly performed by end-to-side anastomosis to recipients' aorta and inferior vena cava. In this model, the donor's kidney was implanted using end-to-end anastomosis to the recipients' renal artery and renal vein. The donor's ureter was anastomosed to the recipient's bladder in an end-to-side 'tunnel' method. This model contributes to better healing of ureter-bladder anastomosis and increases the recipients' survival by avoiding interference with blood supply and venous reflux of the lower body. This model can be used to investigate the mechanisms of acute and chronic immune and pathologic rejection of renal allografts. Here, the study describes the detailed protocols of this orthotopic renal transplantation between rats.

The rat orthotopic renal transplantation model contributes to investigating the mechanism of renal allograft rejection. The current model increases the recipients' survival without interference with blood supply and venous reflux of the lower body using an end-to-end anastomosis of kidney implantation and an end-to-side "tunnel" method of ureter-bladder anastomosis.

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential ...

Liver Transplantation in Rats and Analyzing Allograft Patency

Heterotopic Auxiliary Whole Liver Rat Transplant Model Utilizing a Hepaticoureterostomy for Allograft Rejection Studies

Small animal transplant models are indispensable for organ tolerance studies investigating feasible therapeutic interventions in preclinical studies. Rat liver transplantation (LTx) protocols typically use an orthotopic model where the recipients&#39; native liver is removed and replaced with a donor liver. This technically demanding surgical procedure requires advanced micro-surgical skills and is further complicated by lengthy anhepatic and lower body ischemia times. This prompted the development of a less complicated heterotopic method that can be performed faster with no anhepatic or lower body ischemia time, reducing post-surgery stress for the recipient animal.
This heterotopic LTx protocol includes two main steps: excising the liver from the donor rat and transplanting the whole liver into the recipient rat. During the excision of the donor liver, the surgeon ligates the supra-hepatic vena cava (SHVC) and hepatic artery (HA). On the recipient side, the surgeon removes the left kidney and positions the donor liver with the portal vein (PV), infra-hepatic vena cava (IHVC), and bile duct facing the renal vessels. Further, the surgeon anastomoses the recipient&#39;s renal vein end to end with the IHVC of the liver and arterializes the PV with the renal artery using a stent. A hepaticoureterostomy is utilized for biliary drainage by anastomosing the bile duct to the recipient&#39;s ureter, permitting the discharge of bile via the bladder.
The average duration of the transplantation was 130 min, cold ischemia duration was around 35 min, and warm ischemia duration was less than 25 min. Hematoxylin and eosin histology of the auxiliary liver from syngeneic transplants showed normal hepatocyte structure with no significant parenchymal alterations 30 days post-transplant. In contrast, 8-day post-transplant allogeneic graft specimens demonstrated extensive lymphocytic infiltration with a Banff Schema rejection activity index score of 9. Therefore, this LTx method facilitates a low morbidity rejection model alternative to orthotopic LTx.

The rat heterotopic auxiliary liver transplant protocol described here offers a practical investigational tool for exploring mechanisms of hepatic allograft rejection. This model helps to alleviate the surgical hurdles and animal stress of orthotopic liver transplantation in rats.

Small animal transplant models are indispensable for organ tolerance studies investigating feasible therapeutic interventions in preclinical studies. Rat ...