This work was supported in part by 1R03DK096151.&#160;We acknowledge the UAB-UCSD O&#8217;Brien Center (NIH P30 DK079337) for providing initial technical assistance with setting up this model.

All mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and were housed under pathogen free conditions at the University of Colorado Denver, Barbara Davis Center Animal Facility according to NIH Guidelines and with approval of the University of Colorado Denver IACUC.
1. Donor Kidney Harvest
<ol>
	<li>Sterilize all instruments, wear sterile gloves throughout the procedure and maintain a sterile field. Perform all surgeries with the use of an operating microscope.</li>
	<li>Surgically remove mouse donor kidneys from pentobarbital-anesthetized (60mg/kg IP) donors. Ascertain depth of anesthesia by toe-pinch, and observe the respiratory rate.</li>
	<li>Clip the fur then immobilize the mouse by 4-way restraints. Prepare the skin with povidone-iodine and drape the mouse in a sterile fashion.</li>
	<li>Make a 2 cm midline vertical abdominal incision and enter the abdominal cavity. Retract the bowel superiorly and externalize it on to the chest. Keep the bowel wrapped in sterile moist gauze throughout the procedure.</li>
	<li>Identify the great abdominal vessels and mobilize them, identify any lumbar branches and either cauterize or ligate them with 10/0 nylon suture.</li>
	<li>Immediately distal to the left renal vessels tease the inferior vena cava (IVC) and abdominal aorta (AA) apart by blunt dissection over approximately 2mm. Ligate with 10/0 nylon and divide small arterial and venous branches from the renal vessels.</li>
	<li>Now separate the renal vein from the renal artery carefully by blunt dissection of these structures. This allows for the accurate creation of a Carrel patch at the proximal end of the renal vein, which will be used to form an end-to-side anastomosis to the recipients IVC.</li>
	<li>Identify, ligate and divide the left adrenal vessels. This permits access to the suprarenal aorta and a 6-0 silk suture is placed around the aorta in preparation for ligation, but not tied at this time.</li>
	<li>Mobilize the kidney, vessels and ureter from the surrounding facia. Rotate the kidney to the right and ligate/divide any posterior branches. Then return the kidney to the left.</li>
	<li>Now direct attention to the ureter. Without disturbing the renal hilum free the ureter from the surrounding facia taking care to preserve the ureteric vessels. Divide the ureter at the level of the ductus deferens. The kidney is now ready for recovering.</li>
	<li>Slowly inject 300 units of heparin into the distal IVC thereby heparinizing the donor. Tie down the 6-0 silk suture placed around the suprarenal AA and perfuse the left kidney via the distal abdominal aorta with 0.8ml of heparin saline solution (100 U/ml).</li>
	<li>Once the perfusion has ceased create a venous Carrel patch immediately to eliminate backflow into the kidney. Retract the renal vein toward the kidney revealing the AA and renal artery beneath.</li>
	<li>Divide the aorta adjacent to the renal artery creating a heel and toe cuff as shown in Figure&#160;1. Remove the kidney, vessels and ureter from the donor.</li>
	<li>Remove the kidney directly to a pre-prepared recipient (0 min&#160;cold ischemia time) or stored at 4&#160;oC in a solution of choice for a pre-determined time until time of implantation. Euthanized the donors by exsanguanation and cervical dislocation. Total time to recover donor kidney is approx. 15-20 min.</li>
</ol>
2. Kidney Implant Technique
<ol>
	<li>Anesthetize the recipient mouse with pentobarbital (60mg/kg IP initial dose, 25mg/kg IP supplemental dose if required). Ascertain depth of anesthesia by toe-pinch, and observe the respiratory rate. Clip the fur then immobilize the mouse by 4-way restraints and apply ophthalmic ointment to the eyes. Prepare the skin with povidone-iodine and drape the mouse in a sterile fashion.</li>
	<li>Make a 2 cm midline vertical abdominal incision and enter the abdominal cavity. Retract the bowel superiorly and externalize on to the chest. Keep the bowel wrapped in sterile moist gauze throughout the procedure.</li>
	<li>At this time perform a right nephrectomy. Ligate the renal artery and vein with 6-0 silk sutures. Excise the kidney distal to the sutures. Ligate the ureter with 6-0 silk and then divide proximal to the kidney.</li>
	<li>Isolate the abdominal aorta and inferior vena cava (IVC) below the renal vessels. Place 4-0 cotton ties around the aorta and IVC superior then inferior to the anastomosis site. Identify and ligate any lumbar vessels within the field with 10-0 nylon suture.</li>
	<li>Knot the cotton ties, first the inferior followed by the superior. In this way some blood is retained in the aorta making the aortotomy easier.</li>
	<li>Form the aortotomy with a 30G needle to enter the lumen of the aorta. Extend the incision with fine micro scissors to a length of approximately 2mm.</li>
	<li>Make an end to side anastomosis of the donor aortic heel-and-toe cuff to the recipient aorta in the following fashion. Place a 10-0 nylon suture stay stitch in the donor aorta and to the inferior angle of the incision in the recipient aorta and tie.</li>
	<li>Place a second 10-0 nylon opposite the first in the donor aorta and the superior corner of the incision in the abdominal aorta and tie. Make a running suture line from superior to inferior in the lateral wall of the aorta and tie against the previously placed stay stitch. Then suture the medial wall in a running fashion and tie.</li>
	<li>Make an end to side anastomosis of the donor renal vein to the recipient IVC in the following fashion. Puncture the IVC with a 30G needle and extend the incision with fine micro scissors. Tie the donor renal vein to the inferior corner of the incision in the IVC with 10-0 nylon. Make a running suture line between the renal vein and the IVC and tie. 
	NOTE: It is also vitally important that full thickness passes of the suture needle including the vascular adventitia and the intima are achieved. Eversion of the edges also ensures that there is intima-to-intima contact, which aids in sealing and healing of the anastomoses. While a hemostatic clotting agent can be useful for reducing leaks, we recommend that a surgeon instead rely on good technique.</li>
	<li>Ensure that the anastomoses are &#8220;clean&#8221;. That is, that the opposing walls are not caught when placing stitches. This will cause a significant constriction to flow that will result in a failed graft and in extreme cases to hind limb paralysis. 
	NOTE: Another vitally important factor is ensuring that the tension of the anastomotic suture lines is also optimal. Too loose and there will be irreversible leaking, too tight and stricture to flow will result. If on the arterial side this will result in poor perfusion of the graft, if on the venous side a congested kidney will result.</li>
	<li>Release the distal 4-0 cotton tie re-establishing venous flow. Once hemostasis of the venous anastomosis has been observed gradually loosen the proximal 4-0 cotton tie and observe the arterial anastomosis for hemostasis.</li>
	<li>Remove the cotton ties from the mouse once both anastomoses are considered secure. Pierce the bladder with a 20 G needle creating two holes.</li>
	<li>Pass the tips of curved forceps through the holes and pull the donor ureter through the bladder with the forceps with the proximal end of the ureter being anchored to the bladder wall with two 10-0 nylon sutures. Trim the excess length of ureter protruding from the second hole allowing the ureter to retract within the bladder and close the hole with two 10-0 nylon sutures.</li>
	<li>Return the bowel to the abdomen. Close the abdominal wall in two layers using 5-0 Silk suture in a running fashion.</li>
	<li>Administer a 1.0 ml bolus of sterile, warm normal saline into the abdomen as fluid resuscitation upon closing, and inject 0.8 ml of normal saline subcutaneously post-operatively. No other supportive measures are required during the surgery.</li>
	<li>Recover the animal on a warming blanket. Total implant time is approx. 35-45 min. Administer analgesics such as buprenorphine, 0.05 mg/kg, SC, 0.1-0.2 ml at the beginning of the procedure and every 6-12 hr for 72 hr post-op.</li>
</ol>
3. Contralateral Nephrectomy
<ol>
	<li>Depending on the protocol requirements, several days after the implant procedure, perform a contralateral nephrectomy under isoflurane anesthesia (5% inhaled isoflurane for induction, 1.5-2.0% for maintenance).</li>
	<li>Enter the abdominal cavity and gently retract the bowel to the animal&#8217;s right. Expose the left kidney and blunt dissect from the surrounding fascia.</li>
	<li>Ligate the renal artery, vein and ureter with 6-0 silk sutures and then excise the kidney above the sutures and remove the kidney. Total operative time is approx. 10 min.</li>
</ol>
4. Graft Assessment
<ol>
	<li>Measure serum creatinine using the alkaline picrate method (Jaffe reaction).</li>
</ol>

<ol>
	<li>Skoskiewicz, M., Chase, C., Winn, H. J., Russell, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kidney+transplants+between+mice+of+graded+immunogenetic+diversity.">Kidney transplants between mice of graded immunogenetic diversity.</a> Transplant Proc. 5 (1), 721-725 (1973).</li><li>Kalina, S. L., Mottram, P. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microsurgical+technique+for+renal+transplantation+in+mice.">A microsurgical technique for renal transplantation in mice.</a> Aust N Z J Surg. 63 (3), 213-216 (1993).</li><li>Han, W. R., Murray-Segal, L. J., Mottram, P. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modified+technique+for+kidney+transplantation+in+mice.">Modified technique for kidney transplantation in mice.</a> Microsurgery. 19 (6), 272-274 (1999).</li><li>Ge, F., Gong, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strategies+for+successfully+establishing+a+kidney+transplant+in+a+mouse+model.">Strategies for successfully establishing a kidney transplant in a mouse model.</a> Exp Clin Transplant. 9 (5), 287-294 (2011).</li><li>Martins, P. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Learning+curve,+surgical+results+and+operative+complications+for+kidney+transplantation+in+mice.">Learning curve, surgical results and operative complications for kidney transplantation in mice.</a> Microsurgery. 26 (8), 590-593 (2006).</li><li>Corry, R. J., Winn, H. J., Russell, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primarily+vascularized+allografts+of+hearts+in+mice.+The+role+of+H-2D,+H-2K,+and+non-H-2+antigens+in+rejection.">Primarily vascularized allografts of hearts in mice. The role of H-2D, H-2K, and non-H-2 antigens in rejection.</a> Transplantation. 16 (4), 343-350 (1973).</li><li>Zhang, Z., Chen, H., Qian, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kidney+Transplantation+in+Mice.">Kidney Transplantation in Mice.</a> Experimental Organ Transplantation. , 45-64 (2013).</li><li>Zhang, L., Moskovitz, M., Piscatelli, S., Longaker, M. T., Siebert, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hemodynamic+study+of+different+angled+end-to-side+anastomoses.">Hemodynamic study of different angled end-to-side anastomoses.</a> Microsurgery. 16 (2), 114-117 (1995).</li><li>Liu, Q., Mirc, D., Fu, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+mechanisms+of+thrombosis+in+intact+bent+microvessels+of+rat+mesentery.">Mechanical mechanisms of thrombosis in intact bent microvessels of rat mesentery.</a> J Biomech. 41 (12), 2726-2734 (2008).</li><li>Chesnutt, J. K., Han, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tortuosity+triggers+platelet+activation+and+thrombus+formation+in+microvessels.">Tortuosity triggers platelet activation and thrombus formation in microvessels.</a> J Biomech Eng. 133 (12), 121004 (2011).</li><li>Han, H. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+vessel+buckling+within+soft+surrounding+tissue+generates+tortuosity.">Blood vessel buckling within soft surrounding tissue generates tortuosity.</a> J Biomech. 42 (16), 2797-2801 (2009).</li><li>Gutierrez-Diaz, J. 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=Intraluminal+thrombus+and+neointimal+hyperplasia+after+microvascular+surgery.">Intraluminal thrombus and neointimal hyperplasia after microvascular surgery.</a> Surg Neurol. 24 (2), 153-159 (1985).</li><li>Khouri, R. K., Cooley, B. C., Kenna, D. M., Edstrom, L. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thrombosis+of+microvascular+anastomoses+in+traumatized+vessels:+fibrin+versus+platelets.">Thrombosis of microvascular anastomoses in traumatized vessels: fibrin versus platelets.</a> Plast Reconstr Surg. 86 (1), 110-117 (1990).</li><li>Johnson, P. C., 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=Initial+platelet+deposition+at+the+human+microvascular+anastomosis:+effect+on+downstream+platelet+deposition+to+intact+and+injured+vessels.">Initial platelet deposition at the human microvascular anastomosis: effect on downstream platelet deposition to intact and injured vessels.</a> Plast Reconstr Surg. 90 (4), 650-658 (1992).</li></ol>

The authors have nothing to disclose.

Mastering this transplant technique is difficult, but once accomplished it is a very powerful research tool. The patient surgeon/researcher will be rewarded by attention to detail and consistency of technique, which is the key to mastering any surgical procedure, even more so in small animal models. The technical difficulties of mastering the mouse kidney transplant are many folds, and it is highly probable that experience in other small animal transplant models must be gained before tackling this procedure.
<p class...

Since 1973 the kidney transplant model in mice has been a valuable research tool, but technical issues have hampered its widespread use. Over the years several papers have been published detailing improvements/refinements to this procedure. As a model of primarily vascularized solid organ transplantation this procedure is probably second only to the heterotopic heart transplant model which was also devised by the Russell laboratory in 1973 6. Both models lend themselves to research into allogeneic rejection responses, the development of delayed graft function and ischemia reperfusion injury.
One of the most common issues to be reported with kidney transplantation is the relatively high incidence of arterial thrombosis 4,5,7 which we also experienced in our laboratory. Therefore we set out to perform a literature review of thrombus formation and possibly find the cause of this technical issue and to also devise a possible solution. The most likely cause of thrombosis is the somewhat tortuous path the blood takes from the recipient aorta, into the donor renal aorta then on to the donor renal artery. This path causes turbulence in the renal artery which can lead to platelet activation and thrombus formation. Based on the recent observations and a search of relevant literature 8-14 we came up with a new technique that has reduced thrombosis to 0%.
The technique described here varies from previously reported techniques in the formation of an arterial heel-and-toe cuff which facilities improved blood flow and significantly reduces thrombus formation. The cuff is formed by dividing the infra-renal aorta across the face of the renal arterial ostium at an angle less than 45o to the longitudinal axis of the aorta (Figure 1A &#38; 1B). This results in a cuff approximately 2mm in length. A venous Carrel patch is formed by transecting the renal vein into the IVC thereby increasing the diameter of the cuff. The infra-renal donor abdominal aorta heel-and-toe cuff is end-to-side anastomosed to the recipient abdominal aorta and the donor renal vein/IVC patch is end-to-side anastomosed to the recipient abdominal inferior vena cava (IVC). The ureter is then introduced into and anchored to the bladder as described by Han et al 3.
For this study untreated transplants with warm ischemia times only (i.e., no cold ischemia) are compared. In this case warm ischemia refers to the time from the cessation of blood flow through the donor kidney (step 1.11 below) and reperfusion of the graft in the recipient (step 2.11 below). Cold ischemia refers to the time that the kidney is not perfused and is kept in cold storage until the beginning of the implant procedure.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Instrument</td>
			<td>Roboz #</td>
			<td>Fine Science Tools #</td>
			<td>Arosurgical #</td>
		</tr>
		<tr>
			<td>Straight micro-dissecting forcep #5</td>
			<td>RS-5015</td>
			<td>11295-51</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved micro-dissecting forcep #7</td>
			<td>RS-5047</td>
			<td>11297-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved serrated forcep</td>
			<td>RS-5137</td>
			<td>11052-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas micro-dissecting scissors, short</td>
			<td>RS-5610</td>
			<td></td>
			<td>09.140.08</td>
		</tr>
		<tr>
			<td colspan="2">Micro-dissecting scissors, straight, sharp, long</td>
			<td></td>
			<td>11.602.11</td>
		</tr>
		<tr>
			<td>Micro spring handle needle holder</td>
			<td></td>
			<td></td>
			<td>11.549.15</td>
		</tr>
		<tr>
			<td>Straight mosquito forcep</td>
			<td></td>
			<td>91308-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-dissecting scissors, straight, blunt</td>
			<td>RS-5962</td>
			<td>14078-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-dissecting scissors, curved, blunt</td>
			<td>RS-5981</td>
			<td>14079-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro retractor</td>
			<td>RS-6540</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Instrument tray, 10&#8221; x 6 &#189;&#8221; x &#190;&#8221;</td>
			<td>RT-1350S</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Silk suture, 5/0, 22.5m spool</td>
			<td></td>
			<td>18020-50</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Suture</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10/0 nylon</td>
			<td></td>
			<td></td>
			<td>T4A10Q07</td>
		</tr>
		<tr>
			<td>5/0 silk</td>
			<td></td>
			<td></td>
			<td>E19A05N</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gloves</td>
			<td></td>
			<td>Drapes</td>
			<td></td>
		</tr>
		<tr>
			<td>Biogel from Medex Supply</td>
			<td></td>
			<td>Precept, #64-9012-9</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringes</td>
			<td></td>
			<td>Cotton applicators</td>
			<td></td>
		</tr>
		<tr>
			<td>B-D 1cc insulin, #329424</td>
			<td></td>
			<td colspan="2">Fisher-brand, #23-400-100</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone-Iodine swabs</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PDI, #B40600</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4/0 Cotton ties</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">Domestic cotton autoclaved with instruments</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

murine kidney transplant technique

The first mouse kidney transplant technique was published in 19731 by the Russell laboratory. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique using the donor aorta to form the arterial anastomosis instead of the renal artery was developed and reported in 1993 by Kalina and Mottram 2 with a further advancement coming from the same laboratory in 1999 3. While one can become proficient in this model, a search of the literature reveals that many labs still experience a high proportion of graft loss due to arterial thrombosis. We describe here a technique that was devised in our laboratory that vastly reduces the arterial thrombus reported by others 4,5. This is achieved by forming a heel-and-toe cuff of the donor infra-renal aorta that facilitates a larger anastomosis and straighter blood flow into the kidney.

This surgical technique allows for either simple graft survival/rejection studies, or quite complex experimental protocols. In the figures below we demonstrate the advantages of using this improved arterial anastomosis technique. Using this technique we have significantly reduced the incidence of arterial thrombosis from 35% to 0% thus increasing productivity. We have used this technique for over one year with the same 0% thrombosis result maintained. Figure 1 describes the method for the formation of th...

Watch this Scientific Journal Video about Murine Kidney Transplant Technique at JoVE.com

Murine Kidney Transplant Technique

The goal of this manuscript is to describe the steps required to perform a kidney transplant in a mouse, paying particular attention to the details of the arterial anastomosis.

The first mouse kidney transplant technique was published in 19731 by the Russell laboratory. Although it took some years for other labs to become ...

Research

JoVE Journal

Medicine

15.8K vistas.  University of Colorado, Denver. The goal of this manuscript is to describe the steps required to perform a kidney transplant in a mouse, paying particular attention to the details of the arterial anastomosis. 

Video: Murine Kidney Transplant Technique

A Method for Murine Islet Isolation and Subcapsular Kidney Transplantation

Since the early pioneering work of Ballinger and Reckard demonstrating that transplantation of islets of Langerhans into diabetic rodents could normalize their blood glucose levels, islet transplantation has been proposed to be a potential treatment for type 1 diabetes 1,2. More recently, advances in human islet transplantation have further strengthened this view 1,3. However, two major limitations prevent islet transplantation from being a widespread clinical reality: (a) the requirement for large numbers of islets per patient, which severely reduces the number of potential recipients, and (b) the need for heavy immunosuppression, which significantly affects the pediatric population of patients due to their vulnerability to long-term immunosuppression. Strategies that can overcome these limitations have the potential to enhance the therapeutic utility of islet transplantation.
Islet transplantation under the mouse kidney capsule is a widely accepted model to investigate various strategies to improve islet transplantation. This experiment requires the isolation of high quality islets and implantation of islets to the diabetic recipients. Both procedures require surgical steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol. We also briefly discuss different transplantation models: syngeneic, allogeneic, syngeneic autoimmune, and allogeneic autoimmune.

Transplantation of isolated islets has been proposed to be a potential treatment for type 1 diabetes. Here we describe a method to isolate islets from mouse pancreata and transplant them to the subcapsular space of the kidney.

Since the early pioneering work of Ballinger and Reckard demonstrating that transplantation of islets of Langerhans into diabetic rodents could normalize ...

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are available to perform mechanistic in vivo studies. While heart transplantation models using a suture technique were first successfully developed in rats, the translation into an equally widespread used murine equivalent was never achieved due the technical complexity of the microsurgical procedure. In contrast, non-suture cuff techniques, also developed initially in rats, were successfully adapted for use in mice1-3. This technique for revascularization involves two major steps I) everting the recipient vessel over a polyethylene cuff; II) pulling the donor vessel over the formerly everted recipient vessel and holding it in place with a circumferential tie. This ensures a continuity of the endothelial layer, short operating time and very high patency rates4.
Using this technique for vascular anastomosis we performed more than 1,000 cervical heart transplants with an overall success rate of 95%. For arterial inflow the common carotid artery and the proximal aortic arch were anastomosed resulting in a retrograde perfusion of the transplanted heart. For venous drainage the pulmonary artery of the graft was anastomosed with the external jugular vein of the recipient5.
Herein, we provide additional details of this technique to supplement the video.

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach.
Herein, we provide additional details of this technique to supplement the video.

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are ...

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for liver transplantation die without receiving an organ transplant or are delisted for disease progression. One strategy to increase the donor pool is the utilization of marginal grafts, such as fatty livers, grafts from older donors, or donation after cardiac death (DCD). The current preservation technique of cold static storage is only poorly tolerated by marginal livers resulting in significant organ damage. In addition, cold static organ storage does not allow graft assessment or repair prior to transplantation.
These shortcomings of cold static preservation have triggered an interest in warm perfused organ preservation to reduce cold ischemic injury, assess liver grafts during preservation, and explore the opportunity to repair marginal livers prior to transplantation. The optimal pressure and flow conditions, perfusion temperature, composition of the perfusion solution and the need for an oxygen carrier has been controversial in the past.
In spite of promising results in several animal studies, the complexity and the costs have prevented a broader clinical application so far. Recently, with enhanced technology and a better understanding of liver physiology during ex vivo perfusion the outcome of warm liver perfusion has improved and consistently good results can be achieved.
This paper will provide information about liver retrieval, storage techniques, and isolated liver perfusion in pigs. We will illustrate a) the requirements to ensure sufficient oxygen supply to the organ, b) technical considerations about the perfusion machine and the perfusion solution, and c) biochemical aspects of isolated organs.

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

Marginal grafts, such as fatty livers, grafts from older donors, or livers retrieved after cardiac death (DCD) tolerate conventional, cold static storage only poorly. We developed a novel model of subnormothermic ex vivo liver perfusion for preservation, assessment, and repair of marginal liver grafts prior to transplantation.

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for ...

Murine Heterotopic Heart Transplant Technique

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique was developed and reported in 2001 by Niimi. Described here are the techniques that have evolved over more than a decade in the hands of three surgeons (Plenter, Grazia, Pietra) in our center. These techniques are now being passed on to a younger generation of surgeons and researchers.
Based largely on the Niimi experience, the procedures used have evolved in the fine details - details which we will endeavor to relate here in such a way that others may be able to use this very useful model. Like Niimi, we have found that a video aid to learning is a priceless resource for the beginner.

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become ...

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 ...

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

Kidney transplantation has become a well-established treatment option for patients with end-stage renal failure. The persisting organ shortage remains a serious problem. Therefore, the acceptance criteria for organ donors have been extended leading to the usage of marginal kidney grafts. These marginal organs tolerate cold storage poorly resulting in increased preservation injury and higher rates of delayed graft function. To overcome the limitations of cold storage, extensive research is focused on alternative normothermic preservation methods.
Ex vivo normothermic organ perfusion is an innovative preservation technique. The first experimental and clinical trials for ex vivo lung, liver, and kidney perfusions demonstrated favorable outcomes.
In addition to the reduction of cold ischemic injury, the method of normothermic kidney storage offers the opportunity for organ assessment and repair. This manuscript provides information about kidney retrieval, organ preservation techniques, and isolated ex vivo normothermic kidney perfusion (NEVKP) in a porcine model. Surgical techniques, set up for the perfusion solution and the circuit, potential assessment options, and representative results are demonstrated.

Normothermic Ex Vivo Kidney Perfusion for the Preservation of Kidney Grafts prior to Transplantation

The severe organ shortage has resulted in increased use of marginal kidney grafts for transplantation. This has triggered interest in alternative storage methods, since marginal grafts especially tolerate cold storage poorly. The technique of normothermic ex vivo kidney perfusion (NEVKP) represents a novel preservation method for kidney grafts prior to &#160;transplantation.

Kidney transplantation has become a well-established treatment option for patients with end-stage renal failure. The persisting organ shortage remains a ...

An &#8220;All-laser&#8221; Endothelial Transplant

The &#8220;all laser&#8221; assisted endothelial keratoplasty is a procedure that is performed with a femtosecond laser used to cut the donor tissue at an intended depth, and a near infrared diode laser to weld the corneal tissue. The proposed technique enables to reach the three main goals in endothelial keratoplasty: a precise control in the thickness of the donor tissue; its easy insertion in the recipient bed and a reduced risk of donor lenticule dislocation. The donor cornea thickness is measured in the surgery room with optical coherence tomography (OCT), in order to correctly design the donor tissue dimensions. A femtosecond laser is used to cut the donor cornea. The recipient eye is prepared by manual stripping of the descemetic membrane. The donor endothelium is inserted into a Busin-injector, the peripheral inner side is stained with a proper chromophore (a water solution of Indocyanine Green) and then it is pulled in the anterior chamber. The transplanted tissue is placed in the final and correct location and then diode laser welding is induced from outside the eyeball. The procedure has been performed on more than 15 patients evidencing an improvement in surgery performances, with a good recovery of visual acuity and a reduced donor lenticule dislocation event.

An &amp;#8220;All-laser&amp;#8221; Endothelial Transplant

An all laser procedure is proposed in the endothelial transplant. The surgical technique is based on the use of a femtosecond laser to prepare the donor tissue. Laser welding technique is then used to secure the donor endothelium in the correct position.

The &#8220;all laser&#8221; assisted endothelial keratoplasty is a procedure that is performed with a femtosecond laser used to cut the donor tissue at an ...

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 ...

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

Organ Ischemia-Reperfusion Injury by Simulating Hemodynamic Changes in Rat Liver Transplant Model

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, including ischemia-reperfusion injury (IRI) of extrahepatic organs. This model requires numerous experiments and devices. The duration of anhepatic phase is closely related to the time to develop IRI after transplantation. In this experiment, we used hemodynamic changes to induce extrahepatic organ damage in rats and determined the maximum tolerance time. The time until the most severe organ injury varied for different organs. This method can easily be replicated and can also be used to study IRI of the extrahepatic organs after liver transplantation.

This paper provides a detailed description of how to build an animal model of the anhepatic phase (liver ischemia) in rats to facilitate basic research into ischemia-reperfusion injury after liver transplantation.

Orthotopic liver transplantation (OLT) in rats is a tried and proven animal model used for preoperative, intraoperative, and postoperative studies, ...