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/pubmed?term=((Kidney+transplants+between+mice+of+graded+immunogenetic+diversity%5BTitle%5D)+AND+%22Transplant+Proc%22%5BJournal%5D)">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/pubmed?term=((A+microsurgical+technique+for+renal+transplantation+in+mice%5BTitle%5D)+AND+%22Aust+N+Z+J+Surg%22%5BJournal%5D)">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/pubmed?term=((Modified+technique+for+kidney+transplantation+in+mice%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">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/pubmed?term=((Strategies+for+successfully+establishing+a+kidney+transplant+in+a+mouse+model%5BTitle%5D)+AND+%22Exp+Clin+Transplant%22%5BJournal%5D)">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/pubmed?term=((Learning+curve%2C+surgical+results+and+operative+complications+for+kidney+transplantation+in+mice%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">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/pubmed?term=((Primarily+vascularized+allografts+of+hearts+in+mice.+The+role+of+H-2D%2C+H-2K%2C+and+non-H-2+antigens+in+rejection%5BTitle%5D)+AND+%22Transplantation%22%5BJournal%5D)">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. Kidney Transplantation in Mice. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aZhang%2C+Z+%22Experimental+Organ+Transplantation%22">Experimental Organ Transplantation</a>. Chen, H., Qian, S. , Nova Science Publishers. 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/pubmed?term=((Hemodynamic+study+of+different+angled+end-to-side+anastomoses%5BTitle%5D)+AND+%22Microsurgery%22%5BJournal%5D)">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/pubmed?term=((Mechanical+mechanisms+of+thrombosis+in+intact+bent+microvessels+of+rat+mesentery%5BTitle%5D)+AND+%22J+Biomech%22%5BJournal%5D)">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/pubmed?term=((Tortuosity+triggers+platelet+activation+and+thrombus+formation+in+microvessels%5BTitle%5D)+AND+%22J+Biomech+Eng%22%5BJournal%5D)">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/pubmed?term=((Blood+vessel+buckling+within+soft+surrounding+tissue+generates+tortuosity%5BTitle%5D)+AND+%22J+Biomech%22%5BJournal%5D)">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/pubmed?term=((Intraluminal+thrombus+and+neointimal+hyperplasia+after+microvascular+surgery%5BTitle%5D)+AND+%22Surg+Neurol%22%5BJournal%5D)">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/pubmed?term=((Thrombosis+of+microvascular+anastomoses+in+traumatized+vessels%3A+fibrin+versus+platelets%5BTitle%5D)+AND+%22Plast+Reconstr+Surg%22%5BJournal%5D)">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/pubmed?term=((Initial+platelet+deposition+at+the+human+microvascular+anastomosis%3A+effect+on+downstream+platelet+deposition+to+intact+and+injured+vessels%5BTitle%5D)+AND+%22Plast+Reconstr+Surg%22%5BJournal%5D)">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.
It is important to ensure that the anastomoses are &#8220;clean&#8221;; i.e.,&#160;the opposing vascular walls must not be caught when placing stitches in order to maintain a patent lumen. Otherwise there will be significant constriction to flow that will more than likely result in a failed graft and in extreme cases lead to hind-limb paralysis. It is also vitally important that full thickness passes of the suture needle including the vascular adventitia and the intima are achieved as this results in proper evertion of the edges ensuring 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 upon good technique.
Also attention must be paid to ensuring the tension of the anastomotic suture lines is optimal, too loose and there will be irreversible leaking, to tight and reduced 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. Attaining this correct tension is a matter of practice and experience with other small animal microvascular models will be of great assistance. Above all consistency of technique and unwavering attention to detail will yield excellent mouse and graft survival, and kidney transplant function.
The limitations of this technique are governed only by the uses which can be applied by the investigator. As with any microvascular procedure, so long as good technique is observed the results should be reproducible.
The incidence of arterial thrombosis using this technique has been drastically reduced. This results in at least 1/3 of all kidney transplants being converted from potential technical failures to usable data, resulting in reduced costs and increased productivity.
As a fully vascularized, orthotopic transplant model the future applications will include rejection/tolerance studies, the phenomenon of delayed graft function and investigations into the mechanisms of ischemia/reperfusion injury.

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><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>Micro-dissecting scissors, straight, sharp, long</td><td></td><td>11.602.11</td><td></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&amp;rdquo; x 6 &amp;frac12;&amp;rdquo; x &amp;frac34;&amp;rdquo;</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>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>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>Syringes</td><td></td><td>Cotton applicators</td><td></td></tr><tr><td>B-D 1cc insulin, #329424</td><td></td><td>Fisher-brand, #23-400-100</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>4/0 Cotton ties</td><td></td><td></td><td></td></tr><tr><td>Domestic cotton autoclaved with instruments</td><td></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 the arterial heel-and-toe cuff which is the basis of this new technique. This cuff provides for a longer anastomosis and a straighter blood flow path into the kidney. Both of these points result in significantly lessened turbulence thereby reducing the likelihood of platelet activation and thrombus formation.
<img alt="Figure 1" src="/files/ftp_upload/52848/52848fig1.jpg" /> 
Figure 1.&#160;Schematic representation of the preparation of the donor cuff and the resulting arterial anastomosis. (A) A line diagram depicting the formation of the arterial heel-and-toe cuff. The cuff is formed by dividing the abdominal aorta obliquely across the renal arterial ostium at an angle less than 45o to the longitudinal axis of the aorta. (B) A simplified diagram showing how the infra-renal aorta is divided to form the cuff. (C) The cuff depicting the resulting lumen. (D) The completed anastomosis (venous and ureteric structures not shown for clarity) results in a large cross-sectional lumen and a straight blood flow path.
<img alt="Figure 2" src="/files/ftp_upload/52848/52848fig2.jpg" /> 
Figure 2.&#160;Immunohistochemical analysis. (A) Periodic Acid-Schiff stained section of a control non-transplanted kidney. (B) A syngeneic kidney transplant at POD 8 using the Revised technique demonstrates normal proximal tubular cells, which are cuboidal, with a clear cytoplasm and a round light nucleus in the middle of the cell. There is evidence of mild brush border injury (arrows) but the majority of brush borders are regular and well preserved. Magnification: 400x.
<img alt="Figure 3" src="/files/ftp_upload/52848/52848fig3.jpg" />
Table 1.&#160;Comparison of thrombosis incidence and warm ischemia times between the two techniques. All values are expressed as mean &#177; SD. For single comparisons, normally distributed data are evaluated using unpaired, two-tailed Student t tests, and non-normally distributed data are analyzed by the nonparametric unpaired Mann-Whitney U test. P values of less than 0.05 are considered statistically significant.

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 ...

Nanyang Technological University

Research

JoVE Journal

Medicine

16.0K Views.  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 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

Isolation and Transplantation of Different Aged Murine Thymic Grafts.

The mechanisms that regulate the efficacy of thymic selection remain ill-defined. The method presented here allows in vivo analyses of the development and selection of T cells specific for self and foreign antigens. The approach entails implantation of thymic grafts derived from various aged mice into immunodeficient scid recipients. Over a relatively short period of time the recipients are fully reconstituted with T cells derived from the implanted thymus graft. Only thymocytes seeding the thymus at the time of isolation undergo selection and develop into mature T cells. As such, changes in the nature and specificity of the engrafted T cells as a function of age-dependent thymic events can be assessed. Although technical expertise is required for successful thymic transplantation, this method provides a unique strategy to study in vivo a wide range of pathologies that are due to or a result of aberrant thymic function and/or homeostasis.

This report provides a detailed description of transplanting murine thymi from different aged donor mice under the kidney capsule of immunodeficient mouse recipients. The goal of this approach is to model T cell development and thymic selection events in vivo.

The mechanisms that regulate the efficacy of thymic selection remain ill-defined. The method presented here allows in vivo analyses of the development and ...

Immunology and Infection

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 ...

Fate Mapping of Human Embryonic Stem Cells by Teratoma Formation 

Human embryonic stem cells (hESCs) have an unlimited capacity for self-renewal, and the ability to differentiate into cells derived from all three embryonic germ layers (1). Directed differentiation of hESCs into specific cell types has generated much interest in the field of regenerative medicine (e.g., (2-5)), and methods for determining the in vivo fate of selected or manipulated hESCs are essential to this endeavor. We have adapted a highly efficient teratoma formation assay for this purpose. A small number of specifically selected hESCs is mixed with undifferentiated wild type hESCs and Phaseolus vulgaris lectin to form a cell pellet. This is grafted beneath the kidney capsule in an immunodeficient mouse. As few as 2.5 x 105 hESCs are needed to form a 16 cm3 teratoma within 8-12 weeks. The fate of the originally selected hESCs can then be determined by immunohistochemistry. This method provides a valuable tool for characterizing tissue-specific reagents for cell-based therapy.

Fate Mapping of Human Embryonic Stem Cells by Teratoma Formation

Directed differentiation of hESCs into specific cells has generated much interest in regenerative medicine. We provide a concise, step-by-step protocol for determining the in vivo fate of selected hESCs that provides a valuable tool for characterizing tissue-specific reagents for cell-based therapy.

Human embryonic stem cells (hESCs) have an unlimited capacity for self-renewal, and the ability to differentiate into cells derived from all three ...

Heterotopic Renal Autotransplantation in a Porcine Model: A Step-by-Step Protocol

Kidney transplantation is the treatment of choice for patients suffering from end-stage renal disease. It offers better life expectancy and higher quality of life when compared to dialysis. Although the last few decades have seen major improvements in patient outcomes following kidney transplantation, the increasing shortage of available organs represents a severe problem worldwide. To expand the donor pool, marginal kidney grafts recovered from extended criteria donors (ECD) or donated after circulatory death (DCD) are now accepted for transplantation. To further improve the postoperative outcome of these marginal grafts, research must focus on new therapeutic approaches such as alternative preservation techniques, immunomodulation, gene transfer, and stem cell administration.
Experimental studies in animal models are the final step before newly developed techniques can be translated into clinical practice. Porcine kidney transplantation is an excellent model of human transplantation and allows investigation of novel approaches. The major advantage of the porcine model is its anatomical and physiological similarity to the human body, which facilitates the rapid translation of new findings to clinical trials. This article offers a surgical step-by-step protocol for an autotransplantation model and highlights key factors to ensure experimental success. Adequate pre- and postoperative housing, attentive anesthesia, and consistent surgical techniques result in favorable postoperative outcomes. Resection of the contralateral native kidney provides the opportunity to assess post-transplant graft function. The placement of venous and urinary catheters and the use of metabolic cages allow further detailed evaluation. For long-term follow-up studies and investigation of alternative graft preservation techniques, autotransplantation models are superior to allotransplantation models, as they avoid the confounding bias posed by rejection and immunosuppressive medication.

Porcine models of organ transplantation provide an important platform to study mechanisms of organ preservation. This article describes a heterotopic porcine renal autotransplantation model, which allows investigating new approaches to improve the outcome of transplantation using marginal kidney grafts.

Kidney transplantation is the treatment of choice for patients suffering from end-stage renal disease. It offers better life expectancy and higher quality ...

Production of Humanized Mouse via Thymic Renal Capsule Grafting, CD34+ Cells Injection, and Cytokine Delivery

Animal models provide a vital translation between in vitro and in vivo biomedical research. Humanized mouse models provide a bridge in the representation of human systems, thereby allowing for a more accurate study of pathogenesis, biomarkers, and many other scientific queries. In this method described, immune-deficient NOD-scid IL2R&#947;null (NSG) mice are implanted with autologous thymus, injected with liver-derived CD34+ cells followed by a series of injected cytokine deliveries. In contrast to other models of a similar nature, the model described here promotes an improved reconstitution of immune cells by delivering cytokines and growth factors via transgenes encoded in AAV8 or pMV101 DNA-based vectors. Moreover, it offers long-term stability with reconstituted mice having an average lifespan of 30 weeks after CD34+ injections. Through this model, we hope to provide a stable and impactful method of studying immunotherapy and human disease in a murine model, thus demonstrating the need for predictive preclinical models.

Production of Humanized Mouse via Thymic Renal Capsule Grafting, CD34+ Cells Injection, and Cytokine Delivery

Humanized mouse models provide a more accurate representation of the human immune microenvironment. This manuscript describes the process in which these models are created through a renal graft of human thymus, injection of human CD34+ cells, and the targeted delivery of human cytokine transgenes to promote CD34+ cell proliferation and differentiation.

Animal models provide a vital translation between in vitro and in vivo biomedical research. Humanized mouse models provide a bridge in the representation ...

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 ...

Orthotopic Rat Kidney Transplantation: A Novel and Simplified Surgical Approach

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

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

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

Extracting Intact Kidney from Mouse: A Technique to Obtain Kidney Without Renal Capsule from Murine Models

Source: Grainger, N. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/62040" target="_blank">Isolating and Imaging Live, Intact Pacemaker Regions of Mouse Renal Pelvis by Vibratome Sectioning.</a> J. Vis. Exp. (2021)
In this video, we describe the procedure for kidney extraction from the retroperitoneal space of the mouse. The extracted kidney can further be microdissected to obtain an intact kidney without a renal capsule and the surrounding fat layer.

Source: Grainger, N. et al. Isolating and Imaging Live, Intact Pacemaker Regions of Mouse Renal Pelvis by Vibratome Sectioning. J. Vis. Exp. (2021)
In ...

Murine Kidney Transplant Technique

Samenvatting

Meer video's verkennen