Funding from Kidney Research UK, The Royal College of Surgeons of Edinburgh and The European Society of Organ Transplantation supported this study.

Appropriate national and local institutional ethics should be in place before performing animal experiments. Specifically in the United Kingdom the following experiments were undertaken under the Animals (Scientific Procedures) Act 1986. Where two microsurgeons are available to operate simultaneously the donor surgeon should perform steps 1.1 to 1.16 then 3.1 to 3.5, whilst the recipient surgeon performs 2.1 to 2.8. For a single operator the steps can be followed sequentially.
1. Donor Preparation
Note: The procedure presented here is for a donor C57BL/6BM12 and recipient C57BL/6 male mice aged between 8 to 16 weeks old with a body weight greater than 20 g. However this procedure can be reproducibly performed on a variety of mouse strains. The data presented in the representative results section was obtained from C57BL/6, C57BL/6BM12 and BALB/c mice.
<ol>
	<li>Conduct procedures using sterile surgical instruments and consumables (autoclaved), with endeavors to keep the operating area as sterile as possible. Perform the donor preparation simultaneously with the recipient preparation if two surgeons are available.</li>
	<li>Anesthetize the mouse with an intraperitoneal injection (31G needle) of medetomidine (0.5 mg/kg) and ketamine hydrochloride (200 mg/kg). This results in an anesthetic plane that is maintained for 4 hr providing enough time for the whole procedure to be performed. Supplemental anesthesia is not necessary.</li>
	<li>Confirm that the mouse is anaesthetized (no response to toe pinch).</li>
	<li>Shave the mouse&#8217;s abdomen and remove any loose hair with sticky tape.</li>
	<li>Place the mouse on its back on a sterilely draped heated mat and loosely immobilize the limbs with sterile masking tape.</li>
	<li>Monitor the mouse throughout the procedure for thermal burns. If possible use a non-electric heat source.</li>
	<li>Apply an eye lubricant and sanitize the abdominal wall with a dilute iodine solution.</li>
	<li>Make a midline incision in the abdomen to enter the peritoneal cavity and insert a 3 cm Calibri abdominal retractor.</li>
	<li>Apply warmed saline to keep the intestines and surgical area moist and avoid unnecessary drying of the viscera.</li>
	<li>Cover the mouse with sterile drapes and move the intestines to the operator&#8217;s left (mouse&#8217;s right) to expose the aorta, vena cava and left kidney.</li>
	<li>Apply a ratcheted forcep to the stomach and pull superiorly to expose the major vessels and left kidney fully. Pack damp sterile swabs (2 mm x 2 mm) into the abdomen to retract tissues away from the surgical area, such as the liver lobes, seminal vesicles, and bowel.</li>
	<li>Isolate the left kidney from surrounding adventia, fat and the left adrenal gland in the peritoneal cavity by bluntly dissecting the connective tissue using fine tip forceps. Place the closed tips of the forceps between the areas that need to be separated and slowly allow the forcep tips to open to dissect the space.</li>
	<li>Isolate the left renal vein by ligating and then dividing the left adrenal vein and left gonadal vein with 9/O nylon. Place the suture close to the renal vein.</li>
	<li>Ligate and divide the ureter with 7/O silk suture close to the bladder leaving the suture ends long. These long suture ends will be used when the kidney is harvested and are necessary for the ureteric anastomosis.</li>
	<li>To mobilize and fully dissect the aorta and vena cava superiorly and inferior to the renal artery and vein use fine tip forceps to bluntly dissect lymphatic vessels and fat from the front and sides of the vessels.</li>
	<li>Find the tissue plane between the aorta and vena cava and slowly spread the forceps. Slowly open the tips of the fine tipped forceps to spread the tissue with minimal trauma.</li>
	<li>Tie a loose 7/O silk around the aorta superior and inferior to the renal artery by passing an angled fine tip forceps around the back of the vessels and drawing a suture through. These sutures will be tightened prior to kidney retrieval to allow retrograde perfusion of University of Wisconsin solution.</li>
</ol>
2. Recipient Preparation
As per the donor preparation follow steps 1.2 to 1.8.
<ol>
	<li>Move the intestines to the operator&#8217;s right (mouse&#8217;s left) to expose the aorta, vena cava. Cover the intestines with a sterile saline soaked swab.</li>
	<li>Perform a right nephrectomy by ligating the right renal artery and vein together with 7/0 silk suture and then divide.</li>
	<li>Ligate the right ureter with 7/O silk suture and divide.</li>
	<li>Mobilize and fully dissect the aorta and vena cava inferior to the renal artery and vein as described in step 1.14 and 1.15. Ensure complete dissection between the aorta and vena cava. Take care to preserve the internal spermatic artery which runs anterior to the vena cava and aorta along with the lymphatic bundle.</li>
	<li>Identify lumbar vessels running in the retroperitoneum from the vena cava and vein to the spine, ligate with 9/O silk in continuity, there is no need to divide the vessels.</li>
	<li>Identify sufficient space to place microvascular clamps with space between for vascular anastomoses.</li>
	<li>Administer intravenous heparin (5 units) via dorsal penile vein.</li>
</ol>
3. Donor Kidney Retrieval
Tighten the 7/O silk ties that have been placed around the inferior and superior aorta to isolate the kidney from the arterial circulation.
<ol>
	<li>Infuse retrograde cold 0.2 &#8211; 0.5 ml of University of Wisconsin solution with a needle (31G) into the aorta.</li>
	<li>Divide the aorta within the sutures and divide the renal vein at its junction with the vena cava to remove the kidney and the renal artery with a length of aorta. Divide lumbar vessels arising from the aorta, if they are present, without ligation.</li>
	<li>Place the kidney in cold saline on a sterile swab in a culture dish.</li>
	<li>Euthanize the donor mouse by cervical dislocation.</li>
</ol>
4. Kidney Transplantation &#8211; Kidney Preparation
<ol>
	<li>Create an aortic patch by dividing the aorta wall longitudinally directly opposite the renal artery. Identify any vessel lumens in the patch that need to be ligated or avoided when performing the arterial anastomosis.</li>
	<li>Place a 10/O suture from outside to inside the renal vein lumen superiorly and a second separate suture inferiorly. The sutures used to divide the adrenal vein and gonadal vein can be used to orientate the vessel.</li>
	<li>Place the kidney in the right flank of the recipient (Figure 1.1) and ensure the sutures are well placed to ensure vascular anastomoses can be completed without the sutures becoming entwined in swabs or other instruments.</li>
</ol>
5. Kidney Transplantation &#8211; Vascular Anastomosis
Apply microvascular clamps first inferiorly then superiorly encompassing the vena cava and aorta.
<ol>
	<li>Make a ventomy with a needle (31G) by puncturing the anterior wall of the vena cava. Flush blood from the vena cava by injecting approximately 50 &#956;l of 0.9% NaCl.</li>
	<li>Widen the venotomy using fine tip forceps by opening them within the venotomy so that the length is equivalent to the diameter of the donor renal vein.</li>
	<li>Place the 10/O superior suture (which is already in the lumen of the renal vein) first at the apex of the venotomy (Figure 1.2) and join the back wall of the anastomosis in a running suture until the inferior apex is reached (Figure 1.3).</li>
	<li>Place the 10/O inferior suture and tie first with a single knot (Figure 1.4) then tie to the running suture from the back wall.</li>
	<li>Using the inferior suture create the front wall of the anastomosis (Figure 1.5) with a running suture and tie to the superior suture end at the superior apex (Figure 1.6).</li>
	<li>Create an aortotomy by picking up the aorta wall with forceps and cutting an elliptical patch with scissors (approximately the aortotomy should be a fifth of the circumference of the aorta and three times the renal artery lumen in length).</li>
	<li>Place a 10/O suture at the superior point of the aortic patch from outside to inside (Figure 1.7) and pass through the aorta to tie outside the vessels at the superior apex of the aortotomy.</li>
	<li>Create the arterial anastomosis with a running suture starting superiorly (Figure 1.8) anastomosing the donor patch to the recipient aorta, take care not to constrict the anastomosis by tying the suture too tight (Figure 1.9).</li>
	<li>Remove the inferior vascular clamps first then the superior clamp to reperfuse the kidney (Figure 1.10 and 1.11). Visible peristalsis of the ureter maybe seen if adequate blood supply to the ureter is achieved.</li>
</ol>
6. Kidney Transplantation &#8211; Ureteric Anastomosis
<ol>
	<li>Divide any attachments of the bladder from the abdominal wall.</li>
	<li>Pass a needle (21G) from the left to right of the bladder through both walls.</li>
	<li>Place straight forcep tips in the needle lumen and pass both back through the bladder such that the forceps are now passing from right to left out of the left side of the bladder.</li>
	<li>Draw the suture on the donor ureter through the bladder such that the ureter passes in the left bladder defect then out of the right side.</li>
	<li>Suture the adventitia of the ureter to the adventitia of the bladder with three single interrupted sutures around the entry point with 9/O nylon suture on a round-bodied needle.</li>
	<li>Cut the ureter proximal to the ligature, thus opening the ureter to allow urine to flow and allow the ureter end to retract into the body of the bladder. Visible production of urine may be observed along with bleeding from peri-ureteric vessels.</li>
	<li>Close the right side bladder defect with a single interrupted 9/O silk suture.</li>
</ol>
7. Recovery and Post-operative Care
Replace the gastrointestinal viscera into the abdomen in their original orientation and close the abdominal wall by approximating the rectus muscles with 6/O absorbable suture.
<ol>
	<li>Approximate the skin with metal skin clips.</li>
	<li>Partially reverse the anesthesia with a subcutaneous injection of atipamazole hydrochloride (10 &#956;l/g).</li>
	<li>Administer analgesia by subcutaneous buprenorphine hydrochloride (0.05 mg/kg) and for fluid support injection 1 ml of subcutaneous 0.9% NaCl.</li>
	<li>Recover the mouse in a warming cabinet at 28 &#176;C for 24 to 48 hr. Observe the mouse for illness up to the completion of the experiment.</li>
	<li>Remove metal skin clips 7 &#8211; 10 days post operatively.</li>
	<li>Once the experiment is complete euthanize the mouse by cervical dislocation.</li>
	<li>Remove and collect the contralateral kidney and allograft for histological analysis.</li>
</ol>

<ol>
	<li>Guild, W. R., Harrison, J. H., Merrill, J. P., Murray, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Successful+homotransplantation+of+the+kidney+in+an+identical+twin.">Successful homotransplantation of the kidney in an identical twin.</a> Trans. Am. Clin. Climatol Assoc. 67, 167-173 (1955).</li><li>Wolfe, R. 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=Comparison+of+mortality+in+all+patients+on+dialysis,+patients+on+dialysis+awaiting+transplantation,+and+recipients+of+a+first+cadaveric+transplant.">Comparison of mortality in all patients on dialysis, patients on dialysis awaiting transplantation, and recipients of a first cadaveric transplant.</a> N. Engl. J. Med. 341, 1725-1730 (1999).</li><li>Nankivell, B. J., Alexander, S. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Rejection of the Kidney Allograft. N. Engl. J. Med. 363, 1451-1462 (2010).</li><li>Tse, G. H., Hughes, J., Marson, L. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systematic+review+of+mouse+kidney+transplantation.">Systematic review of mouse kidney transplantation.</a> Transplant International. 26, 1149-1160 (2013).</li><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.+Transplant.">Kidney transplants between mice of graded immunogenetic diversity. Transplant.</a> Proc. 5, 721-725 (1973).</li><li>Zhang, Z., 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=Pattern+of+liver,+kidney,+heart,+and+intestine+allograft+rejection+in+different+mouse+strain+combinations.">Pattern of liver, kidney, heart, and intestine allograft rejection in different mouse strain combinations.</a> Transplantation. 62, 1267-1272 (1996).</li><li>Qi, F., 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=Depletion+of+cells+of+monocyte+lineage+prevents+loss+of+renal+microvasculature+in+murine+kidney+transplantation.">Depletion of cells of monocyte lineage prevents loss of renal microvasculature in murine kidney transplantation.</a> Transplantation. 86, 1267-1274 (2008).</li><li>Dang, Z., Mackinnon, A., Marson, L. P., Sethi, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tubular+atrophy+and+interstitial+fibrosis+after+renal+transplantation+is+dependent+on+galectin-3.">Tubular atrophy and interstitial fibrosis after renal transplantation is dependent on galectin-3.</a> Transplantation. 93, 477-484 (2012).</li><li>Jabs, W. J., 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=Heterogeneity+in+the+Evolution+and+Mechanisms+of+the+Lesions+of+Kidney+Allograft+Rejection+in+Mice.">Heterogeneity in the Evolution and Mechanisms of the Lesions of Kidney Allograft Rejection in Mice.</a> Am. J. Transplant. 3, 1501-1509 (2003).</li><li>Lin, T., 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=Deficiency+of+C4+from+Donor+or+Recipient+Mouse+Fails+to+Prevent+Renal+Allograft+Rejection.">Deficiency of C4 from Donor or Recipient Mouse Fails to Prevent Renal Allograft Rejection.</a> Am. J. Pathol. 168, 1241-1248 (2006).</li></ol>

The authors have nothing to disclose.

The most well described manner to perform the arterial anastomosis is to use the distal aorta of the donor, with the renal artery in continuation, in an end-to-side manner to the recipient aorta. We describe the use of an aortic patch, similar to the &#8216;Carrell patch&#8217; mirroring that performed in human kidney transplantation which we believe to be more convenient. Though reports in the literature of donor and recipient operative time are sparse we believe that utilizing an aortic patch to recipient aorta instead...

Successful renal transplantation for the treatment of renal failure was first described in 1955 between monozygotic twins1, since then it has become a revolutionary treatment for patients with end-stage renal failure throughout the world, offering both improvement in length and quality of life2. However long term graft survival has been hampered by a multitude of pathological processes resulting in chronic allograft damage3.
Rejection of the transplanted kidney in humans remains a major cause of morbidity, despite significant improvements in immunosupporessive regimens. The aim of developing a mouse model of renal transplantation is to closely replicate the process and pathology found in human renal transplantation4. Skoskiewicz et al. first described the mouse model of renal transplantation in 19735. Although advanced microsurgical skills are required, it is a valuable tool for several reasons: the mouse genome has been well characterized and there is a great variety of experimental methods and techniques available for mouse studies.
Many groups using the mouse model of renal transplantation have used the transplanted kidney as a life-supporting organ, however in other studies and in our described methodology one of the recipient mouse&#8217;s native kidneys is left in situ for the duration of the experiment4. The benefit is that the mouse undergoes a single anesthesia and operation thereby reducing the morbidity to the mouse and the risk of death from a second procedure. Additionally the mouse does not suffer from the adverse effects of gradual renal failure.
Although models of allogeneic rejection exist in other organs such as the heart and skin, these are not always directly relevant to renal transplantation. There is evidence that these models elicit different modes and dynamics of rejection, for instance the time course of rejection in cardiac allograft and renal allograft differs significantly in certain strain combinations6. We have described acute renal allograft rejection patterns in BALB/c donors into non-transgenic FVB/nj mice, this model showed cellular mediated injury with accumulation of T cells and macrophages7. Alternatively we have also described a model of chronic allograft damage that exhibits interstitial fibrosis and tubular atrophy, this results from transplanting a kidney from C57BL/6BM12 donors into C57BL/6 recipients, as these mice are characterized by a single MHC class II loci mis-match8.
Multiple aspects of transplantation have been studied using the mouse model of renal transplantation including acute rejection, cellular and humoral rejection, ischemia reperfusion injury, and trialing novel therapeutic agents. We have modified the surgical technique to reduce operating time and improve the ease of surgery. Particularly we have described simultaneous donor and recipient preparation and a simplified vascular anastomosis technique by utilizing a continuous aortic patch anastomosis. This video and manuscript will provide key points to aid in the establishment of this technique.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Surgical Instruments</td>
		</tr>
		<tr>
			<td>Blunt Dissecting Scissors</td>
			<td>Fine Science Tools</td>
			<td>&#160;14072-10</td>
			<td>For skin cutting</td>
		</tr>
		<tr>
			<td>Curved Castoviejo scissors</td>
			<td>Fine Science Tools</td>
			<td>15017-10</td>
			<td>For tissue cutting</td>
		</tr>
		<tr>
			<td>Spring Scissors &#8211; straight</td>
			<td>Fine Science Tools</td>
			<td>15000-08</td>
			<td>For suture cutting</td>
		</tr>
		<tr>
			<td>Toothed forceps 1x2 teeth</td>
			<td>Fine Science Tools</td>
			<td>11021-12</td>
			<td></td>
		</tr>
		<tr>
			<td>2 x Fine Tip forceps (Dumont No.5)</td>
			<td>Fine Science Tools</td>
			<td>11251-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Angled Fine Tip forceps (Dumont No. 5/45)</td>
			<td>Fine Science Tools</td>
			<td>11253-25</td>
			<td>For blunt dissecting</td>
		</tr>
		<tr>
			<td>Curved Fine Tip forcep (Dumont No.7)</td>
			<td>Fine Science Tools</td>
			<td>11273-22</td>
			<td>Useful to pass around vessels</td>
		</tr>
		<tr>
			<td>Curved Crile Haemostat</td>
			<td>Fine Science Tools</td>
			<td>1300-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro clip applicator with lock</td>
			<td>Fine Science Tools</td>
			<td>18056-14</td>
			<td></td>
		</tr>
		<tr>
			<td>2 x Micro serrefines spring width 2mm, jaw length 4mm</td>
			<td>Fine Science Tools</td>
			<td>18055-04</td>
			<td>Microvascular clamps</td>
		</tr>
		<tr>
			<td>2 x Colibri 3cm wire retractor</td>
			<td>Fine Science Tools</td>
			<td>17000-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo needle holder with lock</td>
			<td>Fine Science Tools</td>
			<td>120660-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Wound clip applicator</td>
			<td>Fine Science Tools</td>
			<td>12031-07</td>
			<td></td>
		</tr>
		<tr>
			<td>7mm wound clips</td>
			<td>Fine Science Tools</td>
			<td>12032-07</td>
			<td>Remove 7 to 10 days after surgery</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>OPMI pico microscope</td>
			<td>Carl Zeiss</td>
			<td>S100</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermal cautery unit with fine tip</td>
			<td>Geiger</td>
			<td>150A</td>
			<td></td>
		</tr>
		<tr>
			<td>Heat electronic pad</td>
			<td>Cozee Cumfort</td>
			<td>n/a</td>
			<td></td>
		</tr>
		<tr>
			<td>Euroklav 23-S</td>
			<td>Melag</td>
			<td>n/a</td>
			<td>Autoclave</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Disposable equipment</td>
		</tr>
		<tr>
			<td>7/O Silk braided suture</td>
			<td>Pearsall</td>
			<td>30514</td>
			<td></td>
		</tr>
		<tr>
			<td>10/O Dafilon (polyamide) suture</td>
			<td>B-Braun</td>
			<td>&#160;G1118099</td>
			<td></td>
		</tr>
		<tr>
			<td>6/O Vicryl (plygalectin)</td>
			<td>Ethicon</td>
			<td>W9537</td>
			<td></td>
		</tr>
		<tr>
			<td>Regular bevel needle, 1 inch, 21G</td>
			<td>Bection, Dickinson and Company</td>
			<td>305175</td>
			<td>For ureteric anastamosis</td>
		</tr>
		<tr>
			<td>Regular bevel needle, 5/8 inch, 25G</td>
			<td>Bection, Dickinson and Company</td>
			<td>305122</td>
			<td></td>
		</tr>
		<tr>
			<td>Regular bevel needle, 1/2 inch, 30G</td>
			<td>Bection, Dickinson and Company</td>
			<td>304000</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin needle 1ml, 29G</td>
			<td>Bection, Dickinson and Company</td>
			<td>324827</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin needle 0.3ml, 30G</td>
			<td>Bection, Dickinson and Company</td>
			<td>324826</td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml syringe slip tip</td>
			<td>Bection, Dickinson and Company</td>
			<td>300184</td>
			<td></td>
		</tr>
		<tr>
			<td>5 ml syringe slip tip</td>
			<td>Bection, Dickinson and Company</td>
			<td>302187</td>
			<td></td>
		</tr>
		<tr>
			<td>Wypall paper swabs</td>
			<td>Kimberley-Clark</td>
			<td>L40</td>
			<td>sterilised by autoclave</td>
		</tr>
		<tr>
			<td>Cotton wool buds</td>
			<td>Johnson and Johnson</td>
			<td>n/a</td>
			<td>sterilised by autoclave</td>
		</tr>
		<tr>
			<td>Plain drapes</td>
			<td>Guardian</td>
			<td>CB03</td>
			<td>sterilised by autoclave</td>
		</tr>
		<tr>
			<td>Cell culture dish 60mm x 15mm</td>
			<td>Corning Incorporated</td>
			<td>430166</td>
			<td></td>
		</tr>
		<tr>
			<td>Dispensing Pin</td>
			<td>B-Braun</td>
			<td>DP3500L / 413501</td>
			<td>Used with NaCl 0.9%</td>
		</tr>
		<tr>
			<td>Re-agents and Drugs</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>(Lacri-Lube) White soft paraffin 57.3%, mineral oil 42.5% and lanolin alcohols 0.2%</td>
			<td>Allergan Ltd</td>
			<td>21956GB10X</td>
			<td></td>
		</tr>
		<tr>
			<td>(Videne) Povidone-iodine 10%</td>
			<td>Ecolab Ltd</td>
			<td>PL 04509/0041</td>
			<td></td>
		</tr>
		<tr>
			<td>(Vetalar V) Ketamine hydrochloride</td>
			<td>Pfizer Animal Health</td>
			<td>Vm 42058/4165</td>
			<td>100mg/ml solution (dose 200mg/kg)</td>
		</tr>
		<tr>
			<td>(Domitor) Medetomidine hydrochloride&#160;</td>
			<td>Orion Pharma</td>
			<td>Vm 06043/4003</td>
			<td>1mg/ml (dose 0.5mg/kg)</td>
		</tr>
		<tr>
			<td>(Vetergesic) Bupernorphine hydrochloride&#160;</td>
			<td>Alsto Animal Health</td>
			<td>Vm 00063/4002</td>
			<td>0.3mg/ml (dose 0.05mg/kg)</td>
		</tr>
		<tr>
			<td>(Antisedan) Atipamezole hydrochoride</td>
			<td>Orion Pharma</td>
			<td>Vm 06043/4004</td>
			<td>5mg/ml (dose 2mg/kg)</td>
		</tr>
		<tr>
			<td>University of Wisconsin Solution</td>
			<td>Belzer Bridge to Life</td>
			<td>n/a</td>
			<td>dose approximately 500 microlitres/mouse</td>
		</tr>
		<tr>
			<td>NaCl 0.9%</td>
			<td>Baxter</td>
			<td>FKE1323</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin Sulphate</td>
			<td>non-proprietary</td>
			<td>n/a</td>
			<td>5000units/ml (dose 5units/mouse)</td>
		</tr>
	</tbody>
</table>

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.

Renal allograft rejection can be assessed by histological analysis of methacarn-fixed paraffin-embedded tissue sections of the transplanted kidney (Figure 2). Isograft transplantation of kidneys between syngeneic mice results in renal ischemic reperfusion injury, however by 4 weeks the tubules have recovered and are histologically comparable to native kidneys. Acute rejection can be modeled by C57BL/6 kidney transplantation into BALB/c recipients, within 1 week there is diffuse mononuclear cell infiltrat...

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Mouse Kidney Transplantation: Models of Allograft Rejection

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 ...

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20.4K Views.  The University of Edinburgh. 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.

Video: Mouse Kidney Transplantation: Models of Allograft Rejection

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 ...

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 ...

Mouse Model of Alloimmune-induced Vascular Rejection and Transplant Arteriosclerosis

Vascular rejection that leads to transplant arteriosclerosis (TA) is the leading representation of chronic heart transplant failure. In TA, the immune system of the recipient causes damage of the arterial wall and dysfunction of endothelial cells and smooth muscle cells. This triggers a pathological repair response that is characterized by intimal thickening and luminal occlusion. Understanding the mechanisms by which the immune system causes vasculature rejection and TA may inform the development of novel ways to manage graft failure. Here, we describe a mouse aortic interposition model that can be used to study the pathogenic mechanisms of vascular rejection and TA. The model involves grafting of an aortic segment from a donor animal into an allogeneic recipient. Rejection of the artery segment involves alloimmune reactions and results in arterial changes that resemble vascular rejection. The basic technical approach we describe can be used with different mouse strains and targeted interventions to answer specific questions related to vascular rejection and TA.

We describe a protocol for aortic interposition grafting in mice. The goal of the protocol is to provide a model with which to study pathological processes and therapeutic strategies relevant to alloimmune reactions in arteries and the resultant arterial changes that contribute to organ transplant failure.

Vascular rejection that leads to transplant arteriosclerosis (TA) is the leading representation of chronic heart transplant failure. In TA, the immune ...

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.

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 ...

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 ...

Assessment of Kidney Function in Mouse Models of Glomerular Disease

The use of murine models to mimic human kidney disease is becoming increasingly common. Our research is focused on the assessment of glomerular function in diabetic nephropathy and podocyte-specific VEGF-A knock-out mice; therefore, this protocol describes the full kidney work-up used in our lab to assess these mouse models of glomerular disease, enabling a vast amount of information regarding kidney and glomerular function to be obtained from a single mouse. In comparison to alternative methods presented in the literature to assess glomerular function, the use of the method outlined in this paper enables the glomerular phenotype to be fully evaluated from multiple aspects. By using this method, the researcher can determine the kidney phenotype of the model and assess the mechanism as to why the phenotype develops. This vital information on the mechanism of disease is required when examining potential therapeutic avenues in these models. The methods allow for detailed functional assessment of the glomerular filtration barrier through measurement of the urinary albumin creatinine ratio and individual glomerular water permeability, as well as both structural and ultra-structural examination using the Periodic Acid Schiff stain and electron microscopy. Furthermore, analysis of the genes dysregulated at the mRNA and protein level enables mechanistic analysis of glomerular function. This protocol outlines the generic but adaptable methods that can be applied to all mouse models of glomerular disease.

This protocol describes a full kidney work-up that should be carried out in mouse models of glomerular disease. The methods allow for detailed functional, structural, and mechanistic analysis of glomerular function, which can be applied to all mouse models of glomerular disease.

The use of murine models to mimic human kidney disease is becoming increasingly common. Our research is focused on the assessment of glomerular function ...

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 ...

Optical Clearing and Imaging of Immunolabeled Kidney Tissue

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) imaging. They have received great attention in all research areas for the potential to analyze microscopic multicellular structures that extend over macroscopic distances. Given that kidney tubules, vasculature, nerves, and glomeruli extend in many directions, which have been only partially captured by traditional two-dimensional techniques so far, tissue clearing also opened up many new areas of kidney research. The list of optical clearing methods is rapidly growing, but it remains difficult for beginners in this field to choose the best method for a given research question. Provided here is a simple method that combines antibody labeling of thick mouse kidney slices; optical clearing with cheap, non-toxic and ready-to-use chemical ethyl cinnamate; and confocal imaging. This protocol describes how to perfuse kidneys and use an antigen-retrieval step to increase antibody- binding without requiring specialized equipment. Its application is presented in imaging different multicellular structures within the kidney, and how to troubleshoot poor antibody penetration into tissue is addressed. We also discuss the potential difficulties of imaging endogenous fluorophores and acquiring very large samples and how to overcome them. This simple protocol provides an easy-to-setup and comprehensive tool to study tissue in three dimensions.

The combination of antibody labeling, optical clearing, and advanced light microscopy allows three-dimensional analysis of complete structures or organs. Described here is a simple method to combine immunolabeling of thick kidney slices, optical clearing with ethyl cinnamate, and confocal imaging that enables visualization and quantification of three-dimensional kidney structures.

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) ...

Digital Home-Monitoring of Patients after Kidney Transplantation: The MACCS Platform

The MACCS (Medical Assistant for Chronic Care Service) platform enables secure sharing of key medical information between patients after kidney transplantation and physicians. Patients provide information such as vital signs, well-being, and medication intake via smartphone apps. The information is transferred directly into a database and electronic health record at the kidney transplant center, which is used for routine patient care and research. Physicians can send an updated medication plan and laboratory data directly to the patient app via this secure platform. Other features of the app are medical messages and video consultations. Consequently, the patient is better-informed, and self-management is facilitated. In addition, the transplant center and the patient's local nephrologist automatically exchange notes, medical reports, laboratory values, and medication data via the platform. A telemedicine team reviews all incoming data on a dashboard and takes action, if necessary. Tools to identify patients at risk for complications are under development. The platform exchanges data via a standardized secure interface (Health Level 7 (HL7), Fast Healthcare Interoperability Resources (FHIR)). The standardized data exchange based on HL7 FHIR guarantees interoperability with other eHealth solutions and allows rapid scalability to other chronic diseases. The underlying data protection concept is in concordance with the latest European General Data Protection Regulation. Enrollment started in February 2020, and 131 kidney transplant recipients are actively participating as of July 2020. Two large German health insurance companies are currently funding the telemedicine services of the project. The deployment for other chronic kidney diseases and solid organ transplant recipients is planned. In conclusion, the platform is designed to enable home monitoring and automatic data exchange, empower patients, reduce hospitalizations, and improve adherence, and outcomes after kidney transplantation.

The MACCS platform&#160;is a comprehensive telemedicine concept aiming at better outcomes after kidney transplantation by sharing key medical information between patients and physicians. A telemedicine team reviews incoming data to detect potential complications and to improve adherence in kidney transplant recipients to achieve better long-term outcomes.

The MACCS (Medical Assistant for Chronic Care Service) platform enables secure sharing of key medical information between patients after kidney ...

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 ...