We thank Dr. Tiantian Bai team for help with voice over, Miss Mian Pao for her help in medical illustration. This work was supported in part by the German Research Foundation (DFG) to promote international collaborations (HO2581/4-1 to AH) and the National Science Foundation of China (NSFC; #81760291 to FJ).

All animal experiments were conducted according to the guidelines from the directive 2010/63/EU of the European Parliament on protection of animals used for scientific purposes (Animal ethics card: Lower Saxony Ministry of Food and Drug Safety, #33.9-42502-04-11/0492). Conduct procedures using sterile surgical instruments and consumables (autoclaved) and try to keep the operating area as sterile as possible.
NOTE: C57BL/6J male mice served as donors and recipients (syngeneic transplant model) while Balb/c mice served as kidney allograft recipients (model for studying acute allograft rejection model9). Mice were aged between 8-12 weeks, weighed ~25-30 g at transplantation and were housed under standard conditions. Data reported in this manuscript were generated by four surgeons experienced in mice surgery.
1. Preparatory steps
<ol>
	<li>For surgery, use a set of microscopic instruments, including a micro-scissor, micro-forceps, a needle holder, micro hemostatic clamps, and an electrosurgical pen. Perform sutures using 7/0er, 10/0er, or 4/0er nylon monofilament.</li>
	<li>For anesthesia, place the mouse into the box for inhalation of isoflurane (2%) for about 40-60 s in order to induce unconsciousness.</li>
	<li>Once the mouse is anesthetized, weigh the mouse.</li>
	<li>According to the mouse&#180;s weight, apply an intraperitoneal injection of ketamine (100 mg/kg) + xylazine (10 mg/kg) + acepromazine (2 mg/kg) to anesthetize the mouse13. Confirm that the mouse is anesthetized by observing a lack of response to a toe pinch.</li>
	<li>When anesthesia has taken effect, clip the abdominal fur. Then, fix the mouse on the operation table by loosely immobilizing the limbs with a sterile masking tape.</li>
	<li>Disinfect the mouse&#39;s abdomen after placing the mouse on the operation table. Perform disinfection using alternating scrub of povidone iodide (iodophor) and alcohol, three times (use concentric pattern, start scrub in the middle of the abdomen and move outward), and then properly drape the mouse using a fenestrated surgical towel.</li>
	<li>Apply eye ointment and maintain sterility throughout the procedure. 
	&#8203;NOTE: Antibiotics are not recommended throughout the procedure as these substances may influence immunological responses.</li>
</ol>
2. Donor operation procedure
<ol>
	<li>Use scissors to cut the skin and perform a cross abdominal incision of about 3-4 cm. Cut the muscles of the abdominal wall. Cover and cautiously move away the viscera with a saline imbibed gauze.</li>
	<li>Use a cotton swab to gently remove the intestines, stomach, and spleen toward the right side (from point of view of the mouse), cover and cautiously move away the viscera with a saline imbibed gauze.</li>
	<li>Use micro forceps to expose the left kidney, aorta, and inferior vena cava (IVC).</li>
	<li>Use an electrosurgical pencil to cauterize the left lumbar veins, including their underlying branches and other small vessels along with the left adrenal vessel, carefully.</li>
	<li>Use micro scissors and forceps to dissect the left ureter and cautiously mobilize it from the surrounding tissue. Clean cut it close to the urinary bladder. Mobilize the aortic region between the left and right renal arteries approximately 2 mm in length.</li>
	<li>Use micro forceps to separate the infrarenal inferior vena cava (IVC) and aorta, and then use curved forceps to pass under the aorta to place a loose tie of 7-0 silk suture around this vessel.</li>
	<li>Cross clamp the area of the aorta below the right renal artery and the inferior vena cava (IVC) using two 5 mm microvascular clamps.</li>
	<li>Transect the left renal vein from the vena cava.</li>
	<li>Use a syringe to flush the aorta with 1 mL of heparin saline solution (60 U/mL).</li>
	<li>Use micro forceps to tighten the ligature applied at step 2.5. Then, cut the aorta below the ligature as well as below the proximal clamp. With this, the proximal bifurcations of the renal artery (please note that the arterial opening must be cut neatly, otherwise it will affect the anastomosis) and the ureteral artery are included and transected en bloc. Prepare carefully, so that the delicate ureteral artery is completely preserved.</li>
	<li>Use the electrosurgical pencil and forceps to free the left kidney and associated vessels completely by cautiously cauterizing all vessel surrounding tissue. Remove the kidney and store it in saline solution at 4 &#176;C.</li>
	<li>Euthanize the anesthetized donor mouse by decapitation.</li>
</ol>
3. Recipient operation procedure
<ol>
	<li>Perform the initial surgical steps (including anesthesia and sterilization, see steps 1.1 to 1.7) as described for the donor mouse.</li>
	<li>Use scissors to open the abdomen via a median incision (about 2.5 cm in length), and then cover the abdominal organs with a wet gauze using saline solution.</li>
	<li>Carefully preserve the infrarenal aorta and inferior vena cava (IVC) and make sure every large vessel branch is cauterized. Use the electric cautery as well to dissect the left ureter carefully at a position proximal to the kidney pelvis. Then, remove the left kidney.</li>
	<li>Use micro forceps and cotton buds to expose the abdominal aorta and inferior vena cava and detach them from the surrounding adipose tissue (approximately over 4 mm in length).</li>
	<li>Use two microvascular clamps and position them proximally and distally on both the inferior vena cava and the abdominal aorta simultaneously.</li>
	<li>Use a micro needle holder to guide a 10/0 monofilament (made of synthetic fiber with a smooth surface) suture needle, which is placed through the aorta wall in a proximal to distal manner.</li>
	<li>Achieve an elliptical arteriotomy of approximately 1 mm with a gentle upward traction of the suture, while cutting directly below the lower face of the needle with fine, curved scissors.</li>
	<li>Use micro scissors to cut the inferior vena cava (IVC) longitudinally with sufficient length of approximately 1.5 mm. Position this incision slightly below its aortic counterpart.</li>
	<li>Perform the donor and recipient aorta anastomosis in an end-to-side manner. Place the donor kidney on the right side of the recipient&#39;s inferior vena cava aligning the cuff of the donor&#39;s abdominal aorta with the anastomosis of the recipient&#39;s abdominal aorta.</li>
	<li>Use a micro needle holder and two separate 10-0 sutures to stitch the proximal and distal ends of the anastomosis.</li>
	<li>After tying, leave the two long sutures, including the needle, in place. Sew the left side of the aortal wall of the anastomosis continuously with two evenly spaced stitches in a distal-proximal direction.</li>
	<li>After the last stitch, guide the suture through a partial thickness of the vessel wall above the upper stay suture tie.</li>
	<li>Use micro forceps to simultaneously exert gentle traction to the short end of the lower suture tie. 
	NOTE: In this new knotless technique, the last stitch is not tied to the short end of the upper tie.</li>
	<li>Use micro forceps to turn over the transplanted kidney to its normal position. Now continuously sew the right wall of the aortal anastomosis using three stitches in a proximal to distal manner. 
	NOTE: Compared with the conventional surgical technique7,8 the last suture is merged with the distal tie nearby. Do not tie it to the end of the lower suture, cut it to leave a free length of 2-3 mm instead.</li>
	<li>Perform the venous anastomosis using the same suturing procedure as previously described with the difference that four to five stitches are needed for each side of the anastomosis. The final stitch is left as a free end of similar length similar to the aortal anastomosis described above.</li>
	<li>After completing both anastomoses, use a dry swab to exert gentle pressure toward the sutured area for about 10-20 s.</li>
	<li>Use a clip applicator forceps to remove both clamps, first the lower then the upper. Rinse the abdominal cavity with 0.9% sodium chloride at a temperature of 38.0 &#176;C.</li>
	<li>Observe the reperfusion of the transplanted kidney.</li>
</ol>
4. Ureteral implantation
<ol>
	<li>Use a micro needle holder to penetrate through the recipient&#39;s urine bladder with a 10/0 suture (straight needle) and insert it into a 21 G needle lumen for guidance (see Supplementary Figure 1a).</li>
	<li>Now guide the 21 G needle to stitch a hole at the place of the previous needle application (Supplementary Figure 1b).</li>
	<li>Pull out the 21 G needle (Supplementary Figure 1c).</li>
	<li>Use a micro needle holder and 10/0 suture to stitch (no tie) the trimmed ureter end and perforate the bladder with this 10/0 suture again at the place of its entry (Supplementary Figure 1d).</li>
	<li>Use a micro needle holder to tow the 10/0 filament and the ureter into the urine bladder through the constructed hole (Supplementary Figure 1e).</li>
	<li>Use a micro needle holder and another 10/0 suture to anastomose the donor&#39;s ureter to the recipient&#39;s urine bladder. Here, connect the outer membrane of the ureter to the outer membrane of the bladder wall, and perform intermittent sutures with 3 to 4 stitches. Finally, pull out the traction suture (Supplementary Figure 1f).</li>
	<li>Use forceps to place the intestines back into the abdominal cavity. Perform two-layer sutures (first the abdominal muscles followed by the skin) to close the abdominal wound using a 4/0 filament.</li>
	<li>Place the transplanted mice into an oxygen and temperature-controlled chamber for recovering after surgery.</li>
	<li>For postoperative analgesia, directly give Metamizol 200 mg/kg per os after operation. 
	Four and 16 h after operation give Metamizol 200 mg/kg per os plus Carprofen (5mg/kg) s.c. In the further follow up, apply Carprofen (5 mg/kg) s.c. to the transplanted mice every 24 hours for three consecutive days after operation13. If there are any signs of an insufficient analgesia buprenorphine 0.05 mg/kg is additionally given every 8 h s.c.</li>
</ol>
5. Contralateral nephrectomy and sacrifice of the recipient mouse
NOTE: Perform contralateral nephrectomy of the recipient mouse 5 days after transplantation.
<ol>
	<li>Perform the contralateral nephrectomy of the transplanted mouse 5 days after transplantation under anesthesia. Ligate and cut the recipient&#39;s autologous right renal arteries and veins, remove the right kidney and close the abdominal cavity. The postoperative care and analgesia are the same as described before (see step 4.7).</li>
	<li>Raise and record the state of the mouse. Provide the transplanted mouse postoperative analgesia, food, and water supply.</li>
	<li>Four weeks after transplantation, sacrifice half of the transplanted mice and perform H&#38;E staining for their kidney transplants.</li>
	<li>12 weeks after transplantation, sacrifice the remaining mice and perform Masson Gold staining of these kidney transplants.</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> Transplantation Proceedings. 5 (1), 721-725 (1973).</li><li>Jiang, K., 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=Noninvasive+assessment+of+renal+fibrosis+with+magnetization+transfer+MR+imaging:+Validation+and+evaluation+in+murine+renal+artery+stenosis.">Noninvasive assessment of renal fibrosis with magnetization transfer MR imaging: Validation and evaluation in murine renal artery stenosis.</a> Radiology. 283 (1), 77-86 (2017).</li><li>Tse, G. H., 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=Mouse+kidney+transplantation:+Models+of+allograft+rejection.">Mouse kidney transplantation: Models of allograft rejection.</a> Journal of Visualized Experiments. (92), e52163 (2014).</li><li>Okazaki, M., 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=et+al.Costimulatory+blockade-mediated+lung+allograft+acceptance+is+abrogated+by+overexpression+of+Bcl-2+in+the+recipient.">et al.Costimulatory blockade-mediated lung allograft acceptance is abrogated by overexpression of Bcl-2 in the recipient.</a> Transplantation Proceedings. 41 (1), 385-387 (2009).</li><li>Chuck, N. 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=et+al.Ultra-short+echo-time+magnetic+resonance+imaging+distinguishes+ischemia/reperfusion+injury+from+acute+rejection+in+a+mouse+lung+transplantation+model.">et al.Ultra-short echo-time magnetic resonance imaging distinguishes ischemia/reperfusion injury from acute rejection in a mouse lung transplantation model.</a> Transplant International. 29 (1), 108-118 (2016).</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 (9), 1267-1272 (1996).</li><li>Wang, J., Hockenheimer, S., Bickerstaff, A. A., Hadley, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+renal+transplantation+procedure.">Murine renal transplantation procedure.</a> Journal of Visualized Experiments. (29), e1150 (2009).</li><li>Plenter, R., Jain, S., Ruller, C. M., Nydam, T. L., Jani, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+kidney+transplant+technique.">Murine kidney transplant technique.</a> Journal of Visualized Experiments. (104), e52848 (2015).</li><li>Plenter, R. J., Jain, S., Nydam, T. L., Jani, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Revised+arterial+anastomosis+for+improving+murine+kidney+transplant+outcomes.">Revised arterial anastomosis for improving murine kidney transplant outcomes.</a> Journal of Investigative Surgery. 28 (4), 208-214 (2015).</li><li>Rong, S., Lewis, A. G., Kunter, U., Haller, H., Gueler, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+knotless+technique+for+kidney+transplantation+in+the+mouse.">A knotless technique for kidney transplantation in the mouse.</a> Journal of Transplantation. , 127215 (2012).</li><li>Kreimann, K., 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=Ischemia+reperfusion+injury+triggers+CXCL13+release+and+B-cell+recruitment+after+allogenic+kidney+transplantation.">Ischemia reperfusion injury triggers CXCL13 release and B-cell recruitment after allogenic kidney transplantation.</a> Frontiers in Immunology. 11, 1204 (2020).</li><li>Schmidbauer, M., 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=Diffusion-Weighted+imaging+and+mapping+of+T1+and+T2+relaxation+time+for+evaluation+of+chronic+renal+allograft+rejection+in+a+translational+mouse+model.">Diffusion-Weighted imaging and mapping of T1 and T2 relaxation time for evaluation of chronic renal allograft rejection in a translational mouse model.</a> Journal of Clinical Medicine. 10 (19), 4318 (2021).</li><li>Wu, K., 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=Novel+technique+for+blood+circuit+reconstruction+in+mouse+heart+transplantation+model.">Novel technique for blood circuit reconstruction in mouse heart transplantation model.</a> Microsurgery. 26, 594-598 (2006).</li><li>Haas, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+allograft+nephropathy+or+interstitial+fibrosis+and+tubular+atrophy:+what+is+in+a+name.">Chronic allograft nephropathy or interstitial fibrosis and tubular atrophy: what is in a name.</a> Current Opinion in Nephrology and Hypertension. 23 (3), 245-250 (2014).</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 (5), 477-484 (2012).</li><li>Yin, D., 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=Blood+circuit+reconstruction+in+an+abdominal+mouse+heart+transplantation+model.">Blood circuit reconstruction in an abdominal mouse heart transplantation model.</a> Journal of Visualized Experiments. (172), e62007 (2021).</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=Improved+techniques+for+kidney+transplantation+in+mice.">Improved techniques for kidney transplantation in mice.</a> Microsurgery. 16 (2), 103-109 (1995).</li><li>Mannon, R. B., 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=Chronic+rejection+of+mouse+kidney+allografts.">Chronic rejection of mouse kidney allografts.</a> Kidney International. 55 (5), 1935-1944 (1999).</li><li>Coffman, 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=Improved+renal+function+in+mouse+kidney+allografts+lacking+MHC+class+I+antigens.">Improved renal function in mouse kidney allografts lacking MHC class I antigens.</a> Journal of Immunology. 151 (1), 425-435 (1993).</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></ol>

While the skin transplantation model in mice is simple and easy to perform to study alloimmune rejection events, the surgical techniques for investigating more specifically the alloimmune-related inflammatory alterations after heart16 and kidney transplantation10 has been proven to be complex and very demanding. From the transplant nephrologist&#39;s point of view, the establishment of an effective and long-term stable mouse renal transplantation model has still an irreplac...

Since Sakowitz et al. developed mouse models of kidney transplantation in 1973 for the first time1, it has proven as an important experimental tool to study the mechanisms of transplant ischemic injury and alloimmune rejection as well as for developing new treatments aimed to prolong allograft survival and possibly to achieve immunological tolerance. However, the surgical technique has proven to be complex and very demanding, sometimes having complications such as vascular anastomotic strictures leading to prerenal non-immunological kidney transplant failure2, postrenal failure caused by ischemia and subsequent necrosis of the transplanted ureter, strictures of the anastomosis of the transplanted ureter and/or the recipient&#39;s urine bladder leading to a disruption of the urinary outflow. All of these are reasons why renal transplantation in mice has not been further developed and is therefore not widely used. Establishing an effective and long-term stable mouse kidney transplantation model without vascular and urinary tract complications still has irreplaceable significance for many studies in the transplant field with focus on the renal immune mediated but also infectious diseases3. In addition, compared with other organ transplants in murine models such as lung, heart, and intestinal transplantation4,5, the mouse kidney transplantation model offers a chance for studying long-term survival even in the setting of major histocompatibility antigen disparity3,6. It has also been shown that in the same setting of donor-recipient strain combinations different organ transplants such as heart or kidney are characterized by different dynamics and onsets of allograft rejection3. Furthermore, from the nephrological point of view, it is a more suitable model for studying parenchymal mediated immune regulatory mechanisms in the context of acute and chronic rejection events than simple skin transplant experiments.
On the basis of previous reports on the surgical technique of kidney transplantation in mice3,7,8,9, we here demonstrate the following reliable improvements that have been successfully applied during the past 10 years within our group10,11,12: Firstly, the ureteral artery is safely conserved as the renal artery is resected en bloc together with the respective part of the abdominal aorta. Second, a new, simple, and rapid technique of a knotless vascular anastomosis in which the final stitch of anastomosis is not tied with the end of the upper tie like the traditional approach but remains free. This technique enables to increase or decrease the size of the anastomosis after renal reperfusion to avoid vessel stricture and intraabdominal bleeding. Third, 21 G and 30 G syringe needles were used as an auxiliary puncture guiding tool in order to implant the donor ureter into the recipient&#39;s bladder wall reducing the damage to the recipient&#39;s bladder and facilitating the formation of stricture free anastomosis.
In this report, we also compared the traditional, widely used technique with the modified one that is established in our laboratory and found no significant difference in the degree of renal tubular atrophy and kidney transplant interstitial tissue fibrosis. In previous studies, we additionally compared the results of this new technique with the conventional method in terms of local bleeding, thrombosis, time for performing the vessel anastomosis and survival rate. We found improvements such as significant reductions of local thrombosis events (1.1% versus 6.6%), a reduced time for the anastomosis procedure, and a highly reproducible kidney syngeneic graft long-term survival (95% versus 84% with the classical approach)10.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>30G-needles</td>
			<td>Braun</td>
			<td>456300</td>
			<td>-</td>
		</tr>
		<tr>
			<td>acepromazine</td>
			<td>CP Pharma</td>
			<td>Tranquisol P</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Bepanthen eye ointment</td>
			<td>Haus-Apotheke</td>
			<td>PZN 01578675</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Bonn Micro Forceps</td>
			<td>FST</td>
			<td>11083-07</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Box for insulation and oxygen supply device</td>
			<td>RUSKINN</td>
			<td>INVIV</td>
			<td>-</td>
		</tr>
		<tr>
			<td>C57BL/6J&#160; mice</td>
			<td>Charles River. Germany</td>
			<td>no catalog number</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Carprofen</td>
			<td>Zoetis</td>
			<td>Rimadyl 50 mg/ml</td>
			<td>-</td>
		</tr>
		<tr>
			<td>CATHETER-FEP 26G</td>
			<td>TERUMO</td>
			<td>Surflo-W</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Clip Applicator Forceps Style</td>
			<td>FST</td>
			<td>18057-14</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Curved forceps</td>
			<td>WPI</td>
			<td>14114-G</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Cutasept skin disinfection</td>
			<td>VWR</td>
			<td>BODL980365</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Dehydrator</td>
			<td>DIAPATH</td>
			<td>Donatello</td>
			<td>-</td>
		</tr>
		<tr>
			<td>electrosurgical pen</td>
			<td>Bovie</td>
			<td>CHANGE-A-TIP</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Embedding machine</td>
			<td>Wuhan Junjie Electronics Co., Ltd</td>
			<td>JB-P5</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Sinopharm Group Chemical Reagent Co. LtD</td>
			<td>100092683</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Frozen platform</td>
			<td>Wuhan Junjie Electronics Co., Ltd</td>
			<td>JB-L5</td>
			<td>-</td>
		</tr>
		<tr>
			<td>gauze pads, cotton swabs</td>
			<td>Lohmann-Rauscher</td>
			<td>13353</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Glass slide</td>
			<td>Servicebio</td>
			<td>G6004</td>
			<td>-</td>
		</tr>
		<tr>
			<td>HE dye solution set</td>
			<td>Servicebio</td>
			<td>G1003</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Heating mat</td>
			<td>THERMO MAT PRO 30W</td>
			<td>HTP-30</td>
			<td>-</td>
		</tr>
		<tr>
			<td>hemostatic sponge</td>
			<td>CuraSpon</td>
			<td>J1276A</td>
			<td>-</td>
		</tr>
		<tr>
			<td>heparine-solution</td>
			<td>Haus-Apotheke</td>
			<td>PZN 03029820</td>
			<td>-</td>
		</tr>
		<tr>
			<td>ice box</td>
			<td>PETZ</td>
			<td>No Catalog Number available</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Imaging system</td>
			<td>Nikon</td>
			<td>Nikon DS-U3</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Inhalation anesthesia device</td>
			<td>GROPPLER</td>
			<td>BKGM 0616</td>
			<td>-</td>
		</tr>
		<tr>
			<td>isoflurane</td>
			<td>CP Pharma</td>
			<td>Isofluran CP 1 ml/ml</td>
			<td>-</td>
		</tr>
		<tr>
			<td>ketamine</td>
			<td>Zoetis</td>
			<td>no catalog numer</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Masson dye solution set</td>
			<td>Servicebio</td>
			<td>G1006</td>
			<td>-</td>
		</tr>
		<tr>
			<td>metamizole</td>
			<td>WDT</td>
			<td>no catalog numer</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Micro scissors</td>
			<td>FST</td>
			<td>15000-00,15000-10</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Micro Serrefine ( Clamp ) Angled / 16 mm</td>
			<td>FST</td>
			<td>18055-06</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Leica</td>
			<td>LEICAMZ6</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Microscope light</td>
			<td>SCHOTT</td>
			<td>KL2500LED</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Neutral gum</td>
			<td>SCRC</td>
			<td>10004160</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Oven</td>
			<td>Tianjin Laibo Rui Instrument Equipment Co., Ltd</td>
			<td>GFL-230</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Pathology slicer</td>
			<td>Shanghai Leica Instrument Co., Ltd</td>
			<td>RM2016</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Saline solution (NaCl 0.9 %)</td>
			<td>Haus-Apotheke</td>
			<td>PZN 06178437</td>
			<td>-</td>
		</tr>
		<tr>
			<td>scissors</td>
			<td>Peha Instruments</td>
			<td>991083/4</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Slides</td>
			<td>Servicebio</td>
			<td></td>
			<td>-</td>
		</tr>
		<tr>
			<td>small Petri dish</td>
			<td>Sarstedt</td>
			<td>8,33,900</td>
			<td>-</td>
		</tr>
		<tr>
			<td>straight forceps</td>
			<td>WPI</td>
			<td>14113-G</td>
			<td>-</td>
		</tr>
		<tr>
			<td>surgical tape</td>
			<td>BSN</td>
			<td>4120</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Suture Tying Forceps - 10 cm</td>
			<td>FST</td>
			<td>18025-10</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Sutures(10-0)</td>
			<td>Medtronic</td>
			<td>N2540</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Sutures(4-0)</td>
			<td>ETHILON</td>
			<td>V4940H</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Sutures(7-0)</td>
			<td>ETHILON</td>
			<td>1647H</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Syringe (0,3 mL)</td>
			<td>BD</td>
			<td>324826</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Syringe (1 mL)</td>
			<td>BD</td>
			<td>320801</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Tissue spreader</td>
			<td>Zhejiang Kehua Instrument Co., Ltd</td>
			<td>KD-P</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Upright optical microscope</td>
			<td>Nikon</td>
			<td>Nikon Eclipse E100</td>
			<td>-</td>
		</tr>
		<tr>
			<td>xylazine</td>
			<td>Bayer</td>
			<td>Rompun</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Xylene</td>
			<td>Sinopharm Group Chemical Reagent Co. LtD</td>
			<td>10023418</td>
			<td>-</td>
		</tr>
	</tbody>
</table>

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.

Four weeks after transplantation, both the modified technique as well as the conventional technique displayed moderate signs of renal tubular atrophy14,15 when compared to the native recipient contralateral kidneys (Figure 1). The degree of the renal tubules atrophy demonstrated no significant difference between the two different techniques. Masson Goldner&#39;s trichrome staining14,<sup class="xref...

Watch this Scientific Journal Video about A Modified Surgical Technique for Kidney Transplantation in Mice at JoVE.com

A Modified Surgical Technique for Kidney Transplantation in 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 ...

a-modified-surgical-technique-for-kidney-transplantation-in-mice

Research

JoVE Journal

Immunology and Infection

5.5K Views.  Friedrich-Alexander University Erlangen-Nürnberg. 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.

Video: A Modified Surgical Technique for Kidney Transplantation in Mice

A Method for Labeling Vasculature in Embryonic Mice

The establishment of a functional blood vessel network is an essential part of organogenesis, and is required for optimal organ function. For example, in the thymus proper vasculature formation and patterning is essential for thymocyte entry into the organ and mature T-cell exit to the periphery. The spatial arrangement of blood vessels in the thymus is dependent upon signals from the local microenvironment, namely thymic epithelial cells (TEC). Several recent reports suggest that disruption of these signals results in thymus blood vessel defects 1,2. Previous studies have described techniques used to label the neonatal and adult thymus vasculature 1,2. We demonstrate here a technique for labeling blood vessels in the embryonic thymus. This method combines the use of FITC-dextran or Griffonia (Bandeiraea) Simplicifolia Lectin I (GSL 1 - isolectin B4) facial vein injections and CD31 antibody staining to identify thymus vascular structures and PDGFR-&beta; to label thymic perivascular mesenchyme 3-5. The option of using cryosections or vibratome sections is also provided. This protocol can be used to identify thymus vascular defects, which is critical for defining the roles of TEC-derived molecules in thymus blood vessel formation. As the method labels the entire vasculature, it can also be used to analyze the vascular networks in multiple organs and tissues throughout the embryo including skin and heart 6-10.

This article describes a method for labeling embryonic skin and thymus blood vessels.

The establishment of a functional blood vessel network is an essential part of organogenesis, and is required for optimal organ function. For example, in ...

4D Multimodality Imaging of Citrobacter rodentium Infections in Mice

This protocol outlines the steps required to longitudinally monitor a bioluminescent bacterial infection using composite 3D diffuse light imaging tomography with integrated &mu;CT (DLIT-&mu;CT) and the subsequent use of this data to generate a four dimensional (4D) movie of the infection cycle. To develop the 4D infection movies and to validate the DLIT-&mu;CT imaging for bacterial infection studies using an IVIS Spectrum CT, we used infection with bioluminescent C. rodentium, which causes self-limiting colitis in mice. In this protocol, we outline the infection of mice with bioluminescent C. rodentium and non-invasive monitoring of colonization by daily DLIT-&mu;CT imaging and bacterial enumeration from feces for 8 days.
The use of the IVIS Spectrum CT facilitates seamless co-registration of optical and &mu;CT scans using a single imaging platform. The low dose &mu;CT modality enables the imaging of mice at multiple time points during infection, providing detailed anatomical localization of bioluminescent bacterial foci in 3D without causing artifacts from the cumulative radiation. Importantly, the 4D movies of infected mice provide a powerful analytical tool to monitor bacterial colonization dynamics in vivo.

4D Multimodality Imaging of Citrobacter rodentium Infections in Mice

Multi-modality imaging is a valuable approach for studying bacterial colonization in small animal models. This protocol outlines infection of mice with bioluminescent Citrobacter rodentium and the longitudinal monitoring of bacterial colonization using composite 3D diffuse light imaging tomography with &mu;CT imaging to create a 4D movie of C. rodentium infection.

This protocol outlines the steps required to  longitudinally monitor a bioluminescent bacterial infection using composite 3D diffuse light imaging ...

Long Term Chronic Pseudomonas aeruginosa Airway Infection in Mice

A mouse model of chronic airway infection is a key asset in cystic fibrosis (CF) research, although there are a number of concerns regarding the model itself. Early phases of inflammation and infection have been widely studied by using the Pseudomonas aeruginosa agar-beads mouse model, while only few reports have focused on the long-term chronic infection in vivo. The main challenge for long term chronic infection remains the low bacterial burden by P. aeruginosa and the low percentage of infected mice weeks after challenge, indicating that bacterial cells are progressively cleared by the host.
This paper presents a method for obtaining efficient long-term chronic infection in mice. This method is based on the embedding of the P. aeruginosa clinical strains in the agar-beads in vitro, followed by intratracheal instillation in C57Bl/6NCrl mice. Bilateral lung infection is associated with several measurable read-outs including weight loss, mortality, chronic infection, and inflammatory response. The P. aeruginosa RP73 clinical strain was preferred over the PAO1 reference laboratory strain since it resulted in a comparatively lower mortality, more severe lesions, and higher chronic infection. P. aeruginosa colonization may persist in the lung for over three months. Murine lung pathology resembles that of CF patients with advanced chronic pulmonary disease.
This murine model most closely mimics the course of the human disease and can be used both for studies on the pathogenesis and for the evaluation of novel therapies.

Long Term Chronic Pseudomonas aeruginosa Airway Infection in Mice

We describe the agar-beads method to establish persistent long-term chronic Pseudomonas aeruginosa airway infection in the mouse model.

A mouse model of chronic airway infection is a key asset in cystic fibrosis (CF) research, although there are a number of concerns regarding the model ...

Transplantation of Tail Skin to Study Allogeneic CD4 T Cell Responses in Mice

The study of T cell responses and their consequences during allo-antigen recognition requires a model that enables one to distinguish between donor and host T cells, to easily monitor the graft, and to adapt the system in order to answer different immunological questions. Medawar and colleagues established allogeneic tail-skin transplantation in mice in 1955. Since then, the skin transplantation model has been continuously modified and adapted to answer specific questions. The use of tail-skin renders this model easy to score for graft rejection, requires neither extensive preparation nor deep anesthesia, is applicable to animals of all genetic background, discourages ischemic necrosis, and permits chemical and biological intervention. 
In general, both CD4+ and CD8+ allogeneic T cells are responsible for the rejection of allografts since they recognize mismatched major histocompatibility antigens from different mouse strains. Several models have been described for activating allogeneic T cells in skin-transplanted mice. The identification of major histocompatibility complex (MHC) class I and II molecules in different mouse strains including C57BL/6 mice was an important step toward understanding and studying T cell-mediated alloresponses. In the tail-skin transplantation model described here, a three-point mutation (I-Abm12) in the antigen-presenting groove of the MHC-class II (I-Ab) molecule is sufficient to induce strong allogeneic CD4+ T cell activation in C57BL/6 mice. Skin grafts from I-Abm12 mice on C57BL/6 mice are rejected within 12-15 days, while syngeneic grafts are accepted for up to 100 days. The absence of T cells (CD3-/- and Rag2-/- mice) allows skin graft acceptance up to 100 days, which can be overcome by transferring 2 x 104 wild type or transgenic T cells. Adoptively transferred T cells proliferate and produce IFN-&#947; in I-Abm12-transplanted Rag2-/- mice.

Tail-skin transplantation is a powerful model for studying T cell-dependent rejection and tolerance induction during allogeneic immune responses in mice. The advantages of this protocol are minor invasive surgery, and ease of monitoring with no need to sacrifice the recipient mouse.

The study of T cell responses and their consequences during allo-antigen recognition requires a model that enables one to distinguish between donor and ...

Induction of Paralysis and Visual System Injury in Mice by T Cells Specific for Neuromyelitis Optica Autoantigen Aquaporin-4

While it is recognized that aquaporin-4 (AQP4)-specific T cells and antibodies participate in the pathogenesis of neuromyelitis optica (NMO), a human central nervous system (CNS) autoimmune demyelinating disease, creation of an AQP4-targeted model with both clinical and histologic manifestations of CNS autoimmunity has proven challenging. Immunization of wild-type (WT) mice with AQP4 peptides elicited T cell proliferation, although those T cells could not transfer disease to na&#239;ve recipient mice. Recently, two novel AQP4 T cell epitopes, peptide (p) 135-153 and p201-220, were identified when studying immune responses to AQP4 in AQP4-deficient (AQP4-/-) mice, suggesting T cell reactivity to these epitopes is normally controlled by thymic negative selection. AQP4-/- Th17 polarized T cells primed to either p135-153 or p201-220 induced paralysis in recipient WT mice, that was associated with predominantly leptomeningeal inflammation of the spinal cord and optic nerves. Inflammation surrounding optic nerves and involvement of the inner retinal layers (IRL) were manifested by changes in serial optical coherence tomography (OCT). Here, we illustrate the approaches used to create this new in vivo model of AQP4-targeted CNS autoimmunity (ATCA), which can now be employed to study mechanisms that permit development of pathogenic AQP4-specific T cells and how they may cooperate with B cells in NMO pathogenesis.

Here, we present a protocol to induce paralysis and opticospinal inflammation by transfer of aquaporin-4 (AQP4)-specific T cells from AQP4-/- mice into WT mice. In addition, we demonstrate how to use serial optical coherence tomography to monitor visual system dysfunction.

While it is recognized that aquaporin-4 (AQP4)-specific T cells and antibodies participate in the pathogenesis of neuromyelitis optica (NMO), a human ...

Measuring Deformability and Red Cell Heterogeneity in Blood by Ektacytometry

Decreased red cell deformability is characteristic of several disorders. In some cases, the extent of defective deformability can predict severity of disease or occurrence of serious complications. Ektacytometry uses laser diffraction viscometry to measure the deformability of red blood cells subject to either increasing shear stress or an osmotic gradient at a constant value of applied shear stress. However, direct deformability measurements are difficult to interpret when measuring heterogenous blood that is characterized by the presence of both rigid and deformable red cells. This is due to the inability of rigid cells to properly align in response to shear stress and results in a distorted diffraction pattern marked by an exaggerated decrease in apparent deformability. Measurement of the degree of distortion provides an indicator of the heterogeneity of the erythrocytes in blood. In sickle cell anemia, this is correlated with the percentage of rigid cells, which reflects the hemoglobin concentration and hemoglobin composition of the erythrocytes. In addition to measuring deformability, osmotic gradient ektacytometry provides information about the osmotic fragility and hydration status of erythrocytes. These parameters also reflect the hemoglobin composition of red blood cells from sickle cell patients. Ektacytometry measures deformability in populations of red cells and does not, therefore, provide information on the deformability or mechanical properties of individual erythrocytes. Regardless, the goal of the techniques described herein is to provide a convenient and reliable method for measuring the deformability and cellular heterogeneity of blood. These techniques may be useful for monitoring temporal changes, as well as disease progression and response to therapeutic intervention in several disorders. Sickle cell anemia is one well-characterized example. Other potential disorders where measurements of red cell deformability and/or heterogeneity are of interest include blood storage, diabetes, Plasmodium infection, iron deficiency, and the hemolytic anemias due to membrane defects.

Here we present techniques to measure red cell deformability and cellular heterogeneity by ektacytometry. These techniques are applicable to general investigations of red cell deformability and specific investigations of blood diseases characterized by the presence of both rigid and deformable red cells in circulation, such as sickle cell anemia.

Decreased red cell deformability is characteristic of several disorders. In some cases, the extent of defective deformability can predict severity of ...

Printed Glycan Array: A Sensitive Technique for the Analysis of the Repertoire of Circulating Anti-carbohydrate Antibodies in Small Animals

The repertoire of circulating anti-carbohydrate antibodies of a given individual is often associated with its immunological status. Not only the individual immune condition determines the success in combating internal and external potential threat signals, but also the existence of a particular pattern of circulating anti-glycan antibodies (and their serological level variation) could be a significant marker of the onset and progression of certain pathological conditions. Here, we describe a Printed Glycan Array (PGA)-based methodology that offers the opportunity to measure hundreds of glycan targets with very high sensitivity; using a minimal amount of sample, which is a common restriction present when small animals (rats, mice, hamster, etc.) are used as models to address aspects of human diseases. As a representative example of this approach, we show the results obtained from the analysis of the repertoire of natural anti-glycan antibodies in BALB/c mice. We demonstrate that each BALB/c mouse involved in the study, despite being genetically identical and maintained under the same conditions, develops a particular pattern of natural anti-carbohydrate antibodies. This work claims to expand the use of PGA technology to investigate repertoire (specificities) and the levels of circulating anti-carbohydrates antibodies, both in health and during any pathological condition.

This work shows the potential of printed glycan array (PGA) technology for the analysis of circulating anti-carbohydrate antibodies in small animals.

The repertoire of circulating anti-carbohydrate antibodies of a given individual is often associated with its immunological status. Not only the ...

Renal Subcapsular Transplantation of 2'-Deoxyguanosine-Treated Murine Embryonic Thymus in Nude Mice

The thymus is an important central immune organ, which plays an essential role in the development and differentiation of T cells. Thymus transplantation is an important method for investigating thymic epithelial cell function and T cells maturation in vivo. Here we will describe the experimental methods used within our laboratory to transplant 2&#8217;-deoxyguanosine (to deplete donor&#8217;s lymphocytes) treated embryonic thymus into the renal capsule of an athymic nude mouse. This method is both simple and efficient and does not require special skills or devices. The results obtained via this simple method showed that transplanted thymus can effectively support the recipient&#8217;s T cells production. Additionally, several key points with regards to the protocol will be further elucidated.

We provide a simple and efficient method to transplant 2&#8217;-deoxyguanosine treated E18.5 thymus into the renal capsule of a nude mouse. This method should aide in the study of both thymic epithelial cells function and T cells maturation.

The thymus is an important central immune organ, which plays an essential role in the development and differentiation of T cells. Thymus transplantation ...

An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications have been reported since the first description in 1964. After 30 years of performing heterotopic heart transplantation in rats, we have developed a simplified surgical approach, which can be easily taught and performed without further surgical training or background.
After dissection of the ascending aorta and the pulmonary artery and ligation of superior and inferior caval and pulmonary veins, the donor heart is harvested and subsequently perfused with ice-cold saline solution supplemented with heparin. After clamping and incising the recipient abdominal vessels, the donor ascending aorta and pulmonary artery are anastomosed to the recipient abdominal aorta and inferior vena cava, respectively, using continuous running sutures.
Depending on different donor-recipient combinations, this model allows analyses of either acute or chronic rejection of allografts. The immunological significance of this model is further enhanced by a novel approach of in-ear injection of vital cardiac muscle cells and subsequent analysis of draining cervical lymphatic tissue.

We describe a model of heterotopic abdominal heart transplantation in rats, implying modifications of current strategies, which lead to a simplified surgical approach. Additionally, we describe a novel rejection model by in-ear injection of vital cardiac muscle cells, allowing further transplant immunological analyses in rats.

Heterotopic heart transplantation in rats has been a commonly used model for diverse immunological studies for more than 50 years. Several modifications ...

Optimization of the Cuff Technique for Murine Heart Transplantation

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to improve surgical efficiency. In our lab, we have optimized the cuff technique with two major steps. First, we used the inner tube technique to insert a temporary inner tube into the external jugular vein and carotid artery blood vessel to facilitate eversion of the vessel over the cuff. Second, we performed complete heterotopic cardiac transplantation through the collaboration of two experienced surgeons. These modifications effectively reduced the operation time to 25 minutes, with a success rate of 95%. In this report, we describe these procedures in detail and provide a supplemental video. We believe that this report on the improved cuff technique will offer practical guidance for murine heterotopic heart transplantation and will enhance the utility of this mouse model for basic research.

We introduce an inner tube approach to the cuff technique for mouse cervical heterotopic heart transplantation to help evert the vessel over the cuff. We found that cooperation between two experienced surgeons markedly shortens the operation time.

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to ...