The authors would like to acknowledge the superb efforts of the animal care staff of the University of Western Australia and of the Harry Perkins Institute of Medical Research, whose dedication and expertise contributed to the feasibility and success of these surgeries.

This research was performed in accordance with the Code of Practice for the Care and Use of Animals for Scientific Purposes15 and approved under Animal Ethics Protocols RA/3/100/1568 and AE173 (The University of Western Australia Animal Ethics Committee and The Harry Perkins Institute of Medical Research Animal Ethics Committee, respectively). See the Table of Materials for details regarding all materials, instruments, and animals used in this protocol.
1. Preparation of the animal for surgery
NOTE: Personnel are dedicated to either the role of performing surgery or monitoring anesthesia throughout the procedure.
<ol>
	<li>For preoperative analgesia, administer a dose of buprenorphine (0.05-0.1 mg/kg, diluted to 0.03 mg/mL with sodium chloride 0.9%) subcutaneously to the recipient mouse at least 1 h prior to the initiation of recipient surgery. Enter all details related to the administration of drugs, their dose, the time of administration, and their effects in the anesthetic record. 
	NOTE: This approach is not necessarily required for the donor as it is a non-recovery surgery, where the donor is euthanized under general anesthetic immediately after harvest of the organ.</li>
	<li>Induction of anesthesia
	<ol>
		<li>Place the mouse in the induction chamber of the anesthetic breathing system with an oxygen flow of 1-2 L&#183;min&#8722;1 with 4% isoflurane. Confirm adequate anesthesia by observing recumbency, loss of the righting reflex, and a decreased respiratory rate.</li>
		<li>Once adequately anesthetized, remove the mouse from the induction chamber and closely shave the ventral abdomen using clippers to remove hair. In the case of the donor, shave the area extending from the genitalia to the upper margin of the ventral thorax. In the case of the recipient, shave the area extending from the genitalia to the costal margin. In both cases, ensure that the area shaved reaches the midaxillary line laterally.</li>
	</ol>
	</li>
	<li>To maintain anesthesia, position the mouse in dorsal recumbency to receive anesthetic and oxygen from the nose cone of the breathing system (non-rebreathing), which delivers oxygen at a rate of 1 L&#183;min&#8722;1 and isoflurane (1.5%-2.5%). 
	NOTE: The surgical work surface is a surgical board over a heating pad, and each limb of the mouse is secured using micropore tape.</li>
	<li>Given the difficulty in comprehensively monitoring the physiological changes associated with anesthesia in mice, monitor and record limited parameters. Monitor the temperature, depth of anesthesia, and respiratory rate at least every 5 min for the duration of anesthesia.
	<ol>
		<li>To prevent severe hypothermia and hyperthermia (from active warming by the heat pad), monitor the body temperature throughout the procedure. Insert a clean, lubricated rectal probe into the rectum of the animal and then secure it to the surgical board using micropore tape. 
		NOTE: This probe feeds back to a dynamic system (a feature of the anesthetic delivery system), which modifies the heating pad temperature to manage body temperature.</li>
		<li>Have the person responsible for anesthesia assess the anesthetic depth by observing responses to stimulation of the paw or tail from pressure applied by atraumatic forceps, the palpebral reflex, and muscle tone.</li>
		<li>Measure the respiratory rate by observing movement of the chest wall while observing the respiratory effort subjectively to assess tidal volume. Calculate the respiratory rate by counting breaths over a 10-15 s period and multiplying by 6 or 4, respectively, to determine a breath rate/min.</li>
	</ol>
	</li>
	<li>To prepare the skin, disinfect the surgical site using sterile cotton-tipped applicators. Apply chlorhexidine in a circular, expanding motion working from the center of the surgical site to the edges. Repeat this process 3x (with a new cotton-tipped applicator each time) before a final application of a combination of chlorhexidine and ethanol with a new sterile cotton-tipped applicator in the same pattern, moving from the center of the surgical site to the edge.</li>
	<li>Have the surgeon apply an ethanol-based hand gel prior to donning a sterile surgical gown and sterile surgical gloves.</li>
	<li>To prepare the surgical field, place sterile surgical drapes (precut to 25 cm x 25 cm) on either side of the surgical board, serving as the site to place sterile instruments. Use sterile scissors to fenestrate a wider 25 cm x 40 cm sterile drape to cut a small (slightly longer than the incision site), oval-shaped opening. Lay this drape over the top of the animal such that the fenestration is located at the proposed incision site. Ensure that the lateral ends of this third drape overlap the two smaller drapes on either side to create a continuous surgical field.</li>
</ol>
2. Donor surgery
NOTE: See Supplemental Figure S1 for the key aspects of donor surgery.
<ol>
	<li>Perform the donor surgery with the assistance of a surgical binocular microscope. To begin, use a magnification of 8x, and perform a ventral midline skin incision using a surgical scalpel blade (#23). Ensure that the incision spans from the caudal end of the shaved area to the costal margin with an intact margin of prepared skin at either end. 
	NOTE: The starting magnification of 8x is chosen to enable sufficient visualization of the macrostructure of the subject at the beginning of surgery. From this point on, magnification is at the operator&#39;s discretion and should be selected to provide an appropriate balance between the situational awareness provided by lower magnification and the fine detail that can be visualized with higher magnification.</li>
	<li>Using two sterile cotton-tipped applicators moistened with warmed normal saline, shift the small bowel to expose the abdominal aorta and inferior vena cava (IVC). Use the applicators to bluntly dissect these vessels from the surrounding tissue.</li>
	<li>Use a 3.0 mL syringe with a 30 G, 0.5 in needle to draw up 2.5 mL of 100 IU&#183;mL&#8722;1 of heparinized sodium chloride 0.9% solution (maintained at 4 &#176;C until required during surgery). Using straight-tipped, round-body suture forceps with the non-dominant hand to secure the abdominal aorta in the infra-diaphragmatic area, use the dominant hand to inject 1.5 mL of solution into the aorta in the direction of the heart. Seal the resultant aortotomy with pressure from a cotton-tipped applicator.</li>
	<li>Use straight-tipped microsurgical scissors to transect the IVC to allow exsanguination.</li>
	<li>Perform a thoracotomy using surgical scissors to make two incisions in the bilateral midaxillary lines. At this point, confirm the death of the animal and turn off the isoflurane vaporizer.</li>
	<li>Secure the resultant median segment of the thoracic wall using a micro-bulldog clamp. Pass this to the non-sterile surgical assistant who can secure this to the nose cone using micropore surgical tape. 
	NOTE: The goal is to provide traction on this segment of thorax, which aids with exposure of the cardiac tissues.</li>
	<li>Using round body suture forceps, identify and mobilize the intra-thoracic IVC. 
	NOTE: Ideally, the straight-tipped forceps should be in the non-dominant hand and the curved forceps in the dominant hand.</li>
	<li>With the IVC secured in the non-dominant hand&#39;s forceps, use the dominant hand to inject the remaining 1.5 mL of 100 IU&#183;mL&#8722;1 heparinized sodium chloride 0.9 % solution into the heart.</li>
	<li>Using both sets of forceps, ligate the IVC using 7/0 braided silk of 2 cm length. Use an instrument tie surgeon&#39;s knot with two additional throws for security. Make this knot as proximal along the vessel to the heart as possible.</li>
	<li>Secure the two ends of this knot using artery forceps. Position these forceps so that they provide gentle traction of the heart in the caudal direction to facilitate optimal vessel positioning for subsequent dissection.</li>
	<li>Identify the thymus at the anterosuperior aspect of the heart. Use forceps to dissect this organ from the donor to identify the superior vena cava (SVC).</li>
	<li>Remove the adventitia and associated tissues of the SVC using forceps. Use curved forceps to bluntly dissect and make a small channel posterior to the vessel. Ensure that this channel is as proximal to the heart as possible.</li>
	<li>Pass a piece of 7/0 braided silk of 2 cm length through this channel using forceps and then tie it using the aforementioned technique.</li>
	<li>At a point approximately 2 mm from this ligation (on the side opposite to the heart), divide the SVC using curved microsurgical scissors.</li>
	<li>Using cotton-tipped applicators, flip the heart to the anatomical right.</li>
	<li>Identify the azygous vein on the anatomical left of the heart. Using forceps, bluntly dissect it from surrounding structures. As before, use curved-tipped forceps to create a small channel posterior to this vessel.</li>
	<li>Use a third piece of 7/0 braided silk cut to 2 cm to ligate the azygous vein at maximal proximity to the heart using the same knot-tying technique. Cut the vessel 2 mm from the ligation on the side away from the heart.</li>
	<li>Using cotton-tipped applicators, flip the apex of the heart back to the anatomical left. Use forceps to identify and mobilize the ascending aorta. Pass the curved forceps underneath the aortic arch to create a channel between the ascending and descending aorta.</li>
	<li>Using straight-tipped microsurgical scissors, transect the aortic arch proximal to its branches.</li>
	<li>Using forceps, identify and mobilize the pulmonary artery. Using curved forceps, make a channel posterior to the vessel.</li>
	<li>Using straight-tipped microsurgical scissors, transect the artery at a point just proximal to its bifurcation.</li>
	<li>Use a Rycroft irrigating cannula to gently inject 2 mL of 10 IU&#183;mL&#8722;1 of heparinized sodium chloride 0.9% through the pulmonary artery and ascending aorta to flush any remaining blood from the heart. 
	NOTE: An adequate flush is indicated by the clearance of visible blood from the coronary vessels.</li>
	<li>Using a 3 cm piece of 7/0 braided silk, ligate the remaining posterior vessels (the pulmonary veins) en bloc using a surgeon&#39;s knot with two subsequent throws. Separate the heart from the posterior thoracic wall by carefully cutting using surgical scissors.</li>
	<li>Gently remove the heart from the thorax, submerge it in University of Wisconsin Solution (UWS), and then place it on ice for storage (at 4 &#176;C).</li>
</ol>
3. Recipient surgery
<ol>
	<li>Following preparation of the animal as described in section 1., apply eye lubricant. Inject a weight-based dose (8 mg/kg) of bupivacaine (0.25% diluted to 0.625 mg/mL in sodium chloride 0.9% solution) into the subcutaneous tissue of the ventral abdomen along the planned incision site. Use a 29 G insulin syringe for this injection and look for a straight line of visible blebbing that covers the extent of the planned incision (Supplemental Figure S2A-C). 
	NOTE: Five-seven&#160;minutes should be given to allow time for peak effect of the&#160;local anesthetic.</li>
	<li>With the microscope set to 8x magnification, make a ventral midline skin incision using a sterile surgical scalpel blade (#23). Ensure that the laparotomy spans from the lower abdomen to the costal margin. Insert a retractor to maximize the surgical field (Supplemental Figure S2D).</li>
	<li>Moisten a 5 cm x 5 cm segment of sterile gauze with warmed 0.9% sodium chloride solution and position it at the superior aspect of the surgical site. Using dampened sterile cotton buds, gently eviscerate the intestines, position them on top of this gauze, and wrap the gauze around the organ (Supplemental Figure S3A). 
	NOTE: This procedure helps to reduce insensible fluid loss during surgery and aids in retraction.</li>
	<li>Free and mobilize the abdominal aorta and IVC from surrounding tissues using a blunt dissection technique. Use a combination of cotton-tipped applicators and round-body suture forceps for this step. Ensure that the area of clearance is between the infra-renal aspect of the vessels and just above the bifurcation of the aorta (Supplemental Figure S3B). 
	NOTE: Appropriate visualization at this point will facilitate high-quality vascular anastomoses.</li>
	<li>Identify the posterior abdominal vessels. Using forceps, gently traction the aorta in a direction away from the vertebral column (i.e., longitudinal to the axis of the abdominal vessels). 
	NOTE: It is important that only the aorta and not the IVC is handled in such a manner due to the friability of the latter.</li>
	<li>Ligate each abdominal vessel identified in the planned anastomotic zone. Make a channel on either side of these vessels cephalocaudally by passing the curved forceps posterior to the abdominal vessels on either side. Ligate each vessel identified and mobilized in this manner using lengths of 10/0 nylon tied with instruments into surgeon&#39;s knots with one additional throw (Supplemental Figure S3C).</li>
	<li>Isolate the anastomotic site from circulation. To do so, install a surgical clamp both at the head and then the caudal ends of the abdominal vessels (importantly, in that precise order). Ensure that the clamps cross both vessels to a sufficient degree to ensure complete occlusion.</li>
	<li>Using forceps in the non-dominant hand to steady the aorta, perform an aortotomy using a 30 G needle at the anterior aspect of the aorta. Extend it using straight-tipped microsurgical scissors (Supplemental Figure S3D).</li>
	<li>Perform a venotomy. Using straight forceps, apply gentle anterior traction to the IVC at the point in line with the middle of the aortotomy. Use curved microsurgical scissors with the concave side facing anteriorly to remove a segment of IVC of equal length to the aortotomy (Supplemental Figure S4A).</li>
	<li>Using 10 IU&#183;mL&#8722;1 heparinized sodium chloride solution, wash the interiors of the opened vessels of remaining blood.</li>
	<li>Place the donor heart into the abdomen. Ensure that the positioning is such that the ascending aorta is directly alongside the abdominal aortotomy and the heart is rotated so that the pulmonary artery can be drawn across for the second anastomosis.</li>
	<li>Using 10/0 nylon, place a stay suture between the 12 o&#39;clock position of the aortotomy and the corresponding extremity of the lumen of the ascending aorta. Perform this using straight-tipped forceps and a microsurgical needle holder and tie it using a surgeon&#39;s knot with three subsequent throws. Cut the ends to leave approximately 2 mm of suture.</li>
	<li>Place a second stay suture between the 6 o&#39;clock position of the aortotomy and the corresponding aspect of the ascending aorta. As this suture will also serve as the base for subsequent running sutures, leave at least 10 mm of the tail for the ultimate tie-off.</li>
	<li>Place a continuous running suture of 10/0 nylon in an ascending manner to oppose the anatomical right edge of the aortotomy and the corresponding free edge of the ascending aorta. Use approximately four throws for this line.</li>
	<li>Pass the free end of the suture around the distal stay suture before placing a second continuous running suture down the anatomical left side to affect apposition with the remaining free edge of the aortotomy. Tie off the suture to the tail using a surgeon&#39;s knot with two additional throws.</li>
	<li>Place a stay suture between the 12 o&#39;clock position of the IVC venotomy and the corresponding extremity of the lumen of the pulmonary artery.</li>
	<li>From this anchor point, place a continuous running suture in a descending manner between the anatomical left edge of the pulmonary artery and the corresponding edge of the venotomy. Use an average of four throws for this line followed by one between the 6 o&#39;clock position of the venotomy and the corresponding extremity of the pulmonary artery lumen. Make four more throws to draw the final free edges of the pulmonary artery and venotomy together.</li>
	<li>Tie off the free end of the suture to the anchor end using an instrument tie surgeon&#39;s knot with two additional throws.</li>
	<li>Reposition the heart to sit centrally in the abdomen using cotton buds. Check the vessels for twisting, which would interfere with blood flow.</li>
	<li>Position gel foam over all the suture lines (Supplemental Figure S4B). Place and mold two pieces of approximately 2 mm each around them such that all the visible suture lines are covered.</li>
	<li>Release the vascular clamps: first the caudal clamp, and then the cephalo-clamp. As a small amount of hemorrhage is to be expected, pre-emptively position cotton-tipped applicators over the anastomotic sites to provide pressure.</li>
	<li>Once free from observable leaks, assess the heart for pulsation (Supplemental Figure S4C). If this is not occurring, check to ensure that no twisting of the cardiac vessels has occurred (especially for the IVC).</li>
	<li>Reposition the intestines now over and around the heart. If appearing dry, moisten the peritoneal cavity using warmed sodium chloride solution.</li>
	<li>Close the abdominal cavity using non-absorbable 6/0 prolene monofilament by layers: 
	first the muscular layer, and then the skin (Supplemental Figure S4D). Use the continuous non-interrupted technique.</li>
	<li>Gently remove the recipient from the surgical board and take it off the anesthetic.</li>
	<li>Administer 1 mL of warm saline subcutaneously and place the recipient in a preprepared cage with warming for observation as per postoperative recovery protocols (Supplemental Figure S4E).</li>
</ol>
4. Postoperative care
<ol>
	<li>Immediately after surgery, place the recipient in a clean cage on a heating pad under close observation for at least 3 h. During this period, monitor various parameters (activity, body posture, coat condition, facial expression, gait, ventilation, appearance of the surgical site, presence of a palpable abdominal heartbeat) at least every 30 min. Attribute a score to each parameter (0 = normal, 1 = slightly or intermittently abnormal, 2 = moderately or consistently abnormal). 
	NOTE: Interventions are triggered by the sum of the monitored parameters exceeding&#160;welfare scores specified by the ethics protocol related to this model.</li>
	<li>Move the recipients to a warmed cabinet kept at 25 &#176;C, where they remain until post-op day 7. During the first 3 days, monitor them at least 2x daily. Over the remaining 4 days, monitor them at least 1x daily. For post-operative analgesia, administer a dose of buprenorphine (0.5-0.1 mg/kg, diluted to 0.03 mg/mL with sodium chloride 0.9%) subcutaneously to the recipient mouse on the evening post-surgery and twice daily for the next 3 postoperative days.</li>
	<li>Once removed from the warmed cabinet, monitor the recipients at least 2x weekly until the appropriate experimental endpoint.</li>
</ol>

<ol>
	<li>Mann, F. C., Priestley, J. T., Markowitz, J., Yater, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transplantation+of+the+intact+mammalian+heart.">Transplantation of the intact mammalian heart.</a> Archives of Surgery. 26 (2), 219-224 (1933).</li><li>Neptune, W. B., Cookson, B. A., Bailey, C. P., Appler, R., Rajkowski, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complete+homologous+heart+transplantation.">Complete homologous heart transplantation.</a> A.M.A. Archives of Surgery. 66 (2), 174-178 (1953).</li><li>Downie, H. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Homotransplantation+of+the+dog+heart.">Homotransplantation of the dog heart.</a> A.M.A. Archives of Surgery. 66 (5), 624-636 (1953).</li><li>Blanco, G., Adam, A., Rodriguezperez, D., Fernandez, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Complete+homotransplantation+of+canine+heart+and+lungs.">Complete homotransplantation of canine heart and lungs.</a> A.M.A. Archives of Surgery. 76 (1), 20-23 (1958).</li><li>Abbott, C. P., Lindsey, E. S., Creech, O., Dewitt, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technique+for+heart+transplantation+in+the+rat.">A technique for heart transplantation in the rat.</a> Archives of Surgery. 89, 645-652 (1964).</li><li>Ono, K., Lindsey, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+technique+of+heart+transplantation+in+rats.">Improved technique of heart transplantation in rats.</a> The Journal of Thoracic and Cardiovascular Surgery. 57 (2), 225-229 (1969).</li><li>Corry, R. J., Winn, H. J., Russell, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primarily+vascularized+allografts+of+hearts+in+mice.+The+role+of+H-2D,+H-2K,+and+non-H-2+antigens+in+rejection.">Primarily vascularized allografts of hearts in mice. The role of H-2D, H-2K, and non-H-2 antigens in rejection.</a> Transplantation. 16 (4), 343-350 (1973).</li><li>Joffre, O., 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=Prevention+of+acute+and+chronic+allograft+rejection+with+CD4+CD25+Foxp3++regulatory+T+lymphocytes.">Prevention of acute and chronic allograft rejection with CD4+CD25+Foxp3+ regulatory T lymphocytes.</a> Nature Medicine. 14 (1), 88-92 (2008).</li><li>Gregori, S., 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=Regulatory+T+cells+induced+by+1+alpha,25-dihydroxyvitamin+D3+and+mycophenolate+mofetil+treatment+mediate+transplantation+tolerance.">Regulatory T cells induced by 1 alpha,25-dihydroxyvitamin D3 and mycophenolate mofetil treatment mediate transplantation tolerance.</a> The Journal of Immunology. 167 (4), 1945-1953 (2001).</li><li>Niimi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+technique+for+heterotopic+cardiac+transplantation+in+mice:+Experience+of+3000+operations+by+one+surgeon.">The technique for heterotopic cardiac transplantation in mice: Experience of 3000 operations by one surgeon.</a> The Journal of Heart and Lung Transplantation. 20 (10), 1123-1128 (2001).</li><li>Hasegawa, T., Visovatti, S. H., Hyman, M. C., Hayasaki, T., Pinsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heterotopic+vascularized+murine+cardiac+transplantation+to+study+graft+arteriopathy.">Heterotopic vascularized murine cardiac transplantation to study graft arteriopathy.</a> Nature Protocols. 2 (3), 471-480 (2007).</li><li>Hoogstraten-Miller, S. L., Brown, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Techniques+in+aseptic+rodent+surgery.">Techniques in aseptic rodent surgery.</a> Current Protocols in Immunology. 82, 1-14 (2008).</li><li>Navarro, K. L., 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+anesthesia:+The+art+and+science.">Mouse anesthesia: The art and science.</a> ILAR Journal. , (2021).</li><li>Adams, S., Pacharinsak, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+anesthesia+and+analgesia.">Mouse anesthesia and analgesia.</a> Current Protocols in Mouse Biology. 5 (1), 51-63 (2015).</li><li>Australian code for the care and use of animals for scientific purposes. NHMRC Available from: <a target="_blank" href="https://www.nhmrc.gov.au/about-us/publications/australian-code-care-and-use-animals-scientific-purposes">https://www.nhmrc.gov.au/about-us/publications/australian-code-care-and-use-animals-scientific-purposes</a> (2013)</li><li>Prosser, 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=Dynamic+changes+to+tissue-resident+immunity+after+MHC-matched+and+MHC-mismatched+solid+organ+transplantation.">Dynamic changes to tissue-resident immunity after MHC-matched and MHC-mismatched solid organ transplantation.</a> Cell Reports. 35 (7), 109141 (2021).</li></ol>

The authors have no conflicts of interest to disclose.

The murine orthotopic heart transplant model is a robust preclinical model used primarily to investigate the effects of MHC mismatch on the level and nature of immunological rejection and, more recently, the effect of transplantation on the retention of graft tissue-resident immunity16. While initially closely following the Corry et al.7 protocol, we have refined the protocol to incorporate best-practice standards of aseptic technique, analgesia, and anesthesia. The updatin...

We owe much of our understanding of immunology and transplantation to research based on experimental models of solid organ transplantation using animal subjects. Since the first description of vascularized cardiac transplantation in mammals1, such models have contributed to knowledge in wide-ranging domains, including the therapeutic application of hypothermia2, the benefits of using specialized sutures3, and techniques for total lung and heart homotransplantations4. The development of cardiac transplantation models in rats5,6 provided broader scope for immunological experimentation due to the availability of different breeding lines. The substantially wider range of available inbred and mutant mouse strains led Corry et al.7 to develop a technique of murine heterotopic cardiac transplantation due to the considerable advantages that this range brings to transplantation research. This model has been used widely and has contributed to a greater understanding of graft rejection8 and therapeutics9. Since its first description, however, the technique has remained largely unchanged other than some minor technical details such as adjustments to the position of anastomotic sites10,11.
Since the integration of the technique of Corry et al.7 into our experiments, we have identified areas of promise for improving the protocol, namely those of aseptic technique, anesthesia, and analgesia. Improvements in these areas were expected to offer a positive impact on experimental outcomes and improve animal welfare. This has previously been shown when aseptic technique is used in small animal surgeries as it aids in the reduction of postoperative infections12, which not only impacts morbidity and mortality but can also compromise experiments designed to assess the immune response following transplantation surgery. From an anesthesia and analgesic point of view, the use of a refined regimen helps to reduce the cost to animals and balance the ethical argument of this surgical model by mitigating the pain and suffering of experimental subjects. Further, appropriate anesthesia and analgesia limit the pain-associated stress response, improving the quality of postoperative recovery and, ultimately, increasing the surgical success rate13.
With the aim of improving both animal welfare and experimental outcomes, a protocol was developed with adjustments to bridge these gaps. This protocol has been adapted from that originally described by Corry et al.7 with consultation from a veterinary anesthetist and with due consideration for both the effects and duration of effects of the pharmacological interventions utilized in the anesthetic and analgesic regimen. The approach was based on the principles of balanced anesthesia and multimodal analgesia to ensure appropriate perioperative care14. In addition to the application of aseptic technique, the opioid buprenorphine and the local anesthetic bupivacaine were pre-emptively administered. General anesthesia was performed using the inhalant anesthetic agent isoflurane.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2030 Rycroft irrigating cannula 30 G</td>
			<td>McFarlane</td>
			<td>56005HU</td>
			<td></td>
		</tr>
		<tr>
			<td>Braided surgical silk 7-0</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bulldog clamp curved - 35 mm&#160;</td>
			<td>Roboz</td>
			<td>RS-7441-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Bupivacaine 0.25%&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Castroviejo needle holder catch curved -&#160; 145 mm</td>
			<td>Haag-Streit</td>
			<td>11.62.15</td>
			<td></td>
		</tr>
		<tr>
			<td>Chlorhexidine 5% solution</td>
			<td>Ebos</td>
			<td>JJ61371</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton-tipped applicator - 7.5 cm</td>
			<td>Dove</td>
			<td>SN109510</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol 70% solution</td>
			<td>Ebos</td>
			<td>WH130192EE</td>
			<td></td>
		</tr>
		<tr>
			<td>Gauze 5 x 5 cm white</td>
			<td>Aero</td>
			<td>AGS50</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelfoam 80 mm x 125 mm&#160;</td>
			<td>Pfizer</td>
			<td>7481D</td>
			<td></td>
		</tr>
		<tr>
			<td>Hair clipper</td>
			<td>Wahl</td>
			<td>9860L</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin 1,000 IU in 1 mL</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Iris SuperCut scissors straight - 11.5 cm&#160;&#160;</td>
			<td>Inka Surgical&#160;</td>
			<td>11550.11</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane vaporiser</td>
			<td>Darvall</td>
			<td>9176</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro bulldog clamp - 3.7 cm</td>
			<td>Greman</td>
			<td>14119-G</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro scissors curved 105 mm&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Micropore plain paper surgical tape - 2.5 cm wide</td>
			<td>Ebos</td>
			<td>7810L</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsurgical scissors - curved tip</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Monofilament polyprolene suture - 5/0</td>
			<td>Surgipro</td>
			<td>P-205-X</td>
			<td></td>
		</tr>
		<tr>
			<td>Myweigh i101 Precision Scale 100 g x 0.005 g</td>
			<td>Myweigh</td>
			<td>Kit00053</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle - 30 G x 0.5 inch</td>
			<td>BD</td>
			<td>BD304000</td>
			<td></td>
		</tr>
		<tr>
			<td>Needleholder 15 cm curved &#34;super fine&#34;</td>
			<td>Surgical Specialists</td>
			<td>ST-B-15-8.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylene 10/0 x 15 cm on 3.8 mm 3/8 circle round bodied taper (diam 0.07mm) CV300</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Round body suture forceps curved 0.3 mm 120 mm</td>
			<td>B. Braun</td>
			<td>FD281R</td>
			<td></td>
		</tr>
		<tr>
			<td>Round body suture forceps straight 0.3 mm 120 mm</td>
			<td>B. Braun</td>
			<td>FD280R</td>
			<td></td>
		</tr>
		<tr>
			<td>Round handled vannas spring scissors-str/12.5 cm</td>
			<td></td>
			<td>15400-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring scissors-Cvd Sm blades</td>
			<td></td>
			<td>15001-08</td>
			<td></td>
		</tr>
		<tr>
			<td>Stevens scissors blunt straight 110 mm</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical backboard</td>
			<td></td>
			<td></td>
			<td>Rigid laminated cardboard. 15 x 15 cm</td>
		</tr>
		<tr>
			<td>Surgical drapes</td>
			<td></td>
			<td></td>
			<td>Cut into two sizes. 25 cm x 25 cm, and 25 cm x 40 cm&#160;</td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe - 1 mL</td>
			<td>BD</td>
			<td>592696</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe - 3 mL</td>
			<td>Leica</td>
			<td>M651</td>
			<td></td>
		</tr>
		<tr>
			<td>Toothed forceps</td>
			<td>BD</td>
			<td>309657</td>
			<td></td>
		</tr>
		<tr>
			<td>University of Wisconsin Solution</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Warming pad</td>
			<td></td>
			<td></td>
			<td>Far infrared warming pad 20 x 25 cm</td>
		</tr>
		<tr>
			<td>Westcott spring scissors</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Yasargil clip applier bayonet</td>
			<td>Aesculap</td>
			<td>FE582K</td>
			<td></td>
		</tr>
		<tr>
			<td>Yasargil titanium clip perm 6.6 mm</td>
			<td>Aesculap</td>
			<td>A19FT222T</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse usage</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Strain/SEX/Weight</td>
			<td>Donor</td>
			<td>Recipent</td>
			<td></td>
		</tr>
		<tr>
			<td>BALB/c, female, 19-23 g</td>
			<td>7</td>
			<td>21</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6, female, 17-20 g</td>
			<td>7</td>
			<td>0</td>
			<td></td>
		</tr>
		<tr>
			<td>CD45.1 BALB/c, female, 17-21 g</td>
			<td>5</td>
			<td>0</td>
			<td></td>
		</tr>
	</tbody>
</table>

a modified murine heterotopic heart transplant protocol matching contemporary standards of aseptic technique, anesthesia, and analgesia

The development of experimental models of cardiac transplantation in animals has contributed to many advances in the fields of immunology and solid organ transplantation. While the heterotopic vascularized murine cardiac transplantation model was initially utilized in studies of graft rejection using combinations of mismatched inbred mouse strains, access to genetically modified strains and therapeutic modalities can provide powerful new preclinical insights. Fundamentally, the surgical methodology for this technique has not changed since its development, especially with respect to important factors such as aseptic technique, anesthesia, and analgesia, which make material impacts on postsurgical morbidity and mortality. Additionally, improvements in perioperative management are expected to provide improvements in both animal welfare and experimental outcomes. This paper reports upon a protocol developed in collaboration with a subject matter expert in veterinary anesthesia and describes the surgical technique with an emphasis on perioperative management. Additionally, we discuss the implications of these refinements and provide details on troubleshooting critical surgical steps for this procedure.

To determine the effectiveness of the surgical technique in promoting good outcomes of wound healing and mouse recovery, early experiments in the laboratory determined the survival characteristics of a range of heart grafts of variable immunogenicity to the recipient. These included congenic (n = 5) and syngeneic (n = 5) grafts, which share the same major histocompatibility complex (MHC) markers as the recipient, and major mismatch grafts (n = 9), in which the graft and the recipient have different MHC markers. We used d...

Watch this Scientific Journal Video about A Modified Murine Heterotopic Heart Transplant Protocol Matching Contemporary Standards of Aseptic Technique, Anesthesia, and Analgesia at JoVE.com

A Modified Murine Heterotopic Heart Transplant Protocol Matching Contemporary Standards of Aseptic Technique, Anesthesia, and Analgesia

The present paper describes a modified technique for heterotopic vascularized cardiac transplantation with updated aseptic technique, analgesia, and anesthesia.

The development of experimental models of cardiac transplantation in animals has contributed to many advances in the fields of immunology and solid organ ...

a-modified-murine-heterotopic-heart-transplant-protocol-matching

Universidad de Cartagena UDEC

Research

JoVE Journal

Immunology and Infection

1.5K Views.  The University of Western Australia. The present paper describes a modified technique for heterotopic vascularized cardiac transplantation with updated aseptic technique, analgesia, and anesthesia.

Video: A Modified Murine Heterotopic Heart Transplant Protocol Matching Contemporary Standards of Aseptic Technique, Anesthesia, and Analgesia

Reverse Genetics Mediated Recovery of Infectious Murine Norovirus

Human noroviruses are responsible for most cases of human gastroenteritis (GE) worldwide and are recurrent problem in environments where close person-to-person contact cannot be avoided 1, 2. During the last few years an increase in the incidence of outbreaks in hospitals has been reported, causing significant disruptions to their operational capacity as well as large economic losses. The identification of new antiviral approaches has been limited due to the inability of human noroviruses to complete a productive infection in cell culture 3. The recent isolation of a murine norovirus (MNV), closely related to human norovirus 4 but which can be propagated in cells 5 has opened new avenues for the investigation of these pathogens 6, 7. 
MNV replication results in the synthesis of new positive sense genomic and subgenomic RNA molecules, the latter of which corresponds to the last third of the viral genome (Figure 1). MNV contains four different open reading frames (ORFs), of which ORF1 occupies most of the genome and encodes seven non-structural proteins (NS1-7) released from a polyprotein precursor. ORF2 and ORF3 are contained within the subgenomic RNA region and encode the capsid proteins (VP1 and VP2, respectively) (Figure 1). Recently, we have identified that additional ORF4 overlapping ORF2 but in a different reading frame is functional and encodes for a mitochondrial localised virulence factor (VF1) 8.
Replication for positive sense RNA viruses, including noroviruses, takes place in the cytoplasm resulting in the synthesis of new uncapped RNA genomes. To promote viral translation, viruses exploit different strategies aimed at recruiting the cellular protein synthesis machinery 9-11. Interestingly, norovirus translation is driven by the multifunctional viral protein-primer VPg covalently linked to the 5' end of both genomic and subgenomic RNAs 12-14. This sophisticated mechanism of translation is likely to be a major factor in the limited efficiency of viral recovery by conventional reverse genetics approaches. 
Here we report two different strategies based on the generation of murine norovirus-1 (referred to as MNV herewith) transcripts capped at the 5' end. One of the methods involves both in vitro synthesis and capping of viral RNA, whereas the second approach entails the transcription of MNV cDNA in cells expressing T7 RNA polymerase. The availability of these reverse genetics systems for the study of MNV and a small animal model has provided an unprecedented ability to dissect the role of viral sequences in replication and pathogenesis 15-17.

Noroviruses are a major cause of gastroenteritis yet molecular techniques for their characterisation are still relatively new. Here we report two different reverse genetics approaches for the efficient recovery of murine norovirus (MNV), the only member of this genus which can be propagated in cell culture.

Human noroviruses are responsible for most cases of human gastroenteritis (GE) worldwide and are recurrent problem in environments where close ...

Isolation of Cortical Microglia with Preserved Immunophenotype and Functionality From Murine Neonates

Isolation of microglia from CNS tissue is a powerful investigative tool used to study microglial biology ex vivo. The present method details a procedure for isolation of microglia from neonatal murine cortices by mechanical agitation with a rotary shaker. This microglia isolation method yields highly pure cortical microglia that exhibit morphological and functional characteristics indicative of quiescent microglia in normal, nonpathological conditions in vivo. This procedure also preserves the microglial immunophenotype and biochemical functionality as demonstrated by the induction of morphological changes, nuclear translocation of the p65 subunit of NF-&#954;B (p65), and secretion of the hallmark proinflammatory cytokine, tumor necrosis factor-&#945; (TNF-&#945;), upon lipopolysaccharide (LPS) and Pam3CSK4 (Pam) challenges. Therefore, the present isolation procedure preserves the immunophenotype of both quiescent and activated microglia, providing an experimental method of investigating microglia biology in ex vivo conditions.

One key to successful investigation of microglial biology is the preservation of microglial immunofunction ex vivo during isolation from CNS tissue. Isolating microglia via rotary shaking results in highly pure and immunofunctional cell cultures as assessed by fluorescent imaging, immunocytochemistry, and ELISA following microglia activation with the proinflammatory stimuli lipopolysaccharide (LPS) and Pam3CSK4 (Pam).

Isolation of microglia from CNS tissue is a powerful investigative tool used to study microglial biology ex vivo. The present method details a procedure ...

Isolation, Identification, and Purification of Murine Thymic Epithelial Cells

The thymus is a vital organ for T lymphocyte development. Of thymic stromal cells, thymic epithelial cells (TECs) are particularly crucial at multiple stages of T cell development: T cell commitment, positive selection and negative selection. However, the function of TECs in the thymus remains incompletely understood. In the article, we provide a method to isolate TEC subsets from fresh mouse thymus using a combination of mechanical disruption and enzymatic digestion. The method allows thymic stromal cells and thymocytes to be efficiently released from cell-cell and cell-extracellular matrix connections and to form a single-cell suspension. Using the isolated cells, multiparameter flow cytometry can be applied to identification and characterization of TECs and dendritic cells. Because TECs are a rare cell population in the thymus, we also describe an effective way to enrich and purify TECs by depleting thymocytes, the most abundant cell type in the thymus. Following the enrichment, cell sorting time can be decreased so that loss of cell viability can be minimized during purification of TECs. Purified cells are suitable for various downstream analyses like Real Time-PCR, Western blot and gene expression profiling. The protocol will promote research of TEC function and as well as the development of in vitro T cell reconstitution.

Here we describe an efficient method for isolation, identification, and purification of mouse thymic epithelial cells (TECs). The protocol can be utilized for studies of thymus function for normal T cell development, thymus dysfunction, and T cell reconstitution.

The thymus is a vital organ for T lymphocyte development. Of thymic stromal cells, thymic epithelial cells (TECs) are particularly crucial at multiple ...

Isolation of Murine Lymph Node Stromal Cells

Secondary lymphoid organs including lymph nodes are composed of stromal cells that provide a structural environment for homeostasis, activation and differentiation of lymphocytes. Various stromal cell subsets have been identified by the expression of the adhesion molecule CD31 and glycoprotein podoplanin (gp38), T zone reticular cells or fibroblastic reticular cells, lymphatic endothelial cells, blood endothelial cells and FRC-like pericytes within the double negative cell population. For all populations different functions are described including, separation and lining of different compartments, attraction of and interaction with different cell types, filtration of the draining fluidics and contraction of the lymphatic vessels. In the last years, different groups have described an additional role of stromal cells in orchestrating and regulating cytotoxic T cell responses potentially dangerous for the host. 
Lymph nodes are complex structures with many different cell types and therefore require a appropriate procedure for isolation of the desired cell populations. Currently, protocols for the isolation of lymph node stromal cells rely on enzymatic digestion with varying incubation times; however, stromal cells and their surface molecules are sensitive to these enzymes, which results in loss of surface marker expression and cell death. Here a short enzymatic digestion protocol combined with automated mechanical disruption to obtain viable single cells suspension of lymph node stromal cells maintaining their surface molecule expression is proposed.

Isolation of lymph node stromal cells is a multistep procedure including enzymatic digestion and mechanical disaggregation to obtain fibroblastic reticular cells, lymphatic and blood endothelial cells. In the described procedure, a short digestion is combined with automated mechanical disaggregation to minimize surface marker degradation of viable lymph node stromal cells.

Secondary lymphoid organs including lymph nodes are composed of stromal cells that provide a structural environment for homeostasis, activation and ...

Generation of Two-color Antigen Microarrays for the Simultaneous Detection of IgG and IgM Autoantibodies

Autoantibodies, which are antibodies against self-antigens, are present in many disease states and can serve as markers for disease activity. The levels of autoantibodies to specific antigens are typically detected with the enzyme-linked immunosorbent assay (ELISA) technique. However, screening for multiple autoantibodies with ELISA can be time-consuming and requires a large quantity of patient sample. The antigen microarray technique is an alternative method that can be used to screen for autoantibodies in a multiplex fashion. In this technique, antigens are arrayed onto specially coated microscope slides with a robotic microarrayer. The slides are probed with patient serum samples and subsequently fluorescent-labeled secondary antibodies are added to detect binding of serum autoantibodies to the antigens. The autoantibody reactivities are revealed and quantified by scanning the slides with a scanner that can detect fluorescent signals. Here we describe methods to generate custom antigen microarrays. Our current arrays are printed with 9 solid pins and can include up to 162 antigens spotted in duplicate. The arrays can be easily customized by changing the antigens in the source plate that is used by the microarrayer. We have developed a two-color secondary antibody detection scheme that can distinguish IgG and IgM reactivities on the same slide surface. The detection system has been optimized to study binding of human and murine autoantibodies.

We describe here a method to generate customizable antigen microarrays that can be used for the simultaneous detection of serum IgG and IgM autoantibodies from humans and mice. These arrays allow for high-throughput and quantitative detection of antibodies against any antigens or epitopes of interest.

Autoantibodies, which are antibodies against self-antigens, are present in many disease states and can serve as markers for disease activity. The levels ...

An Optimized Protocol to Analyze Glycolysis and Mitochondrial Respiration in Lymphocytes

Lymphocytes respond to a variety of stimuli by activating intracellular signaling pathways, which in turn leads to rapid cellular proliferation, migration and differentiation, and cytokine production. All of these events are tightly linked to the energy status of the cell, and therefore studying the energy-producing pathways may give clues about the overall functionality of these cells. The extracellular flux analyzer is a commonly used device for evaluating the performance of glycolysis and mitochondrial respiration in many cell types. This system has been used to study immune cells in a few published reports, yet a comprehensive protocol optimized particularly for lymphocytes is lacking. Lymphocytes are fragile cells that survive poorly in ex vivo conditions. Oftentimes lymphocyte subsets are rare, and working with low cell numbers is inevitable. Thus, an experimental strategy that addresses these difficulties is required. Here, we provide a protocol that allows for rapid isolation of viable lymphocytes from lymphoid tissues, and for the analysis of their metabolic states in the extracellular flux analyzer. Furthermore, we provide results of experiments in which the metabolic activities of several lymphocyte subtypes at different cell densities were compared. These observations suggest that our protocol can be used to achieve consistent, well-standardized results even at low cell concentrations, and thus it may have broad applications in future studies focusing on the characterization of metabolic events in immune cells.

The study of metabolism is becoming increasingly relevant to immunological research. Here, we present an optimized method for measuring glycolysis and mitochondrial respiration in mouse splenocytes, and T and B lymphocytes.

Lymphocytes respond to a variety of stimuli by activating intracellular signaling pathways, which in turn leads to rapid cellular proliferation, migration ...

Isolation of Extracellular Vesicles from Murine Bronchoalveolar Lavage Fluid Using an Ultrafiltration Centrifugation Technique

Extracellular vesicles (EVs) are newly discovered subcellular components that play important roles in many biological signaling functions during physiological and pathological states. The isolation of EVs continues to be a major challenge in this field, due to limitations intrinsic to each technique. The differential ultracentrifugation with density gradient centrifugation method is a commonly used approach and is considered to be the gold standard procedure for EV isolation. However, this procedure is time-consuming, labor-intensive, and generally results in low scalability, which may not be suitable for small-volume samples such as bronchoalveolar lavage fluid. We demonstrate that an ultrafiltration centrifugation isolation method is simple and time- and labor-efficient yet provides a high recovery yield and purity. We propose that this isolation method could be an alternative approach that is suitable for EV isolation, particularly for small-volume biological specimens.

Here, we describe two extracellular vesicle isolation protocols, ultrafiltration centrifugation and ultracentrifugation with density gradient centrifugation, to isolate extracellular vesicles from murine bronchoalveolar lavage fluid samples. The extracellular vesicles derived from murine bronchoalveolar lavage fluid by both methods are quantified and characterized.

Extracellular vesicles (EVs) are newly discovered subcellular components that play important roles in many biological signaling functions during ...

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 ...

Flow Cytometric Characterization of Murine B Cell Development

Extensive studies have characterized the development and differentiation of murine B cells in secondary lymphoid organs. Antibodies secreted by B cells have been isolated and developed into well-established therapeutics. Validation of murine B cell development, in the context of autoimmune prone mice, or in mice with modified immune systems, is a crucial component of developing or testing therapeutic agents in mice and is an appropriate use of flow cytometry. Well established B cell flow cytometric parameters can be used to evaluate B cell development in the murine peritoneum, bone marrow, and spleen, but a number of best practices must be adhered to. In addition, flow cytometric analysis of B cell compartments should also complement additional readouts of B cell development. Data generated using this technique can further our understanding of wild type, autoimmune prone mouse models as well as humanized mice that can be used to generate antibody or antibody-like molecules as therapeutics.

We describe herein a simple analysis of the heterogeneity of the murine immune B cell compartment in the peritoneum, spleen, and bone marrow tissues by flow cytometry. The protocol can be adapted and extended to other mouse tissues.

Extensive studies have characterized the development and differentiation of murine B cells in secondary lymphoid organs. Antibodies secreted by B cells ...