DK is supported by National Institutes of Health grants 1P01AI116501, R01HL094601, R01HL151078, Veterans Administration Merit Review grant 1I01BX002730, and The Foundation for Barnes-Jewish Hospital.

All animal handling procedures were conducted in compliance with the NIH Care and Use of Laboratory Animals guidelines and approved by the Animal Studies Committee at Washington University School of Medicine. Hearts from C57BL/6 (B6) and BALB/c mice (weighing 20-25 g) were transplanted into gender-matched B6 recipients (6-8 weeks of age). The mice were obtained from commercial sources (see Table of Materials). Syngeneic transplants were performed to evaluate cellular responses related to ischemia-reperfusion injury, and allogeneic transplants were performed to investigate the immune mechanisms involved in graft tolerance and rejection. B6 lysozyme M-green fluorescent protein (LysM-GFP) reporter mice17, originally obtained from Klaus Ley of La Jolla Institute for Allergy and Immunology, La Jolla, CA, and subsequently bred in our facility, were used as recipients for selected experiments to visualize neutrophil infiltration into cardiac grafts. Survival surgery was performed using aseptic procedures.&#160;
1. Donor procedure
<ol>
	<li>Anesthetize the mice by injecting ketamine (80&#8722;100 mg/kg) and xylazine (8&#8722;10 mg/kg) (see Table of Materials) intraperitoneally into the donor mouse. Confirm surgical plane of anesthesia with toe and tail pinch.</li>
	<li>Prepare the surgical area by shaving the hair from the chest and abdomen using an electric razor.</li>
	<li>Administer 100 units of heparin (see Table of Materials) intravenously into the penile vein (males) or external jugular vein (males or females).</li>
	<li>Place the mice in a supine position with forelimbs overhead. Secure the forelimbs and hindlimbs with surgical tape and disinfect the skin with three alternating scrubs of 0.75% iodine and 70% ethanol.</li>
	<li>Perform an incision, median laparosternotomy, from umbilicus to sternal angle (3-4 cm), followed by a bilateral thoracotomy along each costal margin (2 cm bilaterally). Fold the anterior chest wall over the neck for full exposure of the mediastinum.</li>
	<li>Excise the thymus and expose the intrathoracic inferior vena cava.</li>
	<li>Transect across the width of the abdominal aorta for exsanguination.</li>
	<li>For retrograde perfusion, inject 1.5 mL of 4 &#176;C saline into the intrathoracic inferior vena cava with the needle oriented superiorly toward the graft, as previously described13.</li>
	<li>Ligate the superior vena cava using an 8-0 silk suture and divide distally.</li>
	<li>Repeat the retrograde perfusion by injecting another 1.5 mL of 4 &#176;C saline via the inferior vena cava.</li>
	<li>Ligate the inferior vena cava using an 8-0 silk suture and divide distally.</li>
	<li>Dissect the aortic arch and pulmonary artery trunk for graft harvesting and transect both distally. Ligate the pulmonary veins on the posterior surface of the heart using a 6-0 silk suture and divide distally.</li>
	<li>Perform graft preparation by removing the donor heart from the chest cavity. Place the excised heart in a plastic container filled with 4 &#176;C heparinized saline for 1-2 min. Transfer the graft onto a sterile plastic flask filled with ice for cuff placement (Figure 1A). 
	NOTE: The heart graft needs to remain on the flask for approximately 5 min to place the donor pulmonary artery cuff.</li>
	<li>Place a 1 mm long 20 G angiocatheter (see Table of Materials) cuff over the pulmonary artery for the donor cuff. Using fine forceps, gently fold the edges of the artery back over the cuff. Secure the folded vessel to the cuff using a 10-0 nylon tie, as described previously18 (Figure 1B,C).</li>
	<li>Store the donor heart in heparinized saline or another preservation solution at 4 &#176;C. 
	NOTE: While some may prefer specific preservation solutions (e.g., the University of Wisconsin solution) for prolonged ischemic preservation, it can be costly19. Saline may be a suitable alternative for short periods of ischemia (&#60;1 h)20. Ultimately, the choice of preservation solution depends on the experimental design21.</li>
</ol>
2. Recipient procedure
<ol>
	<li>Inject ketamine (80&#8722;100 mg/kg) and xylazine (8&#8722;10 mg/kg) intraperitoneally into the recipient mouse for anesthesia. Inject sustained-release buprenorphine (0.5-1.0 mg/kg) subcutaneously for analgesia. Confirm surgical plane of anesthesia with toe and tail pinch.</li>
	<li>Prepare the surgical area by shaving the hair from the cervical area using an electric razor. Apply sterile, non-medicated ophthalmic ointment to the eyes to prevent corneal drying.</li>
	<li>Place the animal in a supine position with forelimbs adjacent to the body and the head turned slightly to the left. Secure the forelimbs and hindlimbs with surgical tape. Disinfect the skin with three alternating scrubs of 0.75% iodine and 70% ethanol.</li>
	<li>Make a midline cervical incision from the lower mandible to the sternum.</li>
	<li>Transect the right sternocleidomastoid muscle. Excise the right lobe of the submandibular gland to create space for graft implantation.</li>
	<li>Tie a slipknot over the proximal external jugular vein using a 6-0 silk suture. Ligate the distal external jugular vein and adjacent branches using an 8-0 silk suture. Make a transverse incision across the anterior wall of the external jugular vein.</li>
	<li>Place a 10-0 nylon suture through the edge of the proximal external jugular vein and the underlying tissue to secure the vein during cuff insertion (Figure 1D).</li>
	<li>Ligate the distal right common carotid artery using an 8-0 silk suture just inferior to the carotid bifurcation. Tie a slipknot over the proximal common carotid artery using a 6-0 silk suture. Transect the artery distally between the sutures.</li>
	<li>Similar to the donor cuff, place a 0.6 mm long 24 G angiocatheter cuff over the recipient&#39;s right common carotid artery. Using fine forceps, gently fold the edges of the artery back over the cuff. Secure the folded vessel to the cuff using a 10-0 nylon tie.</li>
	<li>Place the donor heart superior to the right cervical area.</li>
	<li>Drip cold saline onto the heart graft every few minutes during implantation.</li>
	<li>Place a 10-0 nylon suture through the edge of the donor aorta and through a superficial bite of the underlying tissue to secure the graft in place (Figure 1E).</li>
	<li>Flush the donor aorta with 0.5 mL of 0.9% heparinized saline.</li>
	<li>Insert the recipient&#39;s common carotid artery cuff into the donor aorta. Secure the anastomosis with an 8-0 silk tie (Figure 1F). Remove the aortic anchor suture.</li>
	<li>De-air the external jugular vein by flushing the recipient&#39;s external jugular vein with 0.5 mL of 0.9% heparinized saline.</li>
	<li>Perform pulmonary artery anastomosis by inserting the donor pulmonary artery cuff into the recipient&#39;s external jugular vein and secure with an 8-0 silk tie (Figure 1G). Remove the external jugular vein anchor suture and transect the remaining posterior wall of the external jugular vein to free the graft from the underlying tissue. Ensure the graft is properly oriented without kinking or twisting of the anastomoses.</li>
	<li>Release the slipknots on the recipient&#39;s external jugular vein followed by the common carotid artery to initiate heart graft reperfusion (Figure 1H).</li>
	<li>Close the cervical skin incision using an interrupted 6-0 nylon suture.</li>
</ol>
3. Postoperative care
<ol>
	<li>Place the recipient in a warm recovery chamber immediately following surgery and monitor closely until fully recovered from anesthesia (approximately 1 h).</li>
	<li>Continue to closely monitor the animal (every 6-8 h) for at least 72 h after surgery for signs of abnormal behavior, such as lethargy, shaking, rapid breathing, or anorexia.</li>
	<li>For pain control, inject carprofen (5 mg/kg) subcutaneously every 8-12 h for analgesia, in addition to subcutaneous buprenorphine (0.05 mg/kg) every 8-12 h for 24-48 h starting at the end of the surgery.</li>
</ol>
4. Intravital two-photon imaging of leukocyte trafficking in the heart graft
<ol>
	<li>Inject ketamine (80-100 mg/kg) and xylazine (8-10 mg/kg) intraperitoneally into a B6 LysM-GFP recipient mouse17 2 h after graft reperfusion for anesthesia.</li>
	<li>Perform orotracheal intubation using a 20 G angiocatheter, as previously described18.</li>
	<li>Connect the angiocatheter to tubing from a mouse mechanical ventilator and ventilate with room air at a rate of 120 breaths/min and a tidal volume of 0.5 mL18.</li>
	<li>Inject 12 &#181;L of 655 nm nontargeted Quantum dots (see Table of Materials), suspended in 50 &#181;L of PBS intravenously, as described previously13.</li>
	<li>Reopen the neck incision to expose the heart graft. Place the mouse in a stabilization chamber.</li>
	<li>Secure a portion of the free wall of the left ventricle using a thin ring of tissue adhesive (see Table of Materials), applied to a glass coverslip attached to the upper chamber plate.</li>
	<li>Place the chamber under the two-photon microscope objective to acquire images and videos, as described previously13.</li>
</ol>

<ol>
	<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> The 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=Heart+transplantation+in+congenic+strains+of+mice.">Heart transplantation in congenic strains of mice.</a> Transplantation Proceedings. 5 (1), 733-735 (1973).</li><li>Heron, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technique+for+accessory+cervical+heart+transplantation+in+rabbits+and+rats.">A technique for accessory cervical heart transplantation in rabbits and rats.</a> Acta Pathologica Microbiologica Scandinavica Section A Pathology. 79 (4), 366-372 (1971).</li><li>Chen, Z. H. <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+of+cervical+heterotopic+heart+transplantation+in+mice.">A technique of cervical heterotopic heart transplantation in mice.</a> Transplantation. 52 (6), 1099-1101 (1991).</li><li>Matsuura, A., Abe, T., Yasuura, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplified+mouse+cervical+heart+transplantation+using+a+cuff+technique.">Simplified mouse cervical heart transplantation using a cuff technique.</a> Transplantation. 51 (4), 896-898 (1991).</li><li>Yu, Y., 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=Cuff+anastomosis+of+both+renal+artery+and+vein+to+minimize+thrombosis:+a+novel+method+of+kidney+transplantation+in+mice.">Cuff anastomosis of both renal artery and vein to minimize thrombosis: a novel method of kidney transplantation in mice.</a> Journal of Investigative Surgery. 35 (1), 56-60 (2022).</li><li>Tomita, Y., 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+technique+of+heterotopic+cervical+heart+transplantation+in+mice.">Improved technique of heterotopic cervical heart transplantation in mice.</a> Transplantation. 64 (11), 1598-1601 (1997).</li><li>Wang, Q., Liu, Y., Li, X. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simplified+technique+for+heterotopic+vascularized+cervical+heart+transplantation+in+mice.">Simplified technique for heterotopic vascularized cervical heart transplantation in mice.</a> Microsurgery. 25 (1), 76-79 (2005).</li><li>Oberhuber, R., 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=Murine+cervical+heart+transplantation+model+using+a+modified+cuff+technique.">Murine cervical heart transplantation model using a modified cuff technique.</a> Journal of Visualized Experiments. (92), e50753 (2014).</li><li>Ratschiller, 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=Heterotopic+cervical+heart+transplantation+in+mice.">Heterotopic cervical heart transplantation in mice.</a> Journal of Visualized Experiments. (102), e52907 (2015).</li><li>Mao, X., Xian, P., You, H., Huang, G., Li, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modified+cuff+technique+for+mouse+cervical+heterotopic+heart+transplantation+model.">A modified cuff technique for mouse cervical heterotopic heart transplantation model.</a> Journal of Visualized Experiments. (180), e63504 (2022).</li><li>Li, W., 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=Intravital+2-photon+imaging+of+leukocyte+trafficking+in+beating+heart.">Intravital 2-photon imaging of leukocyte trafficking in beating heart.</a> Journal of Clinical Investigation. 122 (7), 2499-2508 (2012).</li><li>Kreisel, 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=In+vivo+two-photon+imaging+reveals+monocyte-dependent+neutrophil+extravasation+during+pulmonary+inflammation.">In vivo two-photon imaging reveals monocyte-dependent neutrophil extravasation during pulmonary inflammation.</a> Proceedings of the National Academy of Sciences of the United States of America. 107 (42), 18073-18078 (2010).</li><li>Li, W., 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=Visualization+of+monocytic+cells+in+regressing+atherosclerotic+plaques+by+intravital+2-photon+and+positron+emission+tomography-based+imaging-brief+report.">Visualization of monocytic cells in regressing atherosclerotic plaques by intravital 2-photon and positron emission tomography-based imaging-brief report.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 38 (5), 1030-1036 (2018).</li><li>Li, W., 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=Lung+transplant+acceptance+is+facilitated+by+early+events+in+the+graft+and+is+associated+with+lymphoid+neogenesis.">Lung transplant acceptance is facilitated by early events in the graft and is associated with lymphoid neogenesis.</a> Mucosal Immunology. 5 (5), 544-554 (2012).</li><li>Faust, N., Varas, F., Kelly, L. M., Heck, S., Graf, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insertion+of+enhanced+green+fluorescent+protein+into+the+lysozyme+gene+creates+mice+with+green+fluorescent+granulocytes+and+macrophages.">Insertion of enhanced green fluorescent protein into the lysozyme gene creates mice with green fluorescent granulocytes and macrophages.</a> Blood. 96 (2), 719-726 (2000).</li><li>Krupnick, A. 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=Orthotopic+mouse+lung+transplantation+as+experimental+methodology+to+study+transplant+and+tumor+biology.">Orthotopic mouse lung transplantation as experimental methodology to study transplant and tumor biology.</a> Nature Protocols. 4 (1), 86-93 (2009).</li><li>Westhofen, 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=The+heterotopic+heart+transplantation+in+mice+as+a+small+animal+model+to+study+mechanical+unloading+-+Establishment+of+the+procedure,+perioperative+management+and+postoperative+scoring.">The heterotopic heart transplantation in mice as a small animal model to study mechanical unloading - Establishment of the procedure, perioperative management and postoperative scoring.</a> PLoS One. 14 (4), 0214513 (2019).</li><li>Ma, Y., 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=Optimization+of+the+cuff+technique+for+murine+heart+transplantation.">Optimization of the cuff technique for murine heart transplantation.</a> Journal of Visualized Experiments. (160), e61103 (2020).</li><li>Latchana, N., Peck, J. R., Whitson, B., Black, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preservation+solutions+for+cardiac+and+pulmonary+donor+grafts:+a+review+of+the+current+literature.">Preservation solutions for cardiac and pulmonary donor grafts: a review of the current literature.</a> Journal of Thoracic Disease. 6 (8), 1143-1149 (2014).</li><li>Hartley, C. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Doppler+velocity+measurements+from+large+and+small+arteries+of+mice.">Doppler velocity measurements from large and small arteries of mice.</a> American Journal of Physiology - Heart and Circulatory Physiology. 301 (2), 269-278 (2011).</li><li>Bovenkamp, P. R., 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=Velocity+mapping+of+the+aortic+flow+at+9.4+T+in+healthy+mice+and+mice+with+induced+heart+failure+using+time-resolved+three-dimensional+phase-contrast+MRI+(4D+PC+MRI).">Velocity mapping of the aortic flow at 9.4 T in healthy mice and mice with induced heart failure using time-resolved three-dimensional phase-contrast MRI (4D PC MRI).</a> MAGMA. 28 (4), 315-327 (2015).</li><li>Wang, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+animal+models+of+xenotransplantation.">Small animal models of xenotransplantation.</a> Methods in Molecular Biology. 885, 125-153 (2012).</li><li>Martins, P. 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The authors have nothing to disclose.

Utilizing this technique, mouse heterotopic cervical cardiac transplantation can be performed in less than 40 min by an experienced microsurgeon and in approximately 60 min by an entry-level microsurgeon. While cervical heart transplantation has been studied in numerous animal models, a mouse model remains the gold standard due to multiple well-defined genetic strains, genetic alteration capabilities, and the availability of numerous reagents, including monoclonal antibodies24. The technique described here provides a unique opportunity for post-transplant monitoring, such as electrocardiography or intravital imaging, including two-photon microscopy (Figure 3) or serial non-invasive PET imaging13,14,15,25. This method provides a superficial location for the heart graft that is easier to stabilize for intravital imaging, thus avoiding the complexity inherent to the abdominal transplantation method due to the deeper location of the graft and the surrounding abdominal organs. Furthermore, this technique is especially useful in the context of re-transplantation. Re-transplantation models represent powerful tools for identifying resident cells in transplanted cardiac grafts that mediate alloimmune responses. While we have previously utilized this technique in a mouse heart re-transplantation model to assess short-term outcomes, this approach can be expanded upon in future experiments to explore long-term outcomes16. To this end, the present investigations thus far have utilized a short period of cold ischemia (approximately 20 min). Future studies could investigate the effect of prolonged cold or warm ischemia on short- and long-term outcomes to more closely mimic clinical transplantation.
Several critical steps of this technique need to be considered. Previous methods involve the cuff insertion on the smaller external jugular vein into the large lumen of the donor pulmonary artery6,8. The placement of the larger cuff on the donor pulmonary artery to establish proper orientation with blood flow makes it slightly more difficult to insert the cuff into the smaller external jugular vein. Fixing the edge of the vein to the underlying tissue and only partially incising the vein&#39;s anterior wall facilitates the cuff insertion. Additionally, cuff placement on the recipient&#39;s common carotid artery can be quite challenging due to the small caliber of the vessel. As such, prior techniques have reported the utilization of smaller cuffs (e.g., 26 G) for this anastomosis12. However, the current approach utilizes a larger 24 G cuff to ensure adequate graft perfusion, which we believe may offer some survival benefits. Selecting larger recipient mice may help novice microsurgeons. Anchor sutures are removed following reperfusion, and the graft is not fixed in a proper orientation as others have described3. Thus, it is important to check that the graft is properly positioned and oriented prior to cervical skin closure to prevent twisting or torsion (Table 1). Excision of the right submandibular gland is performed to provide adequate space for the heart graft, thus avoiding graft compression following skin closure.
The model described here offers several advantages. By placing the cuffs on the donor pulmonary artery trunk and the recipient&#39;s common carotid artery, the cuff orientation aligns with the direction of the blood flow. This decreases the likelihood of turbulent flow and thrombus formation. Second, a larger 20 G pulmonary artery cuff is utilized to ensure ample return of blood flow to the recipient. Third, a larger 24 G cuff is placed on the common carotid artery to ensure adequate perfusion of the graft. Lastly, 10-0 nylon anchor sutures are used to fix the graft to underlying tissues and facilitate cuff insertion. These modifications help in overcoming the technical challenges of the procedure, prevent anastomotic turbulence, and reduce postoperative complications such as thrombus formation.
An important limitation of all mouse heart transplant models is that physiologic blood flow is not restored through the heart&#39;s chambers. Instead, these models rely on circulation through the coronary vessels. The consequences of this retrograde flow pattern on the graft&#39;s cellular injury and immune responses have not been clearly delineated; however, it is possible that mechanical shear forces resulting from this non-physiologic circulation influence immune responses. A surgical model of heart transplantation in mice that restores physiologic blood flow is yet to be developed and would require substantial technical advances. It is observed that a small proportion of mice (&#60;3%) experience transient erratic behavior (e.g., running in circles) following the procedure. This behavior lasts for approximately 1-2 h before resolution. Given that this behavior is not observed after other procedures using the same anesthetic regimen, it may be related to transient cerebral ischemia due to blood flow alterations after cervical heart transplantation. Full recovery has occurred in all mice without any chronic deficits observed.

Abbott et al. published1 the first description of heterotopic abdominal heart transplantation in rats in 1964. These surgical techniques were refined and simplified by Ono et al. in 19692. Corry et al. first described a method for heterotopic abdominal heart transplantation in mice in 1973; similar to the previously reported rat models, this involved engraftment into the host&#39;s abdomen with revascularization by end-to-side anastomoses of the donor&#39;s pulmonary artery and ascending aorta to the recipient&#39;s inferior vena cava and abdominal aorta, respectively3. Heterotopic cervical heart transplantation in rats was described by Heron in 1971 using Teflon cuffs made from 16 G (1.6 mm outer diameter) intravenous catheters4. Chen5 and Matsuura et al.6 later reported heterotopic cervical heart transplantation in mice in 1991, whose techniques differed primarily in their method of re-anastomosis. Chen&#39;s approach involved sutured anastomoses of the donor&#39;s ascending aorta to the carotid artery of the recipient and the donor&#39;s pulmonary artery to the external jugular vein of the recipient5. Due to the advanced technical skill required for these microsurgical sutured anastomoses, a significant amount of time and experience was necessary to achieve a high success rate. Matsuura et al. described a method utilizing a non-suture cuff technique, similar to that used by Heron, which involved end-to-end anastomoses using the extra-luminal placement of cuffs. He fashioned Teflon cuffs from 22 G (0.8 mm outer diameter) and 24 G (0.67 mm outer diameter) intravenous catheters and placed them over the recipient&#39;s external jugular vein and common carotid artery, respectively6. These cuffs were then placed inside the donor&#39;s pulmonary artery and aorta and secured by tying a suture ligature around the connection. This approach translated into an improved success rate. Most importantly, it resulted in a shortening of the time required to complete both cervical anastomoses, thus reducing the warm ischemic time of the graft to less than one-third of that utilizing the abdominal suturing method. Furthermore, since the cuffs are placed around the external surface of the vessel, there is no foreign body exposed to the vessel lumen, which largely reduces the possibility of thrombosis after surgery7. Meanwhile, utilization of the cuff technique provides support around the vessels at the site of the anastomosis without requiring any suturing, which reduces the risk of bleeding after revascularization6.
Numerous revisions of this technique have been proposed. To accommodate the short length of the mouse common carotid artery (approximately 5 mm), Tomita et al.8 developed a modification of this technique with a smaller arterial cuff (0.6 mm outer diameter) while omitting holding sutures and pulling the artery directly through the cuff with fine forceps instead. Wang et al. further simplified this approach by placing 22 G and 24 G cuffs on the donor&#39;s right pulmonary artery and recipient&#39;s right common carotid artery, respectively9. Various reports have described modifications to these approaches, including the use of specialized cuffs, microsurgical clamps, vessel dilators, and cardioplegia10,11,12. Notably, all of these methods involve the retrograde circulation of blood through the heart, with blood flowing from the recipient common carotid artery to the donor aorta, the coronary arteries, the coronary sinus, then emptying into the right atrium and exiting from the pulmonary artery into the recipient external jugular vein.
Compared to engraftment in the abdomen, cervical cardiac transplantation offers multiple advantages. As previously mentioned, cervical exposure allows for quicker revascularization and shorter warm ischemic times6. The cervical method is also less invasive and is associated with shorter postoperative recovery times as it avoids a laparotomy6. Importantly, end-to-end anastomoses with cuffs can be performed instead of end-to-side anastomoses, which decreases the risk of complications such as anastomotic bleeding. The abdominal approach also poses an increased risk of developing thrombotic complications in the abdominal aorta or inferior vena cava, leading to spinal cord ischemia and hindlimb paralysis. The superficial cervical location of the transplant allows for easy access to graft viability assessment by palpation, electrocardiography, and invasive or non-invasive imaging. Although the cervical grafts resume spontaneous cardiac activity following reperfusion, they do not significantly impact the systolic and diastolic parameters of the recipient. This model provides valuable insight for studying cellular responses following transplantation, such as ischemia-reperfusion injury and graft rejection. Furthermore, this model offers an ideal approach to allow for post-transplant imaging, such as intravital two-photon microscopy or positron emission tomography (PET) imaging. To this end, our laboratory has previously reported methods to image moving tissues and organs in the mouse, including beating murine hearts and aortic arch grafts following heterotopic cervical transplantation&#160;to visualize leukocyte trafficking during ischemia-reperfusion injury and within atherosclerotic plaques, respectively13,14,15. Additionally, due to its superficial location and ease of exposure, this model is suitable for cardiac re-transplantation16.
This report describes a technique that allows for laminar blood flow with the external placement of the vascular cuffs on the vessels from which blood flow originates. This allows for a smooth transition of blood flow from one vessel to the next, avoiding the exposure of the distal vessel edge into the vascular lumen. Additionally, the technique utilizes a larger 20 G cuff, instead of previously used 22 G cuffs, for the donor pulmonary artery to ensure ample return of blood flow to the recipient.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>6-0 braided silk ties</td>
			<td>Henry Schein Inc</td>
			<td>7718729</td>
			<td></td>
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		<tr>
			<td>0.75% Providone iosine scrub</td>
			<td>Priority Care Inc</td>
			<td>NDC 57319-327-0</td>
			<td></td>
		</tr>
		<tr>
			<td>10-0 nylon suture</td>
			<td>Surgical Specialties Corporation</td>
			<td>AK-0106</td>
			<td></td>
		</tr>
		<tr>
			<td>655-nm nontargeted Q-dots</td>
			<td>Invitrogen</td>
			<td>Q21021MP</td>
			<td></td>
		</tr>
		<tr>
			<td>70% Ethanol</td>
			<td>Pharmco Products Inc</td>
			<td>111000140</td>
			<td></td>
		</tr>
		<tr>
			<td>8-0 braided silk ties</td>
			<td>Henry Schein Inc</td>
			<td>1005597</td>
			<td></td>
		</tr>
		<tr>
			<td>Adson forceps</td>
			<td>Fine Science Tools Inc</td>
			<td>91127-12</td>
			<td></td>
		</tr>
		<tr>
			<td>BALB/c and C57BL/6 mice (6-8 weeks)</td>
			<td>Jackson Laboratories</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bipolar coagulator</td>
			<td>Valleylab Inc</td>
			<td>SurgII-20, E6008/E6008B</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen (Rimadyl) injection</td>
			<td>Transpharm</td>
			<td>35844</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen (Rimadyl) oral chewable tablet</td>
			<td>Transpharm</td>
			<td>38995/37919</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom-built 2P microscope running ImageWarp acquisition software</td>
			<td>A&#38;B Software</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont no. 5 forceps</td>
			<td>Fine Science Tools Inc</td>
			<td>11251-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine vannas style spring scissors</td>
			<td>Fine Science Tools Inc</td>
			<td>15000-03</td>
			<td></td>
		</tr>
		<tr>
			<td>GraphPad Prism 5.0</td>
			<td>Sun Microsystems Inc.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Halsey needle holder</td>
			<td>Fine Science Tools Inc</td>
			<td>91201-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Halsted-Mosquito clamp curved tip</td>
			<td>Fine Science Tools Inc</td>
			<td>91309-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Harvard Apparatus mouse ventilator model 687</td>
			<td>Harvard Apparatus</td>
			<td>MA1 55-0001</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin solution (100 U/mL)</td>
			<td>Abraxis Pharmaceutical Products</td>
			<td>504031</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaris</td>
			<td>Bitplane</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine (50 mg/kg)</td>
			<td>Wyeth</td>
			<td>206205-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope&#8212;Leica Wild M651 &#215; 6&#8211;40 magnification</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Moria extra fine spring scissors</td>
			<td>Fine Science Tools Inc</td>
			<td>15396-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Ohio isoflurane vaporizer</td>
			<td>Parkland Scientific</td>
			<td>V3000i</td>
			<td></td>
		</tr>
		<tr>
			<td>Qdots</td>
			<td>ThermoFisher</td>
			<td>1604036</td>
			<td></td>
		</tr>
		<tr>
			<td>S&#38;T SuperGrip Forceps angled tip</td>
			<td>Fine Science Tools Inc</td>
			<td>00649-11</td>
			<td></td>
		</tr>
		<tr>
			<td>S&#38;T SuperGrip Forceps straight tip</td>
			<td>Fine Science Tools Inc</td>
			<td>00632-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile normal saline (0.9% (wt/vol) sodium chloride</td>
			<td>Hospira Inc</td>
			<td>NDC 0409-4888-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile Q-tips (tapered mini cotton tipped 3-inch applicators)</td>
			<td>Puritan Medical Company LLC</td>
			<td>823-WC</td>
			<td></td>
		</tr>
		<tr>
			<td>Surflow 20 gauge 1/4-inch Teflon angiocatheter</td>
			<td>Terumo Medical Corporation</td>
			<td>SR-OX2032CA</td>
			<td></td>
		</tr>
		<tr>
			<td>Surflow 24 gauge 3/4-inch Teflon angiocatheter</td>
			<td>Terumo Medical Corporation</td>
			<td>R-OX2419CA</td>
			<td></td>
		</tr>
		<tr>
			<td>ThermoCare Small Animal ICU System (recovery settings 3 L/min O2, 80 &#176;C, 40% humidity)</td>
			<td>Thermocare Inc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>VetBond</td>
			<td>Santa Cruz Biotechnology SC361931</td>
			<td>NC0846393</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine (10 mg/kg)</td>
			<td>Lloyd Laboratories</td>
			<td>139-236</td>
			<td></td>
		</tr>
	</tbody>
</table>

mouse heterotopic cervical cardiac transplantation utilizing vascular cuffs

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

This mouse cervical heterotopic cardiac transplantation model has been utilized to perform over 1,000 transplants in our laboratory, with a survival rate of approximately 97%. The success rate is slightly higher than previous reports using other cervical heterotopic heart transplantation techniques in mice10,11,20. This could potentially be attributed to the larger 20 G cuff placed on the donor pulmonary artery to ensure ample return of blood flow to the recipient (Figure 1B,C). Additionally, the alignment of blood flow with cuff placement in the present technique minimizes the risk of thrombosis and anastomotic turbulence (Figure 1,2). While magnetic resonance imaging (MRI) or ultrasound could assess the turbulence of graft perfusion22,23, we have not yet utilized these techniques in the experiments. Intraoperative death using this technique is rare for experienced microsurgeons. Postoperative mortality is most often due to bleeding complications. The mean recipient operation time was 36.5 &#177; 3.5 min, with an average cold ischemia time of 20 min. For survival studies, cardiac grafts were assessed daily by direct visualization and digital palpation of the heartbeat. Mice are typically sacrificed for graft evaluation around 7-14 days postoperatively. Intravital two-photon imaging is a terminal procedure usually performed early after transplantation to evaluate leukocyte trafficking (Figure 3).
Most syngeneic transplants maintained strong heartbeats until sacrifice, up to 6 months after transplantation. On gross inspection, most syngeneic grafts appeared normal and histologic examination revealed no evidence of rejection. All non-immunosuppressed allogeneic transplants (BALB/c into B6) developed a diminished heartbeat within 1-2 weeks after engraftment. Excised allogeneic grafts from such mice were grossly dilated, and histologic examination showed diffuse infiltration of lymphocytes and areas of myocardial necrosis.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64089/64089fig01.jpg" /> 
Figure 1: Preparation of heart graft for transplantation. (A) The heart is excised from the donor mouse. (B,C) The pulmonary artery trunk is exposed and pulled through a 20 G cuff, folded back, and secured with a 10-0 nylon suture. (D) A 10-0 nylon suture is placed through the edge of the recipient&#39;s external jugular vein and fixed to underlying tissue. (E) A 10-0 nylon suture is placed through the edge of the donor aorta and secured to the underlying tissue adjacent to the recipient carotid artery. (F) The recipient&#39;s common carotid artery cuff is inserted into the donor aorta and secured with an 8-0 silk suture. (G) The donor pulmonary artery cuff is inserted into the recipient&#39;s external jugular vein and secured with an 8-0 silk suture. (H) Proximal slipknot on the recipient&#39;s external jugular vein is released, followed by the release of the common carotid artery slipknot. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64089/64089fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64089/64089fig02.jpg" /> 
Figure 2: Intra-operative view of cardiac graft. A 1 mm 20 G cuff is pulled over the donor&#39;s pulmonary artery and secured with a 10-0 nylon tie. A 0.6 mm 24 G cuff is pulled over the recipient&#39;s right common carotid artery and secured with a 10-0 nylon tie. Anchor sutures (10-0 nylon) are placed in the wall of the donor aorta and the recipient&#39;s right external jugular vein and secured to underlying tissue to prevent movement during cuff insertion. (AO = aorta, PA = pulmonary artery, CCA = common carotid artery, EJV = external jugular vein). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64089/64089fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64089/64089fig03.jpg" /> 
Figure 3: Intravital two-photon imaging of leukocyte dynamics in the heart graft. Intravital two-photon imaging of beating heart transplanted from B6 mouse to B6 LysM-GFP recipient demonstrates trafficking of recipient neutrophils into the cardiac graft tissues between 2-3 h postoperatively. (Green = neutrophils, red = vessels labeled with quantum dots). Scale bar = 20 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64089/64089fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Complication</td>
			<td>Possible Causes</td>
			<td>Solutions</td>
		</tr>
		<tr>
			<td rowspan="2">Recipient death</td>
			<td>Hypothermia</td>
			<td>Heating pad</td>
		</tr>
		<tr>
			<td>Dehydration</td>
			<td>0.9% saline i.p. postoperatively</td>
		</tr>
		<tr>
			<td rowspan="2">Poor graft perfusion</td>
			<td>Carotid artery torsion</td>
			<td>Re-anastomosis, or</td>
		</tr>
		<tr>
			<td>Thrombus or air emboli</td>
			<td>Open arterial anastomosis and flush with heparinized saline</td>
		</tr>
		<tr>
			<td rowspan="2">Venous obstruction</td>
			<td rowspan="2">Thrombus or air emboli</td>
			<td>Re-anastomosis, or</td>
		</tr>
		<tr>
			<td>Open venous anastomosis and flush with heparinized saline</td>
		</tr>
		<tr>
			<td rowspan="3">Postoperative bleeding</td>
			<td>Bleeding jugular vein branches</td>
			<td>Ligate jugular vein branches</td>
		</tr>
		<tr>
			<td></td>
			<td>Cotton swab compression</td>
		</tr>
		<tr>
			<td>Loose cuffs</td>
			<td>Tighten cuffs</td>
		</tr>
		<tr>
			<td>Weak heartbeat</td>
			<td>Cold cardiac graft</td>
			<td>Drip warm saline on surface of heart</td>
		</tr>
		<tr>
			<td>Graft twisting</td>
			<td>Improper graft position</td>
			<td>Ensure graft is properly oriented before skin closure</td>
		</tr>
		<tr>
			<td>Erratic activity (eg. running in circles)</td>
			<td>Cerebral ischemia</td>
			<td>Ligate common carotid artery inferior to carotid bifurcation</td>
		</tr>
	</tbody>
</table>
Table 1: Troubleshooting for complications. Commonly encountered complications with solutions.

Watch this Scientific Journal Video about Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs at JoVE.com

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after ...

mouse-heterotopic-cervical-cardiac-transplantation-utilizing-vascular

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

2.8K Views.  Washington University School of Medicine. Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Video: Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Orthotopic Hind-Limb Transplantation in Rats

Composite tissue allotransplantation (CTA) now represents a valid therapeutic option after the loss of a hand, forearm or digits and has become a novel therapeutic entity in reconstructive surgery. However, long term high-dose multi-drug immunosuppressive therapy is required to ensure graft survival, bearing the risk of serious side effects which halters broader application. Further progression in this field may depend on better understanding of basic immunology and ischemia reperfusion injury in composite tissue grafts.
To date, orthotopic hind limb transplantation in rats has been the preferred rodent model for reconstructive transplantation (RT), however, it is an extremely demanding procedure that requires extraordinary microsurgical skills for reattachment of vasculature, bones, muscles and nerves.
We have introduced the vascular cuff anastomosis technique to this model, providing a rapid and reliable approach to rat hind limb transplantation. This technique simplifies and shortens the surgical procedure and enables surgeons with basic microsurgical experience to successfully perform the operation with high survival and low complication rates. The technique seems to be well suited for immunological as well as ischemia reperfusion injury (IRI) studies.

Here we describe the orthotopic rat hind-limb transplantation procedure, which seems to be the gold standard in vivo model for composite tissue allotransplantation research.

Composite tissue allotransplantation (CTA) now represents a valid therapeutic option after the loss of a hand, forearm or digits and has become a novel ...

Isolation of Functional Cardiac Immune Cells

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5 These immune cells, which include mast cells, T-cells and macrophages; store and release a variety of biologically active mediators including cytokines and proteases such as tryptase.6-8 These mediators have been shown to be key players in extracellular matrix metabolism by activating matrix metalloproteinases or causing collagen accumulation by modulating the cardiac fibroblasts' function.9-11 However, available techniques for isolating cardiac immune cells have been problematic because they use bacterial collagenase to digest the myocardial tissue. This technique causes activation of the immune cells and thus a loss of function. For example, cardiac mast cells become significantly less responsive to compounds that cause degranulation.12 Therefore, we developed a technique that allows for the isolation of functional cardiac immune cells which would lead to a better understanding of the role of these cells in cardiac disease.13, 14
This method requires a familiarity with the anatomical location of the rat's xiphoid process, axilla and falciform ligament, and pericardium of the heart. These landmarks are important to increase success of the procedure and to ensure a higher yield of cardiac immune cells. These isolated cardiac immune cells can then be used for characterization of functionality, phenotype, maturity, and co-culture experiments with other cardiac cells to gain a better understanding of their interactions.

This method for isolating functional immune cells from the heart provides an alternative to the conventional methods of collagenase digestion, which causes unwanted immune cell activation, resulting in a decreased responsiveness of these cells. Our method of isolation yields functional cardiac immune cells by avoiding problems associated with enzymatic digestion.

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5  These immune cells, which ...

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 ...

Murine Full-thickness Skin Transplantation

Murine full-thickness skin transplantation is a well-established in vivo model to study alloimmune response and graft rejection. Despite its limited application to humans, skin transplantation in mice has been widely employed for transplantation research. The procedure is easy to learn and perform, and it does not require delicate microsurgical techniques nor extensive training. Moreover, graft rejection in this model occurs in a very reproducible immunological reaction and is easily monitored by direct inspection and palpation. In addition, secondary skin transplantation with donor-matched or third-party skin grafts can be performed on more complex transplant models as an alternative and uncomplicated method to assess donor-specific tolerance. The complications are low and are in general limited to anesthesia overdose or respiratory distress after the procedure. Graft failure, on the other hand, occurs commonly as a result of poor preparation of the graft, incorrect positioning in the graft bed, or inappropriate placement of the bandage. In this article, we present a protocol for full-thickness skin transplantation in mice and describe the important steps necessary for a successful procedure.

Murine full-thickness skin transplantation is a well-established model to study rejection in an alloimmune setting. Here, we provide a tutorial of each step involved in performing a BALB/c--&#62;C57BL/6 full-thickness skin transplant.

Murine full-thickness skin transplantation is a well-established in vivo model to study alloimmune response and graft rejection. Despite its limited ...

In Vitro and In Vivo Assessment of T, B and Myeloid Cells Suppressive Activity and Humoral Responses from Transplant Recipients

The main concern in transplantation is to achieve specific tolerance through induction of regulatory cells. The understanding of tolerance mechanisms requires reliable models. Here, we describe models of tolerance to cardiac allograft in rat, induced by blockade of costimulation signals or by upregulation of immunoregulatory molecules through gene transfer. Each of these models allowed in vivo generation of regulatory cells such as regulatory T cells (Tregs), regulatory B cells (Bregs) or regulatory myeloid cells (RegMCs). In this manuscript, we describe two complementary protocols that have been used to identify and define in vitro and in vivo regulatory cell activity to determine their responsibility in tolerance induction and maintenance. First, an in vitro suppressive assay allowed rapid identification of cells with suppressive capacity on effector immune responses in a dose dependent manner, and can be used for further analysis such as cytokine measurement or cytotoxicity. Second, the adoptive transfer of cells from a tolerant treated recipient to a newly irradiated grafted recipient, highlighted the tolerogenic properties of these cells in controlling graft directed immune responses and/or converting new regulatory cells (termed infectious tolerance). These methods are not restricted to cells with known phenotypic markers and can be extended to any cell population. Furthermore, donor directed allospecificity of regulatory cells (an important goal in the field) can be assessed by using third party donor cells or graft either in vitro or in vivo. Finally, to determine the specific tolerogenic capacity of these regulatory cells, we provide protocols to assess the humoral anti-donor antibody responses and the capacity of the recipient to develop humoral responses against new or former known antigens. The models of tolerance described can be used to further characterize regulatory cells, to identify new biomarkers, and immunoregulatory molecules, and are adaptable to other transplantation models or autoimmune diseases in rodent or human.

In Vitro and In Vivo Assessment of T, B and Myeloid Cells Suppressive Activity and Humoral Responses from Transplant Recipients

Here, we present a protocol to induce tolerance in transplantation, and assess in vitro and in vivo the suppressive capacity of distinct cell subsets from the recipient and the immune status of the recipient toward donor or exogenous antigens.

The main concern in transplantation is to achieve specific tolerance through induction of regulatory cells. The understanding of tolerance mechanisms ...

Assessment of the Cytotoxic and Immunomodulatory Effects of Substances in Human Precision-cut Lung Slices

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new therapeutics. Additionally, registration of new substances requires appropriate risk assessment with adequate testing systems to avoid the risk of individuals being harmed, for example, in the working environment. Such risk assessments are usually conducted in animal studies. In view of the 3Rs principle and public skepticism against animal experiments, human alternative methods, such as precision-cut lung slices (PCLS), have been evolving. The present paper describes the ex vivo technique of human PCLS to study the immunomodulatory potential of low-molecular-weight substances, such as ammonium hexachloroplatinate (HClPt). Measured endpoints include viability and local respiratory inflammation, marked by altered secretion of cytokines and chemokines. Pro-inflammatory cytokines, tumor necrosis factor alpha (TNF-&#945;), and interleukin 1 alpha (IL-1&#945;) were significantly increased in human PCLS after exposure to a sub-toxic concentration of HClPt. Even though the technique of PCLS has been substantially optimized over the past decades, its applicability for the testing of immunomodulation is still in development. Therefore, the results presented here are preliminary, even though they show the potential of human PCLS as a valuable tool in respiratory research.

In view of the 3Rs principle, respiratory models as alternatives to animal studies are evolving. Especially for risk assessment of respiratory substances, there is a lack of appropriate assays. Here, we describe the use of human precision-cut lung slices for the assessment of airborne substances.

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new ...

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 ...

Visualization of SARS-CoV-2 using Immuno RNA-Fluorescence In Situ Hybridization

This manuscript provides a protocol for in situ hybridization chain reaction (HCR) coupled with immunofluorescence to visualize severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) RNA in cell line&#160;and three-dimensional (3D) cultures of human airway epithelium. The method allows highly specific and sensitive visualization of viral RNA by relying on HCR initiated by probe localization. Split-initiator probes help amplify the signal by fluorescently labeled amplifiers, resulting in negligible background fluorescence in confocal microscopy. Labeling amplifiers with different fluorescent dyes facilitates the simultaneous recognition of various targets. This, in turn, allows the mapping of the infection in tissues to better understand viral pathogenesis and replication at the single-cell level. Coupling this method with immunofluorescence may facilitate better understanding of host-virus interactions, including alternation of the host epigenome and immune response pathways. Owing to sensitive and specific HCR technology, this protocol can also be used as a diagnostic tool. It is also important to remember that the technique may be modified easily to enable detection of any RNA, including non-coding RNAs and RNA viruses that may emerge in the future.

Here, we describe a simple method that combines RNA fluorescence in situ&#160;hybridization (RNA-FISH) with immunofluorescence to visualize severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) RNA. This protocol may increase understanding of the molecular characteristics of SARS-CoV-2 RNA-host interactions at a single-cell level.

This manuscript provides a protocol for in situ hybridization chain reaction (HCR) coupled with immunofluorescence to visualize severe acute respiratory ...

Purification and Expansion of Mouse Invariant Natural Killer T Cells for in vitro and in vivo Studies

Invariant Natural Killer T (iNKT) cells are innate-like T Lymphocytes expressing a conserved semi-invariant T cell receptor (TCR) specific for self or microbial lipid antigens presented by the non-polymorphic MHC class I-related molecule CD1d. Preclinical and clinical studies support a role for iNKT cells in cancer, autoimmunity and infectious diseases. iNKT cells are very conserved throughout species and their investigation has been facilitated by mouse models, including CD1d-deficient or iNKT-deficient mice, and the possibility to unequivocally detect them in mice and men with CD1d tetramers or mAbs specific for the semi-invariant TCR. However, iNKT cells are rare and they need to be expanded to reach manageable numbers for any study. Because the generation of primary mouse iNKT cell line in vitro has proven difficult, we have set up a robust protocol to purify and expand splenic iNKT cells from the iV&#945;14-J&#945;18 transgenic mice (iV&#945;14Tg), in which iNKT cells are 30 times more frequent. We show here that primary splenic iV&#945;14Tg iNKT cells can be enriched through an immunomagnetic separation process, yielding about 95-98% pure iNKT cells. The purified iNKT cells are stimulated by anti-CD3/CD28 beads plus IL-2 and IL-7, resulting in 30-fold expansion by day +14 of the culture with 85-99% purity. The expanded iNKT cells can be easily genetically manipulated, providing an invaluable tool to dissect mechanisms of activation and function in vitro and, more importantly, also upon adoptive transfer in vivo.

We describe a rapid and robust protocol to enrich invariant natural killer T (iNKT) cells from mouse spleen and expand them in vitro to suitable numbers for in vitro and in vivo studies.

Invariant Natural Killer T (iNKT) cells are innate-like T Lymphocytes expressing a conserved semi-invariant T cell receptor (TCR) specific for self or ...

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

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Purification and Expansion of Mouse Invariant Natural Killer T Cells for in vitro and in vivo Studies