We want to thank Britta Trautewig, Corinna L&#246;bbert and Ingrid Meder for their commitment.

All animal experiences have been performed according to the guidelines of the local Ethics Animal Review Board of the regional authorities for consumer protection and food safety of Lower Saxony (LAVES, Oldenburg, Germany) with the approval IDs 12/0768 and 17/2472.
1. Heart explantation and perfusion
NOTE: As graft donors, female or male rats at an age of 7-22 weeks were used.
<ol>
	<li>Anesthetize the donor rat by isoflurane inhalation (induction at 5% and maintenance at 3% with an O2 flow of 1 L/min). Inject 5 mg of Carprofen subcutaneously per kg of bodyweight for perioperative analgesia and check for the absence of the toe pinch withdrawal reflex.</li>
	<li>Apply eye lubricant and remove the abdominal and thoracic fur using a mechanical clipper.</li>
	<li>Place the donor in a supine position, fix the limbs at the base of the operation table with elastic bands and sterilize the skin with 70% ethanol or another sufficient alternative.</li>
	<li>Incise the skin in longitudinal direction and after application of local anesthetic (e.g., lidocaine 0.2%) perform a median laparotomy by using scissors.</li>
	<li>Insert retractors, mobilize the intestine to the left of the donor, and expose the inferior vena cava with sterilized cotton swabs.</li>
	<li>For anticoagulation, inject 500 I.U. of heparin dissolved in 1 mL of ice-cold isotonic saline solution intravenously by puncturing the inferior vena cava. Stop the bleeding at the puncture site by light compression with a cotton swab after retraction of the needle (Figure 3A).</li>
	<li>Incise the diaphragm and perform lateral thoracotomy to both sides of the donor.</li>
	<li>Pin the mobilized ventral wall of the thorax onto the operation table.</li>
	<li>Remove the pericardium and the vagal nerve by blunt preparation using two micro-needle holders.</li>
	<li>Perform transection of abdominal vessels in order to exsanguinate the donor and unload the heart.</li>
	<li>Insert the blunt branch of a probe pointed scissors into the transvers pericardial sinus and separate the ascending aorta and pulmonary artery as distal as possible under light caudal traction of the heart with a wetted compress (Figure 3B).</li>
	<li>Place a single 5-0 ligature around the superior and inferior vena cava and the pulmonary veins and tighten it as dorsal as possible (Figure 3C).</li>
	<li>Sever the tissue dorsal to the ligature and extract the heart (Figure 3D).</li>
	<li>Perfuse the explanted heart with an 18 G cannula from an intravenous catheter through the ascending aorta and the pulmonary artery with 30 mL of ice-cold, isotone saline solution supplemented with 1000 I.U. of heparin and place the heart in a 15 mL tube filled with saline solution on ice (Figure 3E-F).</li>
</ol>
2. Heart implantation
NOTE: As recipients, 10-14 weeks old female or male rats were used. Donors and recipients were approximately weight matched.
<ol>
	<li>Perform anesthesia of the recipient rat by also using isoflurane inhalation (induction at 5% and maintenance at 1.5-2% with an O2 flow of 1 L/min). Inject 5 mg of Carprofen subcutaneously per kg of bodyweight for perioperative analgesia&#160;and check for the absence of the toe pinch withdrawal reflex.</li>
	<li>Apply eye lubricant, remove the abdominal fur, fix the limbs and sterilize the skin analogously to the donor preparation. For optimal postoperative outcome, perform the operation on a heating mat to prevent intraoperative hypothermia.</li>
	<li>After longitudinal incision of the skin, apply a local anesthetic, such as lidocaine (0.2%), on the abdominal fascia. Open the abdominal cavity by median laparotomy and insert retractors.</li>
	<li>Mobilize the intestine to the upper left side of the recipient and place it in a warm, wetted compress.</li>
	<li>After mobilizing the duodenum and proximal jejunum, respectively, using the surgical microscope (or magnifying spectacles) with a 5-7x magnification, expose the abdominal aorta and inferior vena cava by blunt preparation with cotton swabs. Do not separate the abdominal vessels.</li>
	<li>Elevate the abdominal vessels using two micro needle holders without injuring the lumbar veins and position the Cooley vascular clamp (Figure 4A).</li>
	<li>Puncture the abdominal vessels with a 30-45&#176; arched 27 G cannula (Figure 4B).</li>
	<li>Enlarge the puncture site using Potts scissors to create a longitudinal incision that matches the size of the lumen of the donor vessels (Figure 4C-D) and perfuse the recipient vessels with saline solution in order to remove clots and prevent postoperative thrombosis.</li>
	<li>Place the graft into the situs and fixate the donor ascending aorta to the recipient abdominal aorta by two simple interrupted stitches (8-0 monofilament non-resorbable suture) at the cranial and caudal corner of the longitudinal incision (Figure 4E).</li>
	<li>Anastomose the ascending aorta of the donor with the abdominal aorta of the recipient by a running 8-0 monofilament suture in two steps: first, place the graft to the right of the recipient vessels and perform the first half of the anastomosis (Figure 4E). Subsequently, place the graft to the left of the recipient vessels and perform the second half of the anastomosis (Figure 4F).</li>
	<li>Fixate the donor pulmonary artery to the inferior vena cava analogously to the aortal anastomosis (8-0 monofilament non-resorbable suture). Suture the first half of the venous anastomosis from the intraluminal side of the vessel (Figure 4G-H).</li>
	<li>Flush the anastomoses with saline directly before tightening the knots to prevent peripheral embolism.</li>
	<li>Place a hemostatic gauze around both anastomoses and carefully release the Cooley vascular clamp so that the reperfusion of the graft can begin. Handle bleeding along the anastomoses by light compression with sterilized cotton swabs. 
	NOTE: The graft should start beating after around 60 s.</li>
	<li>Replace the intestine in a meander like fashion. Make sure that there are no malrotations of the mesenteric radix to prevent intestinal necrosis or mechanical obstruction.</li>
	<li>Close the abdominal muscles/fascia and skin separately using continuous 3-0 polyfilament running sutures.</li>
</ol>
3. Postoperative care
<ol>
	<li>For postoperative analgesia, supply the recipients with an additional subcutaneous injection of 5 mg of Carprofen per kg of bodyweight on the first postoperative day (POD). Additionally, add 1 g of Metamizol to 500 mL of drinking water until the third POD.</li>
	<li>Start monitoring the heart graft function by daily abdominal palpation on the third POD. 
	NOTE: In case of graft failure before the third POD, a surgical rather than an immunological failure should be considered. However, this of course depends on the chosen strain combination and the respective immunological model (e.g., hyperacute rejection after prior immunization).</li>
	<li>Following graft rejection, extract tissues like the draining retroperitoneal lymph nodes cranial of the anastomoses, the spleen, the blood, the thymus and the graft for further immunological analyses via flow cytometry or immunohistochemistry.</li>
</ol>
4. Enzymatic digestion of the heart and subcutaneous injection of heart cells in the ear
<ol>
	<li>Perform heart explantation and perfusion analogously to heterotopic heart transplantation (see step 1).</li>
	<li>Shred the heart in 3 mm x 3 mm blocks using a sterile scalpel or sterile scissors and incubate it for 30 min at 37 &#176;C in culture medium containing 0.5 mg/mL collagenase. 
	NOTE: It is important to use culture medium containing penicillin, streptomycin and glutamine without fetal calf serum (FCS) particularly as FCS inhibits collagenase digestion.</li>
	<li>Add the digested tissue to a large-pored sieve, while removing the culture medium and mince thoroughly to get a suspension of vital cardiac muscle cells, mostly dead single heart cells and remaining blood cells. Wash the cell suspension twice with sterile isotonic saline solution. 
	NOTE: Centrifugation settings: 10 min, 200 x g, 20 &#176;C</li>
	<li>Filter the suspension using a 40 &#181;m cell strainer and collect the vital cell congeries by flushing the cell strainer with 5-10 mL of isotonic saline solution.</li>
	<li>After centrifugation, resuspend the cardiac muscle cells in saline solution dissolved at a concentration of 5x105 cells/mL and draw the cell solution up into a 1 mL syringe.</li>
	<li>Perform anesthesia analogously to the protocol described for the recipient narcosis (see step 2) for heterotopic heart transplantation.</li>
	<li>Place the recipient in a lateral position and fix the ear with a finger using double sided tape (Figure 5A).</li>
	<li>Inject 20 &#181;L of the cardiac muscle cell solution (containing 1 x 104 cells) via a 27 G cannula s.c. close to the visual capillary vessels into the recipient&#39;s ear (Figure 5B).</li>
	<li>After a defined observation period (depending on the chosen strain combination and strength of rejection), extract the draining cervical lymph nodes and perform further analyses such as flow cytometry or co-cultures (Figure 5C). 
	NOTE: Furthermore, histological analysis of the pinna can be performed to determine cell infiltration.</li>
</ol>

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The authors have nothing to disclose.

The previously described method of heterotopic cardiac transplantation in rats is mainly based on the description of Ono and Lindsey in 19693. Since then, several modifications have been introduced in various species leading to a wide diversity of this model. Combining several of these modifications and introducing our own experience resulting from over 30 years of performing heterotopic heart transplants in the laboratory, we created a feasible surgical approach, which does not require long training periods or surgical background. In the following, we will discuss general limitations of this model and underline critical steps of the protocol. Furthermore, we will emphasize the benefits of combining heterotopic heart transplantation with a novel method of in-ear injection of cardiac muscle cells.
Anesthesia and general complications 
Although it has been reported that isoflurane anesthesia is superior to injection anesthesia regarding early survival after heterotopic heart transplantation, intraoperative respiratory depression still represents one of the most common complications and thus requires a careful narcosis management26. Instead of serial graft explantation and implantation, respectively, we advise beginning the preparation of the recipient and the abdominal vessels immediately after thoracotomy of the donor animal, particularly because operation times of less than one hour are associated with a better outcome regarding graft and recipient survival26,27. Besides already mentioned complications such as hypothermia due to long operating times and missing heating mats, and intestinal necrosis or obstruction by unphysiological placement of the intestine, a paresis of hinder limbs represents a further complication, which can be prevented by atraumatic vascular clamping and thorough flushing of the anastomoses to avoid peripheral embolism28,29.
Ligature of pulmonary and caval veins and bleeding complications 
In order to reduce warm ischemia time, we use one single ligature for both caval and all four pulmonary veins. As a possible complication resulting from a too proximal/ventral placement of the ligature, the disruption of venous backflow by occlusion of the sinus coronarius has to be mentioned. In case of severe bleeding after removal of the Cooley clamp and visible beating of the graft, the ligature has to be checked for insufficiency immediately. We observed this type of complication especially if the dissection of the pulmonary veins was performed too close to the ligature while removing the graft from the thoracic cavity. Otherwise, severe bleeding is mainly caused by insufficiency of the vascular anastomoses. Additionally, a coronary artery running along the ascending aorta which is often severed during explantation is described to cause lethal bleeding upon reperfusion27.
Ischemia time and perfusion 
Regardless of the transplantation model, it is always indispensable to reduce ischemia time, especially as the heart is considered as a vulnerable organ regarding ischemia damage. By performing surgery with two surgeons, we are able to achieve a minimum warm and cold ischemic time and therefore forgo the usage of cardioplegic solutions in order to reduce ischemia-reperfusion damage30. Generally, the perfusion of the graft plays a key role and is essential to cool the graft and remove blood cells, which could result in thrombosis or embolism. Whereas low perfusion pressures lead to insufficient perfusion and thus to an incomplete removal of blood cells, high perfusion pressures can cause endothelial damage31,32. We advise perfusion of the pulmonal artery as well as the ascending aorta until the coronary arteries are visibly flushed clear.
Incision of recipient vessels 
A critical step in the protocol constitutes the incision of the recipient vessels without causing damage to the vessels posterior wall: Schmid et al. described the benefit of aortotomy or venotomy performing a small transverse incision followed by longitudinal enlargement in cranial and caudal direction, which further leads to a decreased stenosis rate of aortal anastomosis33. In our&#160;model, the recipient vessels are punctured using a small cannula. Afterwards, the puncture site is enlarged by using Potts scissors to create a longitudinal incision. For a better feasibility, we recommend bending the tip of the cannula to an angle of 30-45&#176; leading to a decreased risk of damaging the vessels posterior wall. We did not observe clinically relevant vascular stenoses in any of our recipients. Similar benefits of opening recipient vessels by puncturing the abdominal aorta and the inferior vena cava with a cannula have been described by Shan et al.34.
Mastering the model of heterotopic heart transplantation 
During the last decade, the model has been performed by researchers in the department with little or no surgical background. As stated above, we operate in pairs of surgeons, whereas the more experienced researcher is in charge of guaranteeing the success of the transplantation and at the same of gradually improving the skill set of the unexperienced researcher. After a short training period of approximately ten graft explantations and implantations in dead animals, the unexperienced researcher is in charge of performing the heart explantation and assisting the graft implantation in approximately ten live animals. Subsequently, the unexperienced researcher actively performs the vascular anastomoses, so that after approximately ten further transplantations, the former unexperienced researcher is usually capable of performing all critical steps of the model.
Applying this training concept, past publications from our department using heterotopic heart transplantation in rats showed no differences regarding morbidity, mortality or graft function despite several different teams of surgeons13,18,19,20,21,22,23,24.
Of note, performing the vascular anastomoses represents the most critical step in this protocol and solid organ transplant models in general. We, therefore, recommend prolonged training periods using dead animals until anastomoses are performed accurate and fast, especially if an experienced researcher is not available for guidance in live animals.
General advantages and disadvantages of the model 
Whereas spontaneous tolerance induction is often described as a phenomenon in liver transplantation and also observed in kidney transplantation, the heart is considered as a rather immunogenic organ and thus enables reliable rejection in transplant models35,36. Contrary to these findings, we could also notice long-term survival and absence of rejection after heterotopic heart transplantation in certain donor-recipient combinations despite complete major histocompatibility complex disparity.
An often-mentioned criticism of heterotopic heart transplantation is the subjectivity of graft monitoring by manual palpation. Therefore, the model has been extended to femoral anastomosis techniques in order to facilitate the access for palpation and introduce further monitoring techniques such as transfemoral echocardiography15,16. On the other hand, Mottram et al. demonstrated that the monitoring of the graft via palpation correlates well with electrocardiographic measurements37. Thus, manual palpation in heterotopic heart transplants seems sufficient for monitoring graft function in an acute rejection model.
As a consequence of heterotopic placement and left ventricular unloading, the heart does not function under anatomical or physiological conditions assuming that this does not impact immunological analyses. Contrary to this assumption, it had been shown that cardiac remodeling resulting from left ventricular unloading during left ventricular assist device therapy leads to a decreased cytokine release38,39. On the other hand, Tang-Quan et al. described the unloaded setting as a more appropriate approach for immunological analysis, since long-term ischemic damage of the graft resulting from perfusion with partially deoxygenated blood in the left ventricular loaded model was observed40.
Although abdominal placement of the grafts offers surgical benefits in terms of practicability, it is difficult to harvest sufficient numbers of draining retroperitoneal lymph nodes upon graft rejection impairing further analyses. For this reason, we introduced a novel method of in-ear injection of allogenic cardiac muscle cells. Originating from parasitological research, this concept has not been applied for immunological analyses in transplantation, despite its feasibility41,42.The advantage of this model is the possibility of identifying and harvesting a significant number of draining lymph nodes, which offers the possibility of performing complex immunological analyses. Of note, both models could be combined in one recipient offering further insights into the mechanisms of rejection and tolerance in cell and organ transplantation in rats.
Our models of rat heart and cardiac muscle cell transplantation represent a practicable and well-studied approach and can be performed without further surgical training or background. Facing the fact that new cloning technologies for rats have been introduced and developed recently, these models offer vast opportunities for transplant immunological researchers.

Heterotopic heart transplantation is a frequently used experimental model for different investigations regarding transplantation tolerance, acute and chronic allograft rejection, ischemia-reperfusion injury, machine perfusion or cardiac remodelling. Among other advantages, the graft function can be monitored noninvasively by palpation and graft failure does not lead to a vital impairment of the recipient in contrast to other organs, such as kidneys or livers.
In 1964, Abbott et al. initially described heterotopic abdominal heart transplantation in rats1. Later, in 1966, the end-to-side technique for anastomoses was described by Tomita et al.2. The groundwork for the currently used model was reported by Ono and Lindsey in 19693. During the last decades, several modifications have been published to create different types of unloaded, partially loaded or loaded left ventricular heart grafts including combined heterotopic heart-lung transplantation4,5,6. For immunological analyses a non-volume loaded heart graft transplantation is most commonly performed. In this case, the blood flow retrogradely enters the donor ascending aorta and subsequently the coronary arteries. The venous drainage occurs along the coronary sinus into the right atrium and ventricle (Figure 1A-B). Therefore, the left ventricle is excluded from blood flow, apart from marginal amounts of blood from Thebesian veins. This also makes it a useful model for studying the pathophysiological mechanisms during left ventricular assist device therapy7.
Heterotopic heart transplantation has been performed in various species including mice, rabbits, pigs and has even been used as a uni- or biventricular assist device in humans8,9,10,11. The rat still represents a popular experimental animal for transplant models, especially since the graft survival times for different rat strain combinations have been well-defined in the past and a large number of immunological reagents are accessible12,13. Unlike mice, rats are larger making surgery and access to lymphatic tissue for immunological analyses more feasible12. Furthermore, the introduction of commercial cloning technologies in rats in recent years will most likely lead to a recurring interest in experimental rat models14.
In general, heterotopic heart grafts can be attached to the recipient vessels either by performing cervical or abdominal anastomosis. However, a few studies suggest that a femoral anastomosis facilitates improved monitoring due to better access for manual palpation or transfemoral echocardiography and thus allows a more precise detection of graft failure15,16.
It has been shown that there is no difference regarding operation time, complication rate, outcome and graft survival time between both anastomosis techniques17. Clearly, the availability of a sufficient number of draining lymph nodes must be mentioned as a benefit of cervical anastomosis; however, longer training periods are required. In contrast, the abdominal anastomosis is less complicated and equally valuable for immunological investigations, especially when combined with results from a novel method of in-ear injection of allogenic cardiac muscle cells and subsequent cervical lymphadenectomy. A combination of both models offers a broad spectrum of post-interventional immunological analyses.
The following protocol refers to operating in pairs of surgeons in order to reduce ischemia time. However, all experiments can be performed by a single person. The setup of instruments and materials for heart explantation and implantation is displayed in Figure 2A-B.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Anesthesia device (including isoflurane vaporizer)</td>
			<td>Summit Anesthesia Solutions</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Cannula (27 G)</td>
			<td>BD Microlance</td>
			<td>302200</td>
			<td></td>
		</tr>
		<tr>
			<td>Carprofen</td>
			<td>Pfizer</td>
			<td>Rimadyl 50 mg/mL</td>
			<td></td>
		</tr>
		<tr>
			<td>Cellstar Tubes (15 mL)</td>
			<td>GreinerBioOne</td>
			<td>188271</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer (40 &#181;m)</td>
			<td>BD Falcon</td>
			<td>2271680</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase Type CLSII</td>
			<td>Biochrome</td>
			<td>C2-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Compresses 5x5 cm</td>
			<td>Fuhrmann</td>
			<td>31501</td>
			<td></td>
		</tr>
		<tr>
			<td>Compresses 7.5x7.5 cm</td>
			<td>Fuhrmann</td>
			<td>31505</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton swabs</td>
			<td>Heinz Herenz Medizinalbedarf</td>
			<td>1032128</td>
			<td></td>
		</tr>
		<tr>
			<td>Dexpathenol (5 %)</td>
			<td>Bayer</td>
			<td>&#34;Bepanthen&#34;</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS BioWhittaker</td>
			<td>Lonza</td>
			<td>17-512F</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>B. Braun</td>
			<td>Aesculap BD557R</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>B. Braun</td>
			<td>Aesculap BD313R</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>B. Braun</td>
			<td>Aesculap BD35</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating mat</td>
			<td>Gaymar Industries</td>
			<td>&#34;T/Pump&#34;</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemostatic gauze</td>
			<td>Ethicon</td>
			<td>Tabotamp</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin-Natrium 25 000 I.E.</td>
			<td>Ratiopharm</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Isofluran CP</td>
			<td>CP-Pharma</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Large-pored sieve (stainless steel)</td>
			<td>Forschungswerkst&#228;tten Hannover Medical School</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine</td>
			<td>Astra Zeneca</td>
			<td>2 % Xylocain</td>
			<td></td>
		</tr>
		<tr>
			<td>Metamizol-Natrium</td>
			<td>Ratiopharma</td>
			<td>Novaminsulfon 500 mg/mL</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro forceps</td>
			<td>B. Braun</td>
			<td>Aesculap BD3361</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro needle holder</td>
			<td>Codman, Johnson &#38; Johnson Medical</td>
			<td>Codmann 80-2003</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro needle holder</td>
			<td>B. Braun</td>
			<td>Aesculap BD336R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro needle holder</td>
			<td>B. Braun</td>
			<td>Aesculap FD241R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro scissors</td>
			<td>B. Braun</td>
			<td>Aesculap FD101R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro scissors</td>
			<td>B. Braun</td>
			<td>Aesculap FM471R</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>B. Braun</td>
			<td>Aesculap BM221R</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin/Glutamine (100x)</td>
			<td>PAA</td>
			<td>P11-010</td>
			<td></td>
		</tr>
		<tr>
			<td>Peripheral venous catheter (18 G)</td>
			<td>B. Braun</td>
			<td>4268334B</td>
			<td></td>
		</tr>
		<tr>
			<td>Peripheral venous catheter (22 G)</td>
			<td>B. Braun</td>
			<td>4268091B</td>
			<td></td>
		</tr>
		<tr>
			<td>Probe pointed scissors</td>
			<td>B. Braun</td>
			<td>Aesculap BC030R</td>
			<td></td>
		</tr>
		<tr>
			<td>Retractors</td>
			<td>Forschungswerkst&#228;tten Hannover Medical School</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI culture medium</td>
			<td>Lonza</td>
			<td>BE12-702F</td>
			<td></td>
		</tr>
		<tr>
			<td>Saline solution (NaCl 0.9 %)</td>
			<td>Baxter</td>
			<td>No Catalog Number available</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>B. Braun</td>
			<td>Aesculap BC414</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td>Carl-Zeiss</td>
			<td>OPMI-MDM</td>
			<td></td>
		</tr>
		<tr>
			<td>Sutures (anastomoses)</td>
			<td>Catgut</td>
			<td>Mariderm 8-0 monofil</td>
			<td></td>
		</tr>
		<tr>
			<td>Sutures (ligature)</td>
			<td>Resorba</td>
			<td>Silk 5-0 polyfil</td>
			<td></td>
		</tr>
		<tr>
			<td>Sutures (skin, fascia)</td>
			<td>Ethicon</td>
			<td>Mersilene 3-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe (1 mL)</td>
			<td>B. Braun</td>
			<td>9166017V</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe (10 mL)</td>
			<td>B. Braun</td>
			<td>4606108V</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe (20 mL)</td>
			<td>B. Braun</td>
			<td>4606205V</td>
			<td></td>
		</tr>
		<tr>
			<td>Vascular clamp</td>
			<td>B. Braun</td>
			<td>Aesculap FB708R</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

In the past, different immunological issues have been addressed on the basis of the model, which was validated in the work group by more than 500 transplants with a survival rate of more than 95%13,18,19,20,21,22,23,24. Total operating times (including graft explantation and implantation) usually did not exceed 60 minutes, whereas combined cold and warm ischemia times were around 30 minutes. The strain combinations applied were mainly based on Lewis (Lew) background. Syngenic transplants survived up to 100 days without signs of graft failure, but significant weight and size reduction upon graft explantation. Most recently, we performed heterotopic heart transplantation in two different donor-recipient combinations simulating a fast and a prolonged rejection model: Lew.1a &#8594; Lew wt leading to fast rejection (mean survival time of 7.4 days) and Lew.1u-7B&#8594; Lew.1a leading to a more prolonged rejection (mean survival time of 42.5 days) (Figure 6). Macroscopically, the rejected grafts showed a thrombosis accompanied by a livid discoloration and swelling, whereas non-rejected grafts show distinct atrophy, most likely as a consequence of an unloaded left ventricle. Furthermore, we drew up cryostat sections of transplanted hearts in order to detect cell infiltration using an alkaline phosphatase-antialkaline phosphatase (APAAP) staining method. Single frames with a 50x magnification were merged to one composite image, giving an overview of the complete graft and the distribution of infiltrating cells. Histological analysis showed an increased infiltration of (e.g., CD4+, TCR+, or NKR-P1A/B+) immune cells in the allogenic grafts, whereas syngenic grafts were largely free of cell infiltration (Figure 7A-C).
Cervical lymphadenectomy and re-stimulation assays of draining lymph node cells after in ear injection of cardiac muscle cells in the above mentioned strain combinations revealed distinct strain-specific immune responses towards allogenic cardiac tissue and allowed further immunological analyses, such as cytokine profiling (Figure 8A-C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60956/60956fig1.jpg" /> 
Figure 1: Schematic presentation of the end-to-side anastomoses and the resulting blood flow through the heart. After anastomosing the donor ascending aorta end-to-side to the recipient abdominal aorta and analogously the pulmonary artery to the recipient inferior vena cava (A), blood flow enters the coronary arteries via the ascending aorta. The venous drainage occurs via the sinus coronarius into the right atrium and ventricle and through the pulmonary artery into the recipient inferior vena cava (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig1large.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/60956/60956fig2.jpg" /> 
Figure 2: Required surgical instruments and materials. (A) Explantation: 1: elastic limb band, 2: retractors, 3: 5-0 ligature, 4: probe pointed scissors, 5-6: micro needle holders, 7: scissors, 8: surgical forceps, 9: micro forceps, 10: eye lubricant, 11: compresses, 12: cotton swabs, 13: perfusion base, 14: saline solution on ice. (B) Implantation: 1: elastic limb band, 2: retractors, 5-6: micro needle holders, 7: scissors, 8: forceps, 9: micro forceps, 10: eye lubricant, 11: compresses, 12: cotton swabs, 15: micro scissors, 16: micro forceps, 17: Potts scissors, 18: needle holder, 19: arched cannula, 20: Cooley vascular clamp, 21: 8-0 monofilament sutures, 22: hemostatic gauze, 23: 3-0 polyfilament sutures, 24: Petri dish, 25: Carprofen (5 mg/mL), 26: local anesthetic (lidocaine 0.2%), 27: saline solution. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig2large.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/60956/60956fig3.jpg" /> 
Figure 3: Heart explantation. After heparinization (A), thoracotomy is performed and ascending aorta and pulmonary artery are severed as distal as possible (B). With one single ligature pulmonary and both caval veins are occluded (C) and the heart is removed from the thoracic cavity (D). (E) shows the heart before and (F) after perfusion with 30 mL saline solution containing heparin. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60956/60956fig4.jpg" /> 
Figure 4: Heart implantation. After exposition of abdominal vessels and placement of a Cooley vascular clamp (A) the vessels are cannulated (B) and a longitudinal incision is performed using Potts scissors (C-D). The donor ascending aorta is fixated with one knot each at the cranial and caudal corner of the incision of the recipient abdominal aorta (E) and anastomosis is performed by continuously running sutures (E-F). Note that the graft is placed on the right side of the vessels for the first half of the anastomosis (E) and onto the left side of the vessels for the second half of the anastomosis (F) and the subsequent venous anastomosis. The first half of the venous anastomosis is performed by an intraluminal suture (G). After finishing the second half of the venous anastomosis, the graft is ready for reperfusion (H). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60956/60956fig7.jpg" /> 
Figure 5: In-ear injection of allogenic cardiac muscle cells. After fixing the recipient&#39;s ear on a finger using double-sided tape (A) allogenic vital cardiac muscle cells are injected subcutaneously close to visual capillary vessels (B). After an observation period draining cervical lymph nodes (*) are extracted (C). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60956/60956fig5.jpg" /> 
Figure 6: Heart survival in different syngenic and allogenic donor-recipient combinations. Kaplan-Meier analysis shows the survival of syngenic (n=10 fast rejection model; n=5 prolonged rejection model) and allogenic grafts (n=11 fast rejection model; n=14 prolonged rejection model). In the prolonged rejection model six of 14 recipients reached the end of the observation period (60 days) without graft failure, leading to a prolonged graft survival in this group. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/60956/60956fig6.jpg" /> 
Figure 7: Histological analysis of syngenic and allogenic heart grafts. (A-B) show the infiltration of CD4+ cells using the APAAP staining method in a syngenic graft (A) and an allogenic graft upon rejection (B). (C) presents the increase of cell infiltration in allogenic grafts when compared to syngenic grafts (serving as reference group) for different immune cell subsets. The classification applied to quantify cell infiltration was modified from Hirschburger et al.25: 0 = infiltration comparable to syngenic grafts; 0.5 = slight increase of stained cells in isolated tissue sections; 1 = increase of singular stained cells over the whole tissue section; 1.5 = increase of stained cell clusters uniformly distributed over the whole tissue section; 2 = strong; 2.5 = very strong; 3 = strongest increase of stained cell clusters throughout the whole tissue section. The histological sections of the grafts were analyzed using a 50x magnification. Five grafts per group (syngenic and allogenic, respectively) were included in the analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/60956/60956fig8.jpg" /> 
Figure 8: Analysis of draining lymph node cells after in-ear injection of allogenic cardiac muscle cells. Specific re-stimulation (with 2 x 105 splenocytes of the respective donor strain) of 2 x 105 lymphocytes harvested from either draining cervical or mesenteric lymph nodes (LN) of fast rejecting Lew wt and prolonged rejecting Lew.1a recipients showed significantly reduced proliferation of draining lymph node cells in Lew.1a recipients (A), whereas proliferative capacity in general was still observable after unspecific stimulation with CD3/CD28 antibody (B). Surprisingly, cytokine profiling revealed an increase of inflammatory cytokines in the lymph nodes of prolonged rejecting recipients (C). Results are presented as mean &#177; SEM of at least 4 animals per group. Significance is indicated with * for p-values &#8804; 0.05 and **** for p-values &#8804; 0.0001. (This figure has been modified from Beetz et al.24). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60956/60956fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about An Immunological Model for Heterotopic Heart and Cardiac Muscle Cell Transplantation in Rats at JoVE.com

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

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 ...

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Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

7.5K Views.  Hannover Medical School. 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.

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

Murine Model of CD40-activation of B cells

Research on B cells has shown that CD40 activation improves their antigen presentation capacity. When stimulated with interleukin-4 and CD40 ligand (CD40L), human B cells can be expanded without difficulties from small amounts of peripheral blood within 14 days to very large amounts of highly-pure CD40-B cells (&gt;109 cells per patient) from healthy donors as well as cancer patients1-4. CD40-B cells express important lymph node homing molecules and can attract T cells in vitro5. Furthermore they efficiently take up, process and present antigens to T cells6,7. CD40-B cells were shown to not only prime na&iacute;ve, but also expand memory T cells8,9. Therefore CD40-activated B cells (CD40-B cells) have been studied as an alternative source of immuno-stimulatory antigen-presenting cells (APC) for cell-based immunotherapy1,5,10. In order to further study whether CD40-B cells induce effective T cell responses in vivo and to study the underlying mechanism we established a cell culture system for the generation of murine CD40-activated B cells. Using splenocytes or purified B cells from C57BL/6 mice for CD40-activation, optimal conditions were identified as follows: Starting from splenocytes of C57BL/6 mice (haplotype H-2b) lymphocytes are purified by density gradient centrifugation and co-cultured with HeLa cells expressing recombinant murine CD40 ligand (tmuCD40L HeLa)11. Cells are recultured every 3-4 days and key components such as CD40L, interleukin-4, -Mercaptoethanol and cyclosporin A are replenished. In this protocol we demonstrate how to obtain fully activated murine CD40-B cells (mCD40B) with similar APC-phenotype to human CD40-B cells (Fig 1a,b). CD40-stimulation leads to a rapid outgrowth and expansion of highly pure (&gt;90%) CD19+ B cells within 14 days of cell culture (Fig 1c,d). To avoid contamination with non-transfected cells, expression of the murine CD40 ligand on the transfectants has to be controlled regularly (Fig 2). Murine CD40-activated B cells can be used to study B-cell activation and differentiation as well as to investigate their potential to function as APC in vitro and in vivo. Moreover, they represent a promising tool for establishing therapeutic or preventive vaccination against tumors and will help to answer questions regarding safety and immunogenicity of this approach12.

In this video, we demonstrate the procedure of CD40-activation and expansion of murine B cells from splenocytes of C57BL/6 mice, which can be used as a model antigen-presenting cell (APC) to study induction of immunity.

Research on B cells has shown that CD40 activation improves their antigen presentation capacity. When stimulated with interleukin-4 and CD40 ligand ...

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 ...

An Ex vivo Model of an Oligodendrocyte-directed T-Cell Attack in Acute Brain Slices

Death of oligodendrocytes accompanied by destruction of neurons and axons are typical histopathological findings in cortical and subcortical grey matter lesions in inflammatory demyelinating disorders like multiple sclerosis (MS). In these disorders, mainly CD8+ T-cells of putative specificity for myelin- and oligodendrocyte-related antigens are found, so that neuronal apoptosis in grey matter lesions may be a collateral effect of these cells. Different types of animal models are established to study the underlying mechanisms of the mentioned pathophysiological processes. However, although they mimic some aspects of MS, it is impossible to dissect the exact mechanism and time course of &#8216;&#8216;collateral&#8217;&#8217; neuronal cell death. To address this course, here we show a protocol to study the mechanisms and time response of neuronal damage following an oligodendrocyte-directed CD8+ T cell attack. To target only the myelin sheath and the oligodendrocytes, in vitro activated oligodendrocyte-specific CD8+ T-cells are transferred into acutely isolated brain slices. After a defined incubation period, myelin and neuronal damage can be analysed in different regions of interest. Potential applications and limitations of this model will be discussed.

An Ex vivo Model of an Oligodendrocyte-directed T-Cell Attack in Acute Brain Slices

To address mechanisms of demyelination and neuronal apoptosis in cortical lesions of inflammatory demyelinating disorders, different animal models are used. We here describe an ex vivo approach by using oligodendrocyte-specific CD8+ T-cells on brain slices, resulting in oligodendroglial and neuronal death. Potential applications and limitations of the model are discussed.

Death of oligodendrocytes accompanied by destruction of neurons and axons are typical histopathological findings in cortical and subcortical grey matter ...

Normal and Malignant Muscle Cell Transplantation into Immune Compromised Adult Zebrafish

Zebrafish have become a powerful tool for assessing development, regeneration, and cancer. More recently, allograft cell transplantation protocols have been developed that permit engraftment of normal and malignant cells into irradiated, syngeneic, and immune compromised adult zebrafish. These models when coupled with optimized cell transplantation protocols allow for the rapid assessment of stem cell function, regeneration following injury, and cancer. Here, we present a method for cell transplantation of zebrafish adult skeletal muscle and embryonal rhabdomyosarcoma (ERMS), a pediatric sarcoma that shares features with embryonic muscle, into immune compromised adult rag2E450fs homozygous mutant zebrafish. Importantly, these animals lack T cells and have reduced B cell function, facilitating engraftment of a wide range of tissues from unrelated donor animals. Our optimized protocols show that fluorescently labeled muscle cell preparations from &#945;-actin-RFP transgenic zebrafish engraft robustly when implanted into the dorsal musculature of rag2 homozygous mutant fish. We also demonstrate engraftment of fluorescent-transgenic ERMS where fluorescence is confined to cells based on differentiation status. Specifically, ERMS were created in AB-strain myf5-GFP; mylpfa-mCherry double transgenic animals and tumors injected into the peritoneum of adult immune compromised fish. The utility of these protocols extends to engraftment of a wide range of normal and malignant donor cells that can be implanted into dorsal musculature or peritoneum of adult zebrafish.

Here, we present a protocol for cell transplantation of zebrafish skeletal muscle and embryonal rhabdomyosarcoma (ERMS) into adult immune compromised rag2E450fs homozygous mutant zebrafish. This protocol allows for the efficient analysis of regeneration and malignant transformation of muscle cells.

Zebrafish have become a powerful tool for assessing development, regeneration, and cancer. More recently, allograft cell transplantation protocols have ...

An Endothelial Planar Cell Model for Imaging Immunological Synapse Dynamics

Adaptive immunity is regulated by dynamic interactions between T cells and antigen presenting cells (&#39;APCs&#39;) referred to as &#39;immunological synapses&#39;. Within these intimate cell-cell interfaces discrete sub-cellular clusters of MHC/Ag-TCR, F-actin, adhesion and signaling molecules form and remodel rapidly. These dynamics are thought to be critical determinants of both the efficiency and quality of the immune responses that develop and therefore of protective versus pathologic immunity. Current understanding of immunological synapses with physiologic APCs is limited by the inadequacy of the obtainable imaging resolution. Though artificial substrate models (e.g., planar lipid bilayers) offer excellent resolution and have been extremely valuable tools, they are inherently non-physiologic and oversimplified. Vascular and lymphatic endothelial cells have emerged as an important peripheral tissue (or stromal) compartment of &#39;semi-professional APCs&#39;. These APCs (which express most of the molecular machinery of professional APCs) have the unique feature of forming virtually planar cell surface and are readily transfectable (e.g., with fluorescent protein reporters). Herein a basic approach to implement endothelial cells as a novel and physiologic &#39;planar cellular APC model&#39; for improved imaging and interrogation of fundamental antigenic signaling processes will be described.

Adaptive immunity is controlled by dynamic 'immunological synapses' formed between T cells and antigen presenting cells. This protocol describes methods for investigating endothelial cells both as understudied physiologic APCs and as a novel type of 'planar cellular APC model'.

Adaptive immunity is regulated by dynamic interactions between T cells and antigen presenting cells (&#39;APCs&#39;) referred to as &#39;immunological ...

Establishment of a High-throughput Setup for Screening Small Molecules That Modulate c-di-GMP Signaling in Pseudomonas aeruginosa

Bacterial resistance to traditional antibiotics has driven research attempts to identify new drug targets in recently discovered regulatory pathways. Regulatory systems that utilize intracellular cyclic di-GMP (c-di-GMP) as a second messenger are one such class of target. c-di-GMP is a signaling molecule found in almost all bacteria that acts to regulate an extensive range of processes including antibiotic resistance, biofilm formation and virulence. The understanding of how c-di-GMP signaling controls aspects of antibiotic resistant biofilm development has suggested approaches whereby alteration of the cellular concentrations of the nucleotide or disruption of these signaling pathways may lead to reduced biofilm formation or increased susceptibility of the biofilms to antibiotics. We describe a simple high-throughput bioreporter protocol, based on green fluorescent protein (GFP), whose expression is under the control of the c-di-GMP responsive promoter cdrA, to rapidly screen for small molecules with the potential to modulate c-di-GMP cellular levels in Pseudomonas aeruginosa (P. aeruginosa). This simple protocol can screen upwards of 3,500 compounds within 48 hours and has the ability to be adapted to multiple microorganisms.

Establishment of a High-throughput Setup for Screening Small Molecules That Modulate c-di-GMP Signaling in Pseudomonas aeruginosa

This article describes the high-throughput assay that has been successfully established to screen large libraries of small molecules for their potential ability to manipulate cellular levels of cyclic di-GMP in Pseudomonas aeruginosa, providing a new powerful tool for antibacterial drug discovery and compound testing.

Bacterial resistance to traditional antibiotics has driven research attempts to identify new drug targets in recently discovered regulatory pathways. ...

Flow Cytometry-based Assay for the Monitoring of NK Cell Functions

Natural killer (NK) cells are an important part of the human tumor immune surveillance system. NK cells are able to distinguish between healthy and virus-infected or malignantly transformed cells due to a set of germline encoded inhibitory and activating receptors. Upon virus or tumor cell recognition a variety of different NK cell functions are initiated including cytotoxicity against the target cell as well as cytokine and chemokine production leading to the activation of other immune cells. It has been demonstrated that accurate NK cell functions are crucial for the treatment outcome of different virus infections and malignant diseases. Here a simple and reliable method is described to analyze different NK cell functions using a flow cytometry-based assay. NK cell functions can be evaluated not only for the whole NK cell population, but also for different NK cell subsets. This technique enables scientists to easily study NK cell functions in healthy donors or patients in order to reveal their impact on different malignancies and to further discover new therapeutic strategies.

A simple and reliable method is described here to analyze a set of NK cell functions such as degranulation, cytokine and chemokine production within different NK cell subsets.

Natural killer (NK) cells are an important part of the human tumor immune surveillance system. NK cells are able to distinguish between healthy and ...

Isolation and Identification of Extravascular Immune Cells of the Heart

The immune system is an essential component of a healthy heart. The myocardium is home to a rich population of different immune cell subsets with functional compartmentalization both during steady state and during different forms of inflammation. Until recently, the study of immune cells in the heart required the use of microscopy or poorly developed digestion protocols, which provided enough sensitivity during severe inflammation but were unable to confidently identify small &#8212; but key &#8212; populations of cells during steady state. Here, we discuss a simple method combining enzymatic (collagenase, hyaluronidase and DNAse) and mechanical digestion of murine hearts preceded by intravascular administration of fluorescently-labelled antibodies to differentiate small but unavoidable intravascular cell contaminants. This method generates a suspension of isolated viable cells that can be analyzed by flow cytometry for identification, phenotyping and quantification, or further purified with fluorescence-activated cell sorting or magnetic bead separation for transcriptional analysis or in vitro studies. We include an example of a step-by-step flow cytometric analysis to differentiate the key macrophage and dendritic cell populations of the heart. For a medium sized experiment (10 hearts) the completion of the procedure requires 2&#8211;3 h.

This protocol presents a simple and efficient method to isolate, identify and quantify immune cells residing in the myocardium of mice during steady state or inflammation. The protocol combines enzymatic and mechanical digestion for the generation of a single cell suspension that can be further analyzed by flow cytometry.

The immune system is an essential component of a healthy heart. The myocardium is home to a rich population of different immune cell subsets with ...

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 ...

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

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Murine Model of CD40-activation of B cells

Orthotopic Hind-Limb Transplantation in Rats

An Ex vivo Model of an Oligodendrocyte-directed T-Cell Attack in Acute Brain Slices

Normal and Malignant Muscle Cell Transplantation into Immune Compromised Adult Zebrafish

An Endothelial Planar Cell Model for Imaging Immunological Synapse Dynamics

Establishment of a High-throughput Setup for Screening Small Molecules That Modulate c-di-GMP Signaling in Pseudomonas aeruginosa

Flow Cytometry-based Assay for the Monitoring of NK Cell Functions

Isolation and Identification of Extravascular Immune Cells of the Heart

Optimization of the Cuff Technique for Murine Heart Transplantation

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