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 train...

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><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 &amp;micro;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>&quot;Bepanthen&quot;</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>&quot;T/Pump&quot;</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&amp;auml;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 &amp;amp; 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&amp;auml;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. Tota...

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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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

Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant vasculopathy (TVP).
The commonly used animal model for cardiovascular chronic rejection studies is the heterotopic heart transplant model performed in laboratory rodents. This model is used widely in experiments since Ono and Lindsey (3) published their technique. To analyze the findings in the blood vessels, the heart has to be sectioned and all vessels have to be measured.
Another method to investigate chronic rejection in cardiovascular questionings is the aortic transplant model (1, 2). In the orthotopic aortic transplant model, the aorta can easily be histologically evaluated (2). The PVG-to-ACI model is especially useful for CAV studies, since acute vascular rejection is not a major confounding factor and Cyclosporin A (CsA) treatment does not prevent the development of CAV, similar to what we find in the clinical setting (4). A7-day period of CsA is required in this model to prevent acute rejection and to achieve long-term survival with the development of TVP.
This model can also be used to investigate acute cellular rejection and media necrosis in xenogeneic models (5).

This video demonstrates the orthotopic aortic transplant model as a simple model to study the development of transplant vasculopathy (TVP) in rats.

Research models of chronic rejection are essential to investigate pathobiological and pathophysiological processes during the development of transplant ...

Medicine

Heterotopic Heart Transplantation in Mice

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable for studying rejection and immune response now that newer transgenic and gene knockout mice are available, and a large number of immunologic reagents have been developed. The heart transplant model is less stringent than the skin transplant models, although technically more challenging. We have developed a modified technique and have completed over 1000 successful cases of heterotopic heart transplantation in mice. When making anastomosis of the ascending aorta and abdominal aorta, two stay sutures are placed at the proximal and distal apexes of recipient abdominal aorta with the donor s ascending aorta, then using 11-0 suture for anastomosis on both side of aorta with continuing sutures. The stay sutures make the anastomosis easier and 11-0 is an ideal suture size to avoid bleeding and thrombosis.
When making anastomosis of pulmonary artery and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s pulmonary artery. The left wall of the inferior vena cava and donor s pulmonary artery is closed with continuing sutures in the inside of the inferior vena cava after, one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s pulmonary artery are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis. In this article, we provide details of the technique to supplement the video.

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response. However, the techniques involved are still challenging. By modifying standard techniques we have had success with more than 1000 transplants.

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable ...

Biology

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are available to perform mechanistic in vivo studies. While heart transplantation models using a suture technique were first successfully developed in rats, the translation into an equally widespread used murine equivalent was never achieved due the technical complexity of the microsurgical procedure. In contrast, non-suture cuff techniques, also developed initially in rats, were successfully adapted for use in mice1-3. This technique for revascularization involves two major steps I) everting the recipient vessel over a polyethylene cuff; II) pulling the donor vessel over the formerly everted recipient vessel and holding it in place with a circumferential tie. This ensures a continuity of the endothelial layer, short operating time and very high patency rates4.
Using this technique for vascular anastomosis we performed more than 1,000 cervical heart transplants with an overall success rate of 95%. For arterial inflow the common carotid artery and the proximal aortic arch were anastomosed resulting in a retrograde perfusion of the transplanted heart. For venous drainage the pulmonary artery of the graft was anastomosed with the external jugular vein of the recipient5.
Herein, we provide additional details of this technique to supplement the video.

The murine cervical heart transplantation model is well suited for immunological as well as ischemia reperfusion injury studies. We modified the procedure using a non-suture cuff technique and performed more than 1,000 successful transplants with this approach.
Herein, we provide additional details of this technique to supplement the video.

Mouse models are of special interest in research since a wide variety of monoclonal antibodies and commercially defined inbred and knockout strains are ...

Murine Heterotopic Heart Transplant Technique

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become proficient in and utilize this technique, it is now widely used by many laboratories around the world. A significant refinement to the original technique was developed and reported in 2001 by Niimi. Described here are the techniques that have evolved over more than a decade in the hands of three surgeons (Plenter, Grazia, Pietra) in our center. These techniques are now being passed on to a younger generation of surgeons and researchers.
Based largely on the Niimi experience, the procedures used have evolved in the fine details - details which we will endeavor to relate here in such a way that others may be able to use this very useful model. Like Niimi, we have found that a video aid to learning is a priceless resource for the beginner.

The manuscript describes the steps required to perform the heterotopic heart transplant in the mouse.

It is now over forty years since this technique was first reported by Corry, Wynn and Russell. Although it took some years for other labs to become ...

Rat Heterotopic Abdominal Heart/Single-lung Transplantation in a Volume-loaded Configuration

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft involves anastomosis of donor inferior vena cava (IVC) to recipient IVC, and donor ascending aorta (Ao) to recipient abdominal Ao. The right upper and middle lung lobes are preserved and function as conduits for blood flow from right heart to left heart.
There are several advantages to using this technique, and it lends itself to a broad range of applications. Because the graft is transplanted in a configuration that allows for dyamic volume-loading, cardiac function may be directly assessed in vivo. The use of pressure-volume conductance catheters permits characterization of load-dependent and load-independent hemodynamic parameters. The graft may be converted to a loaded configuration by applying a clamp to the recipient&#8217;s infra-hepatic IVC. We describe modified surgical techniques for both donor and recipient operations, and an ideal myocardial protection strategy. Depending on the experimental aim, this model may be adapted for use in both acute and chronic studies of graft function, immunologic status, and variable ventricular loading conditions. The conducting airways to the transplanted lung are preserved, and allow for acute lung re-ventilation. This facilitates analysis of the effects of the mixed venous and arterial blood providing coronary perfusion to the graft.
A limitation of this model is its technical complexity. There is a significant learning curve for new operators, who should ideally be mentored in the technique. A surgical training background is advantageous for those wishing to apply this model. Despite its complexity, we aim to present the model in a clear and easily applicable format. Because of the physiologic similarity of this model to orthotopic transplantation, and its broad range of study applications, the effort invested in learning the technique is likely to be worthwhile.

We describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The transplant configuration results in a partially loaded graft circulation, allowing direct functional assessment. This model may be employed for acute or chronic studies of function and immunologic status of the transplanted graft.

Herein, we describe a novel technique for heterotopic abdominal heart-lung transplantation (HAHLT) in rats. The configuration of the transplant graft ...

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 ...

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression must be pursued. Employing vascularized bone marrow transplantation and co-transplantation of the thymus have shown promise in this regard in various animal models.1-11 Vascularized bone marrow transplantation allows for the uninterrupted transfer of donor bone marrow cells within the preserved donor microenvironment, and the incorporation of thymus tissue with vascularized bone marrow transplantation has shown to increase T-cell chimerism ultimately playing a supportive role in the induction of immune regulation. The combination of solid organ and vascularized composite allotransplantation can uniquely combine these strategies in the form of a novel transplant model. Murine models serve as an excellent paradigm to explore the mechanisms of acute and chronic rejection, chimerism, and tolerance induction, thus providing the foundation to propagate superior allograft survival strategies for larger animal models and future clinical application. Herein, we developed a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique. The experience in syngeneic and allogeneic transplant settings is described for future broader immunological investigations via an instructional manuscript and video supplement.

To study combined solid organ and vascularized composite allotransplantation, we describe a novel heterotopic en bloc chest wall, thymus, and heart transplant model in mice using a cervical non-suture cuff technique.

Exploration of novel strategies in organ transplantation to prolong allograft survival and minimizing the need for long-term maintenance immunosuppression ...

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

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

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

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

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of transplant with the potential to deliver durable heart valves capable of somatic growth with no requirement for anticoagulation. However, the immunobiology of heart valve transplants remains unexplored, highlighting the need for animal models to study this new type of transplant. Previous rat models for heterotopic aortic valve transplantation into the abdominal aorta have been described, though they are technically challenging and costly. For addressing this challenge, a renal subcapsular transplant model was developed in rodents as a practical and more straightforward method for studying heart valve transplant immunobiology. In this model, a single aortic valve leaflet is harvested and inserted into the renal subcapsular space. The kidney is easily accessible, and the transplanted tissue is securely contained in a subcapsular space that is well vascularized and can accommodate a variety of tissue sizes. Furthermore, because a single rat can provide three donor aortic leaflets and a single kidney can provide multiple sites for transplanted tissue, fewer rats are required for a given study. Here, the transplantation technique is described, providing a significant step forward in studying the transplant immunology of heart valve transplantation.

This protocol describes a simple and efficient method for the transplantation of aortic valve leaflets under the renal capsule to allow for the study of alloreactivity of heart valves.

There is an urgent clinical need for heart valve replacements that can grow in children. Heart valve transplantation is proposed as a new type of ...

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for investigating the mechanism of cardiac allograft rejection in preclinical studies because of their high homology with human genes. This understanding would help develop unique approaches to improving patients&#39; long-term survival treated with cardiac allografts. In a mouse model, abdominal donor heart implantation is commonly performed with an end-to-end&#160;anastomosis to the recipient&#39;s aorta and inferior vena cava using stitches. In this model, the donor&#39;s heart is implanted by end-to-end anastomosis to the recipient&#39;s carotid artery and jugular vein by the modified-Cuff technique. The transplantation surgery is performed without stitching and thus may increase the survival of the recipient since there is no interference with the blood supply and venous reflux of the lower body. This mouse model would help investigate the mechanisms underlying the immunological and pathological (acute/chronic) rejection of cardiac allografts.

In the present protocol, a mouse heart transplantation model is used for investigating the mechanism of cardiac allograft rejection. In this heterotopic heart transplantation model, operation efficiency is improved, and the survival of cardiac grafts is ensured by a cervical end-to-end anastomosis of heart implantation using a modified Cuff technique.

Cardiac allograft rejection limits the long-term survival of patients after heart transplantation. A mouse heart transplantation model is ideal for ...

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

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Orthotopic Aortic Transplantation: A Rat Model to Study the Development of Chronic Vasculopathy

Heterotopic Heart Transplantation in Mice

Murine Cervical Heart Transplantation Model Using a Modified Cuff Technique

Murine Heterotopic Heart Transplant Technique

Rat Heterotopic Abdominal Heart/Single-lung Transplantation in a Volume-loaded Configuration

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

A Novel Microsurgical Model for Heterotopic, En Bloc Chest Wall, Thymus, and Heart Transplantation in Mice

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

A Simplified Model for Heterotopic Heart Valve Transplantation in Rodents

A Modified Cuff Technique for Mouse Cervical Heterotopic Heart Transplantation Model