This work was supported by grants from the Canadian Institutes of Health Research and Heart and Stroke Foundation of BC &#38; Yukon (JCC).

All the protocols in this study were reviewed and approved by the Simon Fraser University animal care ethics committee. Use Balb/cYJ (H2d) donor mice and C57Bl/6 (H2b) recipient mice to examine allogeneic reactions. Mice are used for experiments between the ages of 8 to 12 weeks. Use either female or male mice. Syngeneic controls consist of aortic segments from C57Bl/6 donors into C57Bl/6 recipients.
1. Donor and Recipient Preparation
Note: Both the donor and recipient are anesthetized and prepared before the surgery to minimize ischemia of the graft. Injectable anesthetics are used in the protocol to prevent obstruction of the animal by equipment needed for the delivery of inhaled anesthetics. However, if desired, inhaled anesthetic is an appropriate alternative. From the initial injection of the anesthetics, the entire procedure takes approximately 90 min to complete. Ischemic time of the graft is less than 30 min.
<ol>
	<li>Anesthetize the mouse with intraperitoneal injections of ketamine (100 mg/kg; 10 mg/ml) and xylazine (10 mg/kg; 1 mg/ml). The mouse will be sedated within 10 to 15 min. Assess the depth of anesthesia by pinching the fatty part of the animal foot pad. Administer 1/3 of the original dose of the anesthetic cocktail, as needed, until the animal does not exhibit withdrawal reflex. Monitor the respiratory rate closely after every cocktail is administered. During surgery, assess the level of anesthesia every 15 min by pinching the anterior abdominal wall with a pair of forceps. The mice should remain deeply anaesthetized for 60 to 90 min.</li>
	<li>Lubricate the mouse&#8217;s eyes with an ophthalmic ointment to prevent dryness while under anesthesia.</li>
	<li>Shave hair as close to the skin as possible on the abdominal ventral region from the mid-thorax to the pubis. Be particularly careful not to nick any skin. Do not use depilatory cream because it may be absorbed into the skin and could be inflammatory.</li>
	<li>Place the animal in supine position on a towel above the heating table or pad.</li>
	<li>Clean the surgical site with a preliminary scrub with 2% chlorohexidine. Prepare a large working area to maximize the surgical field. Start scrubbing at the center of the surgical site and move to the outside in a linear or circular manner. Dispose of the gauze. Repeat this procedure at least 5 more times. Apply an alcohol prep pad to the same area. Once the alcohol is dry, prepare the clean area with Betadine solution and drape with sterile gauze.</li>
</ol>
2. End-to-end Anastomosis Procedure
<ol>
	<li>Donor Operation
	<ol>
		<li>Maintain aseptic technique throughout the operation. Clean all countertop and surgical table surfaces with 0.5% accelerated hydrogen peroxide solution prior to use. Wrap and autoclave all surgical instruments, gauzes, drapes and gowns prior to use. Verify sterility of the instruments with a steam sterilizer indicator strip placed in each pack. Sterile surgical gloves are used and disposed of between surgeries. For multiple surgeries, sterilize the surgical instruments between uses with the hot glass bead sterilizer.</li>
		<li>Place the donor mouse in a supine position on a clean, thin Plexiglas board wrapped with sterile drape under the operating microscope at 8-30X magnification.</li>
		<li>After ensuring adequate surgical anesthesia as outlined in section 1.1, proceed with surgery.</li>
		<li>Using sterile scissors, make a single midline lower longitudinal abdominal incision, from the pubis to the xyphoid process.</li>
		<li>Using a small retractor, open the abdominal walls to expose the cavity.</li>
		<li>Using sterile cotton tipped applicators, gently retract the intestines superiorly to the animal&#8217;s left and cover with gauze moistened with saline solution. Move the reproductive organs inferiorly and locate the infrarenal aorta and the inferior vena cava (IVC). Moisten the exposed tissues periodically with saline solution.</li>
		<li>Using the medical No. 5 forceps, separate the aorta from the IVC, from the level of the left renal artery to the bifurcation. Use 10-0 polyamide monofilament sutures to ligate the small branches near the aorta.</li>
		<li>Once the donor&#8217;s aorta has been separated from the IVC, saturate the vessel with saline, cover the exposed cavity with moistened gauze and set the donor aside onto a sterile area. Check the status of the donor (respiratory and cardiovascular function, and depth of anesthesia) every 15 min. Begin operating on the recipient, isolating the aorta as described below (Steps 2.2.1 to 2.2.7).</li>
		<li>Once the recipient aorta has been separated from the IVC and set aside, return the donor under the microscope. Cross clamp (proximal and distal to the segment of interest) the donor aorta, approximately 5 mm apart, with two 4 mm microvascular clamps.</li>
		<li>Using the Vannas-Tubingen microscissors, transect a small graft segment (3 to 4 mm in length) of the abdominal aorta.</li>
		<li>Using a 25 G 5/8 needle attached to a syringe, flush the excised aorta with heparinized (100 U/ml) saline solution. Ensure the tip of the needle does not come into contact with the vessel.</li>
		<li>Place aorta in heparinized (100 U/ml) saline solution on ice and set aside. While still under deep anesthesia, release the microvascular clamps. Euthanize the donor by exsanguination.</li>
		<li>Implant the donor vessel within 30 min of excision. Although it is possible to use one donor vessel for multiple recipients by excising a larger length of aorta, keep the ischemic time to less than 30 min.</li>
	</ol>
	</li>
	<li>Recipient Operation
	<ol>
		<li>Maintain aseptic technique throughout the operation, as in donor operation.</li>
		<li>Place the recipient mouse in a supine position on a thin Plexiglas board wrapped with sterile drape under the operating microscope at 8-30X magnification.</li>
		<li>After ensuring adequate anesthesia, proceed with surgery when the animal does not exhibit withdrawal reflex.</li>
		<li>Using sterile scissors, make a single midline lower longitudinal abdominal incision, from the pubis to the xyphoid process.</li>
		<li>Using a small retractor, open the abdominal walls to expose the cavity.</li>
		<li>Using sterile cotton tipped applicators, gently retract the intestines superiorly to the animal&#8217;s left and cover with gauze moistened with saline solution. Move the reproductive organs inferiorly and locate the infrarenal aorta and the IVC. Moisten the exposed tissues periodically with saline solution.</li>
		<li>Separate the aorta from the IVC, from the level of the left renal artery to the bifurcation. If necessary, use 10-0 polyamide monofilament sutures to ligate the small branches near the aorta.</li>
		<li>Cross clamp (proximal and distal to the segment of interest) the aorta, approximately 5 mm apart, with two 4 mm microvascular clamps.</li>
		<li>Using the Vannas-Tubingen microscissors, make a single horizontal aortotomy and resect a small segment (no more than 0.5 mm) of the abdominal aorta to accommodate the donor aortic graft.</li>
		<li>Flush the excised aorta with heparinized (100 U/ml) saline solution. The donor aortic graft should be of appropriate length to connect the recipient&#8217;s transected aortic ends.</li>
		<li>Place the donor aortic graft in the orthotopic position and anastomose the donor&#8217;s graft end to the recipient&#8217;s end, matching the respective graft anatomical orientation with that of the recipient.</li>
		<li>Gently grasp the tunica externa of the vessel and evert it slightly using the medical No.5 forceps. Using the forceps, drive the needle attached to the 10-0 polyamide monofilament suture through the full thickness of the vessel wall in order to secure the donor aortic graft to the recipient&#8217;s resected vessel. Take care to ensure that the vessel opening is not closed off due to inadvertent stitching of the back wall of the vessel.</li>
		<li>For continuous stitches, place stay sutures at 9 o&#8217;clock in both the upper and lower ends of the graft. Starting at the upper end of the graft from 3 o&#8217;clock, anastomose the resected ends with 2 running sutures and secure the suture to the stay suture.
		<ol>
			<li>Flip the graft over and continue the running suture to dorsal part of the vessel, meeting the origin stay suture. Secure without applying much pressure on the vessel. Repeat the suturing for the lower anastomosis. 
			Note: The interrupted sutures start the same way as the continuous suture with the exception that the vessel is anastomosed with three separated stitches between the stay sutures. Anastomosis time is usually 20 min.</li>
		</ol>
		</li>
		<li>Once the anastomosis is complete, release the distal microvascular clamp to allow retrograde blood to flow and check for leakage of the anastomosed sites. If there is a leakage, immediately place a stitch to close the defective site. If there is no bleeding at the sites, then release the proximal clamp.</li>
		<li>Examine the transplant and check that there is no blood obstruction in the graft, and the proximal and distal portion of the recipient&#8217;s vessel. Vigorous pulse pattern in both the donor&#8217;s and recipient&#8217;s vessel is a primary indication that the blood is flowing freely. Using the pair of forceps, gently grasp one end of the stay sutures and slightly evert the vessel to inspect the back wall of the vessel. 
		Note: There should be no puckering of the vessel wall at both ends of the anastomosed sites. Poor blood flow after removal of the clamps is a sign of thrombosis.</li>
		<li>Using cotton tipped applicators, return the intestines into the abdominal cavity.</li>
		<li>With the Castroviejo needle holder and Graefe forceps, close the abdominal wall with 5-0 polypropylene sutures using continuous stitching. Close the skin layer with the same sutures using subcuticular closure.</li>
		<li>Administer Torbugesic (1 mg/kg) i.m. immediately upon completion of the transplant.</li>
		<li>Give immediately, in the order, Atipamezole (1 mg/kg), ketoprofen (5 mg/kg) and warmed Lactated Ringer&#8217;s solution subcutaneously.</li>
		<li>Immediately after surgery, place mice in a heated cage under a water blanket overnight (12 hr). During the anesthetic-recovery period, place the animals alone in a clean, dry unobstructed area. Line the cage (autoclaved) with clean paper towels and adjust the temperature of the water blanket to approximately 20 to 22 &#176;C. Provide dry and wet kibbles ad libitum on the cage floor. 
		Note: An important component of post&#8211;surgical care is the observation of the animal and appropriate intervention, as required, during recovery from anesthesia and surgery. The necessary intensity of monitoring will vary with the animal and might be greater during the immediate anesthetic recovery period as compared to later in postoperative recovery.</li>
		<li>Continually monitor animals having undergone anesthesia monitored until they recover completely. The animal must be able to maintain unassisted sternal recumbency and it must appear calm and free of pain before it can be left unattended.</li>
		<li>Give Buprenorphine (0.1 mg/kg BID) and ketoprofen (5 mg/kg SID) for a period of three days, both subcutaneously.</li>
		<li>Monitor cardiovascular and respiratory function, body temperature, and postoperative pain or discomfort during recovery from anesthesia for a minimum of three days. Additional care may be required, such as administration of analgesics and other drugs, and parenteral fluids to minimize dehydration and electrolyte loss.</li>
		<li>Assess the success of the transplant surgery by observing the motor function of the hind limbs. Complete success of the transplant involves unobstructed blood flow to the hind limb and tail, which should be immediate upon recovery of the animal, and complete recovery with no paralysis of the hind limbs on the second day</li>
	</ol>
	</li>
</ol>
3. Tissue Collection
Note: Pre-determined end points range from 3-60 days depending on the type of analysis desired and the nature of the antigen mismatch. Generally, intimal thickening is robust at day 30 after transplantation across complete MHC mismatched mouse strains.
<ol>
	<li>Anesthetize the mouse with intraperitoneal injections of ketamine (100 mg/kg; 10 mg/ml) and xylazine (10 mg/kg; 1 mg/ml). Assess the depth of anesthesia by pinching the fatty part of the animal foot pad. Proceed with surgery when the animal does not exhibit withdrawal reflex.</li>
	<li>Clean the surgical site with water and ending with 70% alcohol.</li>
	<li>Place the animal in supine position on a tray lined with absorbent pad.</li>
	<li>Open the abdominal cavity with a large midline longitudinal incision. Place a self-retaining retractor to expose the cavity, the IVC and the abdominal aorta.</li>
	<li>Gently separate the transplanted donor aortic segment from the neighboring IVC using a pair of medical No. 5 forceps. Be particularly careful not to strip the adventitia of the transplanted vessel.</li>
	<li>Expose the thoracic cavity by cutting through the ribs along both sides of the thoracic spine all the way to the thoracic inlet. Reflect the anterior chest wall superiorly to expose the pericardium and secure the ribcage with a pair of hemostat.</li>
	<li>Once the heart is exposed, insert a 25 G 5/8 needle (attached to a syringe) into the left ventricle and flush with 0.1 ml of heparinized (100 U/ml) saline solution.</li>
	<li>Using a pair of scissors, remove the right atrium to allow the blood, heparinized saline, and fixative to leave the body during perfusion. Perfuse the animal with 5 ml of 4% paraformaldehyde or until the fluid runs clear. Remove the heart to ensure euthanasia.</li>
	<li>Excise transplanted vessel segment and immerse in 4% paraformaldehyde for no more than one hour. Set the transplanted vessel in OCT (optimal cutting temperature) matrix and flash freeze. Section the vessel under a cryostat set at 8 um thickness. Stain sections with Haematoxylin &#38; Eosin and/or Verhoeff-van Gieson (van Gieson) elastic stain.</li>
</ol>
4. Morphological Analysis of Grafts
Note: TA is characterized by intimal thickening 13. In this model, intimal thickening and a resultant reduction in the size of the lumen are reflective of the severity of immune-mediated vascular injury. Syngraft controls are used to determine the baseline characteristics of vessels in the absence of allogeneic immune responses. These controls also permit evaluation of arterial damage that occurs as a result of the surgical procedure. There should be no intimal thickening in syngrafts.
<ol>
	<li>To examine intimal thickening and luminal occlusion, measure the area within the endothelial cell layer, internal elastic lamina (IEL) and external elastic lamina (EEL) on elastic van Gieson stained artery segments with an image analysis software. The parameters below can be used to determine arterial changes reflective of TA.
	<ol>
		<li>Measure the absolute intimal area: area within the internal elastic lamina - the area within the endothelial cell layer. This provides an absolute measurement of intimal expansion, which is the hallmark of TA.</li>
		<li>Measure the absolute medial area: area within the external elastic lamina - area within the internal elastic lamina. This provides an absolute measurement of medial degradation or growth, which can occasionally occur as a result of immune-mediated changes to this region of the vessel wall.</li>
		<li>Measure the total vessel area as the area within the external elastic lamina. This provides a measurement of vessel remodelling which involves expansion or constriction of artery as a result of immunological damage.</li>
		<li>Measure the intima/Media: (area within the internal elastic lamina &#8211; area within the endothelial cell layer) / (area within the external elastic lamina &#8211; area within the internal elastic lamina). This provides a relative measure of intimal expansion and normalizes for differences in vessel size.</li>
		<li>Measure the % Luminal Narrowing: [ (area within the internal elastic lamina &#8211; area within the endothelial cell layer) / area with the internal elastic lamina) ]*100. This provides a measure of the extent of luminal occlusion, which results from a combination of intimal expansion and remodeling (inward or outward) of the vessel wall.</li>
	</ol>
	</li>
</ol>

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We have described a protocol for aortic interposition grafting in mice that is useful for studying immune-mediated vascular rejection and TA. This model can be used to investigate the causes of TA as well as the development of novel therapeutic strategies. It has been used in the past to establish an essential role of adaptive immunity, cytotoxic T cell responses, cytokine-mediated CD4 T cell effector responses, and antibody-mediated graft damage in TA 14,17-21. Artery transplantation in mice is difficult due ...

Over the past 30+ years, advances in immunosuppressive drugs have diminished graft rejection due to acute rejection but chronic rejection remains a main challenge. The main manifestation of chronic heart transplant rejection is transplant arteriosclerosis (TA) 1,2. This condition is characterized by intimal hyperplasia and vasomotor dysfunction of allograft arteries and develops as a result of immunological targeting of endothelial and smooth muscle cells by the recipient immune system. The specific targeting of the graft vasculature due to recognition of foreign peptide-major histocompatibility complex (MHC) is highlighted by the development of TA exclusively in graft arteries while sparing host vessels 3. In keeping with this is the observation that TA does not occur experimentally when the recipient is genetically identical to the donor or when the recipient lacks T and B cells 4. Immune-mediated vascular injury and dysfunction causes the development of intimal thickening and fibrosis, as well as the aberrant accumulation of lipids and ECM proteins, in TA 5. Intimal thickening tends to be concentric throughout the entire arterial tree 4-6. Graft loss and death usually occur as a result of progressive ischemia resulting from luminal occlusion of allograft arteries 4.
In 1991, Mennander et al. 7 pioneered an aortic interposition model in rats to model TA. Several groups have subsequently adapted this procedure for use in mice. In this model, allograft aortic segments develop lesions that have features comparable to TA observed in clinical transplants. This includes intimal thickening characterized by the accumulation of smooth muscle-like cells and recipient leukocytes 7. Over the past 2 decades this model has been used to generate important insight into the mechanisms of vascular injury, rejection and TA. It can be used to examine questions related to immune and vascular responses during arterial pathology. The choice of antigen mismatch impacts the ability to appropriately address these questions.
Transplantation across complete MHC barriers permits a comprehensive evaluation of immune responses that are known to be involved in organ transplant rejection. This includes direct CD4 and CD8 T cell recognition and targeting of foreign peptide-MHC presented by graft-derived cells, indirect CD4 (and possibly CD8) T cell recognition and targeting of graft-derived alloantigens presented by recipient antigen presenting cells, and antibody-mediated recognition of alloantigens on vascular cell surfaces 8. However, the vascular response to injury in complete MHC-mismatched experiments may be different than that observed clinically. Johnson et al. 9 showed that, in aortic interposition grafts transplanted across a complete MHC mismatch barrier, most of the neointimal cells are of recipient origin and not of donor origin. This is different than that observed in human transplants where most intimal smooth muscle cells are of donor origin 9,10. To account for this limitation, alternate experimental models that involve grafting across minor histocompatibility antigen mismatches have been developed that trigger vascular responses that more closely resemble those observed in clinical transplantation 11. While these alternate models allow for important conclusions to be made regarding the vascular responses that drive the development of TA, the immunological processes that cause vascular rejection in minor histocompatibility antigen mismatched grafts do not completely re-capitulate those which occur in the clinical setting. For instance, minor histocompatibility antigens are recognized poorly by graft reactive antibodies 12. Given the above considerations, it is important to consider the pathological question being examined when choosing the type of antigen mismatch used in an aortic interposition model. Here we describe a detailed protocol for murine aortic interposition grafting. We describe interposition grafting between complete MHC-mismatched mice but the same protocol is used for grafting across other antigen mismatched mouse strains.

<table><tbody><tr><td>C57BL/6J (H-2b)</td><td>Jackson Laboratories, Bar Harbour ME</td><td>Strain# 000664</td><td></td></tr><tr><td>Balb/cBYJ</td><td>Jackson Laboratories, Bar Harbour ME</td><td>Strain# 001026</td><td></td></tr><tr><td>Ketamine Hydrochloride Injection USP 100 mg/ml</td><td>Ketalean</td><td>DIN 00612316</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>Xylazine Injection 20 mg/ml</td><td>Rompum</td><td>DIN 02169592</td><td></td></tr><tr><td>Ketoprofen Injection 100 mg/ml</td><td>Anafen</td><td>DIN 01938126</td><td></td></tr><tr><td>Butorphanol Tartrate&amp;nbsp;injection 10 mg/ml</td><td>Torbugesic</td><td>DIN 008450000</td><td></td></tr><tr><td>Buprenorphine Injection 0.3 mg/ml</td><td>Reckitt Benckiser</td><td>B.N. 5241</td><td></td></tr><tr><td>Atipamezole hydrochloride sterile injectable solution</td><td>Antisedan</td><td>DIN 02237744</td><td></td></tr><tr><td>Heparin Sodium Injection, USP, 1,000 units/ml</td><td>McKesson Distribution</td><td>DIN 02264315</td><td></td></tr><tr><td>Tears naturale ophthalmic ointment</td><td>Alcon</td><td>DIN 02082519</td><td></td></tr><tr><td>Stereomicroscope</td><td>Leica</td><td>M80</td><td></td></tr><tr><td>0.9% Sodium Chloride, sterile</td><td>Baxter Corporation</td><td></td><td></td></tr><tr><td>Lactated Ringer&amp;rsquo;s solution, sterile</td><td>Baxter Corporation</td><td></td><td></td></tr><tr><td>0.9% Sodium Chloride Injection, sterile, 10 ml</td><td>Baxter Corporation</td><td></td><td></td></tr><tr><td>Alcohol Prep Pads</td><td>Loris</td><td></td><td></td></tr><tr><td>Povidone Iodine</td><td>Betadine</td><td></td><td></td></tr><tr><td>Chlorohexidine Gluconate 4% w/v</td><td>Germi-Stat</td><td></td><td></td></tr><tr><td>Black Polyamide Monofilament</td><td>AROSurgical Instruments</td><td>T4A10Q07</td><td></td></tr><tr><td>Suture, 10-0 suture, 70 microns</td><td>Corporation</td><td></td><td></td></tr><tr><td>Blue monofilament suture 5-0, P3 needle</td><td>Ethicon</td><td>8698G</td><td></td></tr><tr><td>1 ml Syringe</td><td>BD</td><td>REF 309659</td><td></td></tr><tr><td>10 ml Syringe</td><td>BD</td><td>REF 309604</td><td></td></tr><tr><td>1cc TB insulin syringe with 28 G 1/2</td><td>BD</td><td>REF 309309</td><td></td></tr><tr><td>25 G 7/8, hypodermic needle</td><td>BD</td><td>REF 305124</td><td></td></tr><tr><td>27 G 1/2, hypodermic needle</td><td>BD</td><td>REF 305109</td><td></td></tr><tr><td>Colibri Retractor- 1.5 cm spread 4 cm</td><td>Fine Science Tools</td><td>17000-04</td><td></td></tr><tr><td>S&amp;amp;T CAF-4 Clip applying forceps, without lock</td><td>Fine Science Tools</td><td>00072-14</td><td></td></tr><tr><td>Supergrip forceps, S&amp;amp;T</td><td>Fine Science Tools</td><td>00632-11</td><td></td></tr><tr><td>Medical No.5 forceps</td><td>Fine Science Tools</td><td>11253-20</td><td></td></tr><tr><td>Lexer Baby Scissors</td><td>Fine Science Tools</td><td>14078-10</td><td></td></tr><tr><td>Micro Adson forceps serrated</td><td>Fine Science Tools</td><td>11018-12</td><td></td></tr><tr><td>Vannas-Tubingen microscissors</td><td>Fine Science Tools</td><td>15003-08</td><td></td></tr><tr><td>Micro clamps, b-1; 3.5mm x 1mm; 7mm length</td><td>Fine Science Tools</td><td>00396-01</td><td></td></tr><tr><td>Graefe-forceps, 10cm 1x2 teeth</td><td>Fine Science Tools</td><td>11054-10</td><td></td></tr><tr><td>Castroviejo with lock and tungsten jaws</td><td>Fine Science Tools</td><td>12565-14</td><td></td></tr><tr><td>Hot glass bead sterilizer</td><td>Inotech 250</td><td>IS-250 - Steri-250</td><td></td></tr><tr><td>Non-woven gauzes</td><td>Progene</td><td></td><td></td></tr><tr><td>Cotton Tipped Applicators</td><td>Puritan</td><td></td><td></td></tr><tr><td>Beard Trimmer</td><td>Wahl</td><td></td><td></td></tr><tr><td>Heating pad</td><td>Sunbeam</td><td></td><td></td></tr></tbody></table>

mouse model of alloimmune-induced vascular rejection and transplant arteriosclerosis

Vascular rejection that leads to transplant arteriosclerosis (TA) is the leading representation of chronic heart transplant failure. In TA, the immune system of the recipient causes damage of the arterial wall and dysfunction of endothelial cells and smooth muscle cells. This triggers a pathological repair response that is characterized by intimal thickening and luminal occlusion. Understanding the mechanisms by which the immune system causes vasculature rejection and TA may inform the development of novel ways to manage graft failure. Here, we describe a mouse aortic interposition model that can be used to study the pathogenic mechanisms of vascular rejection and TA. The model involves grafting of an aortic segment from a donor animal into an allogeneic recipient. Rejection of the artery segment involves alloimmune reactions and results in arterial changes that resemble vascular rejection. The basic technical approach we describe can be used with different mouse strains and targeted interventions to answer specific questions related to vascular rejection and TA.

In this model, the abdominal aorta from a Balb/cYJ mouse is interposed into the infrarenal aorta of a C57Bl/6 recipient. This permits a comprehensive evaluation of alloimmune responses that target allograft arteries. Immune-mediated vascular injury in this model initiates vascular reparative responses that culminate in intimal thickening, luminal narrowing and recruitment of immune cells (Figures 1 and 2). These criteria then serve as a read-out for the severity of alloimmune responses, ...

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Mouse Model of Alloimmune-induced Vascular Rejection and Transplant Arteriosclerosis

We describe a protocol for aortic interposition grafting in mice. The goal of the protocol is to provide a model with which to study pathological processes and therapeutic strategies relevant to alloimmune reactions in arteries and the resultant arterial changes that contribute to organ transplant failure.

Vascular rejection that leads to transplant arteriosclerosis (TA) is the leading representation of chronic heart transplant failure. In TA, the immune ...

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8.4K Views.  Simon Fraser University. We describe a protocol for aortic interposition grafting in mice. The goal of the protocol is to provide a model with which to study pathological processes and therapeutic strategies relevant to alloimmune reactions in arteries and the resultant arterial changes that contribute to organ transplant failure. 

Video: Mouse Model of Alloimmune-induced Vascular Rejection and Transplant Arteriosclerosis

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 ...

Orthotopic Aortic Transplantation in Mice for the Study of Vascular Disease

Vascular procedures involving anastomoses in the mouse are generally thought to be difficult and highly dependent on the skill of the individual surgeon. This is largely true, but there are a number of important principles that can reduce the difficulty of these procedures and enhance reproducibility. Orthotopic aortic transplantation is an excellent procedure in which to learn these principles because it involves only two end-to-end anastomoses, but requires good suturing technique and handling of the vessels for consistent success. This procedure begins with the procurement of a length of abdominal aorta from a donor animal, followed by division of the native aorta in the recipient. The procured aorta is then placed between the divided ends of the recipient aorta and sutured into place using end-to-end anastomoses. To accomplish this objective successfully requires a high degree of concentration, good tools, a steady hand, and an appreciation of how easily the vasculature of a mouse can be damaged, resulting in thrombosis. Learning these important principles is what occupies most of the beginner's time when learning microsurgery in small rodents. Throughout this protocol, we refer to these important points. This model can be used to study vascular disease in a variety of different experimental systems1-8. In the context shown here, it is most often used for the study of post-transplant vascular disease, a common long-term complication of solid organ transplantation in which intimal hyperplasia occurs within the allograft. The primary advantage of the model is that it facilitates quantitative morphometric analyses and the transplanted vessel lies contiguous to the endogenous vessel, which can serve as an additional control9. The technique shown here is most often used for mice weighing 18-25 grams. We have accumulated most of our experience using the C57BL/6J, BALB/cJ, and C3H/HeJ strains.

We describe a technique in which a section of the abdominal aorta from a mouse is transplanted orthotopically to just below the renal arteries in an allogeneic or syngeneic recipient. This technique can be useful in studies in which transplantation of large arteries of uniform size is deemed advantageous.

Vascular procedures involving anastomoses in the mouse are generally thought to be difficult and highly dependent on the skill of the individual surgeon. ...

Mouse Models for Graft Arteriosclerosis

Graft arteriosclerois (GA), also called allograft vasculopathy, is a pathologic lesion that develops over months to years in transplanted organs characterized by diffuse, circumferential stenosis of the entire graft vascular tree. The most critical component of GA pathogenesis is the proliferation of smooth muscle-like cells within the intima. When a human coronary artery segment is interposed into the infra-renal aortae of immunodeficient mice, the intimas could be expand in response to adoptively transferred human T cells allogeneic to the artery donor or exogenous human IFN-&gamma; in the absence of human T cells. Interposition of a mouse aorta from one strain into another mouse strain recipient is limited as a model for chronic rejection in humans because the acute cell-mediated rejection response in this mouse model completely eliminates all donor-derived vascular cells from the graft within two-three weeks. We have recently developed two new mouse models to circumvent these problems. The first model involves interposition of a vessel segment from a male mouse into a female recipient of the same inbred strain (C57BL/6J). Graft rejection in this case is directed only against minor histocompatibility antigens encoded by the Y chromosome (present in the male but not the female) and the rejection response that ensues is sufficiently indolent to preserve donor-derived smooth muscle cells for several weeks. The second model involves interposing an artery segment from a wild type C57BL/6J mouse donor into a host mouse of the same strain and gender that lacks the receptor for IFN-&gamma; followed by administration of mouse IFN-&gamma; (delivered via infection of the mouse liver with an adenoviral vector. There is no rejection in this case as both donor and recipient mice are of the same strain and gender but donor smooth muscle cells proliferate in response to the cytokine while host-derived cells, lacking receptor for this cytokine, are unresponsive. By backcrossing additional genetic changes into the vessel donor, both models can be used to assess the effect of specific genes on GA progression. Here, we describe detailed protocols for our mouse GA models.

We describe protocols for our mouse graft arteriosclerois (GA) models which involve interposition of a mouse vessel segment into a recipient of the same inbred strain. By backcrossing additional genetic changes into the vessel donor, the model can assess the effect of specific genes on GA.

Graft arteriosclerois (GA),  also called allograft vasculopathy, is a pathologic lesion that develops over  months to years in transplanted organs ...

Mouse Kidney Transplantation: Models of Allograft Rejection

Rejection of the transplanted kidney in humans is still a major cause of morbidity and mortality. The mouse model of renal transplantation closely replicates both the technical and pathological processes that occur in human renal transplantation. Although mouse models of allogeneic rejection in organs other than the kidney exist, and are more technically feasible, there is evidence that different organs elicit disparate rejection modes and dynamics, for instance the time course of rejection in cardiac and renal allograft differs significantly in certain strain combinations. This model is an attractive tool for many reasons despite its technical challenges. As inbred mouse strain haplotypes are well characterized it is possible to choose donor and recipient combinations to model acute allograft rejection by transplanting across MHC class I and II loci. Conversely by transplanting between strains with similar haplotypes a chronic process can be elicited were the allograft kidney develops interstitial fibrosis and tubular atrophy. We have modified the surgical technique to reduce operating time and improve ease of surgery, however a learning curve still needs to be overcome in order to faithfully replicate the model. This study will provide key points in the surgical procedure and aid the process of establishing this technique.

Here, we present a protocol to study the immunology of rejection. The surgical model presented reports a short operating time and a concise technique. Depending on the donor-recipient strain combination, the transplanted kidney may develop acute cellular rejection or chronic allograft damage, defined by interstitial fibrosis and tubular atrophy.

Rejection of the transplanted kidney in humans is still a major cause of morbidity and mortality. The mouse model of renal transplantation closely ...

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 ...

Immunology and Infection

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 ...

Orthotopic Hind Limb Transplantation in the Mouse

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and translational research in the field of transplantation medicine. A vast array of reagents is available exclusively in this setting, including mono- and polyclonal antibodies for both diagnostic and interventional applications. In addition, a vast number of genotyped, inbred, transgenic, and knock out strains allow detailed investigation of the individual contributions of humoral and cellular components to the complex interplay of an immune response and make the mouse the gold standard for immunological research. 
Vascularized Composite Allotransplantation (VCA) delineates a novel field of transplantation using allografts to replace "like with like" in patients suffering traumatic or congenital tissue loss. This surgical methodological protocol shows the use of a non-suture cuff technique for super-microvascular anastomosis in an orthotopic mouse hind limb transplantation model. The model specifically allows for comparison between established paradigms in solid organ transplantation with a novel form of transplants consisting of various different tissue components. Uniquely, this model allows for the transplantation of a viable vascularized bone marrow compartment and niche that have the potential to exert a beneficial effect on the balance of immune acceptance and rejection. This technique provides a tool to investigate alloantigen recognition and allograft rejection and acceptance, as well as enables the pursuit of functional nerve regeneration studies to further advance this novel field of transplantation.

This novel model for orthotopic hind limb transplantation in the mouse, applying a non-suture cuff technique for super-microvascular anastomosis, provides a powerful tool for in&#160;vivo mechanistic immunological research related to vascularized composite allotransplantation (VCA).

In vivo animal model systems, and in particular mouse models, have evolved into powerful and versatile scientific tools indispensable to basic and ...

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand and face transplants and, more recently, abdominal wall, uterus, and urogenital transplants. Functional outcomes have exceeded initial expectations, and most recipients enjoy an improved quality of life. However, as clinical experience accumulates, chronic rejection and complications from the immunosuppression must be addressed. In many cases where grafts have failed, the causative pathology has been ischemic vasculopathy. <span style="font-size: 11pt; line-height: 115%; font-family: Arial, sans-serif; color: rgb(41, 43, 49); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">The biological mechanisms of the acute and chronic rejection associated with VCA, especially ischemic vasculopathy, are important areas of research. However, due to the very small number of VCA patients, the evaluation of proposed mechanisms is better addressed in an experimental model. Multiple groups have used animal models to address some of the relevant unsolved questions in VCA rejection and vasculopathy. Several model designs involving a variety of species are described in the literature. Here we present a reproducible model of VCA heterotopic hindlimb osteomyocutaneous flap in the rat that can be utilized for translational VCA research. This model allows for the serial evaluation of the graft, including biopsies and different imaging modalities, while maintaining a low level of morbidity.

Vascularized composite allograft offers life-altering benefits to transplant recipients, but the biological causes of graft rejection and vasculopathy remain poorly understood. The rodent surgical model presented here offers a reproducible, clinically relevant model of transplantation, allowing researchers to evaluate rejection events and potential therapeutic strategies to prevent their occurrence.

Vascularized composite allotransplantation (VCA) is a relatively new field in the reconstructive surgery. Clinical achievements in human VCA include hand ...

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

A Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential model to investigate the mechanism of renal allograft rejection in pre-clinical studies and could aid in the development of novel approaches to improve the long-term survival of renal allografts. Donor kidney implantation in rat orthotopic renal transplantation is commonly performed by end-to-side anastomosis to recipients' aorta and inferior vena cava. In this model, the donor's kidney was implanted using end-to-end anastomosis to the recipients' renal artery and renal vein. The donor's ureter was anastomosed to the recipient's bladder in an end-to-side 'tunnel' method. This model contributes to better healing of ureter-bladder anastomosis and increases the recipients' survival by avoiding interference with blood supply and venous reflux of the lower body. This model can be used to investigate the mechanisms of acute and chronic immune and pathologic rejection of renal allografts. Here, the study describes the detailed protocols of this orthotopic renal transplantation between rats.

The rat orthotopic renal transplantation model contributes to investigating the mechanism of renal allograft rejection. The current model increases the recipients' survival without interference with blood supply and venous reflux of the lower body using an end-to-end anastomosis of kidney implantation and an end-to-side "tunnel" method of ureter-bladder anastomosis.

Renal allograft rejection limits the long-term survival of patients after renal transplantation. Rat orthotopic renal transplantation is an essential ...

Mouse Model of Alloimmune-induced Vascular Rejection and Transplant Arteriosclerosis

Summary

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Chapters in this video

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

Orthotopic Aortic Transplantation in Mice for the Study of Vascular Disease

Mouse Models for Graft Arteriosclerosis

Mouse Kidney Transplantation: Models of Allograft Rejection

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

Orthotopic Hind Limb Transplantation in the Mouse

Modified Heterotopic Hindlimb Osteomyocutaneous Flap Model in the Rat for Translational Vascularized Composite Allotransplantation Research

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

A Rat Orthotopic Renal Transplantation Model for Renal Allograft Rejection