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.

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			<td>Ketoprofen Injection 100 mg/mL</td>
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			<td>Butorphanol Tartrate&#160;injection 10 mg/mL</td>
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			<td>Buprenorphine Injection 0.3 mg/mL</td>
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			<td>Lactated Ringer&#8217;s solution, sterile</td>
			<td>Baxter Corporation</td>
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			<td></td>
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		<tr>
			<td>0.9% Sodium Chloride Injection, sterile, 10 mL</td>
			<td>Baxter Corporation</td>
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			<td>Alcohol Prep Pads</td>
			<td>Loris</td>
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			<td>Povidone Iodine</td>
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			<td>Germi-Stat</td>
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			<td>Black Polyamide Monofilament</td>
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			<td>Suture, 10-0 suture, 70 microns</td>
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			<td>Blue monofilament suture 5-0, P3 needle</td>
			<td>Ethicon</td>
			<td>8698G</td>
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			<td>BD</td>
			<td>REF 309659</td>
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			<td>10 ml Syringe</td>
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			<td>REF 309604</td>
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			<td>BD</td>
			<td>REF 309309</td>
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			<td>27G 1/2, hypodermic needle</td>
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			<td>Fine Science Tools</td>
			<td>14078-10</td>
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			<td>Micro Adson forceps serrated</td>
			<td>Fine Science Tools</td>
			<td>11018-12</td>
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			<td>Fine Science Tools</td>
			<td>15003-08</td>
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			<td>Progene</td>
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			<td>Cotton Tipped Applicators</td>
			<td>Puritan</td>
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			<td>Beard Trimmer</td>
			<td>Wahl</td>
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			<td>Heating pad</td>
			<td>Sunbeam</td>
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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, ...

Watch this Scientific Journal Video about Mouse Model of Alloimmune-induced Vascular Rejection and Transplant Arteriosclerosis at JoVE.com

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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Research

JoVE Journal

Medicine

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

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 ...

Technical Aspects of the Mouse Aortocaval Fistula

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta and inferior vena cava (IVC) are exposed. The proximal infrarenal aorta and the distal aorta are dissected for clamp placement and needle puncture, respectively. Special attention is paid to avoid dissection between the aorta and the IVC. After clamping the aorta, a 25 G needle is used to puncture both walls of the aorta into the IVC. The surrounding connective tissue is used for hemostatic compression. Successful creation of the AVF will show pulsatile arterial blood flow in the IVC. Further confirmation of successful AVF can be achieved by post-operative Doppler ultrasound.

The goal is to produce an arteriovenous fistula that is simple and reproducible. This method does not use sutures or glue adhesive. Therefore the samples can be used with the least amount of foreign materials for analysis.

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta ...

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 ...

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 ...

Ferric Chloride-induced Thrombosis Mouse Model on Carotid Artery and Mesentery Vessel

Severe thrombosis and its ischemic consequences such as myocardial infarction, pulmonary embolism and stroke are major worldwide health issues. The ferric chloride injury is now a well-established technique to rapidly and accurately induce the formation of thrombi in exposed veins or artery of small and large diameter. This model has played a key role in the study of the pathophysiology of thrombosis, in the discovery and validation of novel antithrombotic drugs and in the understanding of the mechanism of action of these new agents. Here, the implementation of this technique on a mesenteric vessel and carotid artery in mice is presented. The method describes how to label circulating leukocytes and platelets with a fluorescent dye and to observe, by intravital microscopy on the exposed mesentery, their accumulation at the injured vessel wall which leads to the formation of a thrombus. On the carotid artery, the occlusion caused by the clot formation is measured by monitoring the blood flow with a Doppler probe.

The FeCl3 induced thrombosis model in mice is described herein. A method to monitor thrombus growth by intravital microscopy observation on a mesenteric vessel and by blood flow measurement in the carotid artery is presented.

Severe thrombosis and its ischemic consequences such as myocardial infarction, pulmonary embolism and stroke are major worldwide health issues. The ferric ...

Intraluminal Drug Delivery to the Mouse Arteriovenous Fistula Endothelium

Delivery of therapeutic agents to enhance arteriovenous fistula (AVF) maturation can be administered either via intraluminal or external routes. The simple murine AVF model was combined with intraluminal administration of drug solution to the venous endothelium at the same time as fistula creation. Technical aspects of this model are discussed. Under general anesthesia, an abdominal incision is made and the aorta and inferior vena cava (IVC) are exposed. The infra-renal aorta and IVC are dissected for clamping. After proximal and distal clamping, the puncture site is exposed and a 25 G needle is used to puncture both walls of the aorta and into the IVC. Immediately after the puncture, a reporter gene-expressing viral vector was infused in the IVC via the same needle, followed by 15 min of incubation. The intraluminal administration method enabled more robust viral gene delivery to the venous endothelium compared to administration by the external route. This novel method of delivery will facilitate studies that explore the role of the endothelium in AVF maturation and enable intraluminal drug delivery at the time of surgical operation.

After puncturing the aorta through the inferior vena cava (IVC) to create an aorto-caval fistula in the mouse, solution containing a drug is infused into the IVC via the same needle, followed by incubation. This method enables more robust drug delivery to the venous endothelium compared to the external route.

Delivery of therapeutic agents to enhance arteriovenous fistula (AVF) maturation can be administered either via intraluminal or external routes. The ...

Vein Interposition Model: A Suitable Model to Study Bypass Graft Patency

Bypass grafting is an established treatment method for coronary artery disease. Graft patency continues to be the Achilles heel of saphenous vein grafts. Research models for bypass graft failure are essential for a better understanding of pathobiological and pathophysiological processes during graft patency loss. Large animal models, such as pigs or sheep, resemble human anatomical structures but require special facilities and equipment. This video describes a rat vein interposition model to investigate vein graft patency loss. Rats are inexpensive and easy to handle. Compared to mouse models, the convenient size of rats permits better operability and enables a sufficient amount of material to be obtained for further diverse analysis. In brief, the inferior epigastric vein of a donor rat is harvested and used to replace a segment of the femoral artery. Anastomosis is conducted via single stitches and sealed with fibrin glue. Graft patency can be monitored non-invasively using duplex sonography. Myointimal hyperplasia, which is the main cause for graft patency loss, develops progressively over time and can be calculated from histological cross sections.

This video demonstrates a model to study the development of myointimal hyperplasia after venous interposition surgery in rats.

Bypass grafting is an established treatment method for coronary artery disease. Graft patency continues to be the Achilles heel of saphenous vein grafts. ...

Using the Sleeve Technique in a Mouse Model of Aortic Transplantation - An Instructional Video

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the aorta in mice using the sleeve method was short and easy averaging 20 min, permitting studies of iso/allo grafts. The following article describes the aortic transplantation procedure used in our laboratory. The mice were anesthetized with a mixture of 1.5% volume isoflurane and 100% oxygen through a face mask. At this point, the segment of the aorta between the renal arteries and its bifurcation was separated from the vena cava, freely prepared and clampedat the proximal and distal segments with a single silk suture. Prior to the removal of the aorta, a saline solution containing heparin was injected into the inferior vena cava. Then the aorta was cut between the clamps and a saline heparin solution was used to flush the lumen. The sleeve technique with monofilament sutures was used in order to transplant the abdominal aorta in the orthotopic position.

We present an orthotopic aortic transplantation model using the sleeve technique in mice. It is a very rapid anastomosis method, which can be employed in studies of vascular disease.

Orthotopic aortic transplantation using the sleeve technique reduces injury to the aorta with failure rate of only 10-20%. The time to anastomose the ...

Mouse Model for Pancreas Transplantation Using a Modified Cuff Technique

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the availability of the widest range of molecular probes and reagents to perform in vivo as well as in vitro studies. Based on our experience with various murine transplantation models, we developed a heterotopic pancreas transplantation model in mice with the intent to analyze mechanisms underlying severe ischemia reperfusion injury-associated early graft damage. In contrast to previously described techniques using suture techniques, herein we describe a new procedure using a non-suture cuff technique. 
In recent years, we have performed more than 300 pancreas transplantations in mice with an overall success rate of &gt;90%, a success rate never described before in mouse pancreas transplantation. The backbone of this non-suture cuff technique for graft revascularization consists of two major steps: (I) pulling the recipient vessel over a polyethylene/polyamide cuff and fixing it with a circumferential ligature, and (II) placing the donor vessel over the everted recipient vessel and fixing it with a second circumferential ligature. The resultant continuity of the endothelial layer results in less thrombogenic lesions with high patency rates and, finally, high success rates.
In this model, arterial anastomosis is achieved by pulling the abdominal aorta of the donor graft over the everted common carotid artery of the recipient animal. Venous drainage of the graft is achieved by pulling the portal vein of the graft over the everted external jugular vein of the recipient. This manuscript provides details and crucial steps of the organ recovery and organ implantation procedures, which will allow researchers with microsurgical skills to perform the transplantation successfully in their laboratories.

Among abdominal solid organ transplantation, pancreatic grafts are prone to develop severe ischemia reperfusion injury-associated graft damage, leading eventually to early graft loss. This protocol describes a model of murine pancreas transplantation using a non-suture cuff technique, ideally suited for analyzing these early, deleterious damages.

Mouse models have several advantages in transplantation research, including easy handling, a variety of genetically well-defined strains, and the ...

Quantitative Micro-CT Analysis of Aortopathy in a Mouse Model of &#946;-aminopropionitrile-induced Aortic Aneurysm and Dissection

Aortic aneurysm and dissection is associated with significant morbidity and mortality in the population and can be highly lethal. While animal models of aortic disease exist, in vivo imaging of the vasculature has been limited. In recent years, micro-computerized tomography (micro-CT) has emerged as a preferred modality for imaging both large and small vessels both in vivo and ex vivo. In conjunction with a method of vascular casting, we have successfully used micro-CT to characterize the frequency and distribution of aortic pathology in &#946;-aminopropionitrile-treated C57/Bl6 mice. Technical limitations of this method include variations in the quality of the perfusion introduced by poor animal preparation, the application of proper methodologies for vessel size quantification, and the non-survivability of this procedure. This article details a methodology for the intravascular perfusion of a lead-based radiopaque silicone rubber for the quantitative characterization of aortopathy in a mouse model of aneurysm and dissection. In addition to visualizing aortic pathology, this method may be used for examining other vascular beds in vivo or vascular beds removed post-mortem.

Quantitative Micro-CT Analysis of Aortopathy in a Mouse Model of &amp;#946;-aminopropionitrile-induced Aortic Aneurysm and Dissection

This article describes a detailed methodology of using a radiopaque lead-based silicone rubber to perfuse the murine vasculature for aortic diameter quantification in a mouse model of aortic aneurysm and dissection.

Aortic aneurysm and dissection is associated with significant morbidity and mortality in the population and can be highly lethal. While animal models of ...