The authors gratefully acknowledge funding through the Department of Defense Congressionally Directed Research Program (DOD CDMRP; W81XWH-16-1-0582) to ABB and RS. The authors also acknowledge funding through the American Heart Association (17IRG33410888), the DOD CDMRP (W81XWH-16-1-0580) and the National Institutes of Health (1R21EB023551-01; 1R21EB024147-01A1; 1R01HL141761-01) to ABB.

Studies involving animals were performed with the approval of the University of Texas at Austin and the UTHealth Science Center at Houston Institutional Animal Care and Use Committee (IACUC), the Animal Care and Use Review Office (ACURO) of The United States Army Medical Research and Materiel Command Office of Research Protections, and in accordance with NIH guidelines for animal care.
1. Induction of diabetes and hyperlipidemia
<ol>
	<li>Transition the New Zealand rabbits (4&#8211;6 months old) from one cup of standard alfalfa chow to 0.1% cholesterol chow over the course of four days. For days 1&#8211;5, use standard chow to cholesterol chow ratios of 1:0, 3:1, 1:1, 1:3, and 0:1, respectively. After two weeks on 0.1% cholesterol chow, induce rabbits to have diabetes using alloxan injection as described in the following steps</li>
	<li>Sedate the rabbits using 35&#8211;75 mg/mL ketamine and 1&#8211;2 mg/mL acepromazine via subcutaneous injection and prep for an IV injection by introducing a catheter into the marginal left ear vein using a 22 g catheter.</li>
	<li>Collect a drop of blood from the rabbits via hub of the ear vein catheter for the baseline blood glucose level (BGL) measurement. Any standard glucometer can be used. Normal glucose levels for a rabbit are typically in the range of 80 to 150 mg/dL.</li>
	<li>Inject alloxan at 100 mg/kg reconstituted in saline to a volume of 8 mL through the ear catheter slowly over an 8-minute period using a syringe pump.</li>
	<li>Check the BGL every hour for the next 12 h using a standard glucometer to monitor for hypoglycemia.
	<ol>
		<li>Place the rabbit in&#160;a restrainer.</li>
		<li>Anesthetize the ear with 2.5% lidocaine/2.5% prilocaine cream.</li>
		<li>Take blood from the lateral ear vein using a 27&#160;G needle and measure BGL using a standard meter.</li>
	</ol>
	</li>
	<li>Measure the BGL two times a day for the first 7 days. Give the rabbits an injection of insulin if the BGL reaches or exceeds 350 mg/dL.</li>
	<li>Prepare a 3-mm stainless steel ball for implantation as a size marker during angiograms prior to the day of surgery.
	<ol>
		<li>Cut a 10-mm circular piece of silastic sheeting out of a larger sheet using a biopsy punch.</li>
		<li>Mount the ball in the center of the sheet using clear silicone sealant.</li>
		<li>Completely cover the ball with the sealant. Allow the sealant to cure for a minimum of 24 h.</li>
		<li>Place the ball in an open 2 inch x 3 inch low density polyethylene bag and place it into a sterilization bag to be sterilized with ethylene oxide gas.</li>
	</ol>
	</li>
</ol>
2. Preparation of rabbit for surgery
<ol>
	<li>Anesthetize the rabbit using 20&#8211;40 mg/kg ketamine and 2 mg/kg midazolam via subcutaneous injection. Place the rabbit on 1.5%&#8211;3% isoflurane (typically 2%) throughout the initial sedation using a mask. Give an injection of alfaxalone to maintain anesthesia via an intramuscular injection of 3 mg/kg.</li>
	<li>Once anesthetized, remove the mask and insert a cuffed endotracheal tube, into the airway and connect to a ventilator. Continue to administer isoflurane at 1.5%&#8211;3%.</li>
	<li>Collect blood from the central artery from either ear for a baseline chemistry panel.</li>
	<li>Place a 22 G ear vein catheter in the lateral ear vein for Lactated Ringer&#39;s Solution drip throughout the surgical procedure. Alternatively, normal saline (0.9% sodium chloride) can be used.</li>
	<li>Using the lateral vein in the opposite ear, place a catheter in the vein and deliver alfaxalone at 6 mg/kg/h. Gradually increase the alfaxalone to 8 mg/kg/h while decreasing isoflurane to 0.6% during the prep period.</li>
	<li>To limit pain and risk of infection, administer buprenorphine (0.01 mg/kg) and enrofloxacin (5 mg/kg) using a subcutaneous injection with a 25 G needle.</li>
	<li>Trim the hair on the neck, right and left inner thighs, and back using clippers (#40 blade). Hair is removed from the back to maintain contact with the grounding pad.</li>
	<li>Place a blood pressure cuff on each of the hind limbs and measure initial blood pressure. Place the cuff just below the knee with the probe just above the hock on the lateral surface.</li>
	<li>Position the rabbit on the surgery table on its back and scrub and drape the surgery sites. This includes the neck for carotid artery access and inner right thigh for femoral artery access. Perform the sterilization scrub with alternating scrubs of 2% chlorhexidine and 70% ethyl alcohol. Repeat this three times, then apply a final spray with 2% chlorhexidine solution.</li>
	<li>Place a 3-mm stainless steel ball that has been sterilized inside a low density polyethyene bag on top of the right (scrubbed) leg near the upper part of the thigh to serve as a size reference during angiogram measurements. Place a sterile drape over the leg until the time of surgery. Leave the ball inside the sterile plastic bag during the first angiogram.</li>
</ol>
3. Angiography
<ol>
	<li>Expose the right common carotid artery
	<ol>
		<li>Make a 4&#8211;5 cm long incision just lateral to the trachea using a scalpel with a #15 blade.</li>
		<li>Use blunt dissection to expose the carotid artery and open the incision using small Weitlaner retractors. Carefully isolate the carotid artery from the jugular vein and vagus nerve. Typically, a curved Metzenbaum scissors and a curved mosquito hemostat are used for the blunt dissection. Be sure to get full separation of the carotid artery from the nerve and jugular vein to make the ligatures only ligate the artery.</li>
	</ol>
	</li>
	<li>Place a ligature using a 4-0 silk suture at the proximal and distal ends of the exposed artery. Tie off the distal end of carotid with a surgeon&#8217;s knot followed by four square knots. On the proximal end, use a ligaloop to allow it to be tightened or loosened as needed. The use of a ligaloop placed at the proximal end of the exposed artery can help secure the introducer and catheter.</li>
	<li>Administer 500 IU of heparin through the IV. Use approximately 0.5 mL of 1% lidocaine applied along the exposed carotid to dilate the vessel. One treatment is usually sufficient, but it can be repeated as needed. Cut approximately half way through the carotid artery using a scalpel or iris scissors, then place&#160;the 4-inch wire insertion tool into the artery.</li>
	<li>Feed a 0.014 inch x 185 cm guidewire through the insertion tool to the aortic bifurcation at the iliac crest in the descending aorta. Remove the insertion tool and insert a 3F pigtail angiographic catheter over the wire.</li>
	<li>Advance the pigtail catheter to be 2 cm proximal to the aortic bifurcation at the iliac crest in the descending aorta.</li>
	<li>Position the tip of the catheter between the seventh lumbar and first sacral vertebrae. Test the location of the catheter by manually injecting a 2&#8211;4 mL of contrast agent.</li>
	<li>Administer an intra-arterial injection of 100 &#956;g nitroglycerin through the catheter to increase vasodilation.</li>
	<li>Administer 0.8&#160;mL of 1% lidocaine to the rabbit through the catheter to assist with vasodilation during the angiogram. Attach the tubing for the injector to the catheter and remove any air bubbles in the line. Inject 8-9 mL of contrast media using automated angiographic injector through the catheter.</li>
	<li>Record serial images of the hind limbs using angiography.
	<ol>
		<li>Set the power injector to inject contrast at 3 mL/sec for a total of 8-9&#160;mL. Perform digital subtraction angiography at 6 frames per second.</li>
		<li>Select the serial images created and alter a photo of each angiogram using approximately -40% setting to minimize appearance of bone and capture a complete picture of the vessel perfusion with contrast. An example angiogram of the vascular flow after femoral artery ligation/excision is shown in Figure 1.</li>
	</ol>
	</li>
</ol>
4. Isolation of the femoral artery
<ol>
	<li>Make a longitudinal incision in the skin over the right femoral artery using a scalpel (#15 blade). Ensure that the incision extends inferiorly from the inguinal ligament ending at the area just proximal to the patella (approximately 6 cm).</li>
	<li>Use blunt dissection with curved Metzenbaum scissors or a curved mosquito hemostat to expose the femoral artery.</li>
	<li>Use Weitlaner retractors to hold the incision open.</li>
	<li>Add 0.5 mL of 1% lidocaine locally to reduce nerve irritation and promote vasodilation.</li>
	<li>Continue blunt dissection of the tissues to free the entire length of the femoral artery along with all branches of the femoral artery, including the inferior epigastric, deep femoral, lateral circumflex, and superficial epigastric arteries (Figure 2A).</li>
	<li>Dissect further along the popliteal and saphenous arteries as well as the external iliac artery (Figure 2A). Periodically moisten the area with saline to protect from tissue damage. If the blunt dissection is performed along the femoral groove (between the muscles) there is no need to cut the muscle.</li>
	<li>Carefully separate the artery from the vein and nerve as shown in Figure 2B,C. Ligate the arteries indicated by the diagram with 4.0 silk sutures by placing two ties with enough space between them to cut the artery. These ties are performed with a Surgeon&#8217;s knot followed by four square knots.</li>
	<li>Cut between the two ties on the ligated arteries using the small Metzenbaum scissors. Excise the femoral artery from its proximal origin as a branch of the external iliac artery to the point distally where it bifurcates to form the saphenous and popliteal arteries.</li>
</ol>
5. Repeat angiography
<ol>
	<li>Use 4-0 silk suture to attach the silastic sheet, with 3-mm stainless steel ball to the upper part of the quadriceps muscle. Pull the skin over the ball after it is in place.</li>
	<li>Administer an intra-arterial injection of 100 &#956;g nitroglycerin through the catheter to increase vasodilation.</li>
	<li>If needed, administer another 0.8&#160;mL of 1% lidocaine to the rabbit through the catheter to assist with vasodilation during the angiogram.</li>
	<li>Inject 8-9 mL of contrast media using an automated angiographic injector.</li>
	<li>Perform angiography as described in step 3.9.</li>
</ol>
6. Wound closure and recovery
<ol>
	<li>Remove the catheter from the right artery. Tie off the artery using the 4-0 silk suture that is already in place around the artery.</li>
	<li>Suture both wounds closed. Close muscle and subcuticular layers using 4-0 polydioxanone or 3-0 polyglactin 910 on a taper needle (see Table of Materials) in a continuous suture pattern. Close the skin using 4-0 polydioxanone or 4-0 polyglactin 910 on a reverse cutting needle (see Table of Materials) in a buried continuous subcuticular suture pattern. 
	NOTE: If available, polydioxanone is preferred for both.</li>
	<li>Administer intradermal injections of 0.25% bupivacaine near the incisions using a syringe with a 25 G needle. Insert the needle and inject 0.5 mL while the needle is pulled back. Give one injection per side of the wound for the incision on the neck (two injections on the neck) and two injections per side of the wound for the incision on the leg (four injections on the leg; six injections in total). The total volume injected is 3 mL (0.5 mL x 6 injections).</li>
	<li>Administer subcutaneous injections of 0.5 mg/kg meloxicam and sustained release buprenorphine at 0.12 mg/kg.</li>
	<li>Monitor the rabbit as it recovers from anesthesia. The rabbit will automatically start to swallow as it wakes up from anesthesia. Once the swallowing response occurs, remove the endotracheal tube. Provide close monitoring and thermal support until the rabbit is able to maintain cardiovascular function and body temperature. Return the rabbit to its enclosure once it is able to ambulate.</li>
	<li>Employ fresh vegetables and/or syringe feeding of a critical care diet along with subcutaneous saline injections if the rabbit does not tolerate chow after the surgery. Cabbage, broccoli, cauliflower, carrots, or other in seasonal vegetables can be used. Shred the vegetables and mix them together to aid in the rabbit returning to eating.</li>
</ol>
7. Monitoring
<ol>
	<li>Anesthetize the rabbits every two weeks to acquire blood pressure on both legs as described in step 2.8. Harvest blood from the central artery of the ear for use in blood chemistry assays. Alternatively, take blood from the saphenous vein or cephalic vein. Take approximately 2 mL at each time point. Use a standard blood chemistry panel for analysis. If needed, add tests for low density lipoprotein (LDL), high density lipoprotein (HDL), or hemoglobin A1c (HbA1c).</li>
	<li>Take a very small amount of blood for BGL measurements.</li>
</ol>
8. Treatment
<ol>
	<li>Prepare ten syringes with treatment, carrier, and crosslinker. Fill each syringe just prior to use with 100 &#956;L of calcium sulfate slurry and then 100 &#956;L of 2% sodium alginate with growth factors or other treatments such that the alginate is nearest the tip of the syringe.</li>
	<li>Administer one prepared injection into the muscle before preparing the next. This reduces the time that the alginate interacts with the calcium sulfate in the syringe. Space the injections evenly along both sides of the femoral artery on the thigh. To achieve uniform injections, create a silicone sheet with holes to guide the injection, as described in other studies19. This can be easily prepared by using a biopsy punch to create holes in commercially available silicone sheeting.</li>
</ol>
9. Endpoint angiography, euthanasia, perfusion fixation and tissue harvest
<ol>
	<li>At the endpoint date, perform angiography as described in step 3 but use the left carotid artery for access.</li>
	<li>After angiography, move the animal to the necropsy table and perform perfusion fixation to preserve the hind limb tissues:
	<ol>
		<li>Increase the isoflurane to 3%&#8211;4% and perform a toe pinch to confirm the anesthesia is sufficiently deep.</li>
		<li>Administer 1000-2000 IU of heparin intravenously.</li>
		<li>Create an incision along the midline of the ribcage and spanning the length of the diaphragm using a scalpel with a #20 blade.</li>
		<li>With the rib cage exposed, cut the ribs just left of the midline using rib cutters. Use Weitlaner retractors to expose the heart.</li>
		<li>Set up the pump with output tubing with an inner diameter of 1/8 inch and a 18G needle at the end. Preload the line with saline and have at least 600 mL of saline and formalin prepared in separate containers for the perfusion.</li>
		<li>Insert the 18 G needle connected to the pump into the left ventricle via the apex of the heart. Insert another 18 G needle (unattached to anything) into the right atrium and allow blood to flow out into the downdraft of the necropsy table.</li>
		<li>Use a perfusion pump to control the flow of approximately 500 mL of saline into the heart. Use a pump setting to flow 110 mL/min.</li>
		<li>Once the fluid coming from the heart is clear, move the tubing from the saline reservoir to one filled with a 10% formalin solution. Twitching will occur in all four limbs if the perfusion is working properly. Pump approximately 500 mL of formalin solution into the left ventricle.</li>
		<li>Turn the pump off and remove the needles from the heart.</li>
	</ol>
	</li>
	<li>Remove both hind limbs at the hip by cutting around the hip joint with a scalpel with #20 blade. Use a small rib cutter to remove the limbs. Use the non-ischemic limb as a control.</li>
	<li>Store the limbs in formalin for 24 h at 4 &#176;C and then stored in 70% ethanol at 4 &#176;C.</li>
	<li>For histological analysis, take multiple biopsies from the limbs. We have used eight 6-mm biopsies taken at regions across the thigh and calf in both limbs. 
	NOTE: While ankle blood pressure measurement and angiography are the most commonly used methods for measuring recovery of blood flow, other methods can be used to track recovery of the animals including Doppler ultrasound, laser Doppler imaging, infrared thermography62, microsphere determined perfusion63,64, computed tomography (CT) imaging, and magnetic resonance imaging (MRI)65.</li>
</ol>

The authors have nothing to disclose.

We have presented a preclinical model for inducing hind limb ischemia in rabbits with diabetes and hyperlipidemia. In many studies, there is ambiguity to the technique used to create hind limb ischemia in rabbits. In mice, the severity and recovery from hind limb ischemia is highly dependent on the location the ligation and technique used to induce ischemia. The significance of the technique presented in this work is that it allows for the consistent induction of ischemia that does not fully recover after 8 weeks in diab...

Peripheral arterial disease (PAD) is a common circulatory disorder in which the progression of atherosclerotic plaque formation leads to a narrowing of blood vessels in the limbs of the body. The recent increase in risk factors for atherosclerosis, including diabetes, obesity, and inactivity, has led to increasing prevalence of vascular disease1. Currently, it is estimated that 12%&#8211;20% of the general population over 60 years old has peripheral arterial disease2. A major consequence of peripheral arterial disease is the development of peripheral ischemia, most commonly found in the lower limbs. In severe cases, patients can develop critical limb ischemia, a state in which there is constant pain due to a lack of blood flow. Patients with critical limb ischemia have a 50% likelihood of having one limb amputated within one year of diagnosis. Furthermore, patients with diabetes have a higher incidence of peripheral arterial disease and poorer outcomes following interventions for revascularization3,4. Current therapies for peripheral ischemia include percutaneous interventions such as atherectomy and stenting or surgical bypass. However, for many patients these treatments only provide short-term benefits and many are not healthy enough for major surgical procedures. In this work, we describe a preclinical animal model for testing new treatments targeting peripheral vascular disease that incorporates the generation of peripheral ischemia in rabbits through surgical ligation in the context of the diabetic disease state.
The hind limb ischemia model in rabbits has been used as a physiological model for obstructive vascular disease and preclinical precursor to human studies for over half a century5,6. Rabbits are often a preferred species for studies on peripheral ischemia due to the developed musculature of the ankle and calf muscle, in contrast to common large animal models that are ungulates (animals with hooves). Several recent reviews have addressed the use of this model and others in modeling peripheral vascular disease in humans7,8. Similar models using hind limb ischemia in rabbits were used in preclinical studies of growth factors9,10,11,12,13,14,15,16,17,18,19,20, gene therapy21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42,43,44, and stem cells45,46,47,48,49,50,51for therapeutic neovascularization in the limbs. Unfortunately, the clinical trials that followed these successful animal studies did not show significant benefits for patients52.
One suggested explanation of the reason for this translational failure is that the condition of peripheral ischemia in human patients is one that includes resistance to angiogenic signals53,54,55,56,57,58,59. Several studies have shown defects in angiogenic signaling pathways in diabetes and hyperglycemia. Diabetes and hyperlipidemia lead to a loss of heparan sulfate proteoglycans and an increase in enzymes that cut heparan sulfate, presenting a potential mechanism for resistance to therapeutic angiogenesis/arteriogenesis with growth factors60,61. Thus, a key feature of a model for peripheral ischemia should include an aspect of therapeutic resistance so that therapies may be evaluated in the context of the disease state present in human patients.
In this work, we describe a rabbit model of peripheral ischemia through surgical ligation of the femoral arteries. A lead-in period with the induction of diabetes and hyperlipidemia is incorporated into the model. We compared this model to another model that incorporates a higher fat diet without diabetes and found that the model with diabetes and lower level of hyperlipidemia was more effective in reducing blood vessel growth. Our model combines advancements that have been used by separate groups, with the goal of providing a practical and standardized method to achieve consistent results in peripheral vascular disease research.

<table>
	<tbody>
		<tr>
			<td>0.9% Sodium Chloride</td>
			<td>Henry Schein Medical</td>
			<td>1537468 / 1531434</td>
			<td>250 mL bag / 1000 mL irrigation btl</td>
		</tr>
		<tr>
			<td>1 mL Syringe</td>
			<td>VWR</td>
			<td>BD309628</td>
			<td></td>
		</tr>
		<tr>
			<td>10 mL Syringe</td>
			<td>VWR</td>
			<td>BD309695</td>
			<td></td>
		</tr>
		<tr>
			<td>10% Formalin</td>
			<td>Fisher-Scientific</td>
			<td>23-245684</td>
			<td></td>
		</tr>
		<tr>
			<td>18G Needle</td>
			<td>VWR</td>
			<td>89219-294</td>
			<td></td>
		</tr>
		<tr>
			<td>20G Needle</td>
			<td>VWR</td>
			<td>89219-340</td>
			<td></td>
		</tr>
		<tr>
			<td>25G Needle</td>
			<td>VWR</td>
			<td>89219-290</td>
			<td></td>
		</tr>
		<tr>
			<td>27G Needle</td>
			<td>VWR</td>
			<td>89219-288</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL Syringe</td>
			<td>VWR</td>
			<td>BD309646</td>
			<td></td>
		</tr>
		<tr>
			<td>5% Dextrose</td>
			<td>Patterson Veterinary</td>
			<td>07-800-9689</td>
			<td></td>
		</tr>
		<tr>
			<td>Acepromazine</td>
			<td>Patterson Veterinary</td>
			<td>VEDC207</td>
			<td></td>
		</tr>
		<tr>
			<td>Alfaxalone</td>
			<td>Patterson Veterinary</td>
			<td>07-891-6051</td>
			<td></td>
		</tr>
		<tr>
			<td>Alginate</td>
			<td>Sigma-Aldrich</td>
			<td>PHR1471-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>Alloxan Monohydrate</td>
			<td>Sigma-Aldrich</td>
			<td>A7413</td>
			<td></td>
		</tr>
		<tr>
			<td>Angiography Equipment</td>
			<td>Toshiba</td>
			<td>Infinix-i</td>
			<td></td>
		</tr>
		<tr>
			<td>Angiography Injector</td>
			<td>Medrad</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-Mouse Ab Alexa 594</td>
			<td>Thermo Fisher Scientific</td>
			<td>A-11032</td>
			<td>Secondary Antibody for IHC</td>
		</tr>
		<tr>
			<td>Anti-Rabbit Ab Alexa 488</td>
			<td>Thermo Fisher Scientific</td>
			<td>A-11008</td>
			<td>Secondary Antibody for IHC</td>
		</tr>
		<tr>
			<td>a-SMA Antibody</td>
			<td>Abcam</td>
			<td>ab5694</td>
			<td>Primary Antibody for IHC</td>
		</tr>
		<tr>
			<td>Baytril</td>
			<td>Bayer Animal Health</td>
			<td>724089904201</td>
			<td>Enrofloxacin</td>
		</tr>
		<tr>
			<td>Blood Chemistry Panel</td>
			<td>IDEXX</td>
			<td>2616</td>
			<td>Rabbit Panel</td>
		</tr>
		<tr>
			<td>Blood Pressure Cuff</td>
			<td>WelchAllyn</td>
			<td></td>
			<td>Flexiport Disposable BP Cuff-infant size 7</td>
		</tr>
		<tr>
			<td>Blood Pressure Monitor</td>
			<td>Vmed Technology</td>
			<td></td>
			<td>Vmed Vet-Dop2</td>
		</tr>
		<tr>
			<td>Bupivacaine</td>
			<td>Henry Schein Medical</td>
			<td>6023287</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Patterson Veterinary</td>
			<td>42023017905</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine SR</td>
			<td>ZooPharm</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Sulfate</td>
			<td>CB Minerals</td>
			<td></td>
			<td>Food and Pharmaceutical Grade USP and FCC</td>
		</tr>
		<tr>
			<td>Chlorhexidine Scrub</td>
			<td>Patterson Veterinary</td>
			<td>07-888-4598</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloroform</td>
			<td>Fisher-Scientific</td>
			<td>C298-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Cholesterol</td>
			<td>Sigma-Aldrich</td>
			<td>C8503</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Thermo Fisher Scientific</td>
			<td>62248</td>
			<td></td>
		</tr>
		<tr>
			<td>Ear Vein Catheter</td>
			<td>Patterson Veterinary</td>
			<td>SR-OX165</td>
			<td>Surflo IV catheters</td>
		</tr>
		<tr>
			<td>Endotracheal tube</td>
			<td>Patterson Veterinary</td>
			<td></td>
			<td>Sheridan Brand, Depends on Rabbit Size</td>
		</tr>
		<tr>
			<td>Glucometer</td>
			<td>Amazon</td>
			<td>B001A67WH2</td>
			<td>Accu-Chek Aviva</td>
		</tr>
		<tr>
			<td>Glucometer Test Strips</td>
			<td>McKesson Medical-Surgical</td>
			<td>788222</td>
			<td>Accu-Chek Aviva Plus</td>
		</tr>
		<tr>
			<td>Guidewire</td>
			<td>Boston Scientific</td>
			<td>39122-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Hair Clippers</td>
			<td>Amazon</td>
			<td>B000CQZI3Q</td>
			<td>Oster #40 blade</td>
		</tr>
		<tr>
			<td>Heating Pad</td>
			<td>Cincinnati Subzero</td>
			<td>273</td>
			<td></td>
		</tr>
		<tr>
			<td>Heating Pad Pump</td>
			<td>Gaymar</td>
			<td></td>
			<td>Gaymar T/Pump</td>
		</tr>
		<tr>
			<td>Hemostat</td>
			<td>Fine Science Tools</td>
			<td>13009-12</td>
			<td>Curved Mosquito Hemostat</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Patterson Veterinary</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Insertion Tool</td>
			<td>Merit Medical Systems</td>
			<td>MAP550</td>
			<td>metal wire insertion tool</td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>HPB Pharmacy</td>
			<td></td>
			<td>Novalin R &#38; Novalin N</td>
		</tr>
		<tr>
			<td>Insulin Syringes</td>
			<td>McKesson Medical-Surgical</td>
			<td>942674</td>
			<td></td>
		</tr>
		<tr>
			<td>Introducer</td>
			<td>Cook Medical</td>
			<td>G28954</td>
			<td>3F Check Flo Performer Introducer</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Henry Schein Medical</td>
			<td>1100734</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Patterson Veterinary</td>
			<td>856440301</td>
			<td></td>
		</tr>
		<tr>
			<td>Lactated Ringers</td>
			<td>McKesson Medical-Surgical</td>
			<td>186662</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine</td>
			<td>McKesson Medical-Surgical</td>
			<td>239936</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine/Prilocaine cream</td>
			<td>McKesson Medical-Surgical</td>
			<td>761240</td>
			<td></td>
		</tr>
		<tr>
			<td>Ligaloop</td>
			<td>V. Mueller</td>
			<td>CH117 / CH116</td>
			<td>White Mini / Yellow Mini</td>
		</tr>
		<tr>
			<td>Mazola Corn Oil</td>
			<td>Amazon</td>
			<td>B0049IIVCI</td>
			<td></td>
		</tr>
		<tr>
			<td>Medrad Syringe</td>
			<td>McKesson Medical-Surgical</td>
			<td>346920</td>
			<td>150 mL</td>
		</tr>
		<tr>
			<td>Meloxicam</td>
			<td>Patterson Veterinary</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Metal ball sutures</td>
			<td>Ethicon-Johnson &#38; Johnson</td>
			<td>K891H</td>
			<td>4-0 silk C-1 30&#34;</td>
		</tr>
		<tr>
			<td>Metzenbaum Scissors</td>
			<td>Fine Science Tools</td>
			<td>14019-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Midazolam</td>
			<td>Henry Schein Medical</td>
			<td>1215470</td>
			<td></td>
		</tr>
		<tr>
			<td>Nitroglycerin</td>
			<td>McKesson Medical-Surgical</td>
			<td>927528</td>
			<td></td>
		</tr>
		<tr>
			<td>PECAM Antibody</td>
			<td>Novus Biologicals</td>
			<td>NB600-562</td>
			<td>Primary Antibody for IHC</td>
		</tr>
		<tr>
			<td>Perfusion Pump</td>
			<td>Masterflex</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pigtail Catheter</td>
			<td>Merit Medical Systems</td>
			<td>1310-21-0053</td>
			<td>3F pigtail</td>
		</tr>
		<tr>
			<td>Polydioxanone (PDS II) suture</td>
			<td>McKesson Medical-Surgical</td>
			<td>129271</td>
			<td>4-0 taper RB-1 (needle comes on suture)</td>
		</tr>
		<tr>
			<td>Polydioxanone (PDS II) suture</td>
			<td>McKesson Medical-Surgical</td>
			<td>129031</td>
			<td>4-0 reverse cutting FS-2</td>
		</tr>
		<tr>
			<td>Polyglactin 910 (Vicryl) suture</td>
			<td>Butler</td>
			<td>7233-41</td>
			<td>3-0 taper RB-1</td>
		</tr>
		<tr>
			<td>Polyglactin 910 (Vicryl) suture</td>
			<td>McKesson</td>
			<td>104373</td>
			<td>4-0 reverse cutting FS-2</td>
		</tr>
		<tr>
			<td>Rabbit Chow (Alfalfa)</td>
			<td>LabDiet</td>
			<td>5321</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit Restrainer</td>
			<td>VWR</td>
			<td>10718-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Rib Cutters</td>
			<td>V. Mueller</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Blade</td>
			<td>Fine Science Tools</td>
			<td>10015-00</td>
			<td>#15 blade</td>
		</tr>
		<tr>
			<td>Silk Sutures</td>
			<td>Ethicon-Johnson &#38; Johnson</td>
			<td>A183H</td>
			<td>4-0 silk ties 18&#34;</td>
		</tr>
		<tr>
			<td>Stainless Steel Ball</td>
			<td>McMaster-Carr</td>
			<td>1598K23</td>
			<td>3-mm diameter</td>
		</tr>
		<tr>
			<td>Surgical Drapes</td>
			<td>Gepco</td>
			<td>8204S</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Pump</td>
			<td>DRE Veterinary</td>
			<td></td>
			<td>Versaflow VF-300</td>
		</tr>
		<tr>
			<td>Visipaque contrast media</td>
			<td>McKesson Medical-Surgical</td>
			<td>509055</td>
			<td></td>
		</tr>
		<tr>
			<td>Weitlaner Retractor</td>
			<td>Fine Science Tools</td>
			<td>17012-13</td>
			<td></td>
		</tr>
	</tbody>
</table>

preclinical model of hind limb ischemia in diabetic rabbits

Peripheral vascular disease is a widespread clinical problem that affects millions of patients worldwide. A major consequence of peripheral vascular disease is the development of ischemia. In severe cases, patients can develop critical limb ischemia in which they experience constant pain and an increased risk of limb amputation. Current therapies for peripheral ischemia include bypass surgery or percutaneous interventions such as angioplasty with stenting or atherectomy to restore blood flow. However, these treatments often fail to the continued progression of vascular disease or restenosis or are contraindicated due to the overall poor health of the patient. A promising potential approach to treat peripheral ischemia involves the induction of therapeutic neovascularization to allow the patient to develop collateral vasculature. This newly formed network alleviates peripheral ischemia by restoring perfusion to the affected area. The most frequently employed preclinical model for peripheral ischemia utilizes the creation of hind limb ischemia in healthy rabbits through femoral artery ligation. In the past, however, there has been a strong disconnect between the success of preclinical studies and the failure of clinical trials regarding treatments for peripheral ischemia. Healthy animals typically have robust vascular regeneration in response to surgically induced ischemia, in contrast to the reduced vascularity and regeneration in patients with chronic peripheral ischemia. Here, we describe an optimized animal model for peripheral ischemia in rabbits that includes hyperlipidemia and diabetes. This model has reduced collateral formation and blood pressure recovery in comparison to a model with a higher cholesterol diet. Thus, the model may provide better correlation with human patients with compromised angiogenesis from the common co-morbidities that accompany peripheral vascular disease.

Following induction of diabetes and initiation of the 0.1% cholesterol diet, the total cholesterol for the rabbits with diabetes and cholesterol diet was 123.3 &#177; 35.1 mg/dL (n = 6 male rabbits) averaged overall time points and rabbits. The BGL level for these rabbits was 248.3 &#177; 50.4 mg/dL (n = 6 male rabbits). A time course for blood chemistries and leg blood pressure ratios in a typical rabbit is shown in Figure 3 in comparison to rabbits under a higher choles...

Watch this Scientific Journal Video about Preclinical Model of Hind Limb Ischemia in Diabetic Rabbits at JoVE.com

Preclinical Model of Hind Limb Ischemia in Diabetic Rabbits

We describe a surgical procedure used to induce peripheral ischemia in rabbits with hyperlipidemia and diabetes. This surgery acts as a preclinical model for conditions experienced in peripheral artery disease in patients. Angiography is also described as a means to measure the extent of introduced ischemia and recovery of perfusion.

Peripheral vascular disease is a widespread clinical problem that affects millions of patients worldwide. A major consequence of peripheral vascular ...

preclinical-model-of-hind-limb-ischemia-in-diabetic-rabbits

Research

JoVE Journal

Medicine

9.5K Views.  University of Texas at Austin. We describe a surgical procedure used to induce peripheral ischemia in rabbits with hyperlipidemia and diabetes. This surgery acts as a preclinical model for conditions experienced in peripheral artery disease in patients. Angiography is also described as a means to measure the extent of introduced ischemia and recovery of perfusion.

Video: Preclinical Model of Hind Limb Ischemia in Diabetic Rabbits

Gene Transfer for Ischemic Heart Failure in a Preclinical Model

Various emerging technologies are being developed for patients with heart failure. Well-established preclinical evaluations are necessary to determine their efficacy and safety.
Gene therapy using viral vectors is one of the most promising approaches for treating cardiac diseases. Viral delivery of various different genes by changing the carrier gene has immeasurable therapeutic potential.
In this video, the full process of an animal model of heart failure creation followed by gene transfer is presented using a swine model. First, myocardial infarction is created by occluding the proximal left anterior descending coronary artery. Heart remodeling results in chronic heart failure. Unique to our model is a fairly large scar which truly reflects patients with severe heart failure who require aggressive therapy for positive outcomes. After myocardial infarct creation and development of scar tissue, an intracoronary injection of virus is demonstrated with simultaneous nitroglycerine infusion. Our injection method provides simple and efficient gene transfer with enhanced gene expression. This combination of a myocardial infarct swine model with intracoronary virus delivery has proven to be a consistent and reproducible methodology, which helps not only to test the effect of individual gene, but also compare the efficacy of many genes as therapeutic candidates.

A method of gene transfer for the treatment of ischemic heart failure is described using a swine model of myocardial infarction. Our simple and reproducible method enables us to readily evaluate the efficacy of various gene transfers with a very simple and reproducible way.

Various emerging technologies are being developed for patients with heart failure. Well-established preclinical evaluations are necessary to determine ...

Longitudinal Evaluation of Mouse Hind Limb Bone Loss After Spinal Cord Injury using Novel, in vivo, Methodology

Spinal cord injury (SCI) is often accompanied by osteoporosis in the sublesional regions of the pelvis and lower extremities, leading to a higher frequency of fractures 1. As these fractures often occur in regions that have lost normal sensory function, the patient is at a greater risk of fracture-dependent pathologies, including death. SCI-dependent loss in both bone mineral density (BMD, grams/cm2) and bone mineral content (BMC, grams) has been attributed to mechanical disuse 2, aberrant neuronal signaling 3 and hormonal changes 4. The use of rodent models of SCI-induced osteoporosis can provide invaluable information regarding the mechanisms underlying the development of osteoporosis following SCI as well as a test environment for the generation of new therapies 5-7 (and reviewed in 8). Mouse models of SCI are of great interest as they permit a reductionist approach to mechanism-based assessment through the use of null and transgenic mice. While such models have provided important data, there is still a need for minimally-invasive, reliable, reproducible, and quantifiable methods in determining the extent of bone loss following SCI, particularly over time and within the same cohort of experimental animals, to improve diagnosis, treatment methods, and/or prevention of SCI-induced osteoporosis.
An ideal method for measuring bone density in rodents would allow multiple, sequential (over time) exposures to low-levels of X-ray radiation. This study describes the use of a new whole-animal scanner, the IVIS Lumina XR (Caliper Instruments) that can be used to provide low-energy (1-3 milligray (mGy)) high-resolution, high-magnification X-ray images of mouse hind limb bones over time following SCI. Significant bone density loss was seen in the tibiae of mice by 10 days post-spinal transection when compared to uninjured, age-matched control (na&iuml;ve) mice (13% decrease, p&lt;0.0005). Loss of bone density in the distal femur was also detectable by day 10 post-SCI, while a loss of density in the proximal femur was not detectable until 40 days post injury (7% decrease, p&lt;0.05). SCI-dependent loss of mouse femur density was confirmed post-mortem through the use of Dual-energy X-ray Absorptiometry (DXA), the current "gold standard" for bone density measurements. We detect a 12% loss of BMC in the femurs of mice at 40 days post-SCI using the IVIS Lumina XR. This compares favorably with a previously reported BMC loss of 13.5% by Picard and colleagues who used DXA analysis on mouse femurs post-mortem 30 days post-SCI 9. Our results suggest that the IVIS Lumina XR provides a novel, high-resolution/high-magnification method for performing long-term, longitudinal measurements of hind limb bone density in the mouse following SCI.

Longitudinal Evaluation of Mouse Hind Limb Bone Loss After Spinal Cord Injury using Novel, in vivo, Methodology

A longitudinal examination of bone loss in the femurs and tibiae of adult mice was performed following spinal cord injury using sequential low-dose X-ray scans. Tibia bone loss was detected throughout the study, while bone loss in the femur was not detected until 40 days post injury.

Spinal cord injury (SCI) is often accompanied by osteoporosis in the sublesional regions of the pelvis and lower extremities, leading to a higher ...

A Modified Heterotopic Swine Hind Limb Transplant Model for Translational Vascularized Composite Allotransplantation (VCA) Research

Vascularized Composite Allotransplantation (VCA) such as hand and face transplants represent a viable treatment option for complex musculoskeletal trauma and devastating tissue loss. Despite favorable and highly encouraging early and intermediate functional outcomes, rejection of the highly immunogenic skin component of a VCA and potential adverse effects of chronic multi-drug immunosuppression continue to hamper widespread clinical application of VCA. Therefore, research in this novel field needs to focus on translational studies related to unique immunologic features of VCA and to develop novel immunomodulatory strategies for immunomodulation and tolerance induction following VCA without the need for long term immunosuppression.
This article describes a reliable and reproducible translational large animal model of VCA that is comprised of an osteomyocutaneous flap in a MHC-defined swine heterotopic hind limb allotransplantation. Briefly, a well-vascularized skin paddle is identified in the anteromedial thigh region using near infrared laser angiography. The underlying muscles, knee joint, distal femur, and proximal tibia are harvested on a femoral vascular pedicle. This allograft can be considered both a VCA and a vascularized bone marrow transplant with its unique immune privileged features. The graft is transplanted to a subcutaneous abdominal pocket in the recipient animal with a skin component exteriorized to the dorsolateral region for immune monitoring.
Three surgical teams work simultaneously in a well-coordinated manner to reduce anesthesia and ischemia times, thereby improving efficiency of this model and reducing potential confounders in experimental protocols. This model serves as the groundwork for future therapeutic strategies aimed at reducing and potentially eliminating the need for chronic multi-drug immunosuppression in VCA.

A Modified Heterotopic Swine Hind Limb Transplant Model for Translational Vascularized Composite Allotransplantation VCA Research

Vascularized Composite Allotransplantations (VCA) have become a clinical reality. However, broad clinical application of VCA is limited by chronic multi-drug immunosuppression. The authors present a reliable and reproducible large animal model to translate novel immunomodulatory strategies that can minimize or potentially eliminate the need of immunosuppression in VCA.

Vascularized Composite Allotransplantation (VCA) such as hand and face transplants represent a viable treatment option for complex musculoskeletal trauma ...

A Preclinical Murine Model of Hepatic Metastases

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, murine pancreatic tumor model of hepatic metastases via a hemispleen injection of syngeneic murine pancreatic tumor cells is described. This model mimics many of the clinical conditions in patients with metastatic disease to the liver. Mice consistently develop metastases in the liver allowing for investigation of the metastatic process, experimental therapy testing, and tumor immunology research.

A preclinical, murine model of hepatic metastases performed via a hemispleen injection technique.

Numerous murine models have been developed to study human cancers and advance the understanding of cancer treatment and development. Here, a preclinical, ...

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 ...

Assessment of Spontaneous Alternation, Novel Object Recognition and Limb Clasping in Transgenic Mouse Models of Amyloid-&#946; and Tau Neuropathology

Here we describe a staged, behavioral testing approach that can be used to screen for compounds that exhibit in vivo efficacy on cognitive and functional motor behaviors in transgenic mouse models of &#946;-amyloidosis and tauopathy. The paradigm includes tests for spontaneous alternation in a Y-maze, novel object recognition, and limb clasping. These tests were selected because they: 1) interrogate function of cognitive or motor domains and the correlate neural circuitry relevant to the human disease state, 2) have clearly defined endpoints, 3) have easily implementable quality control checks, 4) can be run in a moderate throughput format, and 5) require little intervention by the investigator. These methods are designed for investigators looking to screen compounds for activity in short-term and working memory tasks, or functional motor behaviors associated with Alzheimer's disease mouse models. The methods described here use behavioral tests that engage a number of different brain regions including hippocampus and various cortical areas. Investigators that desire cognitive tests that specifically assess cognition mediated by a single brain region could use these techniques to supplement other behavioral tests.

Assessment of Spontaneous Alternation, Novel Object Recognition and Limb Clasping in Transgenic Mouse Models of Amyloid-&amp;#946; and Tau Neuropathology

Described here is a staged, behavioral screening approach that can be used to screen for compounds that exhibit in vivo efficacy on cognitive and functional motor behaviors in transgenic mouse models of &#946;-amyloidosis and tauopathy. These methods are optimized to screen compounds for activity in short-term and working memory tasks.

Here we describe a staged, behavioral testing approach that can be used to screen for compounds that exhibit in vivo efficacy on cognitive and functional ...

An Ex Vivo Tissue Culture Model for Fibrovascular Complications in Proliferative Diabetic Retinopathy

Diabetic retinopathy (DR) is the most common microvascular complication of diabetes and one of the leading causes of blindness in working-age adults. No current animal models of diabetes and oxygen-induced retinopathy develop the full-range progressive changes manifested in human proliferative diabetic retinopathy (PDR). Therefore, understanding of the disease pathogenesis and pathophysiology has relied largely on the use of histological sections and vitreous samples in approaches that only provide steady-state information on the involved pathogenic factors. Increasing evidence indicates that dynamic cell-cell and cell-extracellular matrix (ECM) interactions in the context of three-dimensional (3D) microenvironments are essential for the mechanistic and functional studies towards the development of new treatment strategies. Therefore, we hypothesized that the pathological fibrovascular tissue surgically excised from eyes with PDR could be utilized to reliably unravel the cellular and molecular mechanisms of this devastating disease and to test the potential for novel clinical interventions. Towards this end, we developed a novel method for 3D ex vivo culture of surgically-excised patient-derived fibrovascular tissue (FT), which will serve as a relevant model of human PDR pathophysiology. The FTs are dissected into explants and embedded in fibrin matrix for ex vivo culture and 3D characterization. Whole-mount immunofluorescence of the native FTs and end-point cultures allows thorough investigation of tissue composition and multicellular processes, highlighting the importance of 3D tissue-level characterization for uncovering relevant features of PDR pathophysiology. This model will allow the simultaneous assessment of molecular mechanisms, cellular/tissue processes and treatment responses in the complex context of dynamic biochemical and physical interactions within the PDR tissue architecture and microenvironment. Since this model recapitulates PDR pathophysiology, it will also be amenable for testing or developing new treatments.

Here, we present a protocol to study the pathophysiology of proliferative diabetic retinopathy by using patient-derived, surgically-excised, fibrovascular tissues for three-dimensional native tissue characterization and ex vivo culture. This ex vivo culture model is also amenable for testing or developing new treatments.

Diabetic retinopathy (DR) is the most common microvascular complication of diabetes and one of the leading causes of blindness in working-age adults. No ...

Establishment of a Severe Dry Eye Model Using Complete Dacryoadenectomy in Rabbits

Dry eye disease (DED) is a complex disease with multiple etiologies and variable symptoms, having ocular surface inflammation as its key pathophysiologic step. Despite advances in our understanding of DED, significant knowledge gaps remain. Advances are limited in part due to the lack of informative animal models. The authors recently reported on a method of DED induced by injecting all orbital lacrimal gland (LG) tissues with the lectin concanavalin A. Here, we report a novel model of aqueous-deficient DED based on the surgical resection of all orbital LG (dacryoadenectomy) tissues. Both methods use rabbits because of their similarity to human eyes in terms of the size and structure of the ocular surface. One week after removal of the nictitating membrane, the orbital superior LG was surgically removed under anesthesia, followed by removal of the palpebral superior LG, and finally removal of the inferior LG. Dacryoadenectomy induced severe DED, evidenced by a marked reduction in the tear break up time test and the Schirmer&#39;s tear test, and significantly increased tear osmolarity and rose bengal staining. Dacryoadenectomy-induced DED lasted at least eight weeks. There were no complications and animals tolerated the procedure well. The technique can be mastered relatively easily by those with adequate surgical experience and appreciation of the relevant rabbit anatomy. Since this model recapitulates the features of human aqueous-deficient DED, it is suitable for studies of ocular surface homeostasis, DED, and candidate therapeutics.

A novel approach is presented to induce chronic dry eye disease in rabbits by surgically removing all orbital lacrimal glands. This method, distinct from those previously reported, produces a stable, reproducible model of aqueous deficient dry eye well suited to study tear physiology and pathophysiology and ocular therapeutics.

Dry eye disease (DED) is a complex disease with multiple etiologies and variable symptoms, having ocular surface inflammation as its key pathophysiologic ...

Taking the Next Step: a Neural Coaptation Orthotopic Hind Limb Transplant Model to Maximize Functional Recovery in Rat

Limb transplant in particular and vascularized composite allotransplant (VCA) in general have wide therapeutic promise that have been stymied by current limitations in immunosuppression and functional neuromotor recovery. Many animal models have been developed for studying unique features of VCA, but here we present a robust reproducible model of orthotopic hind limb transplant in rats designed to simultaneously investigate both aspects of current VCA limitation: immunosuppression strategies and functional neuromotor recovery. At the core of the model rests a commitment to meticulous, time-tested microsurgical techniques such as hand sewn vascular anastomoses and hand sewn neural coaptation of the femoral nerve and the sciatic nerve. This approach yields durable limb reconstructions that allow for longer lived animals capable of rehabilitation, resumption of daily activities, and functional testing. With short-term treatment of conventional immunosuppressive agents, allotransplanted animals survived up to 70 days post-transplant, and isotransplanted animals provide long lived controls beyond 200 days post-operatively. Evidence of neurologic functional recovery is present by 30 days post operatively. This model not only provides a useful platform for interrogating immunological questions unique to VCA and nerve regeneration, but also allows for in vivo testing of new therapeutic strategies specifically tailored for VCA.

This protocol presents a robust, reproducible model of vascularized composite allotransplant (VCA) geared toward simultaneous study of immunology and functional recovery. The time invested in meticulous technique in a right mid-thigh hind limb orthotopic transplant with hand sewn vascular anastomoses and neural coaptation yields the ability to study functional recovery.

Limb transplant in particular and vascularized composite allotransplant (VCA) in general have wide therapeutic promise that have been stymied by current ...

Predicting Amputation using Local Circulating Mononuclear Progenitor Cells in Angioplasty-treated Patients with Critical Limb Ischemia

Critical limb ischemia (CLI) represents an advanced stage of the peripheral arterial disease. Angioplasty improves the blood flow to the lower limb; however, some patients irreversibly progress to limb amputation. The extent of vascular damage and the mechanisms of vascular repair are factors affecting post-angioplasty outcome. Mononuclear Progenitor Cells (MPCs) are reactive to vascular damage and repair, with the ability to reflect vascular diseases. The present protocol describes quantification of MPCs obtained from blood circulation from vessel close to the angioplasty site, as well as its relationship with endothelial dysfunction and its predictive ability for limb amputation in the next 30 days after angioplasty in patients with CLI.

Lower limb amputation may occur even after angioplasty of obstructed vessels in Critical Limb Ischemia (CLI). Mononuclear Progenitor Cells (MPCs) reflect vascular repair. The present protocol describes the quantification of MPCs from circulation close to angioplasty, and its relationship with endothelial dysfunction and prediction of lower limb amputation.

Critical limb ischemia (CLI) represents an advanced stage of the peripheral arterial disease. Angioplasty improves the blood flow to the lower limb; ...