We thank Jos&#233; Rino and T&#226;nia Carvalho, heads of the Bioimaging Facility and Histology and Comparative Pathology Laboratory of Instituto de Medicina Molecular Jo&#227;o Lobo Antunes, respectively. We also thank Vyacheslav Sushchyk from the Department of Anatomy of Nova Medical School/Faculdade de Ci&#234;ncias M&#233;dicas, Universidade Nova de Lisboa.
Funding reference: project funded by UID/IC/0306/2016 Funda&#231;&#227;o para a Ci&#234;ncia e a Tecnologia. Paula de Oliveira is supported by a fellowship (SFRH/BD/80483/2011) from Funda&#231;&#227;o para a Ci&#234;ncia e Tecnologia.

All animal procedures are in accordance with Directive 2010/63/EU and have been approved by the Institutional Animal Welfare Body and licensed by DGAV, the Portuguese competent authority for animal protection (license number 023861/2013)
CAUTION: Several of the chemicals used in the protocols are toxic and harmful. Please use all appropriate safety practices and personal protective equipment (gloves, lab coat, full-length pants, and closed-toe shoes)
1. The murine model of hindlimb ischemia
NOTE: All experiments are performed on 22-week-old C57BL/6 female mice.
<ol>
	<li>Preparation of solutions
	<ol>
		<li>Preparation of the anesthetic solution 
		Note: This anesthetic combination (medetomidine and ketamine) provides up to 30 min immobilization and a surgical window of 15 min. The effect may be reverted by the administration of atipamezole.
		<ol>
			<li>With a clean, 1cc syringe, withdraw 1 mL of medetomidine (1 mg/Kg body weight), eliminate air bubbles for accurate measurements, and add it to a 15 mL falcon tube.</li>
			<li>With a clean, 1 cc syringe, withdraw 0.75 mL of Ketamine (75 mg/Kg body weight), eliminate air bubbles for accurate measurements and add it to a 15 mL falcon tube containing the medetomidine.</li>
			<li>Add 8.25 mL of sterile saline (0.9% NaCl), in order to obtain 10 mL of anesthetic solution.</li>
			<li>Identify the tube with the name of the compounds, the date mixed and the expiration date.</li>
			<li>Store in the dark at 4 &#176;C.</li>
		</ol>
		</li>
		<li>Preparation of the anti-sedative solution
		<ol>
			<li>With a clean, 1 cc syringe, withdraw 1 mL of atipamezole (5 mg/Kg body weight), eliminate air bubbles for accurate measurements, and add it to a 15 mL falcon tube.</li>
			<li>Add 9 mL of sterile saline (0.9% NaCl), in order to obtain 10 mL of reversion solution.</li>
			<li>Identify the tube with the name of the compounds, the date mixed and the expiration date.</li>
			<li>Store in the dark at 4 &#176;C.</li>
		</ol>
		</li>
		<li>Preparation of the post-operative analgesia solution
		<ol>
			<li>With a clean, 1 cc syringe, withdraw 0.5 mL of buprenorphine (100 &#181;L/15-30 g body weight), eliminate air bubbles for accurate measurements, and add it to a 15 mL falcon tube.</li>
			<li>Add 9.5 mL of sterile saline (0.9% NaCl), in order to obtain 10 mL of analgesia solution.</li>
			<li>Identify the tube with the name of the compounds, the date mixed and the expiration date.</li>
			<li>Store in the dark at 4 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Surgical induction of hindlimb ischemia
	<ol>
		<li>Surgical tools needed for this intervention: pointed forceps, spring scissors, ophthalmic needle holder, surgical scissors and a needle holder. Sterilize all surgical tools in an autoclave before use. Use cotton swabs for the dissection of subcutaneous fat.</li>
		<li>Load the syringe with the anesthesia solution before holding the mouse.</li>
		<li>Restrain the animal by applying force behind its ears against the cage&#8217;s grid, holding as much skin as possible to keep the mouse immobile.</li>
		<li>Keep the animal tilted with its head lowered and perform an intraperitoneal injection of the anesthesia solution, keeping the syringe at a 45&#176; angle with the mouse&#8217;s body.</li>
		<li>Check for the absence of pedal withdrawal reflexes, to evaluate the depth of anesthesia.</li>
		<li>Remove the hair from the right hindlimb using an electric shaver.&#160;Use a chlorhexidine disinfectant soap scrub for skin preparation. Use a clean paper towel to remove the suds.&#160;Apply a chlorhexidine surgical scrub and protect the skin area with a sterile drape before transporting the mice to the surgical sterile suite.</li>
		<li>Place the animal in dorsal decubitus and restrain it with the four limbs spread apart and secured with tape. 
		CAUTION: Perform the procedure on a heated pad to keep the animal&#8217;s body temperature stable; the surgical procedure is performed using a dissection microscope at 10x or 20x magnification.</li>
		<li>Apply a chlorhexidine antiseptic solution using sterile gauze to finalize skin preparation.&#160;Using a number 11 surgical scalpel blade perform a 1-cm skin incision overlying the thigh of the right hindlimb.</li>
		<li>Blunt dissect subcutaneous fat exposing the neurovascular bundle. Using the pointed forceps, the spring scissors and the ophthalmic needle holder, open the membranous ilio-femoral sheath to expose the distal external iliac and femoral vessels.</li>
		<li>Dissect and separate the vessels (artery and vein) from the nerve. Using a 7.0 non-absorbable polypropylene suture ligate the distal external iliac artery and vein and the distal femoral artery and vein.</li>
		<li>Transect the segment of the ilio-femoral artery and veins between the distal and proximal knots with the spring scissors.</li>
		<li>Close the incision with an absorbable suture (e.g., 5.0 Vicryl suture).</li>
		<li>Load the syringe with the anti-sedative solution and use the same method detailed in 1.2.3 and 1.2.4 to perform an intraperitoneal injection of the anti-sedative solution.</li>
	</ol>
	</li>
	<li>Post-operative monitoring 
	NOTE: Animals are carefully observed every 15-20 min to make sure all animals recover well from the anti-sedative; anesthetized animals are never left unattended.
	<ol>
		<li>Assess the recovery from anesthesia: 1) monitor the rate and depth of respiration; 2) assess animal reflexes (pedal and eye position)</li>
		<li>Perform a subcutaneous injection of the post-operative analgesia every 8-12 h.</li>
		<li>Monitor the animals daily after the surgery recording a brief description of the animal&#8217;s health status and surgery site appearance, ensure all sutures are closed.</li>
		<li>Re-suture the incision, if dehiscence occurs. If unsuccessful, apply a skin barrier film on the incision to protect from urine, fecal excretions, friction and shear of the skin and to help the healing process.</li>
	</ol>
	</li>
</ol>
2. Assessment of the angiogenic effect
NOTE: After ischemia induction, apply the therapeutic agent in the study and achieve the following procedures. Steps undertaken at other time points are detailed under the corresponding section.
<ol>
	<li>Laser Doppler perfusion imaging
	<ol>
		<li>Remove the hair from both hindlimbs one day before the analysis using an electrical shaver followed by depilatory cream.</li>
		<li>Anesthetize the mice using the anesthetic solution.</li>
		<li>Place the animal on a 37 &#176;C heating pad for 5 min to ensure that temperature will not change during the procedure</li>
		<li>Place the animal in the supine position with legs secured in an extended position. Measure the distance between right and left legs and feet for each animal since the positioning of the animal should be similar whenever the procedure must be repeated in order to follow changes in hindlimb perfusion over time.</li>
		<li>Start acquiring data by using a laser Doppler perfusion imager.</li>
		<li>After the acquisition is complete, revert the anesthesia with the anti-sedative solution.</li>
		<li>Open the image analysis software and draw the region of interest (ROI) around the ischemic hindlimb and a comparable ROI around the non-ischemic hindlimb.</li>
		<li>Observe the flux values and determine the perfusion as a ratio of the perfusion in the ischemic limb to that in the non-ischemic limb. Changes in the perfusion ratio are measured over time.&#160;</li>
	</ol>
	</li>
	<li>Immunohistochemistry and capillary density analysis
	<ol>
		<li>After anesthetizing, sacrifice the mice by cervical dislocation.</li>
		<li>To harvest the gastrocnemius muscles of both legs, first, remove the skin from both hindlimbs.</li>
		<li>Identify the Achilles tendon and separate the tendon from the bone using an 11-blade scalpel.</li>
		<li>Hold the Achilles tendon and separate the muscle from the tibia and fibula up to the knee joint with spring scissors.</li>
		<li>Separate the biceps femoris from the gastrocnemius muscle.</li>
		<li>Use spring scissors to cut the insertions of the gastrocnemius muscles at the knee joint level.&#160;</li>
		<li>Place the harvested gastrocnemius muscles in a transverse orientation on a small cork disk with the help of 10% tragacanth.</li>
		<li>Snap freeze the specimens in liquid nitrogen-cooled isopentane and store them at -80 &#176;C until sectioning.</li>
		<li>Cut 7 &#181;m sections of the muscle specimens on a cryostat and mount on glass slides. 
		NOTE: The protocol can be paused here.</li>
		<li>Fix the glass slides in a bottle containing cooled pure acetone (approximately 50 mL) for 10 min, at -20 &#176;C.</li>
		<li>Add hydrogen peroxidase 0.3% diluted in methanol (50 mL) for 30 min at room temperature.</li>
		<li>Wash twice in phosphate-buffered saline (PBS) (50 mL) for a total of 10 min.</li>
		<li>Use a hydrophobic pen around the sections. 
		NOTE: The use of a hydrophobic pen allows reduce the amount of solutions spent during the protocol. For all incubations use 100 &#181;L of all solutions, per section.</li>
		<li>Apply a blocking solution of 5% rabbit serum in PBS for 30 min at room temperature.</li>
		<li>Incubate the specimens for 1 h at room temperature with anti-CD31 rat monoclonal antibody diluted at 1:500 in 1 % bovine serum albumin (BSA) in PBS.</li>
		<li>Wash three times in PBS for a total of 30 min.</li>
		<li>Add a secondary biotinylated rabbit anti-rat IgG antibody at 1:200 in 1% BSA in PBS and 5% rabbit serum for 30 min at room temperature.</li>
		<li>Wash in PBS as before and add labeled avidin-conjugated peroxidase complex for 30 min at room temperature.</li>
		<li>Rinse 3 times in PBS for 5 min and add diaminobenzidine (DAB) peroxidase substrate kit for another 5 min.</li>
		<li>Counterstain the sections with hematoxylin for 10 s before mounting.</li>
		<li>Measure the capillary densities by using an upright brightfield microscope (i.e., number of capillaries per number of myocytes) in 2 different sections of 4 distinct anatomic areas in each specimen.</li>
	</ol>
	</li>
	<li>Contrast agent perfusion 
	<ol>
		<li>Anesthetize the mice&#160;using&#160;the anesthetic solution.</li>
		<li>Perform a median laparotomy using a number 15 scalpel blade.</li>
		<li>Expose the abdominal aorta and the caudal vena cava, after retracting laterally the small bowel and using spring scissors.</li>
		<li>Insert one 26-gauge catheter cranially into the abdominal aorta and another into the caudal vena cava in the same direction.</li>
		<li>Secure catheters in place using a 5/0 silk suture to perform a stay suture.</li>
		<li>Wash the vascular system with a 37 &#176;C saline solution containing heparin (500 units/100 mL) pushed into the abdominal aorta using a 10 mL syringe. Continue gentle perfusion until a clear solution is seen coming out of the catheter placed inside the caudal vena cava. 
		NOTE: Usually 30 to 40 mL of a heparinized saline solution are required to rinse the vascular system of an adult rat.</li>
		<li>Administer a mixture of adenosine (1 mg/L) and papaverine (4 mg/L) for 2 min.</li>
		<li>Inject into the aortic catheter 10 mL of a mixture of 50% barium sulfate and 5% gelatine. 
		NOTE: The solution must be kept at 60 &#176;C to avoid thickening.</li>
		<li>Leave the mice at -&#160;4 &#176;C for at least 4 h, in order to allow the vascular cast to solidify.</li>
		<li>Remove the skin from the lower body of each mouse.</li>
	</ol>
	</li>
	<li>Diaphonization - Spalteholz technique
	<ol>
		<li>Fix the mice by immersion in a 10% formaldehyde solution for at least 72 h.</li>
		<li>Bleach the mice by immersion in a 30% hydrogen peroxide solution for 24 h.</li>
		<li>Dehydrate the mice using the freeze substitution technique. This methodology consists of submitting the mice to successive passages (on average four) of pure acetone at -&#160;20 &#176;C for 24 h each.</li>
		<li>Clarify the mice by immersing it in a solution containing a mixture of 100% benzyl benzoate and 100% methyl salicylate in a 3:5 proportion (this mixture is also known as Spalteholz solution) 13.</li>
		<li>Leave the mice immersed in the Spalteholz solution inside in a vacuum chamber until it becomes diaphanous, which usually takes 5 to 7 days.</li>
		<li>Replace the solution, if it becomes dark before the specimen becomes clear.</li>
	</ol>
	</li>
	<li>Collateral vessels quantification
	<ol>
		<li>Observe the mice suspended in the Spalteholz solution under transillumination using a stereotaxic microscope. Use microsurgery forceps to gently grasp and move it.</li>
		<li>Photograph the entire limbs using a digital camera connected to the microscope.</li>
		<li>Obtain entire limb photographs by using the appropriate software.</li>
		<li>Perform a manual segmentation of the collateral vessels by highlighting them using the appropriate software. 
		NOTE: Collateral vessels are defined according to Longland&#8217;s definition, a defined stem, mid-zone, and re-entrant, with a diameter between 20 and 300 &#181;m, excluding femoral, saphenous and popliteal arteries and all venous structures.</li>
		<li>Define ROIs in every mouse in the same anatomic region in the adductor, surrounding and below the surgical occlusion.</li>
		<li>Quantify the collateral vessel density (CVD) in equivalent ROIs corresponding to 20% of the total limb area. The CVD is calculated as the ratio between the vascular area and the ROIs areas. All density measurements are performed using ImageJ software. 
		NOTE: The CVD value of the non-ischemic limb for each mouse is assumed to correspond to 100% to exclude variations in the anatomy, perfusion or diaphonization procedures. The CVD percentage in the ischemic limb was, therefore, calculated relative to the non-ischemic one.</li>
		<li>Determine the percentage of CVD increase as the difference between the CVD percentage among the ischemic and nonischemic limbs.</li>
	</ol>
	</li>
	<li>Laser capture microdissection of capillaries
	<ol>
		<li>After anesthetizing, sacrifice the mice by cervical dislocation.</li>
		<li>To harvest the gastrocnemius muscles of both legs, remove the skin from mice both hindlimbs.</li>
		<li>Identify the Achilles tendon and separate the tendon from the bone using an 11-blade scalpel.</li>
		<li>Hold the Achilles tendon and separate the muscle from the tibia and fibula up to the knee joint with spring scissors.</li>
		<li>Separate the biceps femoris from the gastrocnemius muscle.</li>
		<li>Use spring scissors to cut the insertions of the gastrocnemius muscles at the knee joint level.&#160;</li>
		<li>Place the harvested gastrocnemius muscles in a transverse orientation on a small cork disk with the help of 10 % tragacanth.</li>
		<li>Snap freeze the specimens in liquid nitrogen-cooled isopentane and store them at -80 &#176;C until sectioning.</li>
		<li>Cut on a cryostat 12 &#181;m sections of the muscle specimens and mount on glass slides. 
		NOTE: For RNA protection take a precooled slide and touch the back-side of the slide with your finger (gloves) to warm only the region for placing the section. Transfer section from the cryostat knife by touching with the warmed area and dry at -20 &#176;C in the cryostat for 2-3 min. The protocol can be paused here.
		<ol>
			<li>Fix the glass slides in a bottle containing cooled pure acetone (approximately 50 mL) for 5 min at -20 &#176;C and air-dry them.</li>
			<li>Use a hydrophobic pen around the sections. 
			NOTE: The use of the hydrophobic pen helps in reduction of the amount of solutions spent during the protocol. For all incubations use 100 &#181;L of all solutions, per section.</li>
			<li>Rehydrate with 2 M NaCl/PBS at 4 &#176;C and incubate the specimens overnight at 4 &#176;C with anti-CD31 rat monoclonal antibody at 1:500 in 2M NaCl/PBS.</li>
			<li>Wash twice in 2M NaCl/PBS for a total of 6 min.</li>
			<li>Add a secondary biotinylated rabbit anti-rat IgG antibody at 1:200 in 2M NaCl/PBS and 5% rabbit serum for 30 min at 4 &#176;C.</li>
			<li>Wash in 2 M NaCl/PBS as before and add labeled avidin-conjugated peroxidase complex for 30 min at 4 &#176;C.</li>
			<li>Rinse and add DAB peroxidase substrate kit for 5 min.</li>
			<li>Dehydrate the sections in ice-cold 90 % ethanol followed by 100 % ethanol and leave them to dry.</li>
			<li>Dissect 10 000 capillaries per mice using a laser microdissection system equipped with a pulsed solid-state 355 nm laser and catapult the dissected capillaries into a microfuge tube adhesive-cap.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>RNA extraction, cDNA synthesis, pre-amplification, and RT-PCR
	<ol>
		<li>Isolate total RNA from the microdissected capillaries using an appropriate kit, the total RNA obtained is between 1.5-3 ng/&#181;L.</li>
		<li>Use a cDNA synthesis kit for synthesis and two rounds of pre-amplification, using the primers described in Table 1.</li>
		<li>Perform RT-PCR for the same targets described in the previous step. Use a program consisting of an initial denaturation step at 95 &#176;C for 10 min, followed by 50 cycles at 95 &#176;C for 15 s and at 60 &#176;C for 1 min.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Becker, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+I:+Definitions,+epidemiology,+clinical+presentation+and+prognosis.">Chapter I: Definitions, epidemiology, clinical presentation and prognosis.</a> European Journal of Vascular and Endovascular Surgery. 42 Suppl 2, S4-S12 (2011).</li><li>Fowkes, F. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+artery+disease:+epidemiology+and+global+perspectives.">Peripheral artery disease: epidemiology and global perspectives.</a> Nature Reviews Cardiology. 14 (3), 156-170 (2017).</li><li>Abu Dabrh, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+natural+history+of+untreated+severe+or+critical+limb+ischemia.">The natural history of untreated severe or critical limb ischemia.</a> Journal of Vascular Surgery. 62 (6), 1642-1651 (2015).</li><li>Lejay, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+murine+model+of+sustainable+and+durable+chronic+critical+limb+ischemia+fairly+mimicking+human+pathology.">A new murine model of sustainable and durable chronic critical limb ischemia fairly mimicking human pathology.</a> European Journal of Vascular and Endovascular Surgery. 49 (2), 205-212 (2015).</li><li>Sprengers, R. W., Lips, D. J., Moll, F. L., Verhaar, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progenitor+cell+therapy+in+patients+with+critical+limb+ischemia+without+surgical+options.">Progenitor cell therapy in patients with critical limb ischemia without surgical options.</a> Annals of Surgery. 247 (3), 411-420 (2008).</li><li>Lotfi, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Towards+a+more+relevant+hind+limb+model+of+muscle+ischaemia.">Towards a more relevant hind limb model of muscle ischaemia.</a> Atherosclerosis. 227 (1), 1-8 (2013).</li><li>Masaki, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Angiogenic+gene+therapy+for+experimental+critical+limb+ischemia:+acceleration+of+limb+loss+by+overexpression+of+vascular+endothelial+growth+factor+165+but+not+of+fibroblast+growth+factor-2.">Angiogenic gene therapy for experimental critical limb ischemia: acceleration of limb loss by overexpression of vascular endothelial growth factor 165 but not of fibroblast growth factor-2.</a> Circulation Research. 90 (9), 966-973 (2002).</li><li>Limbourg, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+postnatal+arteriogenesis+and+angiogenesis+in+a+mouse+model+of+hind-limb+ischemia.">Evaluation of postnatal arteriogenesis and angiogenesis in a mouse model of hind-limb ischemia.</a> Nature Protocols. 4 (12), 1737-1746 (2009).</li><li>Hellingman, A. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variations+in+surgical+procedures+for+hind+limb+ischaemia+mouse+models+result+in+differences+in+collateral+formation.">Variations in surgical procedures for hind limb ischaemia mouse models result in differences in collateral formation.</a> European Journal of Vascular and Endovascular Surgery. 40 (6), 796-803 (2010).</li><li>Brevetti, L. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exercise-induced+hyperemia+unmasks+regional+blood+flow+deficit+in+experimental+hindlimb+ischemia.">Exercise-induced hyperemia unmasks regional blood flow deficit in experimental hindlimb ischemia.</a> Journal of Surgical Research. 98 (1), 21-26 (2001).</li><li>Ministro, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Low-dose+ionizing+radiation+induces+therapeutic+neovascularization+in+a+pre-clinical+model+of+hindlimb+ischemia.">Low-dose ionizing radiation induces therapeutic neovascularization in a pre-clinical model of hindlimb ischemia.</a> Cardiovascular Research. 113 (7), 783-794 (2017).</li><li>Pereira, A. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Therapeutic+angiogenesis+induced+by+human+umbilical+cord+tissue-derived+mesenchymal+stromal+cells+in+a+murine+model+of+hindlimb+ischemia.">Therapeutic angiogenesis induced by human umbilical cord tissue-derived mesenchymal stromal cells in a murine model of hindlimb ischemia.</a> Stem Cell Research Therapy. 7 (1), 145 (2016).</li><li>Azaripour, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+survey+of+clearing+techniques+for+3D+imaging+of+tissues+with+special+reference+to+connective+tissue.">A survey of clearing techniques for 3D imaging of tissues with special reference to connective tissue.</a> Progress in Histochemistry and Cytochemistry. 51 (2), 9-23 (2016).</li></ol>

The authors have nothing to disclose.

Murine models of CLI have mostly consisted in ligation of the femoral artery just distal to the origin of the profunda femoris 4,5,6,7,8,9. This has shown to leave most of the collateral circulation intact, which restores blood flow to the limb within 7 days 9. Removal of the collateral bed can be achi...

Peripheral arterial disease (PAD) affects predominantly the lower limbs. PAD is caused by atherosclerosis, an artery obstruction that can cause severe restriction to the blood flow in the lower limbs1. Intermittent claudication is the first manifestation of PAD and refers to muscle pain when walking. CLI is the most severe stage of PAD, being diagnosed in patients that show ischemic rest pain, ulcers or gangrene2. Patients with CLI have a high risk of amputation, especially if untreated3. Lower limb revascularization (either by open surgery or an endovascular procedure) is currently the only way to achieve limb salvage. However, around 30% of CLI patients are not suited for these procedures, for reasons that include the location of the lesions, the pattern of arterial occlusion and extensive comorbidity4,5. Therefore, new therapies are needed for these otherwise untreatable patients, with the promotion of angiogenesis being the strategy under more intense investigation.
Before testing in humans, the effectiveness&#160;and safety of new therapies&#160;in vivo&#160;must be considered in animal models. Several models have been developed for the study of CLI, mostly by inducing hindlimb ischemia (HLI) in mice6,7,8,9,10.&#160;However, these models differ in several aspects including the nature of the&#160;arteries that are ligated and/or excised&#160;and whether&#160;the veins and nerves surrounding are dissected as well6,7,8,9,10. Taken together, these aspects will affect the severity of the ischemia-reperfusion injury in each animal,&#160;making the results difficult to be compared. Therefore, it is critical to develop an effective protocol in which the procedure to induce ischemia and the evaluation of different targets should be standardized to assess whether a given angiogenic therapy will be effective. An experimental protocol designed to cover all these aspects would provide a comprehensive understanding of the mechanisms by which angiogenic therapies exert their effects and a measure of their efficacy at each of their outcomes. Two distinct works recently published by our team are a good example11,12, in which different approaches to induce therapeutic angiogenesis were assessed using the same protocol that will be described&#160;with more detail&#160;in this protocol.
The overall goal of this protocol is to describe a reproducible experimental model that can mimic the effects of CLI and lay the experimental foundation for a comprehensive assessment of the functional, histologic and molecular effects of putative angiogenic agents.

<table border="0" cellpadding="0" cellspacing="0" style="width:840px;" width="839">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="38">
			<td height="38" style="height:39px;width:251px;">7500 Fast Real-Time PCR</td>
			<td style="width:235px;">Applied Biosystems</td>
			<td style="width:132px;"></td>
			<td style="width:223px;">Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Acetone</td>
			<td>Merk</td>
			<td>1000141000</td>
			<td>Reagent; Caution - highly flammable</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Adenosine</td>
			<td>Valdepharm</td>
			<td></td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Atipamezole</td>
			<td>OrionPharma</td>
			<td></td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Barium sulphate (Micropaque)</td>
			<td>Guebert</td>
			<td>8671404 (ref. Infarmed)</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Buprenorphine</td>
			<td>RichterPharma</td>
			<td></td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Carl Zeiss Opmi-1 FC Surgical Microscope</td>
			<td>Carl Zeiss Microscopy, Germany</td>
			<td></td>
			<td>Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">cDNA RT2 PreAMP cDNA Synthesis kit</td>
			<td>Qiagen</td>
			<td style="width:132px;">7335730</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Cryostat Leica CM</td>
			<td>Leica Microsystems</td>
			<td>3050S</td>
			<td>Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">DAB peroxidase substrate kit</td>
			<td>DAKO;Vector Laboratories</td>
			<td>K3468</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">hydrogen peroxidase</td>
			<td>Merk</td>
			<td>1072090250</td>
			<td>Reagent; Caution - nocif</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">hydrophobic pen</td>
			<td>Dako</td>
			<td>411121</td>
			<td>Reagent; Caution - toxic</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Ketamidor</td>
			<td style="width:235px;">Richterpharma</td>
			<td>CN:580393,7 630/01/12 Dfvf</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Laser Doppler perfusion imager moorLDI2-HIR</td>
			<td style="width:235px;">MoorLDI-V6.0, Moor Instruments Ltd, Axminster, UK</td>
			<td>5710</td>
			<td>Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Leica DM2500 upright brightfield microscope</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td>Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Medetor</td>
			<td>Virbac</td>
			<td>037/01/07RFVPT</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">methanol</td>
			<td>VWR</td>
			<td>UN1230</td>
			<td>Reagent; Caution - toxic and highly flammable</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Papaverine</td>
			<td>Labesfal</td>
			<td></td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Pentano Isso</td>
			<td>Merk</td>
			<td>1060561000</td>
			<td>Reagent; Caution - highly flammable</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Power SYBR&#174; Green</td>
			<td>Applied Biosystems</td>
			<td style="width:132px;">4309155</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Purified rat anti-mouse CD31</td>
			<td>Pharmingen</td>
			<td>550274</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">RNeasy Micro kit</td>
			<td>Qiagen</td>
			<td>74004</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Surgic-Pro 6.0</td>
			<td>Medtronic (Coviden)</td>
			<td>VP733X</td>
			<td>Suture</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;width:251px;">VECTASTAIN ABC HRP Kit (Peroxidase, Rat IgG)</td>
			<td>Vectastain ABC kit; Vector Laboratories</td>
			<td>PK-4004</td>
			<td>Reagent</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Vicryl5.0/ Vicryl 6.0</td>
			<td>Medtronic (Covidien)</td>
			<td>UL202/ UL101</td>
			<td>Suture</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Zeiss PALM MicroBeam Laser Microdissection System</td>
			<td>Carl Zeiss Microscopy, Germany</td>
			<td>1023290916</td>
			<td>Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Stereotaxic microscope</td>
			<td>Carl Zeiss Microscopy, Germany</td>
			<td></td>
			<td>Instrument</td>
		</tr>
		<tr height="38">
			<td height="38" style="height:39px;">Digital camera</td>
			<td>Linux</td>
			<td></td>
			<td>Instrument</td>
		</tr>
	</tbody>
</table>

assessing therapeutic angiogenesis in a murine model of hindlimb ischemia

Critical limb ischemia (CLI) is a serious condition that entails a high risk of lower limb amputation. Despite revascularization being the gold-standard therapy, a considerable number of CLI patients are not suited for either surgical or endovascular revascularization. Angiogenic therapies are emerging as an option for these patients but are currently still under investigation. Before application in humans, those therapies must be tested in animal models and its mechanisms must be clearly understood. An animal model of hindlimb ischemia (HLI) has been developed by the ligation and excision of the distal external iliac and femoral arteries and veins in mice. A comprehensive panel of tests was assembled to assess the effects of ischemia and putative angiogenic therapies at functional, histologic and molecular levels. Laser Doppler was used for the flow measurement and functional assessment of perfusion. Tissue response was evaluated by the analysis of capillary density after staining with the anti-CD31 antibody on histological sections of gastrocnemius muscle and by measurement of collateral vessel density after diaphonization. Expression of angiogenic genes was quantified by RT-PCR targeting selected angiogenic factors exclusively in endothelial cells (ECs) after laser capture microdissection from mice gastrocnemius muscles. These methods were sensitive in identifying differences between ischemic and non-ischemic limbs and between treated and non-treated limbs. This protocol provides a reproducible model of CLI and a framework for testing angiogenic therapies.

Using the described protocol, umbilical cord mesenchymal stem cells and low-dose ionizing radiation (LDIR) were tested as putative angiogenic therapies 11,12. Laser Doppler perfusion readings were obtained before ischemia induction and at pre-specified timepoints ranging from immediately after ischemia induction to 45 days post-ischemia. Tissue perfusion readings by laser Doppler were recorded as color-coded images, with no perfusion displayed as dark blue and th...

Watch this Scientific Journal Video about Assessing Therapeutic Angiogenesis in a Murine Model of Hindlimb Ischemia at JoVE.com

Assessing Therapeutic Angiogenesis in a Murine Model of Hindlimb Ischemia

Here, a critical hindlimb ischemia experimental model is presented followed by a battery of functional, histologic and molecular tests&#160;to assess the effectiveness of angiogenic therapies.&#8203;

Critical limb ischemia (CLI) is a serious condition that entails a high risk of lower limb amputation. Despite revascularization being the gold-standard ...

assessing-therapeutic-angiogenesis-murine-model-hindlimb

Research

JoVE Journal

Medicine

8.6K Views.  Lisbon School of Medicine of the Universidade de Lisboa. Here, a critical hindlimb ischemia experimental model is presented followed by a battery of functional, histologic and molecular tests to assess the effectiveness of angiogenic therapies.​

Video: Assessing Therapeutic Angiogenesis in a Murine Model of Hindlimb Ischemia

Assessing Teratogenic Changes in a Zebrafish Model of Fetal Alcohol Exposure

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, including mental retardation due to disordered and damaged brain development. Fetal alcohol spectrum disorder (FASD) is a term used to cover a continuum of birth defects that occur due to maternal alcohol consumption, and occurs in approximately 4% of children born in the United States. With 50% of child-bearing age women reporting consumption of alcohol, and half of all pregnancies being unplanned, unintentional exposure is a continuing issue2. In order to best understand the damage produced by ethanol, plus produce a model with which to test potential interventions, we developed a model of developmental ethanol exposure using the zebrafish embryo. Zebrafish are ideal for this kind of teratogen study3-8. Each pair lays hundreds of eggs, which can then be collected without harming the adult fish. The zebrafish embryo is transparent and can be readily imaged with any number of stains. Analysis of these embryos after exposure to ethanol at different doses and times of duration and application shows that the gross developmental defects produced by ethanol are consistent with the human birth defect. Described here are the basic techniques used to study and manipulate the zebrafish FAS model.

In order to understand the molecular mechanisms of the ethanol-induced developmental damage, we have developed a zebrafish model of ethanol exposure and are exploring the physical, cellular, and genetic alterations that occur after ethanol exposure1. We then seek to find potential interventions and rapidly test them in this animal model.

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, ...

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ischemia and reperfusion. Immune response includes cytokine expression and release or secretion of endogenous ligands of innate immune receptors. Activation of innate immunity can potentially modulate infarct size. We have modified an existing murine closed-chest model using hanging weights which could be useful for studying myocardial pre- and postconditioning and the role of innate immunity in myocardial ischemia and reperfusion. This model allows animals to recover from surgical trauma before onset of myocardial ischemia.
Volatile anesthetics have been intensely studied and their preconditioning effect for the ischemic heart is well known. However, this protective effect precludes its use in open chest models of coronary artery ligation. Thus, another advantage could be the use of the well controllable volatile anesthetics for instrumentation in a chronic closed-chest model, since their preconditioning effect lasts up to 72 hours. Chronic heart diseases with intermittent ischemia and multiple hit models are other possible applications of this model.
For the chronic closed-chest model, intubated and ventilated mice undergo a lateral blunt thoracotomy via the 4th intercostal space. Following identification of the left anterior descending a ligature is passed underneath the vessel and both suture ends are threaded through an occluder. Then, both suture ends are passed through the chest wall, knotted to form a loop and left in the subcutaneous tissue. After chest closure and recovery for 5 days, mice are anesthetized again, chest skin is reopened and hanging weights are hooked up to the loop under ECG control.
At the end of the ischemia/reperfusion protocol, hearts can be stained with TTC for infarct size assessment or undergo perfusion fixation to allow morphometric studies in addition to histology and immunohistochemistry.

Surgical trauma induces an inflammatory response. Cytokines and endogenous ligands are known to modulate myocardial infarct size following ischemia and reperfusion. We present a modified closed-chest model of murine ischemia and reperfusion using hanging weights to minimize effects of thoracotomy.

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ...

Retinal Detachment Model in Rodents by Subretinal Injection of Sodium Hyaluronate

Subretinal injection of sodium hyaluronate is a widely accepted method of inducing retinal detachment (RD). However, the height and duration of RD or the occurrence of subretinal hemorrhage can affect photoreceptor cell death in the detached retina. Hence, it is advantageous to create reproducible RDs without subretinal hemorrhage for evaluating photoreceptor cell death. We modified a previously reported method to create bullous and persistent RDs in a reproducible location with rare occurrence of subretinal hemorrhage. The critical step of this modified method is the creation of a self-sealing scleral incision, which can prevent leakage of sodium hyaluronate after injection into the subretinal space. To make the self-sealing scleral incision, a scleral tunnel is created, followed by scleral penetration into the choroid with a 30 G needle. Although choroidal hemorrhage may occur during this step, astriction with a surgical spear reduces the rate of choroidal hemorrhage. This method allows a more reproducible and reliable model of photoreceptor death in diseases that involve RD such as rhegmatogenous RD, retinopathy of prematurity, diabetic retinopathy, central serous chorioretinopathy, and age-related macular degeneration (AMD).

Creating experimental retinal detachments with a reproducible and sustained height of detachment, and without subretinal hemorrhage, is important for studying the pathophysiology of photoreceptor cell loss in retinal disease and evaluating potential therapeutic interventions. Here, we report such a method in detail.

Subretinal injection of sodium hyaluronate is a widely accepted method of inducing retinal detachment (RD). However, the height and duration of RD or the ...

Reproducable Paraplegia by Thoracic Aortic Occlusion in a Murine Model of Spinal Cord Ischemia-reperfusion

Background 
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered advancements to decrease the occurrence this injury.&#160;The current model demonstrates reproducible lower&#160;extremity paralysis following thoracic aortic occlusion.
Methods 
Adult male C57BL6 mice were anesthetized with isoflurane. Through a cervicosternal incision the aorta was exposed. The descending thoracic aorta and left subclavian arteries were identified without entrance into pleural space. Skeletonization of these arteries was followed by immediate closure (Sham) or occlusion for 4 min (moderate ischemia) or 8 min (prolonged ischemia). The sternotomy and skin were closed and the mouse was transferred to warming bed for recovery.&#160; Following recovery, functional analysis was obtained at 12 hr intervals until 48 hr.
Results 
Mice that underwent sham surgery showed no observable hind&#160;limb deficit. Mice subjected to moderate ischemia for 4 min had minimal functional deficit at 12 hr followed by progression to complete paralysis at 48 hr. Mice subjected to prolonged ischemia had an immediate paralysis with no observable hind-limb movement at any point in the postoperative period. There was no observed intraoperative or post&#160;operative mortality.
Conclusion 
Reproducible lower extremity paralysis whether immediate or delayed can be achieved in a murine model. Additionally, by using a median sternotomy and careful dissection, high survival rates, and reproducibility can be achieved.

The lack of mechanistic understanding of spinal cord ischemia-reperfusion injury has hindered further adjuncts to prevent paraplegia following high risk aortic operations. Thus, the development of animal models is imperative. This manuscript demonstrates reproducible lower extremity paralysis following thoracic aortic occlusion in a murine model.

Background
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered ...

A Murine Model of Myocardial Ischemia-reperfusion Injury through Ligation of the Left Anterior Descending Artery

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms at work during MI and developing effective therapeutic approaches requires methodology to reproducibly simulate the clinical incidence and reflect the pathophysiological changes associated with MI. Here, we describe a surgical method to induce MI in mouse models that can be used for short-term ischemia-reperfusion (I/R) injury as well as permanent ligation. The major advantage of this method is to facilitate location of the left anterior descending artery (LAD) to allow for accurate ligation of this artery to induce ischemia in the left ventricle of the mouse heart. Accurate positioning of the ligature on the LAD increases reproducibility of infarct size and thus produces more reliable results. Greater precision in placement of the ligature will improve the standard surgical approaches to simulate MI in mice, thus reducing the number of experimental animals necessary for statistically relevant studies and improving our understanding of the mechanisms producing cardiac dysfunction following MI. This mouse model of MI is also useful for the preclinical testing of treatments targeting myocardial damage following MI.

We introduce a surgical method to induce experimental ischemia/reperfusion (I/R) injury to simulate myocardial infarction (MI) in mouse models that allows for more clarity in positioning of the ligation on the left anterior descending artery (LAD) to increase the reproducibility of MI experiments in mice.

Acute or chronic myocardial infarction (MI) are cardiovascular events resulting in high morbidity and mortality. Establishing the pathological mechanisms ...

Murine Model of Intestinal Ischemia-reperfusion Injury

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, hypotension, necrotizing enterocolitis, bowel transplantation, trauma and chronic inflammation. Intestinal ischemia-reperfusion (IR) injury is a consequence of acute mesenteric ischemia, caused by inadequate blood flow through the mesenteric vessels, resulting in intestinal damage. Reperfusion following ischemia can further exacerbate damage of the intestine. The mechanisms of IR injury are complex and poorly understood. Therefore, experimental small animal models are critical for understanding the pathophysiology of IR injury and the development of novel therapies.
Here we describe a mouse model of acute intestinal IR injury that provides reproducible injury of the small intestine without mortality. This is achieved by inducing ischemia in the region of the distal ileum by temporally occluding the peripheral and terminal collateral branches of the superior mesenteric artery for 60 min using microvascular clips. Reperfusion for 1 hr, or 2 hr after injury results in reproducible injury of the intestine examined by histological analysis. Proper position of the microvascular clips is critical for the procedure. Therefore the video clip provides a detailed visual step-by-step description of this technique. This model of intestinal IR injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Here we describe the detailed procedure of intestinal ischemia-reperfusion in mice which results in reproducible injury without mortality to encourage the standardization of this technique across the field. This model of intestinal ischemia-reperfusion injury can be utilized to study the cellular and molecular mechanisms of injury and regeneration.

Intestinal ischemia is a life-threatening condition associated with a broad range of clinical conditions including atherosclerosis, thrombosis, ...

Methods for Acute and Subacute Murine Hindlimb Ischemia

Peripheral artery disease (PAD) is a leading cause of cardiovascular morbidity and mortality in developed countries, and animal models that reliably reproduce the human disease are necessary to develop new therapies for this disease. The mouse hindlimb ischemia model has been widely used for this purpose, but the standard practice of inducing acute limb ischemia by ligation of the femoral artery can result in substantial tissue necrosis, compromising investigators&#39; ability to study the vascular and skeletal muscle tissue responses to ischemia. An alternative approach to femoral artery ligation is the induction of gradual femoral artery occlusion through the use of ameroid constrictors. When placed around the femoral artery in the same or different locations as the sites of femoral artery ligation, these devices occlude the artery over 1 - 3 days, resulting in more gradual, subacute ischemia. This results in less substantial skeletal muscle tissue necrosis, which may more closely mimic the responses seen in human PAD. Because genetic background influences outcomes in both the acute and subacute ischemia models, consideration of the mouse strain being studied is important in choosing the best model. This paper describes the proper procedure and anatomical placement of ligatures or ameroid constrictors on the mouse femoral artery to induce subacute or acute hindlimb ischemia in the mouse.

Surgical induction of hindlimb ischemia in the mouse is useful to examine angiogenesis, however this is compromised in certain inbred mouse strains that display marked ischemia-induced tissue necrosis. Methods are described to induce subacute limb ischemia using ameroid constrictors to circumvent this problem through the induction of gradual arterial occlusion.

Peripheral artery disease (PAD) is a leading cause of cardiovascular morbidity and mortality in developed countries, and animal models that reliably ...

Intravital Microscopy of Monocyte Homing and Tumor-Related Angiogenesis in a Murine Model of Peripheral Arterial Disease

The therapeutic goal for peripheral arterial disease and ischemic heart disease is to increase blood flow to ischemic areas caused by hemodynamic stenosis. Vascular surgery is a viable option in selected cases, but for patients without indications for surgery such as progression to rest pain, critical limb ischemia, or major disruptions to life or work, there are few possibilities for mitigating their disease. Cell therapy via monocyte-enhanced perfusion through the stimulation of collateral formation is one of a few non-invasive options.
Our group examines arteriogenesis after monocyte transplantation into mice using the hindlimb ischemia model. Previously, we have demonstrated improvement in hindlimb perfusion using tetanus-stimulated syngeneic monocyte transplantation. In addition to the effects on the collateral formation, tumor growth could be affected by this therapy as well. To investigate these effects, we use a basement membrane-like matrix mouse model by injecting the extracellular matrix of the Engelbreth-Holm-Swarm sarcoma into the flank of the mouse, after occlusion of the femoral artery.
After the artificial tumor studies, we use intravital microscopy to study in vivo tumor-angiogenesis and monocyte homing within collateral arteries. Previous studies have described the histological examination of animal models, which presupposes subsequent analysis to post-mortem artifacts. Our approach visualizes monocyte homing to areas of collateralization in real time sequences, is easy to perform, and investigates the process of arteriogenesis and tumor angiogenesis in vivo.

Monocytes are important mediators of arteriogenesis in the context of peripheral arterial disease. Using a basement membrane-like matrix and intravital microscopy, this protocol investigates monocyte homing and tumor-related angiogenesis after monocyte injection in the femoral artery ligation murine model.

The therapeutic goal for peripheral arterial disease and ischemic heart disease is to increase blood flow to ischemic areas caused by hemodynamic ...

In Vitro Model of Coronary Angiogenesis

Here, we describe an in vitro culture assay to study coronary angiogenesis. Coronary vessels feed the heart muscle and are of clinical importance. Defects in these vessels represent severe health risks such as in atherosclerosis, which can lead to myocardial infarctions and heart failures in patients. Consequently, coronary artery disease is one of the leading causes of death worldwide. Despite its clinical importance, relatively little progress has been made on how to regenerate damaged coronary arteries. Nevertheless, recent progress has been made in understanding the cellular origin and differentiation pathways of coronary vessel development. The advent of tools and technologies that allow researchers to fluorescently label progenitor cells, follow their fate, and visualize progenies in vivo have been instrumental in understanding coronary vessel development. In vivo studies are valuable, but have limitations in terms of speed, accessibility, and flexibility in experimental design. Alternatively, accurate in vitro models of coronary angiogenesis can circumvent these limitations and allow researchers to interrogate important biological questions with speed and flexibility. The lack of appropriate in vitro model systems may have hindered the progress in understanding the cellular and molecular mechanisms of coronary vessel growth. Here, we describe an in vitro culture system to grow coronary vessels from the sinus venosus (SV) and endocardium (Endo), the two progenitor tissues from which many of the coronary vessels arise. We also confirmed that the cultures accurately recapitulate some of the known in vivo mechanisms. For instance, we show that the angiogenic sprouts in culture from SV downregulate COUP-TFII expression similar to what is observed in vivo. In addition, we show that VEGF-A, a well-known angiogenic factor in vivo, robustly stimulates angiogenesis from both the SV and Endo cultures. Collectively, we have devised an accurate in vitro culture model to study coronary angiogenesis.

In vitro models of coronary angiogenesis can be utilized for the discovery of the cellular and molecular mechanisms of coronary angiogenesis. In vitro explant cultures of sinus venosus and endocardium tissues show robust growth in response to VEGF-A and display a similar pattern of COUP-TFII expression as in vivo.

Here, we describe an in vitro culture assay to study coronary angiogenesis. Coronary vessels feed the heart muscle and are of clinical importance. Defects ...

Delayed Intramyocardial Delivery of Stem Cells after Ischemia Reperfusion Injury in a Murine Model

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, cardiac stem cell therapy is studied by delivering SCs concurrently with the induction of myocardial injury. However, this approach presents two significant limitations: the early hostile pro-inflammatory ischemic environment may affect the survival of transplanted SCs, and it does not represent the subacute infarction scenario where SCs will likely be used. Here we describe a two-part series of surgical procedures for the induction of ischemia-reperfusion injury and delivery of mesenchymal stem cells (MSCs). This method of stem cell administration may allow for the longer viability and retention around damaged tissue by circumventing the initial immune response. A model of ischemia reperfusion injury was induced in mice accompanied by the delivery of mesenchymal stem cells (3.0 x 105), stably expressing the reporter gene firefly luciferase under the constitutively expressed CMV promoter, intramyocardially 7 days later. The animals were imaged via ultrasound and bioluminescent imaging for confirmation of injury and injection of cells, respectively. Importantly, there was no added complication rate when performing this two-procedure approach for SC delivery. This method of stem cell administration, collectively with the utilization of state-of-the-art reporter genes, may allow for the in vivo study of viability and retention of transplanted SCs in a situation of chronic ischemia commonly seen clinically, while also circumventing the initial pro-inflammatory response. In summary, we established a protocol for the delayed delivery of stem cells into the myocardium, which can be used as a potential new approach in promoting regeneration of the damaged tissue.

Stems cells are continuously investigated as potential treatments for individuals with myocardial damage, however, their decreased viability and retention within injured tissue can impact their long-term efficacy. In this manuscript we describe an alternative method for stem cell delivery in a murine model of ischemia reperfusion injury.

There is significant interest in the use of stem cells (SCs) for the recovery of cardiac function in individuals with myocardial injuries. Most commonly, ...