The authors would like to thank Keith Michel of the MD Anderson Small Animal Imaging Facility for his technical assistance with micro-CT imaging, Edward T.H. Yeh, MD, for surgical assistance, and Rebecca Bartow, PhD, for editorial assistance. This work was supported in part by the American Heart Association.

1. Priming the Osmotic Pump
<ol>
	<li>Use sterile techniques (e.g. gloves, laminar flow hood) when preparing the pumps. Remove the pump and flow moderator from the packaging. Weigh the empty pump and record the weight (mg).</li>
	<li>Fill the pump slowly, to avoid creating air bubbles, by using a small 1cc syringe and blunt-tipped 27G filling tube. When the solution reaches the top of the pump, stop filling.</li>
	<li>Wipe off the excess solution and weigh the filled pump. For most aqueous solutions, the weight in mg is the same as the volume in &mu;l. The difference in empty versus filled weight should be more than 90% of the specified mean fill volume. If not, evacuate the pump with the syringe and refill. If you experience technical difficulty please refer to the troubleshooting section of the pump instructions at http://www.alzet.com/products/guide_to_use/filling.html </li>
	<li>Use a pair of forceps to break the white flange from the flow moderator; be careful not to bend or crush the tube. Attach a P50 catheter to either end of the moderator, covering about 3-4 mm. Use a syringe to fill the catheter and attached moderator and leave the syringe attached to the distal part of the catheter.</li>
	<li>Insert the flow moderator into the pump until the catheter is flush with the pump surface. Then, remove the syringe from the distal end of the catheter. Place the pump in a sterile 0.9% saline or PBS solution in a 37 &deg;C incubator for a minimum of 6 hr (preferably O/N) to prime the pump before implanting it into the mouse. The pump delivery rate varies with the model number (i.e. Model 1002 = 0.25 &mu;l/hr for 14 days).</li>
</ol>
2. Creating Unilateral Hindlimb Ischemia in the Mouse
<ol>
	<li>Anesthetize the mice with isoflurane set between 2-4% in 100% O2 at a flow rate of 0.4 L/min and remove the hair (preferably the day before surgery) on the hindlimb and back area with a depilatory cream.</li>
	<li>Sterilize all surgical tools before surgery. Place the mouse in the induction anesthesia chamber. Use the same anesthesia settings described in Section 2.1.</li>
	<li>Wait until the mouse is non-responsive to outside stimuli and then place it in the prone position onto the operating table with a fitted nose cone that delivers a continuous flow of isoflurane (1.5-2.0%).</li>
	<li>Inject the recommended dose of anesthetic (Bupivacaine; 0.25% (&lt;1 ml/kg)) at the incision site; treat the skin with a povidone-iodine solution followed by 70% ethanol. Make a 4-6 mm incision on the back (at the side of the spinal column, in the scapulae region). Separate the subcutaneous membrane with fine forceps; then turn the mouse to the supine position treat the skin with a povidone-iodine solution followed by 70% ethanol and make 0.8-1 cm incision with micro-surgical scissors above the knee towards the medial thigh, following the path of the femoral artery.</li>
	<li>Tunnel up the side of the torso with small sheath tubing (6-French) from the medial thigh to the incision on the back.</li>
	<li>Place the mouse in the prone position. Remove the primed osmotic pump/catheter from the PBS solution and guide the catheter, through the sheath tubing, to the thigh.</li>
	<li>Position the pump inside the back and close the back incision with 6-0 prolene suture (treat the skin with a povidone-iodine solution followed by 70% ethanol). Place the mouse back in the supine position; remove the sheath tubing, leaving the catheter exposed through the incision on the thigh. Secure the leg with surgical tape.</li>
	<li>Separate the subcutaneous fat tissue to reveal the vasculature. To better visualize the vasculature, use a retractor to open the incision.</li>
	<li>Pierce the membranous femoral sheath gently by using fine-pointed forceps to access the vascular bundle. Dissect and separate the femoral artery from the femoral vein and nerve near the groin, taking care not to cause excessive bleeding.</li>
	<li>Place two 8-0 prolene adjacent sutures to interrupt the femoral artery 2-3 mm proximal to the femoral bifurcation (the origin of the profunda femoris and epigastrica arteries).</li>
	<li>Cut the catheter to the appropriate length and place the tip to the anterolateral region of the thigh, securing it with a 6-0 prolene suture to the superficial fascia to ensure targeted delivery of the substance. Close the leg incision with a 6-0 running suture. Administer a final subcutaneous injection of analgesic (Buprenorphine; 0.1 mg/kg), treat the skin with a povidone-iodine solution followed by 70% ethanol and return the mouse to the cage. Use a heating lamp during recovery.</li>
	<li>After a 30-min recovery period, prepare the mouse for laser Doppler imaging to confirm the creation of hindlimb ischemia.</li>
	<li>Monitor the animal twice daily for 5 days after surgery. Mobility of the animal will be observed to analyze pain; waste in the cage will serve as an indicator of eating. Also, during the monitor period determine if the animal is experiencing pain or discomfort and give a subcutaneous injection of Buprenorphine (0.1mg/kg) twice daily.</li>
</ol>
3. Imaging Specimen Preparation
<ol>
	<li>Anesthetize the mouse as stated in Section 2.1. Transfer the mouse onto the operating table in the supine position. Inject 1,000 IU of heparin subcutaneously to prevent clotting in the vasculature. While the mouse is attached to the nosecone, make an incision large enough to expose the thoracic cavity and the lower abdomen. Move the abdominal organs to the left side of the animal to expose the infrarenal abdominal aorta (IAA). Carefully separate the aorta from the inferior cava vein.</li>
	<li>Ligate the proximal IAA to reduce bleeding during catheter insertion. Then, make a small incision on the IAA. Insert a 27&frac12; G blunt needle (attached to a P10 catheter) into the IAA and secure it with a 6-0 suture.</li>
	<li>Fill two 20cc syringes with heparinized saline and 10% neutral buffered formalin (NBF). Fill a 5cc syringe with a contrast radiopaque silicone rubber compound. Mix the three ingredients of MV-compound: MV-diluent: MV-curing agent in working ratios of 1:1.25:0.5. Since the mixture (Microfil) only has a working time of 20 min before the agent polymerizes, it is recommended not to add the MV-curing agent until right before injection into the vasculature.</li>
	<li>Make a cut in the right atrium and administer an overdose of isoflurane to induce euthanasia and to allow for drainage of excess fluid. Attach the saline-filled syringe to the needle tip and begin to flush the vasculature for 5 min. Make sure all the blood is flushed out of the vessels; turn off the isoflurane and disconnect the nosecone once the flushing begins. Next, attach the formalin-filled syringe to the catheter detailed above (3.2) and pressure perfuse for an additional 5 min. Finally, perfuse the last syringe, with the contrast agent, slowly into the mouse for 5 min.</li>
	<li>Optional: Photograph the Microfil in the vasculature with a camera to ensure that only the arterial structure is perfused.</li>
	<li>Place the mouse in a container filled with 10% NBF and store at 4 &deg;C O/N for contrast agent polymerization. After 24 hr, remove the skin on the lower half of the torso (distal to the lower abdominal aorta ligation) by using dissection tools, refresh the 10% NBF, and place the mouse back into the container for 4 more days at RT.</li>
	<li>Dissect the contralateral and ischemic hindlimbs, and place the specimens in a formic acid solution (Cal-Ex II) for 48 hr to facilitate decalcification of the bone. Then, rinse the specimens under running water for 1 hr and return to 10% NBF and proceed with Faxitron (x-ray system used to ensure the bone is decalcified) and micro-CT imaging.</li>
</ol>

<ol>
	<li>Epstein, S. E., Fuchs, S., Zhou, Y. F., Baffour, R., Kornowski, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Therapeutic+interventions+for+enhancing+collateral+development+by+administration+of+growth+factors:+basic+principles,+early+results+and+potential+hazards.">Therapeutic interventions for enhancing collateral development by administration of growth factors: basic principles, early results and potential hazards.</a> Cardiovasc. Res. 49, 532-542 (2001).</li><li>Esaki, J., Marui, A., Tabata, Y., Komeda, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controlled+release+systems+of+angiogenic+growth+factors+for+cardiovascular+diseases.">Controlled release systems of angiogenic growth factors for cardiovascular diseases.</a> Expert Opin. Drug Deliv. 4, 635-649 (2007).</li><li>Bentley, M. D., Ortiz, M. C., Ritman, E. L., Romero, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+microcomputed+tomography+to+study+microvasculature+in+small+rodents.">The use of microcomputed tomography to study microvasculature in small rodents.</a> Am. J. Physiol. Regul. Integr. Comp. Physiol. 282, 1267-1279 (2002).</li><li>Golomb, B. A., Dang, T. T., Criqui, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+arterial+disease:+morbidity+and+mortality+implications.">Peripheral arterial disease: morbidity and mortality implications.</a> Circulation. 114, 688-699 (2006).</li><li>Hiatt, W. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacologic+therapy+for+peripheral+arterial+disease+and+claudication.">Pharmacologic therapy for peripheral arterial disease and claudication.</a> J. Vasc. Surg. 36, 1283-1291 (2002).</li><li>Hirsch, A. T., 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+arterial+disease+detection,+awareness,+and+treatment+in+primary+care.">Peripheral arterial disease detection, awareness, and treatment in primary care.</a> JAMA. 286, 1317-1324 (2001).</li><li>McDonald, D. M., Choyke, P. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+of+angiogenesis:+from+microscope+to+clinic.">Imaging of angiogenesis: from microscope to clinic.</a> Nat. Med. 9, 713-725 (2003).</li><li>Hirsch, A. T., 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=ACC/AHA+Guidelines+for+the+Management+of+Patients+with+Peripheral+Arterial+Disease+(lower+extremity,+renal,+mesenteric,+and+abdominal+aortic):+a+collaborative+report+from+the+American+Associations+for+Vascular+Surgery/Society+for+Vascular+Surgery,+Society+for+Cardiovascular+Angiography+and+Interventions,+Society+for+Vascular+Medicine+and+Biology,+Society+of+Interventional+Radiology,+and+the+ACC/AHA+Task+Force+on+Practice+Guidelines+(writing+committee+to+develop+guidelines+for+the+management+of+patients+with+peripheral+arterial+disease)--summary+of+recommendations.">ACC/AHA Guidelines for the Management of Patients with Peripheral Arterial Disease (lower extremity, renal, mesenteric, and abdominal aortic): a collaborative report from the American Associations for Vascular Surgery/Society for Vascular Surgery, Society for Cardiovascular Angiography and Interventions, Society for Vascular Medicine and Biology, Society of Interventional Radiology, and the ACC/AHA Task Force on Practice Guidelines (writing committee to develop guidelines for the management of patients with peripheral arterial disease)--summary of recommendations.</a> J. Vasc. Interv. Radiol. 17, 1383-1397 (2006).</li><li>Zagorchev, L., Mulligan-Kehoe, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+imaging+of+vessels+in+mouse+models+of+disease.">Molecular imaging of vessels in mouse models of disease.</a> Eur. J. Radiol. 70, 305-311 (2009).</li><li>Losordo, D. W., Dimmeler, S. <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+and+vasculogenesis+for+ischemic+disease.+Part+I:+angiogenic+cytokines.">Therapeutic angiogenesis and vasculogenesis for ischemic disease. Part I: angiogenic cytokines.</a> Circulation. 109, 2487-2491 (2004).</li><li>Waters, R. E., Terjung, R. L., Peters, K. G., Annex, B. 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The authors declare no conflict of interest. All animal procedures were conducted in accordance with the Guidelines for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Texas Health Science Center at Houston.

Here we present a method for osmotic drug/substance delivery in a mouse model of hindlimb ischemia. In addition, we describe a casting technique in which we have used Microfil to produce a 3D reconstruction for analysis of the vascular network. 
 The level or severity of ischemia varies according to where the arterial ligation/excision is made. We created a double ligation in the common femoral artery proximal to the origin of the profunda femoris and epigastrica arteries; this approac...

Peripheral arterial disease (PAD) is an atherosclerotic disease that causes insufficient blood supply in the legs4. It affects 8 to 12 million Americans, and current medical treatments offer only limited relief5,6. Novel therapeutic agents that improve blood circulation in legs would not only restrain disease progression but also enhance quality of life. The incidence of PAD is higher in people over the age of 50 years, so local pharmacologic therapy is a more desirable treatment modality because the reduced kidney and liver function often seen in older patients can decrease drug metabolism and increase side effects with systemic administration.
Thus, we created a mouse model of PAD to examine whether locally administered agents alleviate hindlimb ischemia by promoting angiogenesis and microvascular remodeling. Specifically, we used a catheterized osmotic pump to continuously deliver the therapeutic agent to the ischemic thigh muscle of mice. Using our delivery system, we were able to maintain optimal concentrations of the drug in the local environment; this approach allows for appropriate drug bioactivity, avoids possible systemic side effects, and overcomes the disadvantage of limited local drug access associated with systemic administration. Additionally, to assess whether locally administered agents promote revascularization, we used advanced casting and high-definition imaging techniques that enable quantification of changes in the microvasculature. Collectively, the combination of methodologies used in this video article is useful in preclinical studies to aid in understanding pharmacologically induced revascularization in PAD patients7-9.

<table><tbody><tr><td colspan="4">&lt;strong&gt;Reagent/Material&lt;/strong&gt;</td></tr><tr><td>Surgical tools</td><td>Fine Science Tools</td><td></td><td>Type: Tool</td></tr><tr><td>Puritan sterile cotton swabs</td><td>Fisher Scientific</td><td>22-029-499</td><td>Type: Tool</td></tr><tr><td>Betadine (povidone-iodine)</td><td>Fisher Scientific</td><td>19-065534</td><td>Type: Reagent</td></tr><tr><td>70% Alcohol pads</td><td>Fisher Scientific</td><td>NC9926371</td><td>Type: Reagent</td></tr><tr><td>Phosphate buffered saline</td><td>Lonza</td><td>17-516F</td><td>Type: Reagent</td></tr><tr><td>6-0 prolene suture</td><td>Cardinal Health</td><td>8709</td><td>Type: Tool</td></tr><tr><td>8-0 prolene suture</td><td>Cardinal Health</td><td>2775</td><td>Type: Tool</td></tr><tr><td>Depilatory cream</td><td>Nair</td><td></td><td>Type: Tool</td></tr><tr><td>Osmotic pump</td><td><a href="http://www.alzet.com" target="_blank"><img src="/img/new_window.png" /> ALZET</a></td><td>1002</td><td>Type: Tool, 14 day release</td></tr><tr><td>Vinyl catheter</td><td><a href="http://www.alzet.com" target="_blank"><img src="/img/new_window.png" /> ALZET</a></td><td>7760</td><td>Type: Tool</td></tr><tr><td>Heparinized saline (0.9%)</td><td>Baxter</td><td>2B0944</td><td>Type: Reagent</td></tr><tr><td>Neutral buffered formalin</td><td>Richard-Allan Scientific</td><td>5705</td><td>Type: Reagent</td></tr><tr><td>Microfil (silicone rubber contrast agent)</td><td>Flowtech</td><td>MV-112</td><td>Type Reagent, Microfil White</td></tr><tr><td>Cal-Ex II (formic acid solution)</td><td>Fisher Scientific</td><td>CS511-1D</td><td>Type: Reagent</td></tr><tr><td>Buprenex</td><td>CIII</td><td>7571</td><td>Type: Analgesic</td></tr><tr><td>Bupivicaine</td><td>Hospira, Inc.</td><td>381</td><td>Type: Analgesic</td></tr><tr><td colspan="4">&lt;strong&gt;Equipment&lt;/strong&gt;</td></tr><tr><td>Dissecting microscope</td><td>Carl Zeiss Microimaging</td><td>Zeiss Stemi 2000-C</td><td>Type:Equipment</td></tr><tr><td>Laser Doppler perfusion imager</td><td>Perimed Inc.</td><td>Periscan PIM3</td><td>Type:Equipment</td></tr><tr><td>Micro-CT imaging system</td><td>GE Healthcare</td><td>Explore Locus SP</td><td>Type:Equipment</td></tr></tbody></table>

osmotic drug delivery to ischemic hindlimbs and perfusion of vasculature with microfil for micro-computed tomography imaging

Preclinical research in animal models of peripheral arterial disease plays a vital role in testing the efficacy of therapeutic agents designed to stimulate microcirculation. The choice of delivery method for these agents is important because the route of administration profoundly affects the bioactivity and efficacy of these agents1,2. In this article, we demonstrate how to locally administer a substance in ischemic hindlimbs by using a catheterized osmotic pump. This pump can deliver a fixed volume of aqueous solution continuously for an allotted period of time. We also present our mouse model of unilateral hindlimb ischemia induced by ligation of the common femoral artery proximal to the origin of profunda femoris and epigastrica arteries in the left hindlimb. Lastly, we describe the in vivo cannulation and ligation of the infrarenal abdominal aorta and perfusion of the hindlimb vasculature with Microfil, a silicone radiopaque casting agent. Microfil can perfuse and fill the entire vascular bed (arterial and venous), and because we have ligated the major vascular conduit for exit, the agent can be retained in the vasculature for future ex vivo imaging with the use of small specimen micro-CT3.

The animation demonstrates the osmotic pump insertion and hindlimb ischemia surgery detailed in the protocol. Figure 1 shows laser Doppler images of perfusion of the hindlimb, confirming ischemia. After vascular casting with Microfil, a 3D micro-CT image of the vascular network shows that Microfil can fill the vessels effectively (Figure 2A) but that discontinuities can occur (Figure 2B) due to various procedural factors (e.g., air bubbles, lack of pressure, vis...

Watch this Scientific Journal Video about Osmotic Drug Delivery to Ischemic Hindlimbs and Perfusion of Vasculature with Microfil for Micro-Computed Tomography Imaging at JoVE.com

Osmotic Drug Delivery to Ischemic Hindlimbs and Perfusion of Vasculature with Microfil for Micro-Computed Tomography Imaging

We show here the in vivo insertion of an osmotic pump for constant local drug delivery and the creation of hindlimb ischemia in a mouse model. Moreover, the hindlimb vasculature is perfused with Microfil, a silicone radiopaque agent, to prepare for micro-computed tomography (micro-CT) imaging.

Preclinical research in animal models of peripheral arterial disease plays a vital role in testing the efficacy of therapeutic agents designed to ...

osmotic-drug-delivery-to-ischemic-hindlimbs-perfusion-vasculature

Research

JoVE Journal

Medicine

12.3K Views.  The Texas Heart Institute at St. Luke's Episcopal Hospital. We show here the in vivo insertion of an osmotic pump for constant local drug delivery and the creation of hindlimb ischemia in a mouse model. Moreover, the hindlimb vasculature is perfused with Microfil, a silicone radiopaque agent, to prepare for micro-computed tomography (micro-CT) imaging.

Video: Osmotic Drug Delivery to Ischemic Hindlimbs and Perfusion of Vasculature with Microfil for Micro-Computed Tomography Imaging

Contrast Enhanced Vessel Imaging using MicroCT

Microscopic computed tomography (microCT) offers high-resolution volumetric imaging of the anatomy of living small animals. However, the contrast between different soft tissues and body fluids is inherently poor in micro-CT images 1. Under these circumstances, visualization of blood vessels becomes a nearly impossible task. To overcome this and to improve the visualization of blood vessels exogenous contrast agents can be used. Herein, we present a methodology for visualizing the vascular network in a rodent model. By using a long-acting aqueous colloidal polydisperse iodinated blood-pool contrast agent, eXIA 160XL, we optimized image acquisition parameters and volume-rendering techniques for finding blood vessels in live animals. Our findings suggest that, to achieve a superior contrast between bone and soft tissue from vessel, multiple-frames (at least 5-8/ frames per view), and 360-720 views (for a full 360&deg; rotation) acquisitions were mandatory. We have also demonstrated the use of a two-dimensional transfer function (where voxel color and opacity was assigned in proportion to CT value and gradient magnitude), in visualizing the anatomy and highlighting the structure of interest, the blood vessel network. This promising work lays a foundation for the qualitative and quantitative assessment of anti-angiogenesis preclinical studies using transgenic or xenograft tumor-bearing mice.

Contrast enhanced small animal vessel imaging by microCT is a rapid, cost-effective and high-throughput technique for serial in situ examination for tumor development, for analyzing the network of blood vessels that nourish them, and for following the response of tumors to preclinical therapeutic intervention(s).

Microscopic computed tomography (microCT) offers high-resolution volumetric imaging of the anatomy of living small animals. However, the contrast between ...

Retrograde Perfusion and Filling of Mouse Coronary Vasculature as Preparation for Micro Computed Tomography Imaging

Visualization of the vasculature is becoming increasingly important for understanding many different disease states. While several techniques exist for imaging vasculature, few are able to visualize the vascular network as a whole while extending to a resolution that includes the smaller vessels1,2. Additionally, many vascular casting techniques destroy the surrounding tissue, preventing further analysis of the sample3-5. One method which circumvents these issues is micro-Computed Tomography (&mu;CT). &mu;CT imaging can scan at resolutions &lt;10 microns, is capable of producing 3D reconstructions of the vascular network, and leaves the tissue intact for subsequent analysis (e.g., histology and morphometry)6-11. However, imaging vessels by ex vivo &mu;CT methods requires that the vessels be filled with a radiopaque compound. As such, the accurate representation of vasculature produced by &mu;CT imaging is contingent upon reliable and complete filling of the vessels. In this protocol, we describe a technique for filling mouse coronary vessels in preparation for &mu;CT imaging.
Two predominate techniques exist for filling the coronary vasculature: in vivo via cannulation and retrograde perfusion of the aorta (or a branch off the aortic arch) 12-14, or ex vivo via a Langendorff perfusion system 15-17. Here we describe an in vivo aortic cannulation method which has been specifically designed to ensure filling of all vessels. We use a low viscosity radiopaque compound called Microfil which can perfuse through the smallest vessels to fill all the capillaries, as well as both the arterial and venous sides of the vascular network. Vessels are perfused with buffer using a pressurized perfusion system, and then filled with Microfil. To ensure that Microfil fills the small higher resistance vessels, we ligate the large branches emanating from the aorta, which diverts the Microfil into the coronaries. Once filling is complete, to prevent the elastic nature of cardiac tissue from squeezing Microfil out of some vessels, we ligate accessible major vascular exit points immediately after filling. Therefore, our technique is optimized for complete filling and maximum retention of the filling agent, enabling visualization of the complete coronary vascular network &#x2013; arteries, capillaries, and veins alike.

Visualization of the coronary vessels is critical to advancing our understanding of cardiovascular diseases. Here we describe a method for perfusing murine coronary vasculature with a radiopaque silicone rubber (Microfil), in preparation for micro-Computed Tomography (&mu;CT) imaging.

Visualization of the vasculature is becoming increasingly important for understanding many different disease states. While several techniques exist for ...

Computed Tomography and Optical Imaging of Osteogenesis-angiogenesis Coupling to Assess Integration of Cranial Bone Autografts and Allografts

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that implantation of a bone autograft would induce greater bone regeneration by abundant blood vessel formation. To investigate the effect of the graft on neovascularization at the defect site, we developed a micro&#8211;computed tomography (&#181;CT) approach to characterize newly forming blood vessels, which involves systemic perfusion of the animal with a polymerizing contrast agent. This method enables detailed vascular analysis of an organ in its entirety. Additionally, blood perfusion was assessed using fluorescence imaging (FLI) of a blood-borne fluorescent agent. Bone formation was quantified by FLI using a hydroxyapatite-targeted probe and &#181;CT analysis. Stem cell recruitment was monitored by bioluminescence imaging (BLI) of transgenic mice that express luciferase under the control of the osteocalcin promoter. Here we describe and demonstrate preparation of the allograft, calvarial defect surgery, &#181;CT scanning protocols for the neovascularization study and bone formation analysis (including the in vivo perfusion of contrast agent), and the protocol for data analysis.
The 3D high-resolution analysis of vasculature demonstrated significantly greater angiogenesis in animals with implanted autografts, especially with respect to arteriole formation. Accordingly, blood perfusion was significantly higher in the autograft group by the 7th day after surgery. We observed superior bone mineralization and measured greater bone formation in animals that received autografts. Autograft implantation induced resident stem cell recruitment to the graft-host bone suture, where the cells differentiated into bone-forming cells between the 7th and 10th postoperative day. This finding means that enhanced bone formation may be attributed to the augmented vascular feeding that characterizes autograft implantation. The methods depicted may serve as an optimal tool to study bone regeneration in terms of tightly bounded bone formation and neovascularization.

Implantation of autologous and allogeneic bone grafts constitute accepted approaches to treat major craniofacial bone loss. Yet the effect of graft composition on the interplay among neovascularization, cell differentiation and bone formation is unclear. We present a multimodal imaging protocol aimed to elucidate the angiogenesis-osteogenesis interdependence at the graft proximity.

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that ...

Bioengineering

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 ...

Real-Time Intravital Multiphoton Microscopy to Visualize Focused Ultrasound and Microbubble Treatments to Increase Blood-Brain Barrier Permeability

The blood-brain barrier (BBB) is a key challenge for the successful delivery of drugs to the brain. Ultrasound exposure in the presence of microbubbles has emerged as an effective method to transiently and locally increase the permeability of the BBB, facilitating para- and transcellular transport of drugs across the BBB. Imaging the vasculature during ultrasound-microbubble treatment will provide valuable and novel insights on the mechanisms and dynamics of ultrasound-microbubble treatments in the brain.
Here, we present an experimental procedure for intravital multiphoton microscopy using a cranial window aligned with a ring transducer and a 20x objective lens. This set-up enables high spatial and temporal resolution imaging of the brain during ultrasound-microbubble treatments. Optical access to the brain is obtained via an open-skull cranial window. Briefly, a 3-4 mm diameter piece of the skull is removed, and the exposed area of the brain is sealed with a glass coverslip. A 0.82 MHz ring transducer, which is attached to a second glass coverslip, is mounted on top. Agarose (1% w/v) is used between the coverslip of the transducer and the coverslip covering the cranial window to prevent air bubbles, which impede ultrasound propagation. When sterile surgery procedures and anti-inflammatory measures are taken, ultrasound-microbubble treatments and imaging sessions can be performed repeatedly over several weeks. Fluorescent dextran conjugates are injected intravenously to visualize the vasculature and quantify ultrasound-microbubble induced effects (e.g., leakage kinetics, vascular changes). This paper describes the cranial window placement, ring transducer placement, imaging procedure, common troubleshooting steps, as well as advantages and limitations of the method.

This protocol describes the surgical and technical procedures that enable real-time in vivo multiphoton fluorescence imaging of the rodent brain during focused ultrasound and microbubble treatments to increase blood-brain barrier permeability.

The blood-brain barrier (BBB) is a key challenge for the successful delivery of drugs to the brain. Ultrasound exposure in the presence of microbubbles ...

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

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

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

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

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.

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 ...

Laser Doppler Perfusion Imaging in the Mouse Hindlimb

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is a common, noninvasive, repeatable method for assessing blood flow recovery. The technique calculates overall blood flow in the sampled tissue from the Doppler shift in frequency caused when a laser hits moving red blood cells. Measurements are expressed in arbitrary perfusion units, so the contralateral non-intervened upon leg is usually used to help control measurements. Measurement depth is in the range of 0.3-1 mm; for hindlimb ischemia, this means that dermal perfusion is assessed. Dermal perfusion is dependent on several factors&#8212;most importantly skin temperature and anesthetic agent, which must be carefully controlled to result in reliable readings. Furthermore, hair and skin pigmentation can alter the ability of the laser to either reach or penetrate to the dermis. This article demonstrates the technique of LDPI in the mouse hindlimb.

Here, we present a protocol that demonstrates the technique and necessary controls for Laser Doppler perfusion imaging to measure blood flow in the mouse hindlimb.

Blood flow recovery is a critical outcome measure after experimental hindlimb ischemia or ischemia-reperfusion. Laser Doppler perfusion imaging (LDPI) is ...

Real-Time Assessment of Spinal Cord Microperfusion in a Porcine Model of Ischemia/Reperfusion

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its incidence is still considerably high and therefore, influences patient outcome. Microcirculation plays a key role in tissue perfusion and oxygen supply and is often dissociated from macrohemodynamics. Thus, direct evaluation of spinal cord microcirculation is essential for the development of microcirculation-targeted therapies and the evaluation of existing approaches in regard to spinal cord microcirculation. However, most of the methods do not provide real-time assessment of spinal cord microcirculation. The aim of this study is to describe a standardized protocol for real-time spinal cord microcirculatory evaluation using laser-Doppler needle probes directly inserted in the spinal cord. We used a porcine model of ischemia/reperfusion to induce deterioration of the spinal cord microcirculation. In addition, a fluorescent microsphere injection technique was used. Initially, animals were anesthetized and mechanically ventilated. Thereafter, laser-Doppler needle probe insertion was performed, followed by the placement of cerebrospinal fluid drainage. A median sternotomy was performed for exposure of the descending aorta to perform aortic cross-clamping. Ischemia/reperfusion was induced by supra-celiac aortic cross-clamping for a total of 48 min, followed by reperfusion and hemodynamic stabilization. Laser-Doppler Flux was performed in parallel with macrohemodynamic evaluation. In addition, automated cerebrospinal fluid drainage was used to maintain a stable cerebrospinal pressure. After completion of the protocol, animals were sacrificed, and the spinal cord was harvested for histopathological and microsphere analysis. The protocol reveals the feasibility of spinal cord microperfusion measurements using laser-Doppler probes and shows a marked decrease during ischemia as well as recovery after reperfusion. Results showed comparable behavior to fluorescent microsphere evaluation. In conclusion, this new protocol might provide a useful large animal model for future studies using real-time spinal cord microperfusion assessment in ischemia/reperfusion conditions.

Spinal cord microcirculation plays a pivotal role in spinal cord injury. Most methods do not allow real-time assessment of spinal cord microcirculation, which is essential for the development of microcirculation-targeted therapies. Here, we propose a protocol using Laser-Doppler-Flow Needle probes in a large animal model of ischemia/reperfusion.

Spinal cord injury is a devastating complication of aortic repair. Despite developments for the prevention and treatment of spinal cord injury, its ...

Sample Preparation for Computed Tomography-based Three-dimensional Visualization of Murine Hind-limb Vessels

Blood vessels are complex networks with tree-like structures, and vascular networks are essential for maintaining both circulation and maintaining organ function. Clarifying the mechanism of blood vessel formation is therefore extremely useful for elucidating developmental processes and pathological mechanisms. Murine hind-limb vessels are often used as a model for physiological and pathological angiogenesis. Evaluation is mainly performed via a two-dimensional method using tissue sections. However, methods for evaluating three-dimensional (3D) vascular morphology are particularly limited. This paper introduces a method for visualizing murine hind-limbs using computed tomography (CT). Radiation-opaque resin is injected through the descending aorta, and whole vessels are filled with dye. By adjusting the time of dye injection, arterial-specific filling is also possible, and samples can be obtained with any micro-X-ray CT device. This contrast method provides a basic technique for the 3D evaluation of murine blood vessels in the lower extremities. Furthermore, this method can be used to visualize all blood vessels below the diaphragm and evaluate blood vessels in the abdominal organs.

Here, we describe a visualization and quantification method for murine hind-limb vessels using micro-X-ray computed tomography.

Blood vessels are complex networks with tree-like structures, and vascular networks are essential for maintaining both circulation and maintaining organ ...