We thank Dr. Kelly Stevens for initial trials of the protocol, Dr. Michael Simons, Dr. Kip Hauch, and members of both of their labs for general discussion.
This work is support by NIH grants HL087513 and P01 HL094374.

1. Preparations before starting
<ol> 
 <li>Fill each side of the pressure perfusion apparatus with Vasodilator buffer (4mg/L Papaverine + 1g/L Adenosine in PBS) or 4% Paraformaldehyde (PFA) in PBS, respectively.</li>
 <li>Prepare a 1/2cc Insulin Syringe (with a permanently attached 29G &frac12;" needle) by filling it with 0.1 ml of 1:100 Heparin (5000U/ml stock) and bending the needle to ~120 degree angle with the bevel up. Do the same with a 1 ml syringe (with a 26G &frac12;" needle) filled with 0.3 ml saturated KCl solution.</li>
</ol>
2. Exposing the heart and cannulating the aorta
<ol>
 <li>Anesthetize the mouse using your anesthetic of choice. (We use an overdose of a Ketamine/Xylazine mixture: IP injection of 130 mg/kg Ketamine and 8.8 mg/kg Xylazine in saline.)</li>
 <li>Pin the anesthetized mouse onto the dissecting tray, ventral side up. Open the abdominal cavity with a midline incision and retract the skin to expose the organs. Move the intestines to one side to expose a region of the Posterior Vena Cava (PVC).</li>
 <li>Inject Heparin solution into the PVC. As you extract the needle, cover the needle hole with a cotton-tipped applicator to prevent leaking and hold it for a few seconds until the PVC wall clots and seals. Wait 2-3 minutes for Heparin to disperse throughout the mouse circulation.</li>
 <li>Dissect the diaphragm and rib cage so you can observe the beating heart. Slowly inject KCl solution in the PVC until the heart arrests.</li>
 <li>Remove all organs below the diaphragm and excise the posterior portion of the mouse, leaving the region anterior of the diaphragm intact. Remove the diaphragm, being careful to cut the PVC near the diaphragm so the portion proximal to the heart is easy to locate in subsequent steps.</li>
 <li>Locate the cut end of aorta. Place one long length of 6-0 braided silk suture underneath the aorta a few millimeters anterior from the cut end such that the suture is doubled back on itself. Cut this longer suture in half so there are 2 pieces of suture under the aorta. Insert the angiocatheter into the cut end of the aorta (Fig 1A, B) and tie each suture with a double-knot to hold the angiocatheter in place and prevent any back-pressure within the aorta from leaking out. </li>
</ol>
3. Perfusion and Microfil injection
<ol>
 <li>Connect the angiocatheter to the pressure perfusion apparatus (Fig 2) and begin perfusing the vessels with vasodilator buffer (Fig 1C) by pumping the perfusion apparatus to a driving pressure of 100-110 mmHg. Double check that buffer is perfusing through the coronaries by ensuring liquid is exiting from the PVC. Continue to perfuse for at least 3 minutes or until the fluid exiting the PVC is clear. (Continue with the next steps while perfusing.)</li>
 <li>Dissect the ribs and pin back (or remove) the ribcage to expose the heart. Once exposed, be careful not to let the heart dry out by squeezing drops of buffer onto the heart from a buffer-soaked piece of gauze. Clear away the thymus to expose the aortic arch. Ligate the three major aortic branches using 6-0 braided silk suture to ensure fluid is diverted through the coronaries rather than through these larger, low resistance vessels (Fig 1D).</li>
 <li>Perfuse the heart with fixative for 15 minutes, then rinse with Vasodilation buffer for at least 2 minutes. Meanwhile, ligate both Anterior Vena Cavae to prevent Microfil from leaking out of the heart after injection (Fig 1E). Place sutures around the PVC and the aorta but do not tighten them until after filling.</li>
 <li>Prepare the Microfil (as specified in table of reagents) and load it into a 1 ml syringe. Fill the dissection tray with enough water to cover the catheter (so as to prevent the introduction of air bubbles when switching from perfusion tubing to the Microfil syringe). Disconnect the perfusion apparatus from the catheter and connect the prepared Microfil syringe.</li>
 <li>Inject the Microfil into the aorta until good filling of the coronary arteries is evident (Figs 1F, 3A): the arteries will fill first, and then the Microfil will "spill" into the capillaries as the tissue flushes with the color of the Microfil. Once on the venous side, the hydrophobic nature of the Microfil causes it to initially appear as independent spheres as it emerges from the smaller vessels. Continue to inject the Microfil until a continuous column fills the veins. Complete filling will be evident when the Microfil is continuous within the vessels, and it is exiting through the PVC.</li>
 <li>After filling is complete, quickly tighten the sutures that were previously placed around the PVC and aorta to prevent the elastic nature of the cardiac tissue from squeezing the Microfil out of the vessels.</li>
 <li>Cover the heart with wet gauze (soaked with water from the dissecting tray) to prevent its drying out, and let it sit for approximately 1 hour at room temperature until the Microfil has polymerized. Avoid any external pressures on the heart during the polymerization process, such as lifting or turning the heart in an attempt to get an early view of the filled vessels in the back of the heart. This may squeeze Microfil from some vessels into a more elastic area of the heart, causing breaks in the Microfil.</li>
 <li>Remove the heart and post-fix it in 4% PFA overnight at 4&deg;C. Then store in 70% ethanol at 4 &deg;C. The heart vasculature is now ready for &mu;CT imaging.</li>
</ol>
4. Representative Results
Vessels which are effectively perfused by Microfil will have continuous, unbroken Microfil throughout the vessels (Fig. 3A). The extent of filling of the coronary vessels can be judged by eye; veins are epicardially located18, and can be easily observed (Fig 3A, arrowhead); arteries, which are more intramyocardial18, are also visible through the surface of the heart (Fig 3A, arrow). Capillary filling is also evident, as cardiac tissue has a very high density of capillaries, and therefore, when the capillaries fill, the cardiac tissue will flush with the color of the Microfil (Fig. 3A, star). Thus, any vascular networks that failed to fill will be noticeable due to the lack of Microfil (Fig. 3B, C). 
Discontinuities in the Microfil (asterisks in Fig 3B) often appear because the hydrophobic nature of the Microfil will cause it to contract into itself and cause "breaks" within filled vessels. These "breaks" can be reduced if pressure within the vessels is maintained through proper tie-offs of the vascular exit points from the heart. Other discontinuities can be caused by air bubbles within the microfil. To prevent the introduction of air, make sure the angiocatheter is fully submerged in water when switching from the perfusion apparatus to the Microfil syringe. If an air bubble is introduced, it can often be removed simply by continuing the Microfil perfusion until the bubble has been pushed through and out of the coronary vessels. 

Vascular networks may not fill completely if a portion of the vascular bed is blocked (Fig. 3B, arrow). While Heparin inhibits the formation of blood clots, occasional blockages may still occur due to incomplete Heparin perfusion prior to beginning the procedure, or due to other unknown factors. If a blockage occurs, there is, to our knowledge, no method for dislodging the blockage to complete the vascular fill. Incomplete filling can also result if too little pressure is used during filling, as the Microfil will not be forced into all the vascular beds and capillary networks (Fig. 3C). Conversely, too much pressure can cause the capillaries to burst and extravasate Microfil into the surrounding tissue (Fig. 3D). 
<img src="/files/ftp_upload/3740/3740fig1.jpg" alt="Figure 1"/> 
Figure 1. Overview of the Microfil perfusion scheme. (A) The aorta and the PVC are cut at approximately the level of the diaphragm. (B) The ascending aorta is cannulated with an angiocatheter. (C) Vasodilation buffer is perfused through the vessels, driven by the pressure perfusion apparatus (not pictured), while (D) the three main branches off the aortic arch are ligated. (E) 4% PFA is perfused through the coronaries while both Anterior Vena Cavas are ligated. (F) Using a syringe, Microfil is perfused through the coronaries until it is observed exiting from the PVC.

<img src="/files/ftp_upload/3740/3740fig2.jpg" alt="Figure 2"/> 
Figure 2. Perfusion Apparatus. Two Erlenmeyer flasks, each filled with either Vasodilation buffer or 4% PFA, are joined and pressurized through tubes connected to their sidearms. The system is pressurized through manual pumping of the bulb, and a pressure gauge is connected to one of the flasks to allow monitoring and maintenance of pressure. Small tubes extend through rubber stoppers and down into the fluid in each flask. Pressure entering from the sidearms pumps the fluid from each flask out these smaller tubes. The tubes then merge at a stopcock which only allows fluid to flow from one flask at a time. 
<img src="/files/ftp_upload/3740/3740fig3.jpg" alt="Figure 3"/> 
Figure 3. Sample Microfilled hearts. (A) Vessels that are filled well will have few (if any) breaks in the Microfil, and the heart tissue will be tinged the color of the Microfil due to the filled capillaries (star, and compare with C). Both arteries (arrow &#x2013; Left Anterior Descending Artery) and veins (arrowhead &#x2013; Left Coronary Vein) are visible through the heart surface. (B) A heart with breaks in the microfil (asterisks) as well as blockages in some vessels that prevented complete Microfil penetration. The blocked vessels remain red (arrow), as the blood was not flushed out during the perfusion process. (C) A heart with vessels that were incompletely filled. Notice the tissue has not taken on the yellow color of the Microfil, indicating the Microfil did not penetrate into the capillaries. (D) A heart where the capillaries burst during filling, causing the Microfil to leak into the surrounding tissue (arrow).

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Mice were handled with methods approved by the Institutional Animal Care and Use Committee of the University of Washington and in accordance with the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996).

Cardiac tissue has a very high metabolic demand, and therefore requires a constant supply of nutrients and oxygen from the blood delivered by the coronary vasculature. Diseases of the coronary vessels, which decrease coronary function due to vessel stenosis and blockage, can lead to tissue hypoxia and ischemia, and put affected patients at risk for myocardial infarction and irreparable damage to the heart muscle. A better understanding of the diseased state of these vessels is necessary, and critical to our ability to st...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
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 <td><a href="http://www.bd.com/" target="_blank">Becton Dickinson</a></td>
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 <td>1/2cc insulin syringes with permanently attached 29G &frac12;' needles</td>
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 <td>24G &frac34;" Angiocath IV catheter</td>
 <td><a href="http://www.bd.com/" target="_blank">Becton Dickinson</a></td>
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 <td>26G &frac12;"gauge needles</td>
 <td><a href="http://www.bd.com/" target="_blank">Becton Dickinson</a></td>
 <td>BD-305111</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Adenosine</td>
 <td><a href="http://www.sigmaaldrich.com/" target="_blank">Sigma</a></td>
 <td>A9251</td>
 <td>1g/L in PBS for Vasodilation Buffer (with Papaverine)</td>
 </tr>
 <tr>
 <td>Angled Graefe Forceps</td>
 <td><a href="http://www.finescience.com/" target="_blank">Fine Science Tools</a></td>
 <td>11052-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Cotton-tipped applicators: 6" non-sterile</td>
 <td><a href="http://www.cardinal.com/" target="_blank">Cardinal Health</a></td>
 <td>C15055-006</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Curved Surgical Scissors</td>
 <td><a href="http://www.finescience.com/" target="_blank">Fine Science Tools</a></td>
 <td>14085-09</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dissecting stereoscope and light source</td>
 <td><a href="http://www.nikon.com/" target="_blank">Nikon</a></td>
 <td>NA</td>
 <td>NA</td>
 </tr>
 <tr>
 <td>Dissecting Tray, 11.5 x 7.5 inches</td>
 <td><a href="http://www.coleparmer.com/" target="_blank">Cole-Parmer</a></td>
 <td>YO-10915-12</td>
 <td>Filled with tar for pinning down the mouse</td>
 </tr>
 <tr>
 <td>Fine Curved Forceps</td>
 <td><a href="http://www.aesculapusa.com/" target="_blank">Aesculap</a></td>
 <td>FD281R</td>
 <td>Need two</td>
 </tr>
 <tr>
 <td>Heparin, 5000 U/ml stock</td>
 <td><a href="http://www.apppharma.com/" target="_blank">APP Pharmaceuticals LLC</a></td>
 <td>NDC 63323-047-10</td>
 <td>1:100 dilution in water</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td><a href="http://www.fishersci.com/" target="_blank">Fisher</a></td>
 <td>P217</td>
 <td>Saturated solution in H2O</td>
 </tr>
 <tr>
 <td>Ketamin&nbsp; (Ketaset), 100 mg/ml stock</td>
 <td>Fort Dodge, Overland Park, KS, USA</td>
 <td>NDC 0856-2013-01</td>
 <td>Mixed as 130 mg/kg body weight, with Xylazine in 0.9% saline</td>
 </tr>
 <tr>
 <td>Microfil</td>
 <td><a href="http://www.flowtech-inc.com/" target="_blank">Flow Tech</a></td>
 <td>MV-122 (yellow). Other color options are also available.</td>
 <td>Mix 1:1 by weight, with 10% by volume of curing agent. Prepare just before injection, and vortex to ensure it is well mixed</td>
 </tr>
 <tr>
 <td>Non-sterile Suture: 6-0, braided silk</td>
 <td><a href="http://www.harvardapparatus.com/" target="_blank">Harvard Apparatus</a></td>
 <td>723287</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Papaverine</td>
 <td><a href="http://www.americanregent.com/" target="_blank">American Regent Inc.</a></td>
 <td>NDC 0517-4010-01</td>
 <td>4mg/L in PBS for Vasodilation Buffer (with Adenosine)</td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td><a href="http://www.sigmaaldrich.com/" target="_blank">Sigma</a></td>
 <td>P6148</td>
 <td>Prepared as 4% solution</td>
 </tr>
 <tr>
 <td>Perfusion Apparatus</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>See figure 2</td>
 </tr>
 <tr>
 <td>Spring Scissors</td>
 <td><a href="http://www.finescience.com/" target="_blank">Fine Science Tools</a></td>
 <td>15018-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Xylazine (Anased), 20 mg/gl stock</td>
 <td><a href="http://www.lloydlab.com/" target="_blank">Lloyd Labs</a></td>
 <td>NADA #139-236</td>
 <td>Mixed as 8.8 mg/kg body weight, with Ketamin in 0.9% saline</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Retrograde Perfusion and Filling of Mouse Coronary Vasculature as Preparation for Micro Computed Tomography Imaging at JoVE.com

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

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 ...

retrograde-perfusion-filling-mouse-coronary-vasculature-as

Research

JoVE Journal

Medicine

32.4K visualizações.  University of Washington. Visualização dos vasos coronários é fundamental para avançar na compreensão das doenças cardiovasculares. Descrevemos aqui um método para a perfusão vasculatura coronária murino com uma borracha de silicone radiopaco (Microfil), em preparação para imagiologia Tomografia micro-computadorizada (μCT).

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

Segmentation and Measurement of Fat Volumes in Murine Obesity Models Using X-ray Computed Tomography

Obesity is associated with increased morbidity and mortality as well as reduced metrics in quality of life.1 Both environmental and genetic factors are associated with obesity, though the precise underlying mechanisms that contribute to the disease are currently being delineated.2,3 Several small animal models of obesity have been developed and are employed in a variety of studies.4 A critical component to these experiments involves the collection of regional and/or total animal fat content data under varied conditions.
Traditional experimental methods available for measuring fat content in small animal models of obesity include invasive (e.g. ex vivo measurement of fat deposits) and non-invasive (e.g. Dual Energy X-ray Absorptiometry (DEXA), or Magnetic Resonance (MR)) protocols, each of which presents relative trade-offs. Current invasive methods for measuring fat content may provide details for organ and region specific fat distribution, but sacrificing the subjects will preclude longitudinal assessments. Conversely, current non-invasive strategies provide limited details for organ and region specific fat distribution, but enable valuable longitudinal assessment. With the advent of dedicated small animal X-ray computed tomography (CT) systems and customized analytical procedures, both organ and region specific analysis of fat distribution and longitudinal profiling may be possible. Recent reports have validated the use of CT for in vivo longitudinal imaging of adiposity in living mice.5,6 Here we provide a modified method that allows for fat/total volume measurement, analysis and visualization utilizing the Carestream Molecular Imaging Albira CT system in conjunction with PMOD and Volview software packages.

Fat content analysis is routinely conducted in studies utilizing murine obesity models. Emerging methods in small animal CT imaging and analysis are providing for longitudinal detail rich fat content analysis. Here we detail step by step procedures for performing small animal CT imaging, analysis, and visualization.

Obesity is associated with increased morbidity and mortality as well as reduced metrics in quality of life.1 Both environmental and genetic factors are ...

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.

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 ...

Assessment of Viability of Human Fat Injection into Nude Mice with Micro-Computed Tomography

Lipotransfer is a vital tool in the surgeon&#8217;s armamentarium for the treatment of soft tissue deficits of throughout the body. Fat is the ideal soft tissue filler as it is readily available, easily obtained, inexpensive, and inherently biocompatible.1 However, despite its burgeoning popularity, fat grafting is hampered by unpredictable results and variable graft survival, with published retention rates ranging anywhere from 10-80%. 1-3
To facilitate investigations on fat grafting, we have therefore developed an animal model that allows for real-time analysis of injected fat volume retention. Briefly, a small cut is made in the scalp of a CD-1 nude mouse and 200-400 &#181;l&#160;of processed lipoaspirate is placed over the skull. The scalp is chosen as the recipient site because of its absence of native subcutaneous fat, and because of the excellent background contrast provided by the calvarium, which aids in the analysis process. Micro-computed tomography (micro-CT) is used to scan the graft at baseline and every two weeks thereafter. The CT images are reconstructed, and an imaging software is used to quantify graft volumes.
Traditionally, techniques to assess fat graft volume have necessitated euthanizing the study animal to provide just a single assessment of graft weight and volume by physical measurement ex vivo. Biochemical and histological comparisons have likewise required the study animal to be euthanized. This described imaging technique offers the advantage of visualizing and objectively quantifying volume at multiple time points after initial grafting without having to sacrifice the study animal. The technique is limited by the size of the graft able to be injected as larger grafts risk skin and fat necrosis. This method has utility for all studies evaluating fat graft viability and volume retention. It is particularly well-suited to providing a visual representation of fat grafts and following changes in volume over time.

Fat grafting is an essential technique for reconstructing soft tissue deficits. However, it remains an unpredictable procedure characterized by variable graft survival. Our goal was to devise a mouse model that utilizes a novel imaging method to compare volume retention between differing techniques of fat graft preparation and delivery.

Lipotransfer is a vital tool in the surgeon&#8217;s armamentarium for the treatment of soft tissue deficits of throughout the body. Fat is the ideal soft ...

Using Micro-computed Tomography for the Assessment of Tumor Development and Follow-up of Response to Treatment in a Mouse Model of Lung Cancer

Lung cancer is the most lethal cancer in the world. Intensive research is ongoing worldwide to identify new therapies for lung cancer. Several mouse models of lung cancer are being used to study the mechanism of cancer development and to experiment with various therapeutic strategies. However, the absence of a real-time technique to identify the development of tumor nodules in mice lungs and to monitor the changes in their size in response to various experimental and therapeutic interventions hampers the ability to obtain an accurate description of the course of the disease and its timely response to treatments. In this study, a method using a micro-computed tomography (CT) scanner for the detection of the development of lung tumors in a mouse model of lung adenocarcinoma is described. Next, we show that monthly follow-up with micro-CT can identify dynamic changes in the lung tumor, such as the appearance of additional nodules, increase in the size of previously detected nodules, and decrease in the size or complete resolution of nodules in response to treatment. Finally, the accuracy of this real-time assessment method was confirmed with end-point histological quantification. This technique paves the way for planning and conducting more complex experiments on lung cancer animal models, and it enables us to better understand the mechanisms of carcinogenesis and the effects of different treatment modalities while saving time and resources.

We describe a method for the detection of tumor nodule development in the lungs of an adenocarcinoma mouse model using micro-computed tomography and its use for monitoring changes in nodule size over time and in response to treatment. The accuracy of the assessment was confirmed with end-point histological quantification.

Lung cancer is the most lethal cancer in the world. Intensive research is ongoing worldwide to identify new therapies for lung cancer. Several mouse ...

Combined Near-infrared Fluorescent Imaging and Micro-computed Tomography for Directly Visualizing Cerebral Thromboemboli

Direct thrombus imaging visualizes the root cause of thromboembolic infarction. Being able to image thrombus directly allows far better investigation of stroke than relying on indirect measurements, and will be a potent and robust vascular research tool. We use an optical imaging approach that labels thrombi with a molecular imaging thrombus marker&#160;&#8212; a Cy5.5 near-infrared fluorescent (NIRF) probe that is covalently linked to the fibrin strands of the thrombus by the fibrin-crosslinking enzymatic action of activated coagulation factor XIIIa during the process of clot maturation. A micro-computed tomography (microCT)-based approach uses thrombus-seeking gold nanoparticles (AuNPs) functionalized to target the major component of the clot: fibrin. This paper describes a detailed protocol for the combined in vivo microCT and ex vivo NIRF imaging of thromboemboli in a mouse model of embolic stroke. We show that in vivo microCT and fibrin-targeted glycol-chitosan AuNPs (fib-GC-AuNPs) can be used for visualizing both in situ thrombi and cerebral embolic thrombi. We also describe the use of in vivo microCT-based direct thrombus imaging to serially monitor the therapeutic effects of tissue plasminogen activator-mediated thrombolysis. After the last imaging session, we demonstrate by ex vivo NIRF imaging the extent and the distribution of residual thromboemboli in the brain. Finally, we describe quantitative image analyses of microCT and NIRF imaging data. The combined technique of direct thrombus imaging allows two independent methods of thrombus visualization to be compared: the area of thrombus-related fluorescent signal on ex vivo NIRF imaging vs. the volume of hyperdense microCT thrombi in vivo.

This protocol describes the application of combined near-infrared fluorescent (NIRF) imaging and micro-computed tomography (microCT) for visualizing cerebral thromboemboli. This technique allows the quantification of thrombus burden and evolution. The NIRF imaging technique visualizes fluorescently labeled thrombus in excised brain, while the microCT technique visualizes thrombus inside living animals using gold-nanoparticles.

Direct thrombus imaging visualizes the root cause of thromboembolic infarction. Being able to image thrombus directly allows far better investigation of ...

Identifying Coronary Artery Calcification on Non-gated Computed Tomography Scans

Coronary artery calcification (CAC) provides an objective measure of coronary artery disease and can readily be identified on non-gated computed tomography (CT) scans with a high correlation with gated cardiac CT scans. This standardized protocol takes a step-wise approach to not only optimizing an image for the identification of calcification but also to distinguishing CAC from other common causes of calcification in the cardiac silhouette. Recognition of CAC on non-gated CT scans helps to identify a very powerful prognostic factor that can influence therapeutic interventions or downstream diagnostic testing without requiring a gated cardiac scan. These non-gated CT scans are often acquired as part of the routine care of the patient, and this data is readily available without another dose of&#160;ionizing radiation. This protocol allows for the precise and accurate extraction of this data for the purposes of retrospective data analysis in clinical research studies, but also in the clinical evaluation and management of patients.

Here, we present a protocol to reliably and systematically identify coronary artery calcification (CAC) on non-gated computed tomography (CT) scans of the chest or abdomen. CAC provides an objective measure of coronary artery disease for both research and clinical purposes.

Coronary artery calcification (CAC) provides an objective measure of coronary artery disease and can readily be identified on non-gated computed ...

Continuous Blood Sampling in Small Animal Positron Emission Tomography/Computed Tomography Enables the Measurement of the Arterial Input Function

For quantitative analysis and bio-kinetic modeling of positron emission tomography/computed tomography (PET/CT) data, the determination of the temporal blood time-activity concentration also known as arterial input function (AIF) is a key point, especially for the characterization of animal disease models and the introduction of newly developed radiotracers. The knowledge of radiotracer availability in the blood helps to interpret PET/CT-derived data of tissue activity. For this purpose, online blood sampling during the PET/CT imaging is advisable to measure the AIF. In contrast to manual blood sampling and image-derived approaches, continuous online blood sampling has several advantages. Besides the minimized blood loss, there is an improved resolution and a superior accuracy for the blood activity measurement. However, the major drawback of online blood sampling is the costly and time-consuming preparation to catheterize the femoral vessels of the animal. Here, we describe an easy and complete workflow for catheterization and continuous blood sampling during small animal PET/CT imaging and compared it to manual blood sampling and an image-derived approach. Using this highly-standardized workflow, the determination of the fluorodeoxyglucose ([18F]FDG) AIF is demonstrated. Further, this procedure can be applied to any radiotracer in combination with different animal models to create fundamental knowledge of tracer kinetic and model characteristics. This allows a more precise evaluation of the behavior of pharmaceuticals, both for diagnostic and therapeutic approaches in the preclinical research of oncological, neurodegenerative and myocardial diseases.

Here a protocol for continuous blood sampling during PET/CT imaging of rats to measure the arterial input function (AIF) is described. The catheterization, the calibration and setup of the system and the data analysis of the blood radioactivity are demonstrated. The generated data provide input parameters for subsequent bio-kinetic modeling.

For quantitative analysis and bio-kinetic modeling of positron emission tomography/computed tomography (PET/CT) data, the determination of the temporal ...

Tissue Preparation Techniques for Contrast-Enhanced Micro Computed Tomography Imaging of Large Mammalian Cardiac Models with Chronic Disease

Structural remodeling is a common consequence of chronic pathological stresses imposed on the heart. Understanding the architectural and compositional properties of diseased tissue is critical to determine their interactions with arrhythmic behavior. Microscale tissue remodeling, below the clinical resolution, is emerging as an important source of lethal arrhythmia, with high prevalence in young adults. Challenges remain in obtaining high imaging contrast at sufficient microscale resolution for preclinical models, such as large mammalian whole hearts. Moreover, tissue composition-selective contrast enhancement for three-dimensional high-resolution imaging is still lacking. Non-destructive imaging using micro-computed tomography shows promise for high-resolution imaging. The objective was to alleviate sufferance from X-ray over attenuation in large biological samples. Hearts were extracted from healthy pigs (N = 2), and sheep (N = 2) with either induced chronic myocardial infarction and fibrotic scar formation or induced chronic atrial fibrillation. Excised hearts were perfused with: a saline solution supplemented with a calcium ion quenching agent and a vasodilator, ethanol in serial dehydration, and hexamethyldisilizane under vacuum. The latter reinforced the heart structure during air-drying for 1 week. Collagen-dominant tissue was selectively bound by an X-ray contrast-enhancing agent, phosphomolybdic acid. Tissue conformation was stable in air, permitting long-duration microcomputed tomography acquisitions to obtain high-resolution (isotropic 20.7 &#181;m) images. Optimal contrast agent loading by diffusion showed selective contrast enhancement of the epithelial layer and sub-endocardial Purkinje fibers in healthy pig ventricles. Atrial fibrillation (AF) hearts showed enhanced contrast accumulation in the posterior walls and appendages of the atria, attributed to greater collagen content. Myocardial infarction hearts showed increased contrast selectively in regions of cardiac fibrosis, which enabled the identification of interweaving surviving myocardial muscle fibers. Contrast-enhanced air-dried tissue preparations enabled microscale imaging of the intact large mammalian heart and selective contrast enhancement of underlying disease constituents.

Here, we present a protocol to obtain high resolution micro computed tomography images of healthy and pathological large mammalian whole hearts with collagen-selective contrast enhancement.

Structural remodeling is a common consequence of chronic pathological stresses imposed on the heart. Understanding the architectural and compositional ...

DUCT: Double Resin Casting followed by Micro-Computed Tomography for 3D Liver Analysis

The liver is the biggest internal organ in humans and mice, and high auto-fluorescence presents a significant challenge for assessing the three-dimensional (3D) architecture of the organ at the whole-organ level. Liver architecture is characterized by multiple branching lumenized structures, which can be filled with resin, including vascular and biliary trees, establishing a highly stereotyped pattern in the otherwise hepatocyte-rich parenchyma. This protocol describes the pipeline for performing double resin casting micro-computed tomography, or &#34;DUCT&#34;. DUCT entails injecting the portal vein and common bile duct with two different radiopaque synthetic resins, followed by tissue fixation. Quality control by clearing one lobe, or the entire liver, with an optical clearing agent, allows for pre-screening of suitably injected samples. In the second part of the DUCT pipeline, a lobe or the whole liver can be used for micro-computed tomography (microCT) scanning, (semi-)automated segmentation, and 3D rendering of the portal venous and biliary networks. MicroCT results in 3D coordinate data for the two resins allowing for qualitative as well as quantitative analysis of the two systems and their spatial relationship. DUCT can be applied to postnatal and adult mouse liver and can be further extended to other tubular networks, for example, vascular networks and airways in the lungs.

Double resin casting micro-computed tomography, or DUCT, enables visualization, digitalization, and segmentation of two tubular systems simultaneously to facilitate 3D analysis of organ architecture. DUCT combines ex vivo injection of two radiopaque resins followed by micro-computed tomography scanning and segmentation of the tomographic data.

The liver is the biggest internal organ in humans and mice, and high auto-fluorescence presents a significant challenge for assessing the ...

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 ...