This research was supported in part by the NHLBI Intramural Research Program (DIR HL-006247). We would like to thank the NIH Mouse Imaging Facility for guidance in image acquisition and analysis.

All methods described here have been approved by the Institutional Animal Care and Use Committee (ACUC) of the National Heart Lung and Blood Institute.
1. Preparation
<ol>
	<li>Inject the mouse intraperitoneally with heparin (1 unit/g mouse body weight) and allow it to ambulate for 2 min.</li>
	<li>Euthanize the animal in a CO2 chamber.</li>
	<li>Arrange the mouse in a supine position on a surgical board and secure all four limbs to the board with tape. Use magnification for fine dissection.</li>
</ol>
2. Exposing lungs and trachea
<ol>
	<li>Spray the ventral side of the mouse with 70% ethanol to minimize hair interference.</li>
	<li>Grasp the abdominal skin with forceps and make a small incision with scissors in the umbilical region. Slide the tips of scissors into the fascial layer between the abdominal musculature and skin and begin separating the two layers. Work rostrally, removing the skin from the abdomen, ribcage, and neck.</li>
	<li>Open the abdominal musculature with scissors and cut laterally on both sides until the diaphragm is exposed.</li>
	<li>Gently grasp the xiphoid process and slightly lift the ribcage maximizing the view of the caudal lungs through the thin, semitransparent diaphragm. Carefully make a small incision in the diaphragm just beneath the xiphoid process. The lungs will collapse and retract away from the diaphragm. Dissect the diaphragm away from the ribcage, taking care not to nick the lung parenchyma.</li>
	<li>Locate and sever the inferior vena cava (IVC) and esophagus where they pass through the diaphragm. Use gauze to clean up any pooling blood in the thoracic cavity, avoiding contact with the lungs.</li>
	<li>Grasp the xiphoid once again and gently lift. Cut the ribcage bilaterally (roughly at the midaxillary line) avoiding contact with the lungs. Remove the anterior ribcage entirely, making the final cut along the sternal angle just before the manubrium.</li>
	<li>Using a prefilled syringe, liberally wet the lungs with phosphate-buffered saline (PBS, pH 7.4) to prevent drying out. Continue this routine throughout the procedure.</li>
	<li>Using forceps, grasp the manubrium and gently elevate away from the body. Using scissors, cut 1-2 mm lateral to the manubrium, severing clavicles, and remove. This will expose the thymus underneath.</li>
	<li>Grasp each lobe of the thymus, pull apart, and remove. Repeat this procedure with the submandibular gland. Finally, remove the muscular tissue overlaying the trachea. 
	NOTE: Following the dissection, the heart, ascending aorta (AA), pulmonary arterial trunk (PAT), and trachea should be visible. Ensure the primary arterial branches off the trunk are not divided or injured.</li>
</ol>
3. PA catheterization and blood perfusion
<ol>
	<li>To assemble Unit 1, thread 15 cm of PE-10 tubing onto the hub of a 30 G needle and attach to a 1 mL syringe prefilled with 10-4 M sodium nitroprusside (SNP) in PBS. Prime the tubing by advancing the plunger until all the air is purged from this unit ( Figure 1). 
	CAUTION: SNP is toxic if swallowed. Avoid contact with the skin and eyes. Wash skin thoroughly after handling. Wear appropriate personal protective equipment.
	<ol>
		<li>Alternatively, assemble Unit 2. For mice postnatal day 7 (P7) and younger, use a hemostat to detach an additional 30 G needle from its hub and thread the needle onto the open end of the tubing of Unit 1 (Figure 1).</li>
	</ol>
	</li>
	<li>Instead of a needle, use curved sharp forceps to grasp one end of a 10 cm length of 7-0 silk. Penetrate the apex of the heart entering from one side and passing the tips of the forceps through the muscle and out of the other side. Grasp the silk with another set of forceps and pull approximately a 2 cm length through and tie off. Take the remaining 8 cm end of the suture, tugging the heart caudally, and tape the end to the surgical board. 
	NOTE: This will create tension, further exposing the great vessels and tethering the heart in place, allowing for easier placement of the catheter in the pulmonary artery.</li>
	<li>Hook the tips of curved forceps under both the AA and PAT. Pull a 3 cm length of 7-0 silk back through the opening and create a single-throw loose suture.</li>
	<li>Using scissors make a 1-2 mm incision toward the apex of the heart, penetrating the thin-walled right ventricle (RV), to allow for the insertion of the catheter (Unit 1). Prior to the insertion, confirm there is no air in the system. Introduce the primed tubing into the right ventricle and gently advance into the semitransparent thin-walled PAT.
	<ol>
		<li>Visually verify that the catheter has not advanced into either the left or right pulmonary branches and does not abut the pulmonary artery branchpoint. Using tape, secure the distal portion of the tubing to the surgical board. 
		NOTE: To identify the RV, use forceps to pinch the right side of the heart. Unlike the left ventricle, the relatively thin free wall of the RV should be easily grasped.</li>
		<li>For mice younger than P7, attach Unit 2 to a micromanipulator and introduce the needle end of the unit into the PAT as described above using the manipulator.</li>
	</ol>
	</li>
	<li>Gently tighten the loose suture around both great vessels and cut the 8 cm length of suture created in step 3.2 to return the heart to a natural resting position. The catheter is now firmly secured within the PAT.</li>
	<li>Clip the left auricle of the heart to allow perfusate to exit the system.</li>
	<li>Secure SNP-containing syringe (Unit 1 or Unit 2, size dependent) in the syringe pump and perfuse the solution at a rate of 0.05 mL/min to flush the blood and maximally dilate the vasculature. Blood/perfusate will exit via the clipped auricle. Continue perfusion until perfusate runs clear (~200 &#181;L in an adult mouse, less for younger animals). 
	NOTE: When perfusing the low viscosity PBS/SNP, a relatively higher infusion rate was used in the interest of saving time. The more viscous polymer compound is infused at a slower rate to prevent overfilling, rupture, and maximize control over distal endpoints.</li>
</ol>
4. Tracheostomy and lung inflation
<ol>
	<li>Construct the lung inflation unit (Figure 2).
	<ol>
		<li>Connect a flexible plastic 24 G intravenous (IV) catheter (needle removed)/butterfly infusion set to a stopcock, attached to an open 50 mL syringe (no plunger). Hang the syringe from a ring stand.</li>
		<li>Add 10% buffered formalin to the syringe. Open the stopcock, allowing formalin to enter the tubing and purge all air from the system. Close the stopcock and raise the syringe until the meniscus is 20 cm above the trachea8. 
		CAUTION: Formalin is flammable, carcinogenic, acutely toxic when ingested, and causes skin irritation, serious eye damage, skin sensitization, and germ cell mutagenicity. Avoid ingestion and contact with skin and eyes. Avoid inhalation of the vapor or mist. Keep away from sources of ignition. Wear appropriate personal protective equipment.</li>
	</ol>
	</li>
	<li>Place two loose sutures inferior to the cricoid cartilage 2-4 mm apart.</li>
	<li>Using scissors, make a small incision in the cricothyroid ligament superior to the sutures.</li>
	<li>Insert the IV catheter into the opening and advance the tip beyond the two loose sutures.</li>
	<li>Tighten the sutures around the trachea and open the stopcock. Allow the formalin to enter the lungs by gravity and wait for 5 min for the lungs to fully inflate. If the lungs adhere to the ribcage during inflation, grasp the outside of the ribcage with blunt tipped forceps and move in all directions to assist in freeing the lobes. Do not make direct contact with lungs.</li>
	<li>After 5 min, back the IV catheter beyond the first suture and ligate. Repeat for the second suture. The lungs are now inflated in a closed, pressurized state.</li>
</ol>
5. Casting the vasculature
<ol>
	<li>&#160;In a 1.5 mL tube, prepare 1 mL of an 8:1:1 solution8 of polymer:diluent:curing agent and gently invert several times to ensure good mixing.</li>
	<li>Remove the plunger from a 1 cc syringe, cover the opposite end with a gloved finger, and pour the polymer compound into the syringe. Carefully reinsert the plunger, invert, and advance the plunger to remove all air and form a meniscus at the tip of the syringe.</li>
	<li>Remove the SNP/PBS syringe from the hub of the needle and drip additional PBS into the hub to create a meniscus. Carefully check the hub for trapped air, dislodge if necessary, and reform the meniscus. Join the hub to the syringe filled with the polymer compound. 
	NOTE: Creating a meniscus on both ends significantly reduces the chance for air to enter the system.</li>
	<li>Attach the polymer compound filled syringe to the syringe pump and infuse at 0.02 mL/min. 
	NOTE: For smaller lungs, a slower rate can be helpful to prevent overfilling but, is not essential.</li>
	<li>Monitor the compound as it freely moves down the PE tubing and note the syringe volume as it enters the PAT. Continue filling until all lobes are filled completely down to the capillary level and stop the syringe pump. Check the syringe volume again. 
	NOTE: After several runs an estimated volume can be used to gauge an approximate endpoint (~35 &#956;L for an adult mouse and ~5 &#181;L for a P1 pup). After the pump is halted, the residual pressure in the system will continue to push the polymer compound into the pulmonary arteries. All lung lobes should fill at a similar rate.</li>
	<li>Cover the lungs with a fiber optic cleaning wipe, liberally apply PBS, and allow the carcass to sit undisturbed for 30-40 min at room temperature. During this period, the polymer compound will cure and harden.</li>
	<li>Remove the catheter, sever the arms/lower half of the mouse, and place the head/thorax into a 50 mL conical filled with 10% buffered formalin overnight.</li>
	<li>After fixation, grasp the trachea and gently separate the heart/lung unit from the remaining rib cage and thorax. Place the heart/lung block in a formalin filled scintillation vial. Discard the rest.</li>
</ol>
6. Alternative vascular beds for casting (Table 1)
NOTE: Each target vascular bed may require different catheter placements, infusion rates, and optimal filling times. Thus, multiple animals will be necessary to cast multiple organs.
<ol>
	<li>For systemic vascular beds superior or inferior to the diaphragm follow steps 1.1-2.5 as above. See additional notes on the portal system and diaphragm (Table 1).</li>
	<li>Grasp the xiphoid process with a hemostat and cut the ribcage bilaterally (roughly in the midaxillary line) just prior to the internal thoracic arteries.</li>
	<li>Fold the still connected ribcage over such that it is resting on the animal&#39;s neck/head, fully exposing the chest cavity.</li>
	<li>Follow step 3.1 above, then remove the lungs. Once the thoracic aorta (TA) is visible, hook the tips of curved forceps underneath it, ~10 mm superior to the diaphragm. Grasp a 3 cm length of 7-0 silk, pull back through the opening under the TA, and create a single-throw loose suture. Repeat this procedure ~8 mm above the diaphragm.</li>
	<li>For structures superior to the diaphragm, use spring scissors to create a small hole (~30% of the total circumference) on the ventral portion of the TA, ~2 mm inferior to the loose sutures placed in step 6.4.
	<ol>
		<li>For structures inferior to the diaphragm, instead, create a small hole ~2 mm superior to the loose sutures.</li>
	</ol>
	</li>
	<li>Depending on the animal size, introduce Unit 1 or 2 into the vessel, advance beyond loose sutures, and gently ligate the vessel.</li>
	<li>Follow step 3.7, setting the syringe pump at a rate of 1.0 mL/min and perfusing a minimum of 5 mL. Perfusate will exit via the IVC.</li>
	<li>Follow steps 5.1 - 5.4 adjusting the infusion rate to 0.05 mL/min, visually monitoring the target tissue in real-time. 
	NOTE: Infusion volume will be organ and animal age specific. The volume can be further limited by ligating arterial branches leading to non-target vascular beds (i.e., brain, liver, kidney, intestine).</li>
	<li>Follow 5.6 then remove target tissue and place in formalin.</li>
</ol>
7. Sample mount, scan, and reconstruction for micro-CT
<ol>
	<li>Using paraffin film, create a flat surface on the scanning bed and center the wet sample on this surface (Figure 3A). 
	NOTE: If motion artifact is detected, the sample may require further stabilization.</li>
	<li>Lightly tent/cover sample with additional paraffin film to prevent dehydration. Take special care not to rest the paraffin film on the sample causing deformation to the tissue (Figure 3B).</li>
	<li>Scan the sample using settings outlined in Table 2 and standardize these parameters within a given experiment. 
	NOTE: This is experiment/endpoint dependent. Standardize the chosen parameters for the ease of comparison between samples.</li>
	<li>Transfer the reconstructed scans for post-processing and analysis.</li>
</ol>

The authors have nothing to disclose

Executed properly, this method yields striking images of pulmonary arterial networks, allowing for comparison and experimentation in rodent models. Several critical steps along the way ensure success. First, investigators must heparinize the animal in the preparatory stage to prevent blood clots from forming in the pulmonary vasculature and chambers of the heart. This allows for the complete arterial transit of polymer compound. Second, when puncturing the diaphragm and removing the ribcage, take care to protect the lungs from inadvertent damage, cuts or injury. Any leak in the airway will prevent complete inflation and render comparisons between samples inaccurate. Third, tethering the heart at the apex aids catheter placement. Fourth, the use of a strong vasodilator such as SNP will assist in both the removal of blood and complete filling of arterioles and capillaries5,8. Fifth, when placing the catheter into the PAT, take care not to bury the tip into the bifurcation. This will cause an imbalance in flow, shunting polymer compound to either the left or right side, yielding an unequal pressure gradient. Sixth, the use of a syringe pump will allow the user to control the rate and titer the volume to both mouse strain and age. Lastly, leave the heart/lungs attached to the remainder of the thoracic cavity, fix overnight, and remove the following day. The lungs will be well fixed and the potential for deflation due to accidental nicks during separation will be minimized.
While this methodology achieves the desired results, alternative techniques may be helpful to some users. To aid in the placement of the catheter, a micromanipulator may be employed. We chose a version with a small profile and magnetic base to minimize encroachment in an already limited working area while providing a stable base (if using a magnetic base make sure to place a steel plate under the working space to allow the magnet to engage). This allows the user to precisely place the tip of the catheter in the PAT at an angle that follows the natural trajectory of the artery. Additionally, the catheter is secure and at less risk of being dislodged. Another option is the use of a trumpeted catheter tip8. While not trivial to create, a trumpeted catheter is far more secure and less inclined to accidentally slide out of the PAT. Changing the ratio of polymer:diluent alters the viscosity and the ease with which small vessels are filled. Depending on the target vasculature and experimental endpoints this can be a valuable consideration. Euthanasia via CO2 may cause pulmonary hemorrhage in a small percentage of animals and is strain dependent22. Consider an alternative euthanasia protocol should this impact experimental endpoints. When inflating the lungs, the use of formalin aids fixation of the organ in place at the given pressure. A physiologically neutral buffer can be substituted should peripheral vessels need to be filled in an unfixed state. If infusion rate and control are of less importance to a given experiment, perfusion by hand is also possible. Hand injection requires practice and real-time monitoring under magnification to avoid overfilling or vessel rupture8. Finally, the tissue mount/conditions, scanning parameters, and minimal post-processing we employed for this paper should serve merely as a starting point. Different scanners, tissues, experimental endpoints/user needs may demand alternative parameters.
While the vascular images generated from this technique are impressive, there are limitations. Primarily, the above method is suboptimal for measuring vascular caliber due to the inability to monitor and control intravascular pressure during the infusion. Other groups have managed to somewhat address these pressure concerns in systemic vasculature by monitoring driving pressure4,23, however, such concerns are further amplified on the pulmonary side due to the relatively thin pulmonary artery walls that are easily distensible with small changes in pressures24 and the inability to precisely measure and statically control pulmonary intravascular pressure.
A second limitation to this method is that it remains a postmortem, single timepoint experiment, limiting its utility in studies that require truly physiologic conditions or a time course. Other, live animal measures, such as CT pulmonary angiography (CTPA) or contrast-enhanced &#181;CT (CE-CT) offer the possibility of functional and morphologic measures. Repeated scans/longitudinal studies as well as measurements at different points in the cardiac/pulmonary cycle, can be explored10,25,26,27,28. These methods can be reliably used, in addition to echocardiography, to measure the arterial caliber. However, both CTPA and echocardiography measures are currently limited to the assessment of the proximal vasculature. For echocardiogram, the assessment is limited to the pulmonary trunk while CTPA allows adequate calculation of the branch pulmonary artery caliber potentially 1-2 orders further, but resolution is limited, obscuring distal portions of the vasculature7. Radiation dosage is also a concern that should be carefully monitored when using CT especially in multi-scan longitudinal studies29,30. For either of these applications, &#181;CT equipment, scan time, and analysis software may be expensive and require specialized staff training. Animal imaging core facilities at some institutions may ease this burden.
As an alternative to this compound, some groups utilize traditional corrosion casting techniques accompanied by soft tissue removal31,32. These methods yield results similar to this polymer compound, but the end product is brittle, leading to potential artifact15. In addition, the removal of soft tissue eliminates the potential for future histology33. Another option is to leave the soft tissue intact and perform a follow-up step wherein the soft tissue is &#34;cleared&#34; rendering the sample virtually transparent34,35. Tissue clearing gives the user some ability to see deeper within a sample but, on the whole, remains inferior to &#181;CT as it cannot provide the same 3D visualization. Serial histologic sectioning and array tomography are methods that offer exceptionally high resolution. While this technique opens the door to exciting new possibilities, the workload is exponentially higher and not particularly conducive to large cohorts11,12. 3D x-ray histology is a non-destructive approach that couples both &#181;CT and traditional histology or even EM36,37,38. It takes a more high level view of pathology by utilizing &#181;CT to globally identify and accurately scout regions of interest that are then followed up with routine histology39. Substituting lower resolution contrast agents (or in some cases no contrast) with polymer compound into the vasculature might serve to elevate both techniques when possible. Another non-destructive approach that is computationally intensive yet, potentially enhances the contrast, is phase-retrieval &#181;CT imaging40,41. This method can be valuable when employed on noisy data where contrast is weak or not possible42. The polymer compound employed in this technique, however, does not suffer from this limitation. That said, phase-retrieval may be useful where the polymer compound is possibly diluted, for example in distal vasculature43. Finally, stereology has been a standard in lung quantitative structural analysis for years44. It uses random, systematic sampling on cross sections of tissue to make 3-D inferences assuming that the chosen samples are sufficiently representative. While a powerful tool, it has the potential to lead to error and bias. Combining CT imaging with stereology, however, holds great promise45.
The outlined method is relatively straightforward and with training a success rate of &#62;90% is achievable. Once mastered, it allows for the complete and reliable casting of lung vasculature. In fixative, tissue and polymer remain stable indefinitely for future scans, potential histology, or EM46,47. We&#39;ve shown that this technique can be used in animals as young as P1 through adulthood and believe embryonic casting, via the pulmonary artery, is within reach. It should be noted that this technique can be applied to virtually any other vascular bed by simply altering the catheter entry point and determining appropriate endpoints.

The focus of this technique is the visualization of pulmonary arterial architecture using a polymer-based compound in mice. While extensive work has been performed on systemic vascular beds such as brain, heart, and kidney1,2,3,4,5, less information is available regarding the preparation and filling of the pulmonary arterial network. The aim of this study, therefore, is to expand upon previous work6,7,8 and provide a detailed written and visual reference that investigators can easily follow to produce high-resolution images of the pulmonary arterial tree.
While numerous methods exist for labeling and imaging lung vasculature, such as magnetic resonance imaging, echocardiography, or CT angiography9,10, many of these modalities fail to adequately fill and/or capture the small vessels, limiting the scope of what can be studied. Methods such as serial sectioning and reconstruction provide high resolution but are time/labor-intensive11,12,13. Surrounding soft tissue integrity is compromised in traditional corrosion casting10,13,14,15,16. Even animal age and size become factors when attempting to introduce a catheter or, the resolution is lacking. The polymer injection technique, on the other hand, fills arteries to the capillary level and when combined with &#181;CT, allows for unparalleled resolution5. Samples from mouse lungs as young as postnatal day 14 have been successfully casted8 and processed in a matter of hours. These can be rescanned indefinitely, or even sent for histological preparation/electron microscopy (EM) without compromising the existing soft tissue17. The main limitations to this method are the upfront cost of CT equipment/software, challenges with accurately monitoring intravascular pressure, and the inability to acquire data longitudinally in the same animal.
This paper builds on existing work to further optimize the pulmonary artery injection technique and push age/size related boundaries down to postnatal day 1 (P1) to yield striking results. It is most useful for teams that want to study arterial vascular networks. Accordingly, we provide new guidance for catheter placement/stabilization, increased control over fill rate/volume, and highlight notable pitfalls for increased casting success. Resulting casts can then be used for future characterization and morphologic analysis. Perhaps more importantly, this is the first visual demonstration, to our knowledge, that walks the user through this intricate procedure.

<table><tbody><tr><td>1cc syringe</td><td>Becton Dickinson</td><td>309659</td><td></td></tr><tr><td>20ml Glass Scintillation Vials</td><td>Fisher</td><td>03-340-25P</td><td></td></tr><tr><td>30G Needle</td><td>Becton Dickinson</td><td>305106</td><td></td></tr><tr><td>50mL conical tubes</td><td>Cornin</td><td>352098</td><td>For sample Storage and scanning</td></tr><tr><td>60cc syringe</td><td>Becton Dickinson</td><td>309653</td><td></td></tr><tr><td>7-0 silk suture</td><td>Teleflex</td><td>103-S</td><td></td></tr><tr><td>Analyze 12.0 Software</td><td>AnalyzeDirect Inc.</td><td>N/A</td><td>Primary Software</td></tr><tr><td>Amira 6.7 Software</td><td>Thermo Scientific</td><td>N/A</td><td>Alternative Sofware</td></tr><tr><td>CeramaCut Scissors 9cm</td><td>Fine Science tools</td><td>14958-09</td><td></td></tr><tr><td>Ceramic Coated Curved Forceps</td><td>Fine Science tools</td><td>11272-50</td><td></td></tr><tr><td>CO2 Tank</td><td>Robert&#039;s Oxygen Co.</td><td>n/a</td><td></td></tr><tr><td>Dual syringe pump</td><td>Cole Parmer</td><td>EW-74900-10</td><td></td></tr><tr><td>Dumont Mini-Forceps</td><td>Fine Science tools</td><td>11200-14</td><td></td></tr><tr><td>Ethanol</td><td>Pharmco</td><td>111000200</td><td></td></tr><tr><td>Formalin</td><td>Sigma - Life Sciences</td><td>HT501128</td><td></td></tr><tr><td>Gauze</td><td>Covidien</td><td>441215</td><td></td></tr><tr><td>Hemostat</td><td>Fine Science tools</td><td>13013-14</td><td></td></tr><tr><td>Heparin (1000USP Units/ml)</td><td>Hospira</td><td>NDC 0409-2720-01</td><td></td></tr><tr><td>Horos Software</td><td>Horos Project</td><td>N/A</td><td>Alternative Sofware</td></tr><tr><td>induction chamber</td><td>n/a</td><td>n/a</td><td></td></tr><tr><td>Kimwipe</td><td>Fisher</td><td>06-666</td><td>fiber optic cleaning wipe</td></tr><tr><td>Labelling Tape</td><td>Fisher</td><td>15966</td><td></td></tr><tr><td>Magnetic Base</td><td>Kanetec</td><td>N/A</td><td></td></tr><tr><td>Micro-CT system</td><td>PerkinElmer</td><td>Quantum GX</td><td></td></tr><tr><td>Microfil (Polymer Compound)</td><td>Flowech Inc.</td><td>Kit B - MV-122</td><td>8 oz. of MV compound; 8 oz. of diluent; MV-Curing Agent</td></tr><tr><td>Micromanipulator</td><td>Stoelting</td><td>56131</td><td></td></tr><tr><td>Monoject 1/2 ml Insulin Syringe</td><td>Covidien</td><td>1188528012</td><td></td></tr><tr><td>Octagon Forceps Straight Teeth</td><td>Fine Science tools</td><td>11042-08</td><td></td></tr><tr><td>Parafilm</td><td>Bemis company, Inc.</td><td>#PM999</td><td></td></tr><tr><td>PE-10 tubing</td><td>Instech</td><td>BTPE-10</td><td></td></tr><tr><td>Phospahte buffered Saline</td><td>BioRad</td><td>#161-0780</td><td></td></tr><tr><td>Ring Stand</td><td>Fisher</td><td>S13747</td><td>Height 24in.</td></tr><tr><td>Sodium Nitroprusside</td><td>sigma</td><td>71778-25G</td><td></td></tr><tr><td>Steel Plate</td><td>N/A</td><td>N/A</td><td>16 x 16 in. area, 1/16 in thick</td></tr><tr><td>Straight Spring Scissors</td><td>Fine Science tools</td><td>15000-08</td><td></td></tr><tr><td>SURFLO 24G Teflon I.V. Catheter</td><td>Santa Cruz Biotechnology</td><td>360103</td><td></td></tr><tr><td>Surgical Board</td><td>Fisher</td><td>12-587-20</td><td>This is a converted slide holder</td></tr><tr><td>Universal 3-prong clamp</td><td>Fisher</td><td>S24280</td><td></td></tr><tr><td>Winged Inf. Set 25X3/4, 12&quot; Tubing</td><td>Nipro</td><td>PR25G19</td><td></td></tr><tr><td>Zeiss Stemi-508 Dissection Scope</td><td>Zeiss</td><td>n/a</td><td></td></tr></tbody></table>

vascular casting of adult and early postnatal mouse lungs for micro-ct imaging

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and behave by observing the surface of a tissue. This method provides a means to visualize the complex 3-dimensional vascular architecture of the lung.
To accomplish this, a catheter is inserted into the pulmonary artery and the vasculature is simultaneously flushed of blood and chemically dilated to limit resistance. Lungs are then inflated through the trachea at a standard pressure and the polymer compound is infused into the vascular bed at a standard flow rate. Once the entire arterial network is filled and allowed to cure, the lung vasculature may be visualized directly or imaged on a micro-CT (&#181;CT) scanner.
When performed successfully, one can appreciate the pulmonary arterial network in mice ranging from early postnatal ages to adults. Additionally, while demonstrated in the pulmonary arterial bed, this method can be applied to any vascular bed with optimized catheter placement and endpoints.

A successful cast will exhibit uniform filling of the entire pulmonary arterial network. We demonstrate this in C57Bl/6J mice ranging in age: Postnatal day P90 (Figure 4A), P30 (Figure 4B), P7 (Figure 4C), and P1 (Figure 4D). By controlling the rate of flow and visually monitoring the fill in real-time, reliable endpoints of the most distal vasculature were achieved (Figure 5A).
Common challenges include damage to the lungs, incomplete filling, underfilling, or overfilling, wedging the catheter, and animal size.
If there is damage to the lung/airway, small leaks will prevent the lungs from holding pressure (Figure 5B,C). In the absence of complete inflation, it becomes difficult to make accurate quantitative and spatial comparisons across samples. To minimize the risk to the lung parenchyma, avoid cutting too closely to the lungs when removing the ribcage and keep the lungs moist with PBS throughout the procedure to avoid dehydration and adherence to surrounding structures. If a lobe adheres to the rib cage during inflation, gently grasp the outside of the ribcage (away from the lung) with forceps and move it in a direction to free the lobes. Alternatively, a blunt instrument, such as a spatula, with a smooth edge can be used to lift or push the inflated lung away from the ribcage. When inflating the lungs, adhere to suggested pressure parameters and avoid over-inflation as this can lead to rupture of the airway. Finally, do not remove lungs from thoracic cavity until post-fixation is complete. The trachea, lungs, and heart should be removed en bloc from the remaining portions of the thoracic cavity.
Patchy (Figure 5D) or incomplete (Figure 5E) filling can arise from an &#34;airlock&#34;, in which air is introduced into the vascular system via the catheter, blocking downstream flow of the compound. To minimize the chance of an airlock, vigilantly purge air from the tip of the catheter prior to insertion (Step 3.4) and during the syringe transition from SNP/PBS to polymer compound. If the fill remains patchy or incomplete, it could be an indication of increased vascular resistance as a result of focal/long segment stenosis or tortuosity. Blood clots can also lead to incomplete filling and are easily avoided by using heparin prior to the procedure.
Improper injection volume will lead to underfilling or overfilling. Underfilling occurs when too little compound is introduced into the vasculature (Figure 5F). Alternatively, overfilling, or introducing too much polymer compound too rapidly can cause either arterial rupture (Figure 5G) or, more commonly, venous transit (Figure 5H). Both problems can be alleviated by using a syringe pump. Investigators should carefully adhere to the proposed rate and volume restrictions or establish their own rates based upon their specific model and optimization. Monitoring polymer compound perfusion in real time under magnification is critical, and filling of small arterioles/capillaries should be used as an endpoint.
Advancing the catheter too far down the pulmonary trunk can cause the tip to wedge into one pulmonary artery branch and create an imbalance in the flow. As a result, one side fills faster than the other (Figure 5I), which frequently leads to overfilling in one lung and underfilling in the other. While catheter wedging is the most likely reason in this scenario, &#34;airlock&#34; and lack of heparin can also be contributing factors.
Finally, smaller animals present their own set of additional obstacles. Younger animals demand steady hands and small mistakes are less forgiving. High quality instruments, specifically designed for microsurgery, become more important at early postnatal timepoints. Use of a micromanipulator assists greatly in not only placement but preventing catheter dislocation. It is also essential to utilize the syringe pump on small animals to accurately control and manage endpoints.
While specifically shown for the pulmonary vasculature, this procedure can easily be applied to systemic target vascular beds as well (Table 1). In addition to the challenges listed above, choosing the right entry point is crucial. Casting via the thoracic aorta produces excellent results for most vascular beds. It should be noted, however, that inserting the catheter as proximal to the target site as possible and ligating non-target vasculature assists in flow and volume control. These refinements combined with appropriate direct monitoring of distal vascular endpoints (Figure 6A-F) and standard infusion rates optimize the filling. Many examples of such casting methods exist in the literature and are too numerous for complete referencing. However, additional details may be found in organ specific text such as these4,5,7,18,19,20,21.
After casting, samples can be processed for &#181;CT scanning (Figure 7A,B). For post-processing, a commercial software package (see Table of Materials) produced a 3D volume rendering of the pulmonary vascular tree presented as still images (Figure 7C), or movies. Further statistical analyses exploring vascular characteristics such as segment length and number, tortuosity, order (generation or rank), volume, and arcade length can also be performed. In addition to &#181;CT scanning, the casted samples can also be cleared to obtain gross images or processed and cut for histological analysis8.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61242/61242fig1.jpg" /> 
Figure 1: Catheter and needle setup. Syringes are shown with attached tubing and needles (Unit1 and Unit2). Inset: closeup of needle and tubing. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61242/61242fig2.jpg" /> 
Figure 2: Lung inflation setup. Ring stand, clamp, a syringe filled with formalin, and tubing with a catheter attached. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61242/61242fig3.jpg" /> 
Figure 3: Micro-CT sample preparation pre-scan. (A) Here the specimen was centered on a paraffin film base, (B) Here the sample was centered and covered on the parafilm base. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61242/61242fig4.jpg" /> 
Figure 4: Vascular-casted lungs at varying developmental stages from 3 months to 1 day old. Dorsal view of lungs, (A) P90, (B) P30, (C) P7, and (D) P1 <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61242/61242fig5.jpg" /> 
Figure 5: Examples of ideal filling and common errors during polymer compound infusion. (A)&#160;When filling endpoint was reached, a robust and fine vascular network was observed. (B) Fully inflated formalin perfused lungs are represented by a white dashed line, (C) Underinflated/deflated lungs are shown. This was observed due to a compromised pulmonary airway. The original inflated position is represented by a white dashed line and the deflated position is represented by a black dotted line, (D) Patchy filling: the vasculature of portions of the lobe remains unfilled while other areas were entirely filled, (E) Incomplete filling: the polymer compound failed to penetrate entire sections of lung, (F) Underfilling: the polymer compound failed to fill distal vasculature, (G) Rupture: the arrow is pointing to the polymer compound extruded from vasculature, (H) Venous filling: note the arrow pointing to the arterial segments entirely filled and extending into the venous system. Veins and venules were of significantly larger caliber, (I) Catheter wedge: Here the catheter was shunted into one artery preventing the vasculature of the right lobes from filling completely while the left lobe was overfilled. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61242/61242fig6.jpg" /> 
Figure 6. Vascular casting and endpoints in additional organs. (A) Kidney: the punctate appearance of polymer compound in the glomerulus provided the endpoint. (B) Liver: note the small vessels visible at the edges of the organ. (C) Stomach: small vessels were visible and fully filled. (D) Large intestine: Small vessels are easily identifiable and filled. (E) Diaphragm: the muscle here is thin and translucent with small filled vessels apparent. (F). Brain: small vessels were visible in the cortex. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61242/61242fig7.jpg" /> 
Figure 7. CT images and 3D volume rendering of polymer compound&#160;filled lungs. (A) A single grey-scaled reconstructed lung slice, (B) This was a maximum intensity projection of a CT scan produced from polymer filled lungs, (C) A 3D volume rendering of the vascular arcade was generated using commercially available software (see Table of Materials). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61242/61242fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Target Arterial Vascular Bed</td>
			<td>Catheter placement&#160;</td>
			<td>Infusion direction</td>
			<td>Infusion rate</td>
			<td>Notes</td>
		</tr>
		<tr>
			<td>Brain</td>
			<td>Thoracic aorta pointing cranially</td>
			<td>Retrograde into the carotids</td>
			<td>.05ml/min</td>
			<td>Cannulate thoracic aorta, flip mouse to the prone position, open scalp, and visually monitor progress of polymer through skull.&#160;</td>
		</tr>
		<tr>
			<td>Diaphragm</td>
			<td>Left Ventrical</td>
			<td>Anterograde into internal thoracic, phrenic, and intercostal&#160;</td>
			<td>.05ml/min</td>
			<td>Open a window in the side of the ribcage, leaving the majority of the ribcage and the diaphragm intact.&#160; Cannulate left ventrical, clip right atrium, and monitor progress from the caudal side of the diaphragm.</td>
		</tr>
		<tr>
			<td>Upper limb musculature</td>
			<td>Thoracic aorta pointing cranially</td>
			<td>Retrograde into the brachiocephalic and left subclavian</td>
			<td>.02ml/min</td>
			<td>To optimize limb flow, tie off the carotid arteries and remove limb skin to allow visual monitoring of polymer transit into the limb musculature.</td>
		</tr>
		<tr>
			<td>Kidney</td>
			<td>Thoracic aorta pointing caudally</td>
			<td>Anterograde into renal arteries</td>
			<td>.05ml/min</td>
			<td>The internal vasculature is filled blindly.&#160; To avoid venous transit, stop injecting when polymer is visible in a uniform punctate pattern across kidney.</td>
		</tr>
		<tr>
			<td>Portal System</td>
			<td>&#160;Portal vein</td>
			<td>Anterograde into portal system</td>
			<td>.02ml/min</td>
			<td>Gently fold liver up to expose the portal vein.&#160;&#160;&#160;</td>
		</tr>
		<tr>
			<td>Hepatic&#160;</td>
			<td>Thoracic aorta pointing caudally</td>
			<td>Anterograde into the hepatic artery</td>
			<td>.05ml/min</td>
			<td>Tie off portal vein prior to infusion to avoid venous transit from gut flowing into liver.&#160;</td>
		</tr>
		<tr>
			<td>Stomach/ Intestine</td>
			<td>Thoracic aorta pointing caudally</td>
			<td>Anterograde into the celiac, superior mesenteric and/or inferior mesenteric</td>
			<td>.05ml/min</td>
			<td>Some regions of the gut are supplied by multiple arteries and may fill at different times.&#160; To avoid venous transit, tie off arteries not required for areas of interest and visually monitor the progress of the polymer.</td>
		</tr>
		<tr>
			<td>Intra-abdominal fat pads</td>
			<td>Thoracic aorta pointing caudally</td>
			<td>Anterograde but vessel depends on fat pad being studied</td>
			<td>.05ml/min</td>
			<td>Fat pads are supplied by multiple arteries and may fill at different times.&#160; To avoid venous transit, tie off arteries not required for precise area of interest and visually monitor the progress of the polymer.</td>
		</tr>
		<tr>
			<td>Lower limb musculature</td>
			<td>Infrarenal aorta pointing caudally</td>
			<td>Anterograde into the femoral arteries</td>
			<td>.02ml/min</td>
			<td>Remove limb skin to allow visual monitoring of polymer transit into the limb musculature.</td>
		</tr>
	</tbody>
</table>
Table 1. Casting alternative vascular beds.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">CT settings</td>
		</tr>
		<tr>
			<td>kVp</td>
			<td>90</td>
		</tr>
		<tr>
			<td>Target Material</td>
			<td>Tungsten</td>
		</tr>
		<tr>
			<td>Power</td>
			<td>8W</td>
		</tr>
		<tr>
			<td>Filtration</td>
			<td>Cu 0.06 mm + Al 0.5 mm</td>
		</tr>
		<tr>
			<td>Projection Number</td>
			<td>6424</td>
		</tr>
		<tr>
			<td>Detector Size</td>
			<td>Flat panel CMOS - 2944 x 2352 pixels</td>
		</tr>
		<tr>
			<td>Field of View (FOV)</td>
			<td>36 mm</td>
		</tr>
		<tr>
			<td>Voxel Size</td>
			<td>72 &#181;m</td>
		</tr>
		<tr>
			<td>Spatial Resolution</td>
			<td>voxel size x 1.5</td>
		</tr>
		<tr>
			<td>Acquisition Time</td>
			<td>14 min</td>
		</tr>
		<tr>
			<td>Reconstruction</td>
			<td>FBP and commercial algorithm</td>
		</tr>
		<tr>
			<td>Binning</td>
			<td>1x1</td>
		</tr>
	</tbody>
</table>
Table 2. &#181;CT Scanning Parameters.

Watch this Scientific Journal Video about Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging at JoVE.com

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

The aim of this technique is ex vivo visualization of pulmonary arterial networks of early postnatal and adult mice through lung inflation and injection of a radio-opaque polymer-based compound via the pulmonary artery. Potential applications for casted tissues are also discussed.

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and ...

vascular-casting-adult-early-postnatal-mouse-lungs-for-micro-ct

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

7.4K Views.  National Institutes of Health. Dit protocol geeft nauwkeurige, stapsgewijze richtlijnen voor het gieten van de longslagader vasculatuur in zowel volwassen als vroege postnatale muizen. Deze techniek stelt gebruikers in staat om het hele vasculaire netwerk volledig te casten voor visualisatie door een verscheidenheid aan methoden, waaronder micro-CT, met behoud van de integriteit van weke delen. Deze techniek is vooral handig voor het kijken naar long-vasculaire morfologie en architectuur, maar kan gemakkelijk worden aangep...

Video: Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography (&mu;CT).
High resolution &mu;CT is a very versatile yet accurate (1-2 microns of resolution) technique for 3D examination of ex-vivo biological samples1, 2. As opposed to electron tomography, the &mu;CT allows the examination of up to 4 cm thick samples. This technique requires only few hours of measurement as compared to weeks in histology. In addition, &mu;CT does not rely on 2D stereologic models, thus it may complement and in some cases can even replace histological methods3, 4, which are both time consuming and destructive. Sample conditioning and positioning in &mu;CT is straightforward and does not require high vacuum or low temperatures, which may adversely affect the structure. The sample is positioned and rotated 180&deg; or 360&deg;between a microfocused x-ray source and a detector, which includes a scintillator and an accurate CCD camera, For each angle a 2D image is taken, and then the entire volume is reconstructed using one of the different available algorithms5-7. The 3D resolution increases with the decrease of the rotation step. The present video protocol shows the main steps in preparation, immobilization and positioning of the sample followed by imaging at high resolution.

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography ( CT).

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized ...

Bioengineering

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 ...

Medicine

Cerebrovascular Casting of the Adult Mouse for 3D Imaging and Morphological Analysis

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). Animal models are necessary for studying the etiopathology and potential therapies of cerebrovascular diseases. Imaging the vasculature in large animals is relatively easy. However, developing vessel imaging methods of murine brain disease models is desirable due to the cost and availability of genetically-modified mouse lines. Imaging the murine cerebral vascular tree is a challenge. In humans and larger animals, the gold standard for assessing the angioarchitecture at the macrovascular (conductance) level is x-ray catheter contrast-based angiography, a method not suited for small rodents.
In this article, we present a method of cerebrovascular casting that produces a durable skeleton of the entire vascular bed, including arteries, veins, and capillaries that may be analyzed using many different modalities. Complete casting of the microvessels of the mouse cerebrovasculature can be difficult; however, these challenges are addressed in this step-by-step protocol. Through intracardial perfusion of the vascular casting material, all vessels of the body are casted. The brain can then be removed and clarified using the organic solvent methyl salicylate. Three dimensional imaging of the brain blood vessels can be visualized simply and inexpensively with any conventional brightfield microscope or dissecting microscope. The casted cerebrovasculature can also be imaged and quantified using micro-computed tomography (micro-CT)1. In addition, after being imaged, the casted brain can be embedded in paraffin for histological analysis.
The benefit of this vascular casting method as compared to other techniques is its broad adaptation to various analytic tools, including brightfield microscopic analysis, CT scanning due to the radiopaque characteristic of the material, as well as histological and immunohistochemical analysis. This efficient use of tissue can save animal usage and reduce costs. We have recently demonstrated application of this method to visualize the irregular blood vessels in a mouse model of adult BAVM at a microscopic level2, and provide additional images of the malformed vessels imaged by micro-CT scan. Although this method has drawbacks and may not be ideal for all types of analyses, it is a simple, practical technique that can be easily learned and widely applied to vascular casting of blood vessels throughout the body.

In this article, we present a simple, practical technique for cerebrovascular casting that is easy to perform and can be utilized to image the vascular tree of the adult mouse brain.

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). ...

Neuroscience

A Permanent Window for Investigating Cancer Metastasis to the Lung

Metastasis, accounting for ~90% of cancer-related mortality, involves the systemic spread of cancer cells from primary tumors to secondary sites such as the bone, brain, and lung. Although extensively studied, the mechanistic details of this process remain poorly understood. While common imaging modalities, including computed tomography (CT), positron emission tomography (PET), and magnetic resonance imaging (MRI), offer varying degrees of gross visualization, each lacks the temporal and spatial resolution necessary to detect the dynamics of individual tumor cells. To address this, numerous techniques have been described for intravital imaging of common metastatic sites. Of these sites, the lung has proven especially challenging to access for intravital imaging owing to its delicacy and critical role in sustaining life. Although several approaches have previously been described for single-cell intravital imaging of the intact lung, all involve highly invasive and terminal procedures, limiting the maximum possible imaging duration to 6-12 h. Described here is an improved technique for the permanent implantation of a minimally invasive thoracic optical Window for High-Resolution Imaging of the Lung (WHRIL). Combined with an adapted approach to microcartography, the innovative optical window facilitates serial intravital imaging of the intact lung at single-cell resolution across multiple imaging sessions and spanning multiple weeks. Given the unprecedented duration of time over which imaging data can be gathered, the WHRIL can facilitate the accelerated discovery of the dynamic mechanisms underlying metastatic progression and numerous additional biologic processes within the lung.

Here, we present a protocol for the surgical implantation of a permanently indwelling optical window for the murine thorax, which enables high-resolution, intravital imaging of the lung. The permanence of the window makes it well-suited to the study of dynamic cellular processes in the lung, especially those that are slowly evolving, such as metastatic progression of disseminated tumor cells.

Metastasis, accounting for ~90% of cancer-related mortality, involves the systemic spread of cancer cells from primary tumors to secondary sites such as ...

Cancer Research

Real-time X-ray Imaging of Lung Fluid Volumes in Neonatal Mouse Lung

At birth, the lung undergoes a profound phenotypic switch from secretion to absorption, which allows for adaptation to breathing independently. Promoting and sustaining this phenotype is critically important in normal alveolar growth and gas exchange throughout life. Several in vitro studies have characterized the role of key regulatory proteins, signaling molecules, and steroid hormones that can influence the rate of lung fluid clearance. However, in vivo examinations must be performed to evaluate whether these regulatory factors play important physiological roles in regulating perinatal lung liquid absorption. As such, the utilization of real time X-ray imaging to determine perinatal lung fluid clearance, or pulmonary edema, represents a technological advancement in the field. Herein, we explain and illustrate an approach to assess the rate of alveolar lung fluid clearance and alveolar flooding in C57BL/6 mice at post natal day 10 using X-ray imaging and analysis. Successful implementation of this protocol requires prior approval from institutional animal care and use committees (IACUC), an in vivo small animal X-ray imaging system, and compatible molecular imaging software.

We present a protocol to assess the rate of alveolar fluid clearance or pulmonary edema in neonatal mouse lung using X-ray imaging technology.

At birth, the lung undergoes a profound phenotypic switch from secretion to absorption, which allows for adaptation to breathing independently. Promoting ...

A Multimodal Imaging Approach Based on Micro-CT and Fluorescence Molecular Tomography for Longitudinal Assessment of Bleomycin-Induced Lung Fibrosis in Mice

Idiopathic pulmonary fibrosis (IPF) is a fatal lung disease characterized by the progressive and irreversible destruction of lung architecture, which causes significant deterioration in lung function and subsequent death from respiratory failure.
The pathogenesis of IPF in experimental animal models has been induced by bleomycin administration. In this study, we investigate an IPF-like mouse model induced by a double intratracheal bleomycin instillation. Standard histological assessments used for studying lung fibrosis are invasive terminal procedures. The goal of this work is to monitor lung fibrosis through noninvasive imaging techniques such as Fluorescent Molecular Tomography (FMT) and Micro-CT. These two technologies validated with histology findings could represent a revolutionary functional approach for real time non-invasive monitoring of IPF disease severity and progression. The fusion of different approaches represents a step further for understanding the IPF disease, where the molecular events occurring in a pathological condition can be observed with FMT and the subsequent anatomical changes can be monitored by Micro-CT.

We describe a non-invasive multimodal imaging approach based on Micro-CT and fluorescence molecular tomography for longitudinal assessment of the mouse lung fibrosis model induced by double intratracheal instillation of bleomycin.

Idiopathic pulmonary fibrosis (IPF) is a fatal lung disease characterized by the progressive and irreversible destruction of lung architecture, which ...

Immunology and Infection

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 ...

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 ...

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 ...

Non-invasive Skeletal Muscle Quantification in Small Animals Using Micro-computed Tomography

Skeletal muscle size, mass, and composition are critical properties for studying metabolic and muscle-related diseases, as they directly impact the understanding of disease progression and treatment outcomes. Quantifying a live animal's lean, adipose, and skeletal mass is important in metabolic, physiology, pharmacologic, and geroscience studies. However, obtaining accurate body composition measurements, especially of lean mass, remains challenging due to the inherent limitations of conventional assessment techniques. Micro-computed tomography (micro-CT) is a non-invasive radiological technique that enables high-resolution visualization of internal structures in small animal models. A standardized micro-CT method can significantly enhance translational research with more reliable and impactful results, particularly during aging studies or at different time points within the same animal. Despite its potential, the lack of standardization in image acquisition and analysis methods significantly hinders the comparability of results across different studies. Herein, we present a comprehensive and detailed low-cost protocol for lean mass analysis using micro-CT to address these challenges and promote consistency in research involving small animal models.

This protocol uses micro-computed tomography to enable the cost-effective quantification of lean mass, including skeletal muscle and visceral tissue, adipose tissue, and skeletal tissue in small animals. It distinguishes between lean and adipose tissue, which offers significant advantages for biomedical research, particularly in translational research in small animals.

Skeletal muscle size, mass, and composition are critical properties for studying metabolic and muscle-related diseases, as they directly impact the ...

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

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