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 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 (µCT) 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.
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 µCT, 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.
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
2. Exposing lungs and trachea
3. PA catheterization and blood perfusion
4. Tracheostomy and lung inflation
5. Casting the vasculature
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.
7. Sample mount, scan, and reconstruction for micro-CT
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 "airlock", 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, "airlock" 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 µCT 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 µCT scanning, the casted samples can also be cleared to obtain gross images or processed and cut for histological analysis8.
Figure 1: Catheter and needle setup. Syringes are shown with attached tubing and needles (Unit1 and Unit2). Inset: closeup of needle and tubing. Please click here to view a larger version of this figure.
Figure 2: Lung inflation setup. Ring stand, clamp, a syringe filled with formalin, and tubing with a catheter attached. Please click here to view a larger version of this figure.
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. Please click here to view a larger version of this figure.
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 Please click here to view a larger version of this figure.
Figure 5: Examples of ideal filling and common errors during polymer compound infusion. (A) 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. Please click here to view a larger version of this figure.
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. Please click here to view a larger version of this figure.
Figure 7. CT images and 3D volume rendering of polymer compound 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). Please click here to view a larger version of this figure.
Target Arterial Vascular Bed | Catheter placement | Infusion direction | Infusion rate | Notes |
Brain | Thoracic aorta pointing cranially | Retrograde into the carotids | .05ml/min | Cannulate thoracic aorta, flip mouse to the prone position, open scalp, and visually monitor progress of polymer through skull. |
Diaphragm | Left Ventrical | Anterograde into internal thoracic, phrenic, and intercostal | .05ml/min | Open a window in the side of the ribcage, leaving the majority of the ribcage and the diaphragm intact. Cannulate left ventrical, clip right atrium, and monitor progress from the caudal side of the diaphragm. |
Upper limb musculature | Thoracic aorta pointing cranially | Retrograde into the brachiocephalic and left subclavian | .02ml/min | To optimize limb flow, tie off the carotid arteries and remove limb skin to allow visual monitoring of polymer transit into the limb musculature. |
Kidney | Thoracic aorta pointing caudally | Anterograde into renal arteries | .05ml/min | The internal vasculature is filled blindly. To avoid venous transit, stop injecting when polymer is visible in a uniform punctate pattern across kidney. |
Portal System | Portal vein | Anterograde into portal system | .02ml/min | Gently fold liver up to expose the portal vein. |
Hepatic | Thoracic aorta pointing caudally | Anterograde into the hepatic artery | .05ml/min | Tie off portal vein prior to infusion to avoid venous transit from gut flowing into liver. |
Stomach/ Intestine | Thoracic aorta pointing caudally | Anterograde into the celiac, superior mesenteric and/or inferior mesenteric | .05ml/min | Some regions of the gut are supplied by multiple arteries and may fill at different times. To avoid venous transit, tie off arteries not required for areas of interest and visually monitor the progress of the polymer. |
Intra-abdominal fat pads | Thoracic aorta pointing caudally | Anterograde but vessel depends on fat pad being studied | .05ml/min | Fat pads are supplied by multiple arteries and may fill at different times. To avoid venous transit, tie off arteries not required for precise area of interest and visually monitor the progress of the polymer. |
Lower limb musculature | Infrarenal aorta pointing caudally | Anterograde into the femoral arteries | .02ml/min | Remove limb skin to allow visual monitoring of polymer transit into the limb musculature. |
Table 1. Casting alternative vascular beds.
CT settings | |
kVp | 90 |
Target Material | Tungsten |
Power | 8W |
Filtration | Cu 0.06 mm + Al 0.5 mm |
Projection Number | 6424 |
Detector Size | Flat panel CMOS – 2944 x 2352 pixels |
Field of View (FOV) | 36 mm |
Voxel Size | 72 µm |
Spatial Resolution | voxel size x 1.5 |
Acquisition Time | 14 min |
Reconstruction | FBP and commercial algorithm |
Binning | 1×1 |
Table 2. µCT Scanning Parameters.
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 µCT (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, µCT 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 "cleared" 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 µCT 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 µCT and traditional histology or even EM36,37,38. It takes a more high level view of pathology by utilizing µCT 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 µCT 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 >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'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 authors have nothing to disclose.
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.
1cc syringe | Becton Dickinson | 309659 | |
20ml Glass Scintillation Vials | Fisher | 03-340-25P | |
30G Needle | Becton Dickinson | 305106 | |
50mL conical tubes | Cornin | 352098 | For sample Storage and scanning |
60cc syringe | Becton Dickinson | 309653 | |
7-0 silk suture | Teleflex | 103-S | |
Analyze 12.0 Software | AnalyzeDirect Inc. | N/A | Primary Software |
Amira 6.7 Software | Thermo Scientific | N/A | Alternative Sofware |
CeramaCut Scissors 9cm | Fine Science tools | 14958-09 | |
Ceramic Coated Curved Forceps | Fine Science tools | 11272-50 | |
CO2 Tank | Robert's Oxygen Co. | n/a | |
Dual syringe pump | Cole Parmer | EW-74900-10 | |
Dumont Mini-Forceps | Fine Science tools | 11200-14 | |
Ethanol | Pharmco | 111000200 | |
Formalin | Sigma – Life Sciences | HT501128 | |
Gauze | Covidien | 441215 | |
Hemostat | Fine Science tools | 13013-14 | |
Heparin (1000USP Units/ml) | Hospira | NDC 0409-2720-01 | |
Horos Software | Horos Project | N/A | Alternative Sofware |
induction chamber | n/a | n/a | |
Kimwipe | Fisher | 06-666 | fiber optic cleaning wipe |
Labelling Tape | Fisher | 15966 | |
Magnetic Base | Kanetec | N/A | |
Micro-CT system | SkyScan | 1172 | |
Microfil (Polymer Compound) | Flowech Inc. | Kit B – MV-122 | 8 oz. of MV compound; 8 oz. of diluent; MV-Curing Agent |
Micromanipulator | Stoelting | 56131 | |
Monoject 1/2 ml Insulin Syringe | Covidien | 1188528012 | |
Octagon Forceps Straight Teeth | Fine Science tools | 11042-08 | |
Parafilm | Bemis company, Inc. | #PM999 | |
PE-10 tubing | Instech | BTPE-10 | |
Phospahte buffered Saline | BioRad | #161-0780 | |
Ring Stand | Fisher | S13747 | Height 24in. |
Sodium Nitroprusside | sigma | 71778-25G | |
Steel Plate | N/A | N/A | 16 x 16 in. area, 1/16 in thick |
Straight Spring Scissors | Fine Science tools | 15000-08 | |
SURFLO 24G Teflon I.V. Catheter | Santa Cruz Biotechnology | 360103 | |
Surgical Board | Fisher | 12-587-20 | This is a converted slide holder |
Universal 3-prong clamp | Fisher | S24280 | |
Winged Inf. Set 25X3/4, 12" Tubing | Nipro | PR25G19 | |
Zeiss Stemi-508 Dissection Scope | Zeiss | n/a |