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Method Article

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

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DOI:

10.3791/61242

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June 20th, 2020

 ,  ,  ,  ,  ,  ,  , 
1Translational Vascular Medicine Branch, National Heart Lung and Blood Institute, National Institutes of Health, 2Mouse Imaging Facility, National Institute of Neurological Disorders and Stroke, National Institutes of Health, 3Division of Pediatric Surgery, Department of Surgery, University of North Carolina at Chapel Hill

In This Article

Summary

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.

Abstract

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.

Introduction

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.

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Protocol

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

  1. Inject the mouse intraperitoneally with heparin (1 unit/g mouse body weight) and allow it to ambulate for 2 min.
  2. Euthanize the animal in a CO2 chamber.
  3. 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.

2. Exposing lungs and trachea

  1. Spray the ventral side of the mouse with 70% ethanol to minimize hair interference.
  2. 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.
  3. Open the abdominal musculature with scissors and cut laterally on both sides until the diaphragm is exposed.
  4. 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.
  5. 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.
  6. 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.
  7. 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.
  8. 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.
  9. 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.

3. PA catheterization and blood perfusion

  1. 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.
    1. 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).
  2. 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.
  3. 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.
  4. 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.
    1. 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.
    2. 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.
  5. 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.
  6. Clip the left auricle of the heart to allow perfusate to exit the system.
  7. 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 µL 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.

4. Tracheostomy and lung inflation

  1. Construct the lung inflation unit (Figure 2).
    1. 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.
    2. 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.
  2. Place two loose sutures inferior to the cricoid cartilage 2-4 mm apart.
  3. Using scissors, make a small incision in the cricothyroid ligament superior to the sutures.
  4. Insert the IV catheter into the opening and advance the tip beyond the two loose sutures.
  5. 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.
  6. 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.

5. Casting the vasculature

  1.  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.
  2. 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.
  3. 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.
  4. 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.
  5. 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 μL for an adult mouse and ~5 µL 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.
  6. 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.
  7. 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.
  8. 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.

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.

  1. 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).
  2. Grasp the xiphoid process with a hemostat and cut the ribcage bilaterally (roughly in the midaxillary line) just prior to the internal thoracic arteries.
  3. Fold the still connected ribcage over such that it is resting on the animal's neck/head, fully exposing the chest cavity.
  4. 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.
  5. 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.
    1. For structures inferior to the diaphragm, instead, create a small hole ~2 mm superior to the loose sutures.
  6. Depending on the animal size, introduce Unit 1 or 2 into the vessel, advance beyond loose sutures, and gently ligate the vessel.
  7. 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.
  8. 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).
  9. Follow 5.6 then remove target tissue and place in formalin.

7. Sample mount, scan, and reconstruction for micro-CT

  1. 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.
  2. 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).
  3. 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.
  4. Transfer the reconstructed scans for post-processing and analysis.

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Results

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 (

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Discussion

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 lung...

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Disclosures

The authors have nothing to disclose

Acknowledgements

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.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1cc syringeBecton Dickinson309659
20ml Glass Scintillation VialsFisher03-340-25P
30G NeedleBecton Dickinson305106
50mL conical tubesCornin352098For sample Storage and scanning
60cc syringeBecton Dickinson309653
7-0 silk sutureTeleflex103-S
Analyze 12.0 SoftwareAnalyzeDirect Inc.N/APrimary Software
Amira 6.7 SoftwareThermo ScientificN/AAlternative Sofware
CeramaCut Scissors 9cmFine Science tools14958-09
Ceramic Coated Curved ForcepsFine Science tools11272-50
CO2 TankRobert's Oxygen Co.n/a
Dual syringe pumpCole ParmerEW-74900-10
Dumont Mini-ForcepsFine Science tools11200-14
EthanolPharmco111000200
FormalinSigma - Life SciencesHT501128
GauzeCovidien441215
HemostatFine Science tools13013-14
Heparin (1000USP Units/ml)HospiraNDC 0409-2720-01
Horos SoftwareHoros ProjectN/AAlternative Sofware
induction chambern/an/a
KimwipeFisher06-666fiber optic cleaning wipe
Labelling TapeFisher15966
Magnetic BaseKanetecN/A
Micro-CT systemPerkinElmerQuantum GX
Microfil (Polymer Compound)Flowech Inc.Kit B - MV-1228 oz. of MV compound; 8 oz. of diluent; MV-Curing Agent
MicromanipulatorStoelting56131
Monoject 1/2 ml Insulin SyringeCovidien1188528012
Octagon Forceps Straight TeethFine Science tools11042-08
ParafilmBemis company, Inc.#PM999
PE-10 tubingInstechBTPE-10
Phospahte buffered SalineBioRad#161-0780
Ring StandFisherS13747Height 24in.
Sodium Nitroprussidesigma71778-25G
Steel PlateN/AN/A16 x 16 in. area, 1/16 in thick
Straight Spring ScissorsFine Science tools15000-08
SURFLO 24G Teflon I.V. CatheterSanta Cruz Biotechnology360103
Surgical BoardFisher12-587-20This is a converted slide holder
Universal 3-prong clampFisherS24280
Winged Inf. Set 25X3/4, 12" TubingNiproPR25G19
Zeiss Stemi-508 Dissection ScopeZeissn/a

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Pulmonary ArteryMouse LungPolymer InfusionLung InflationCatheter InsertionVascular NetworkSodium Nitroprusside3D Visualization
Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging
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