Precision-cut Mouse Lung Slices to Visualize Live Pulmonary Dendritic Cells

Animal experimental procedures described in this paper were approved by the NIEHS Animal Care and Use Committee (IACUC).

1. Lung Preparation

1. House mice between 6 and 12 weeks of age in specific pathogen-free conditions in accordance with the guidelines provided by the Institutional Animal Care and Use Committees.
   NOTE: Imaged mice can be either naïve, or treated, depending on each researcher's specific interests. Here, we describe immune cell localization in naïve mice and in mice treated with 100 µg ovalbumin (OVA) and 0.1 µg lipopolysaccharide (LPS), using phosphate buffered saline (PBS) as a vehicle in a total volume of 50 µL. To oropharyngeally instill OVA/LPS into the airways, animals were anesthetized with isoflurane inhalation and vertically suspended by their teeth with a rubber band. The tongue was gently grasped with forceps and held to one side to prevent swallowing, and 50 µL of the OVA/LPS solution deposited at the back of the oral cavity as previously described¹⁷.
2. Euthanize the mouse with intraperitoneal injection of sodium pentobarbital (100 mg/kg), and pin the animal to a polystyrene base.
3. Open the abdominal cavity with scissors by cutting the skin and peritoneum from the middle of the abdomen up to the jaw. Pull the intestines aside with forceps or the dull edge of the scissors, and snip the inferior vena cava to drain blood away from lungs. Puncture the diaphragm with the sharp tip of the scissors to allow expansion of the rib cage, being careful not to cut the lungs.
4. Clear salivary glands and other tissue away from the trachea by grabbing the tissue and manually pulling it away from the underlying trachea with forceps. Make a small incision in the trachea on the anterior side of the thickest band of cartilage using fine forceps or scissors, being careful not to cut all the way though the trachea. The incision will be just large enough to allow a 20 G needle to pass through.
5. Slide a 1.5 inch, 20 G needle onto a section of polyethylene tubing, leaving approximately 1 cm of tubing beyond the end of the needle. Cut the tubing at an angle of approximately 45° to bevel the end, attach the needle to a 1 mL syringe, and load the syringe by drawing 0.8 mL of warm (40 °C) 2% low melting point agarose up through the needle.
6. Place the end of the tubing into the incision of the trachea and slowly inject the agarose into the lungs. Without moving the needle or syringe, tape the syringe to the polystyrene base to ensure the needle remains in the trachea.
   NOTE: After injection, the lungs will be larger and inflated within the chest. The right lobes expand first, then the left lobe. If only one lobe expands, the needle has been inserted too deeply (past the bronchus), and it will be necessary to pull it out slightly before proceeding. This part of the procedure needs to be done relatively quickly, before the agarose starts to solidify.
7. Place the mouse, with the base and the syringe still inserted in the trachea, in a cold room or refrigerator for at least 10 min. If necessary, the animal can be kept there for up to several hours, even if proceeding to image cell migration by video microscopy.
8. Once the mouse is cold, carefully excise the agarose-inflated lungs using scissors, and place them in a 3 cm dish with ice cold PBS. Keep on ice.
9. Choose the desired lobe for tissue slicing (e.g., the right superior lobe). Make sure the interior part of the lung (where it connects to trachea) is face down in the dish.
   NOTE: The lobe used and its orientation on the plunger will determine the size of vessels and airways that will be visible. The orientation described here is ideal for visualization of large airways, and parenchyma. Mice have one large left lobe and four smaller lobes on the right. For PCLS imaging following introduction of allergens into the lungs via oropharyngeal or intranasal aspiration, the right superior lobe is typically sectioned, in part because it is a convenient size, but also because inhaled agents disperse uniformly within it. However, other lobes, especially the left lobe, can also be studied. When comparing mouse strains or treatments, it is important to compare the same lobe.

2. Lung Slicing

NOTE: Make slices using an automated slicer, metal cooling block, plunger and metal syringe, according to the manufacturer's instructions.

1. Put a small drop of all-purpose, no-run gel superglue onto the plunger and spread the glue in a circular motion, taking care to not let the glue touch the sides of the metal syringe. If the glue touches the sides of the syringe, it will glue the syringe and plunger together.
2. Immediately after covering the plunger with glue, gently grasp the excised lobe with forceps with the trachea side down, dab it on a tissue to remove excess liquid, and carefully place it on top of the plunger. Trim off any extra tissue extending beyond the edge of the plunger.
3. Move the plunger down so that the sides of the metal syringe move up and over the tissue, creating a well with the lung glued at the bottom, no more than several centimeters deep. Tape around the bottom of the metal syringe to hold this position in place.
4. Carefully pour 2% low melt agarose at 40 °C into the well so it just covers the top of the lobe.
5. Surround the metal syringe with the ice-cold chilling block and cool the lobe and agarose for 1 – 2 min.
   NOTE: A shorter time cooling of the agarose surrounding the tissue could lead to disintegration of the agarose during slicing, which might result in an unevenly cut PCLS.
6. Load the specimen syringe into the automated slicer and fill the buffer tank with ice cold PBS. Align the step motor drive with the specimen syringe and remove the piece of tape from the bottom. Turn the switch to the fast forward (FF) position until the step motor drive just touches the back of the plunger.
7. Align the fresh blade with the specimen syringe. Set tissue thickness, continuous/single slice, oscillation and speed in the slicer. For example, tissue thickness: 150 µm, continuous cutting (cont.), oscillation: 9, and speed: 3 – 4. Press start. The above settings need to be adjusted to the specific automated slicer specifications.
8. Using a thin spatula or paintbrush, collect the PCLS one at a time as they fall into the buffer tank and place them in a 24-well plate containing ice cold PBS.
   NOTE: It is important to keep the slices in order to maintain consistency between experiments. Although every lung is different depending on age, gender and weight, the 10^th slice, for example, generally yields similarly sized airways.

3. Antibody Staining

1. Design a panel of fluorescently tagged antibodies based on the cell type of interest, taking into account potential fluorescent spectral overlap and the detection capabilities of the microscope.
   NOTE: As an example, a panel for DC subset detection is as follows: CD11c/alpha X integrin – BrilliantViolet (BV) 605 (Excitation: 405 nm, Peak emission: 605 nm), CD324/E-cadherin – Alexa 488 (AF488) (Excitation: 488 nm, Peak emission: 519 nm), CD88/C5Ra1 – Phycoerythrin (PE) (Excitation: 561 nm, Peak emission: 578 nm), CD103/alpha E integrin – Allophycocyanin (APC) (Excitation: 633 nm, Peak emission: 660 nm). Fluorescence spectral viewer tools (see Materials List) are useful to design panels.
2. Make an antibody cocktail containing the desired antibodies.
   1. For static PCLS imaging, add 800 µL staining buffer, 100 µL Fc blocker, 50 µL normal mouse serum, and 50 µL normal rat serum to a 1.5 mL microcentrifugation tube for a total volume of 1 mL. For live cell imaging, use 800 µL Leibovitz's medium (1x) containing 10% FBS (Leibovitz-10) instead of staining buffer.
   2. Add antibodies to adjust a desired final concentration of each antibody. Transfer the antibody solution to a 3 cm dish, and keep in the dark until ready to use.
      NOTE: The final concentrations need to be optimized for each individual antibody (usually 1 – 5 µg/mL).
3. Choose a PCLS with an anatomical area of interest, such as large airways or periphery of the lung, and place in the 3-cm dish containing 1 mL antibody solution.
4. Stain PCLS in the dark, on ice, rocking slowly for 30 or 60 min.
5. For static PCLS imaging, proceed to the section 4. For live cell imaging, proceed to section 5.

4. Static Imaging of PCLS

1. After 60 min incubation of PCLS with antibodies, remove the antibody solution from the dish and rinse the slices twice with 1 mL ice cold PBS.
2. Pipette 50 µL PBS onto a 3 cm round glass-bottom dish. Using a spatula or paintbrush, transfer the stained PCLS to the drop, gently manipulating the slice until it is flat and spread out. Remove the PBS with a pipette. The slice should lie as flat as possible. The above procedures need to be performed quickly to avoid photobleaching.
3. Place 1 drop of room temperature mounting medium onto the slice and gently drop a glass coverslip into the well of the plate. Keep PCLS in the dark at 4 °C for up to an hour before imaging.
4. When viewing under a confocal microscope, use a 20X objective (see Materials List). Use the "tile" tool to locate and mark the edges of the tissue in the "convex hull" setting. This will help reduce the time of the tile scan.
5. While setting the z-stack range, verify at multiple areas of the slice, especially along the edges, that the set ranges are appropriate.
   NOTE: The z-range will likely need to be greater than the thickness of the tissue itself because it is very difficult to get the tissue to lay completely flat. For example, with a 150 µm thick PCLS, the z-stack range will likely need to be set to 200 – 250 µm to accommodate bumps and curves in the slice. Depending on the z-stack range and the size of the tissue, each full lung scan will take between 7 – 12 h with a 20X objective. Each lung is unique and the settings must be adjusted for every experiment.

5. Live Cell Imaging of PCLS

1. After 30 min incubation of PCLS with antibodies, remove the antibody solution from the dish and rinse the slices twice with 1 mL cold Leibovitz-10.
2. Pipette fresh 50 µL Leibovitz-10 into one slot of a chambered coverglass (8 slots). Use a spatula to transfer the PCLS to the medium, gently manipulating it until the slice is flat and spread out. The above procedures need to be performed quickly to avoid photobleaching.
3. Remove the 50 µL medium using a pipette. The slice is to be lying as flat as possible.
   NOTE: It is acceptable if the edges of the slice are flipped up vertically against the wall of the chamber. Furthermore, if available, a metal platinum weight can be used to anchor the slice before proceeding to the next step. See the attached Materials List for one example of platinum wire that can be cut and bent into small weights.
4. In a 4 °C cold room, mix 100 µL of growth factor-reduced, phenol red-free extracellular matrix with 100 µL Leibovitz-10. Use precooled pipette tips for this, or the extracellular matrix will solidify in the tip.
   NOTE: This ratio of matrix and medium (1:1) creates a gel dense enough to hold the PCLS down in tissue-culture conditions.
5. Gently pipette the matrix to mix, and carefully pipette on top of the PCLS, making sure that the gel goes on top of the lung slice, and that the slice does not float up on top of the gel. The gel should work as a weight, anchoring the PCLS down onto the bottom of the plate.
6. Carefully transfer the chambered coverglass to a 37 °C incubator and let the matrix solidify for ~5 min.
7. Transfer the entire chambered coverglass onto the pre-warmed stage of a confocal microscope. Maintain the chamber at 37 °C throughout imaging. CO₂ supply is not required during the cell incubation when using Leibovitz's medium.
8. Set the z-stack range and tile range based on the desired interval between frames. When viewing under a confocal microscope, use a 20X objective (see Materials List). Use the "tile" tool to denote the area of interest in the tissue, using the "centered grid" setting (e.g., a 2×2 centered grid).
9. While setting the z-stack range, verify at multiple areas of the slice that the set ranges are appropriate.
   NOTE: The z-range will likely equal the thickness of the tissue. Each lung is unique and the settings must be adjusted for every experiment. 2×2 tiles and 10 z-stacks will generally result in ~1 frame/2 min. Ensure the auto-focus function is activated before starting the experiment to minimize x-y and z-drift during the imaging period.