Characterization of Immune Cells and Proinflammatory Mediators in the Pulmonary Environment

All protocols and procedures performed were approved by the Institutional Animal Care and Use Committee (IACUC) of West Virginia University. The mice were housed under specific-pathogen-free conditions in the vivarium at West Virginia University.  

1. Preparation of solutions

1. Prepare perfusion buffer (phosphate buffered saline, PBS). Use approximately two 10 mL aliquots of ice-cold PBS per mouse.
2. Prepare lung cell medium/FACS buffer. FACS buffer contains PBS supplemented with 1% fetal bovine serum (FBS). Keep the medium cold for the entire process of lung excision and transfer. Prepare approximately 8 mL per lung sample. Prepare fresh medium/FACS buffer prior to the experiments.
3. Prepare dissociation buffer for single cell isolation. Buffer contains 1 mg/mL collagenase D and 200 µg/mL DNase I in Hank’s buffered salt solution (HBSS). Prepare fresh from the stock solution (100 mg/mL collagenase D and 10 mg/mL DNase I) prior to the experiments. Approximately 6 mL per lung sample is needed. Ensure buffer reaches room temperature prior to the use.
4. Prepare homogenization buffer for lung tissue homogenization. Buffer contains PBS and 1x proteinase and phosphatase inhibitor cocktail (stock = 100x). Freshly prepare prior to the experiments. Keep the buffer cold during the entire homogenization process. Approximately 200 µL of the buffer is sufficient for homogenizing a single right lobe of the lungs.

2. Transient middle cerebral artery occlusion (tMCAO)

NOTE: Procedures for tMCAO via monofilament insertion to the middle cerebral artery were documented in detail previously¹⁰. In these experiments, 8- to 12-week-old male C57BL/6J mice, weighing 25-30 g, were used.

1. In brief, subcutaneously administer 2 mg/kg meloxicam to the mice before surgery. Deeply anesthetize the mice in an induction chamber using 5% isoflurane. Confirm deep anesthetization using the toe-pinch method. Maintain anesthesia with 1-2% isoflurane during the surgery using a nose cone. Maintain the body temperature at 36.5 – 38 °C throughout the procedure by placing a warm blanket underneath the mice. All surgical tools should be autoclave-sterilized a day prior to the surgery.
2. Shave and prepare the skin on the ventral neck using 70% ethanol followed by chlorhexidine scrub. Subcutaneously administer 2 mg/kg bupivacaine over the incision area prior to making the first incision. Use eye ointment to cover the eyes to prevent dryness. Make a midline neck incision. Gently pull aside soft tissues around the trachea.
3. Identify the left common carotid artery and the external carotid artery.
4. Apply a temporary suture (size 6/0) to the common carotid artery to stem the flow of the blood. Make an incision in the external carotid artery.
5. Insert a silicone rubber-coated monofilament (size 6-0) into the external carotid artery, then advance the monofilament to the middle cerebral artery. Leave the monofilament in the middle cerebral artery for 60 min (occlusion).
6. Monitor the rate of blood flow using Laser Doppler Flowmetry. A reduction in the flow rate to the middle cerebral artery that is > 80% indicates a successful occlusion.
7. Following the 60 min of occlusion, remove the monofilament to allow for the reperfusion to occur. Close the incision.
8. In sham-operated mice, perform steps 2.1-2.6 without the insertion of the monofilament. In these mice, the rate of blood flow should remain steady during the entire procedure.
9. Monitor the mice daily and measure the neurological deficits using standard scoring criteria¹¹. 0 – no neurological deficit; 1 – retracts contralateral forepaw when lifted by the tail; 2- circles to the contralateral when lifted by the tail; 3- falling to the contralateral while walking; 4 – does not walk spontaneously or is comatose; 5 – dead.
10. Provide subcutaneous injections of normal saline and analgesic (meloxicam, 2 mg/kg) every 24 hours for 3 days following the surgery.
11. Isolate the lungs 24 and 72 h following tMCAO for the downstream analysis.

3. Harvesting the lung tissues

1. Deeply anesthetize the mouse by intraperitoneal (i.p.) injection of 100 mg/kg of ketamine and 10 mg/kg of xylazine. Confirm deep anesthetization using the toe-pinch method.
2. To perform the whole body transcardial perfusion¹², pin the mouse to the surgical platform and wet fur with 70% ethanol to reduce fur distribution into the body cavity. Use fine dissection scissors to make a transverse incision in the caudal abdomen and then a midline incision of the peritoneum, stopping at the thoracic cavity prior to piercing the diaphragm.
   1. Do not cut through the muscle/hide into the peritoneal cavity and do not continue incision past abdomen.
3. Stop the incision as soon as the thoracic cavity has been reached before puncturing the diaphragm. Take care not to damage the heart and lungs which may compromise perfusion quality or cell recovery
4. Using fine dissection scissors make a midline incision of the peritoneum stopping at the thoracic cavity prior to piercing the diaphragm.
5. Grasp the mouse by the distal end of the sternum using forceps and lift upward while cutting through the diaphragm being certain not to damage the heart, lungs, or vasculature. Once organs are visible make an incision through the rib cage and separate and pin back the rib case so that the heart and lungs are fully exposed.
6. Carefully make a small incision in the right atrium near the aortic arch. This will serve as the outflow for the perfusion.
7. Immediately after, gently insert a 25 G x 5/8 needle attached to a syringe filled with 10 mL of cold PBS into the distal superior surface of the left ventricle. Use care not to insert through the ventricular septum into the right ventricle. Needle should be barely inserted into the left ventricle.
   NOTE: If ventricular septum is pierced, fluid will rush out from the mouse’s nose. This will decrease the perfusion quality and result in cell loss.
8. If desired, a clamp may be used to hold the needle securely in place in the left ventricle during the perfusion.
9. Steadily but gently push 10 mL of cold PBS into the heart. Mouse tissue should begin to perfuse and clear off the blood as fluid exits from the right atrium.
   1. Push a second aliquot of 10 mL of PBS until sufficient blood clearance has occurred. Liver will have a light brown appearance and lungs will transition from a reddish pink to mostly white in color.
10. Using forceps and fine dissection scissors, carefully remove all surrounding tissues including the heart, trachea, esophagus, thymus, connective tissue, lymph nodes, and large bronchials.
11. Separate individual lung lobes and place the lobes into a Petri dish on ice containing a 1-2 mL aliquot of cold lung cell medium.

4. Homogenization of lung tissue for multiplex bead arrays using bead homogenizer

1. Pre-chill a 2 mL conical screw cap tube containing 3 sterile 2.3 mm zirconia/silica beads. Add 200 µL of cold homogenization buffer to the tube.
2. Weigh the lobe, transfer the lobe to the pre-chilled conical tube containing the beads and homogenization buffer.
   NOTE: Approximately 50-100 mg of tissue homogenized in 200 µL of buffer will be sufficient for detecting cytokine and chemokine expression in the sample.
   1. Homogenize the lobe using a bead-based homogenizer (see Table of Materials) at 4,000 rpm for 2 min.
   2. Ensure the tissue is completely homogenized. Increase the time if needed.
3. Following homogenization, centrifuge the tube in a pre-chilled (4 °C) micro-centrifuge at 15,870 x g for 3 min.
4. Transfer the supernatants to a pre-chilled 1.5 mL microtube.
5. Place the sample directly on ice for immediate use or store the sample at -80 °C for future use. Avoid multiple freeze-thaw cycles.

5. Lung tissue dissociation and single cell isolation

1. Pour the lung cell medium off, of the remaining lung lobes to avoid dilution of the dissociation buffer. Then, carefully insert a 1 mL syringe with 25G and 5/8 needle containing 1 mL of the dissociation buffer into lung tissue.  
2. Inject small fractions (approximately 1/4^th to 1/5^th) of the dissociation buffer into each lung lobe and gently inflate.
   NOTE: Most of the buffer will rush out soon after inflating as the lobe has been excised.
3. Repeat the injection of dissociation buffer 1-2x.
4. Incubate the lung lobes with dissociation buffer for 2 min at room temperature.
5. Finely mince the lung lobes into small pieces using fine dissection scissors.
6. Transfer the minced lung pieces with the dissociation buffer to a 15 mL centrifuge tube.  Supplement with additional dissociation buffer to a total of 6 mL. Vortex vigorously for 1 min.
7. Incubate the sample for 45 min at 37 °C. Vortex vigorously every 7-8 min.
8. After 45 min of incubation, vortex the sample vigorously for 1 min for optimal results.
9. Further dissociate the sample by passing through a 100 µm cell strainer to remove residual, undesired connective, and interstitial tissue.
10. Centrifuge the sample for 10 min at 380 x g, discard the supernatant.
11. Add 5 mL of cold lung cell medium to the sample. Resuspend the cell pellet by gentle vortexing.
12. Centrifuge the sample for 10 min at 380 x g, discard the supernatant.
13. Add 1 mL of cold lung cell medium. Resuspend the cell pellet by gentle vortexing.
14. Pass through a 100 µm cell strainer, and count cells.

6. Flow cytometric analysis for the lung immune cell niche

1. Seed 1-2 x 10⁶ cells/antibody set into a 96 well round bottom plate (3 sets in total). Centrifuge the cells for 3 min at 830 x g. Discard supernatants.
2. Resuspend the cell pellet in 50 µL of cold PBS containing fixable live/dead stain. Prepare the stain according to the manufacturer’s instructions.
3. Incubate the cells at 4 °C for 20 min protected from light. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.  
4. Resuspend the cell pellet with 25 µL of cold FACS buffer containing 5 µg/mL of anti-mouse CD16/32 antibody. Incubate at 4 °C for 10-15 min.
5. Add 25 µL of cold FACS buffer containing antibody combinations listed in Table 1 to the cells. Mix by gentle pipetting.
   NOTE: Total volume of FACS buffer for the antibody staining is 50 µL. Therefore, when preparing a master mix of antibodies in 25 µL, the concentration of antibodies should be double the final concentration.  
6. Incubate the cells at 4 °C for 20 min protected from light. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.
7. Resuspend the cells with 100 µL of cold FACS buffer. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.
8. Resuspend the cells with 100 µL of cold FACS buffer. Transfer to a polystyrene round-bottom 5 mL FACS tube containing 150 µL of additional FACs buffer. Analyze the sample immediately using a flow cytometer.
   NOTE: Cell fixation using the following steps allows samples to be analyzed later.
9. Following step 6.7., resuspend the sample with 100 µL of 1% paraformaldehyde in PBS.
10. Incubate the sample at 4 °C for 15 min protected from light. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.
11. Resuspend the cells with 100 µL of cold FACS buffer. Centrifuge the cells for 3 min at 830 x g. Discard supernatants.
12. Resuspend the cells with 100 µL of cold FACS buffer. Store the cells at 4 °C, protected from light. Analyze the sample within 72 h.

7. Multiplex bead arrays for cytokine and chemokine detection

NOTE: Commercially available proinflammatory chemokine and inflammation multiplex panels (see Table of Materials) were used to determine the expression of chemokines and cytokines in the lungs following tMCAO induction according to manufacturer protocol, which has been described in detail¹³.

1. In brief, thaw the analytes on ice if they were stored at -80 °C after tissue homogenization. Centrifuge the sample at 590 x g for 2 min at 4 °C before use to remove any residual tissue.
2. Generate a standard curve by serially diluting the standard cocktail (C7) which is at a known concentration of 10 ng/mL into 7 standards (C1-6) with the last standard being assay buffer alone (C0).
3. Once all reagents have warmed to room temperature, add 25 µL of each standard and 25 µL of assay buffer to a V bottom 96 well plate. 
4. To each sample well, add 25 µL of lung homogenate (analyte) and 25 µL of assay buffer to a V bottom 96 well plate
5. Vortex the pre-coated beads vigorously, then add 25 µL of pre-coated beads to each of the standard and sample wells.
6. Seal the plate and protect from light by covering with foil. Shake the plate at 110 rpm for 2 h at room temperature.
7. Centrifuge at 230 x g for 5 min. Add 200 µL of wash buffer at 1x (stock solution is 20x, dilute to 1x with water). Incubate for 1 min.
8. Centrifuge at 230 x g for 5 min. Resuspend the standard and sample with 25 µL detection antibodies.
9. Seal and cover the plate to protect from light. Shake plate at 110 rpm for 1 h at room temperature.
10. Add 25 µL of streptavidin-phycoerythrin (SA-PE) to each standard and sample well.
11. Seal and cover the plate to protect from light. Shake at 110 rpm for 30 min at room temperature.
12. Spin the plate at 230 x g for 5 min. Resuspend the standard and samples in 1x wash buffer.
13. Transfer to a well-labeled polystyrene round-bottom 5 mL FACS tube containing an additional 150 µL of 1x wash buffer.
14. Determine the PE signal fluorescence intensity for the standards and samples using a flow cytometer.
15. Construct a standard curve for each chemokine and cytokine using the mean fluorescence intensity (MFI) of PE against the pre-determined concentrations.
16. Determine the concentration of each chemokine and cytokine using the standard curve and the MFI from each sample. The multiplex panel provides data analysis software that can be used to determine the concentration of each analyte. The weight of the apical/superior lobe is used to determine the concentration of cytokines or chemokine per milligram of tissue.