Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) and the Institutional Review Board (IRB) at St Joseph's Hospital and Medical Center.

1. Isolation of AMs from Murine Bronchoalveolar Lavage (BAL) Fluid

1. Anaesthetize an eight-week-old C57BL/6 mouse with ketamine (87.5 mg/kg body weight) and xylazine (12.5 mg/kg body weight) cocktail via an intraperitoneal injection. Proceed when mouse attains surgical anesthesia with loss of reflexes and relaxation of muscles.
2. Place the mouse on a dissection surface with the ventral side facing up. Apply ophthalmic vet ointment on eyes to prevent dehydration and wipe the entire ventral surface of mouse with sterile alcohol prep pads saturated with 70% isopropanol to disinfect.
3. Mount the mouse with all four legs restrained.
4. Gently open the abdominal cavity with micro dissection tools. Care must be taken to open as much area as possible without damaging any visceral organs or creating overhanging tissue.
5. Gently open the thoracic cavity by slowly incising the rib cage through the mediastinum. Ideally, the rib cage can be excised from side to side. Extreme care must be taken not to injure the pleura of lungs while opening the thoracic cavity.
6. Locate the inferior vena cava and make an incision to bleed the mouse. Use absorbent cotton pads to soak up blood.
7. Inject 10 mL ice-cold phosphate buffered saline (PBS) into the right ventricle (using 10 mL syringe and 25G needle) to flush circulating blood cells. Absorb the flow of blood with absorbent cotton pads. Once completely perfused, the lungs will appear blanched and are then ready for lavage.
8. Gently cut open the skin and muscle in the neck to expose the airway. Carefully excise the adjoining muscles, cartilages, and fat tissues without damaging the trachea.
9. Make a small incision (<2 mm) on the trachea posterior to the larynx. This incision should be just enough to insert a catheter into while the trachea is still attached to the larynx. Insert a 1-inch 22G catheter without a needle into the trachea towards the lungs. Secure the catheter with a silk braided suture (4-0; non-absorbable) with a square knot.
   NOTE: The catheter needs to be trimmed based on the mouse size. Extremely short or long catheters tend to leak and/or tear the airway during lavage. Catheter length should be such that when fully inserted the tip remains well within the trachea and not reach the primary bronchi.
10. Attach a 1-mL syringe filled with ice-cold BAL buffer (Ca²⁺ and Mg²⁺ free PBS + 1 mM Ethylenediaminetetraacetic acid, EDTA) to the catheter and slowly instill the buffer into the lungs. This will inflate the lungs. Keep the syringe attached to the catheter for five seconds and then aspirate the lavage fluid by gently pulling the piston. This will deflate the lungs. Collect the fluid in a 15 mL conical tube placed on ice.
11. Repeat Step 1.10 for nine more times and pool the lavage fluid. Do not exceed 1 mL buffer per flush, as that may exceed the lung capacity and lead to its rupture.
12. Euthanize the mouse with CO₂ overdose by IACUC approved protocol.
13. Centrifuge the 10 mL BAL fluid at 250 x g at 4 °C for 10 minutes. The cell pellet contains BAL cells.
    Caution: The pellet may be very small, so exercise caution when aspirating the supernatant.
14. Resuspend the BAL cells in 100 μL of flow/sorting buffer (PBS + 2% Heat inactivated fetal bovine serum (FBS) + 1 mM EDTA + 25 mM N-2-Hydroxyethylpiperazine-N'-2-ethanesulphonic acid (HEPES)).
15. Block the nonspecific antibody binding to Fc receptor by adding mouse Fc block (clone 2.4G2, 10 μg/mL) and incubating for 20 minutes on ice.
16. Perform antibody staining along with fluorescence minus one (FMO) and single-stain controls^11,25. Evaluate the BAL cells by a cell analyzer (see Table of Materials) followed by single-cell flow cytometry analysis by appropriate analysis software (see Table of Materials).
17. Isolate AMs by a fluorescence-activated cell sorter (see Table of Materials) directly into 1 mL mouse AM culture medium when AMs are intended for in vitro culture or in vivo cell transfer. Alternatively, sort AMs into 1 mL Lysis buffer (see Table of Materials) supplemented with 10 μL/mL β-Mercaptoethanol for RNA extraction.

2. Isolation of AMs from Mouse Lung, Single-cell Suspension

1. Anaesthetize an eight-week-old C57BL/6 mouse with ketamine (87.5 mg/kg body weight) and xylazine (12.5 mg/kg body weight) cocktail via an intraperitoneal injection. Proceed when mouse attains surgical anesthesia with loss of reflexes and relaxation of muscles.
2. Place the mouse on a dissection surface with the ventral side facing up. Apply ophthalmic vet ointment on eyes to prevent dehydration and wipe the entire ventral surface of mouse with sterile alcohol pads saturated with 70% isopropanol to disinfect.
3. Mount the mouse with all four legs restrained.
4. Gently open the abdominal cavity with micro dissection tools. Care must be taken to open as much area as possible without damaging any visceral organs or creating overhanging tissue.
5. Gently open the thoracic cavity by slowly incising the rib cage through the mediastinum. Ideally, the rib cage can be excised from side to side. Extreme care must be taken not to injure the pleura of lungs while opening the thoracic cavity.
6. Locate the inferior vena cava and make an incision to bleed the mouse. Use absorbent cotton pads to soak up blood.
7. Inject 10 mL ice-cold PBS into the right ventricle (using 10 mL syringe and 25G needle) to flush circulating blood cells. Once completely perfused, the lungs will appear blanched and are ready for harvest.
8. Dissect out the lungs by severing the trachea, blood vessels and ligaments into a 15 mL conical tube with 5 mL of cold Dulbecco's Modified Eagle's medium (DMEM). Maintain it on ice until the next step.
9. Euthanize the mouse with CO₂ overdose by IACUC approved protocol.
10. Transfer the perfused lungs into a sterile and pyrogen-free 60 mm culture dish with 3 mL DMEM. With help of dissecting forceps tease out the airways and other hard non-lung tissue material, if present. Mince the lung tissue with a scalpel to <1 mm size.
11. Add 300 µg/mL Liberase TL and 5 U/mL DNase I, mix by pipetting and incubate at 37° C in an incubator for 25 minutes. Gently mix once by pipetting after 10 minutes.
12. Pass the dissociated lungs through a 100 µm cell strainer installed on a 50 mL conical tube. Use the back of a plunger from 1 mL syringe to mash up cell clumps on the filter. Wash the filter with 20 mL wash buffer (PBS + 2% Heat inactivated FBS + 2 mM EDTA).
13. Centrifuge at 500 x g at 4 °C for 5 minutes. Discard the supernatant and resuspend the cell pellet in 20 mL wash buffer.
14. Repeat steps 2.12 and 2.13 twice, changing the filter and tube each time. Pre-soaking the filter in wash buffer for more than five minutes increases the AM yield. Prepare single-cell suspension by adding 100 μL flow/sorting buffer to the cell pellet.
15. Block the nonspecific antibody binding to the Fc receptor by adding mouse Fc block (clone 2.4G2, 10 μg/mL) and incubating for 20 minutes on ice.
16. Perform antibody staining along with FMO and single-stain controls^11,25. Evaluate the lung cells by a cell analyzer (see Table of Materials) followed by single-cell flow cytometry analysis by analysis software.
17. Sort AMs by a cell sorter (see Table of Materials) directly into 1 mL mouse AM culture medium when AMs are intended for in vitro culture or in vivo cell transfer. Alternatively, sort AMs into 1 mL Lysis buffer supplemented with 10 μL/mL β-Mercaptoethanol for RNA extraction.

3. Isolation of AMs from Human BAL Fluid

1. Arrange transfer of the human BAL fluid from the clinic to the laboratory at 4 °C.
2. Centrifuge the BAL fluid (10 – 50 mL) at 250 x g at 4 °C for 10 minutes. The cell pellet contains BAL cells¹¹.
3. Wash the pellet with 20 mL wash buffer (PBS + 2% Heat inactivated FBS + 2 mM EDTA) and centrifuge at 250 x g at 4 °C for 10 minutes.
4. Resuspend the BAL cells are in 100 μL of flow/sorting buffer (PBS + 2% Heat inactivated FBS + 1 mM EDTA + 25 mM HEPES).
5. Block the nonspecific antibody binding to the Fc receptor by adding human Fc block (clone Fc1.3070, 25 μg/mL) and incubating for 20 minutes on ice.
6. Perform antibody staining along with FMO and single-stain controls^11,24. Evaluate the BAL cells by a cell analyzer (see Table of Materials) followed by single-cell flow cytometry analysis by analysis software.
7. Sort AMs by a cell sorter (see Table of Materials) directly into 1 mL human AM culture medium or 1 mL Lysis buffer supplemented with 10 μL/mL β-Mercaptoethanol.

4. In vitro Culture of AMs

1. Following the cell sorting, harvest the cells by centrifuging at 250 x g at 4 °C for 10 min.
2. Resuspend human AMs in 1 mL human AM culture medium [Roswell Park Memorial Institute medium (RPMI) 1640 supplemented with 10% FBS, 20% L-929 culture supernatant (as a source of macrophage colony stimulating factor (M-CSF), 100 U/mL final concentration), 1 mM sodium pyruvate, 10 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulphonic acid (HEPES), and 1x penicillin/streptomycin (optional)]. Similarly, resuspend mouse AMs in 1 mL mouse AM culture medium [DMEM supplemented with 10% FBS, 20% L-929 culture supernatant (as a source of M-CSF, 100 U/mL final concentration), 1 mM Sodium Pyruvate, 10 mM HEPES, and 1x penicillin/streptomycin (optional)].
3. Enumerate the trypan blue-excluding viable cells by a cell counter and adjust the viable cell concentration at 1 x 10⁶/mL.
   NOTE: Ideally, up to 5 x 10⁵ AMs can be isolated from BAL fluid collected from an 8 – 10 week old C57BL/6 male mouse. The AM yield from the mouse lung digest is variable and may be slightly less than that of the mouse BAL fluid. AM yield from human BAL fluid depends on the volume of the BAL fluid, the total volume of saline used in the lavage, and patient's clinical condition.
4. Based on the yield and experimental necessity, seed the cells in chamber slides, culture dishes or culture flasks. Ideally, seed the AMs at 1 x 10⁵/mL with 10 mL medium in a T25 tissue culture flask.
5. Incubate the cells in a 37 °C humidified incubator with 5% CO₂ atmosphere.
   NOTE: AMs will grow as adherent cells and are extremely slow-growing (doubling time >7 days).
6. At 24 hours post-seeding, AMs should be adherent and will not be detached by a gentle change of culture medium. Remove the medium by aspiration from one end and replenish with fresh medium pre-warmed to 37 °C.
   NOTE: The cells can now be used to assess AM physiology or the effects of stimuli. AMs grown in chamber slides can be conveniently stained with appropriate antibodies and are ideal for microscopic visualization. For flow cytometric evaluation, AMs are harvested by incubation with non-enzymatic cell dissociating solution.
7. Aspirate the culture medium and add dissociating solution enough to cover the culture surface (approximately 2 mL per T25 flask) and incubate at 37 °C for 5 – 10 minutes. No scraping will be necessary, and it is not recommended to scrape the cells mechanically. Gently tap the sides, add 1 – 2 mL flow/sorting buffer and gently pipette up and down to detach the majority of adherent cells.
8. For RNA studies, lyse the cells directly on the culture dish by adding Lysis buffer.