Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

The overall goal of this procedure is to isolate human and mice alveolar macrophages from lungs and bronchoalveolar lavage fluid and develop the method for their in vitro culture. This method can help answer some key questions in the field of pulmono immunology, such as the role of alveolar macrophages in respiratory infection, inflammation and autoimmunity. The main advantage of this technique is to be able to distinguish the alveolar macrophages from the other lung leukocytes and obtain a population of highly purified alveolar macrophages for experimental purposes.

Place an anesthetized mouse on a dissection surface with the ventral side facing up. Apply ophthalmic vet ointment on the eyes to prevent dehydration. Wipe the entire ventral surface of the mouse with sterile alcohol prep pads, saturated with 70%isopropanol, to disinfect.

Mount the mouse with all four legs retained. 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.

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 the lungs while opening the thoracic cavity.

Locate the inferior vena cava and make an incision to bleed the mouse. Use absorbent cotton pads to soak up blood. Inject 10 milliliters of ice cold phosphate buffered saline into the right ventricle 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. 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. Next, make a small incision 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.

The single most critical step in collecting BAL fluid is to securely attach a catheter to the trachea that will minimize leaks during lavage. Insert a one inch 22 gauge catheter without a needle into the trachea towards the lungs. Secure the catheter with a silk braided suture and a square knot.

Having sturdy hands and for following this step, slowly and gently prevents rupture of the trachea and lungs. Attach a one milliliter syringe, filled with ice cold BAL buffer 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 milliliter conical tube, placed on ice.

Repeat the inflation and deflation nine more times, and pool the lavage fluid. Do not exceed one milliliter of buffer per flush, as that may exceed the lung capacity and lead to its rupture. Centrifuge the 10 milliliters of BAL fluid at 250 times g and four degrees Celsius for 10 minutes.

The cell pellet contains BAL cells. Resuspend the BAL cells in 100 microliters of flow sorting buffer. Proceed to stain and sort the cells as described in the text protocol.

Prepare an anesthetized mouse for dissection, perform incisions and bleed the mouse as before. Inject 10 milliliters of ice cold PBS into the right ventricle to flush circulating blood cells. Once completely perfused, the lungs will appear blanched and are ready for harvest.

Dissect out the lungs by severing the trachea, blood vessels and ligaments into a 15 milliliter conical tube, with five milliliters of cold DMEM. Maintain it on ice until the next step. Transfer the perfused lungs into a sterile and pyrogen-free 60 millimeter culture dish with three milliliters of DMEM.

With the 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 less than one millimeter size. Add 300 micrograms per milliliter of Liberase TL, and five units per milliliter of DNAse one.

Mix by pipetting and incubate at 37 degrees Celsius in an incubator for 25 minutes. After 10 minutes, gently mix once by pipetting. Pass the dissociated lungs through a 100 micron cell strainer installed on a 50 milliliter conical tube.

Use the back of a plunger from a one milliliter syringe to mash up cell clumps on the filter. Wash the filter with 20 milliliters of wash buffer. Centrifuge at 500 times g and four degrees Celsius for five minutes.

Discard the supernatant, and resuspend the cell pellet in 20 milliliters of wash buffer. Repeat the straining and centrifugation steps twice, changing the filter and tube each time. Finally, prepare the single cell suspension by adding 100 microliters of flow sorting buffer to the cell pellet.

Perform antibody staining and cell sorting, as described in the text protocol. Following the cell sorting, harvest the cells by centrifuging at 250 times g and four degrees Celsius for 10 minutes. Resuspend the mouse AMs in one milliliter of mouse AM culture medium.

Based on the yield and experimental necessity, seed the cells in chamber slides, culture dishes, or culture flasks. Ideally, seed the AMs at 100, 000 per milliliter with 10 milliliters of medium in a T 25 tissue culture flask. Incubate the cells in a 37 degree Celsius humidified incubator with 5%carbon dioxide atmosphere.

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 degrees Celsius. Finally, observe the flask under an inverted microscope.

Flow cytometric assessment of alveolar macrophages in mouse broncho alveolar lavage fluid is shown. The alveolar macrophages are positive identified for being CD45 positive, CD11c positive, and Siglec-F positive cells. Shown here, is a representative result of mouse alveolar macrophages cultured in vitro for 24 hours.

The cells have become adherent and a gentle change of culture medium will not detach them. Once mastered, this technique can be done in approximately two hours, if it is performed properly. After watching this video, you should have a good understanding of how to distinguish the mouse and human alveolar macrophages from other lung cells.

Also, as the procedures for obtaining a beautiful population of alveolar macrophages for downstream experiments. While attempting this procedure, it's important to remember that not all lung resident macrophages or phagocytes are alveolar macrophages. The alveolar macrophages are a type of specialized cells present in the lungs since birth.

For isolation of alveolar macrophages, the focus should be on viable extraction methods from the alveolar micro environment. The in vitro culture method developed here supports growth of alveolar macrophages in a laboratory condition that can be utilized in molecular and cellular assays. Following this procedure, other methods like adoptive cell transfer, antigen presentation and evaluating response to inflammatory stimuli can be performed in order to answer additional questions on the biology of alveolar macrophages.