Quality-Controlled Sputum Analysis by Flow Cytometry

This protocol resolves common challenges experienced with sputum processing that have made it difficult in the past to use with flow cytometry for high-throughput clinical diagnostic purposes. This protocol is designed to save time by using sputum weight to determine antibody and dye staining volumes. During the staining incubation, cell count can be performed for the downstream steps.

It is important to achieve an accurate cell count to determine the correct re-suspension volume to prevent any problems when running on the flow cytometer. This protocol can provide insight into areas of research that investigate lung health. For example, illnesses such as COPD, asthma, lung cancer, or the effects of COVID could be studied.

Demonstrating the procedure will be Dr.Michael Lai, a research and development scientist from my laboratory. To begin with, weigh the sputum sample to determine the volumes of dissociation reagents. Transfer the sample to the appropriately sized vessel based on the sputum weight, then add one milliliter per gram of 0.5%NAC and four milliliters per gram of 0.1%DTT.

Vortex the samples at maximum speed for 15 seconds and rock at maximum speed for 15 minutes at room temperature. Dilute this sample with 4X Hanks balanced salt solution or HBSS to neutralize the NAC and DTT. And after quick vortexing, rock the sample at room temperature for five minutes.

Filter cell suspension through a 100 micrometer nylon mesh cell strainer into one or more 50 milliliter conical centrifuge tubes to create a single-cell suspension. Pellet down the cells. After aspirating the supernatant, combine pellets in one 15 milliliter conical tube and wash with HBSS under the same conditions.

Resuspend the cell pellet in a volume of buffer determined by the initial weight of the sputum sample. Take an aliquot from the cell suspension and mix 10 microliters of the sputum dilution with 30 microliters of 0.4%trypan blue, then load into the counting chambers of a hemocytometer for a live/dead cell count. Label sputum in compensation tubes.

Add HBSS antibody and dye to each sputum cell tube and compensation tubes as indicated in the text. Add the sputum cells to the assay tubes and add compensation beads to the compensation tubes as mentioned in the text. Incubate all the tubes on ice protected from light for 35 minutes.

Then after filling the tubes with ice cold HBSS, centrifuge at four degrees Celsius for 10 minutes at 800 times G.For compensation tubes, aspirate the supernatant as close to the pellets as possible and flick the pellets to loosen. Next, add 0.5 milliliters of cold HBSS to the compensation tubes and store the tubes on the ice in a light protected area until needed for flow cytometry analysis. Then aspirate the supernatant from the sputum unstained, isotype, blood, and epithelial tubes.

Loosen the pellets by flicking the tubes. Add two milliliters of 1%PFA to the unstained and isotype tubes and 10 milliliters to the blood and epithelial tubes. Incubate the tubes on ice in a light protected manner for 30 minutes, then quickly vortex at maximum speed and again incubate for another 30 minutes.

Then fill the tubes with ice cold HBSS. Next, centrifuge the tubes at four degrees Celsius for 10 minutes at 1, 600 times G.Aspirate as much supernatant as possible without disturbing the cell pellet and flick the tube with the fingers to loosen the cells, then add 200 microliters of cold HBSS to the unstained and isotype tubes. Calculate and add the required volume of HBSS for resuspension of the blood and epithelial tube according to the total cell count.

Store all the samples in compensation tubes protected from light on the ice at four degrees Celsius until flow cytometry analysis is performed. Apply appropriate startup procedures for the flow cytometer being used. Use the mixture of National Institute of Standards and Technology or NIST beads to ensure that forward scatter and side scatter voltages are set to place the NIST beads to span the plot without placing the beads too close to the axes.

Set the flow rate to medium for LSRII or high for Navios EX.Adjust the voltages for each parameter used for the scatter and fluorescent parameters to place the cell populations on scale. Use the figures with the gating strategy as guidance to adjust the voltages accordingly. Acquire data for the unstained sputum sample first, followed by the isotype stained sample, and then the blood tube and the epithelial tube.

Sputum weight was used to determine antibody or dye volumes for staining. The correlation between sputum weight and cell yield was not strong. Sputum samples were divided into four weight categories, which clustered in the distribution of cell yield.

Each antibody used was titrated in a concentration on the plateau phase with the highest staining index was chosen as the working concentration. In all weight categories, the average SEC contamination was less than 20%with the percentage of viable cells shown. Presented is the gating strategy used for analysis.

NIST beads were used to set a gate which was applied to the sputum sample to eliminate debris. Small debris was further eliminated by the width gate. Viability dye was used to eliminate dead cells, while the singlet gate removed doublets.

The anti-CD45 antibody labeling allowed the separation of live single sputum cells into a blood cell and a non-blood cell compartment. The profiles obtained from Navios EX, LSRII, and Lyric were compared. The gating strategy for eliminating the debris, dead cells, doublets, and comparing blood and non-blood profiles is shown.

This technique allowed us to apply these methods to detect lung cancer risk for use as a non-invasive clinical test.