Best Way to Quantitatively Measure Autophagic Flux

By Christina Towers, PhD

Autophagy is a stress-induced cellular recycling process that plays an important physiological role in many diseases. It is induced by a variety of stimuli, both intracellular and extracellular, and is a tightly regulated process with over 30 core autophagy proteins necessary for efficient autophagosome formation and fusion with lysosomes to facilitate degradation¹. One of the most common ways to monitor autophagy is by measuring the protein levels of LC3-II, which is incorporated into autophagosomes and then degraded in the lysosome. There are many different ways to measure LC3 protein levels, including Western blot and immunofluorescent microscopy². These methods, however, are complicated by the fact that LC3 levels increase with autophagy induction due to increased autophagosomal formation but also decrease as these forming autophagosomes are turned over, and the only way to accurately monitor LC3 levels is with a flux assay using an autophagy inhibitor like Bafilomycin-A1 or Chloroquine.

Tandem constructs have been designed that couple pH-stable fluorophores like RFP or mCherry with the pH sensitive fluorophore, GFP, to monitor LC3 turn over in the lysosome. Attempts to quantify LC3 foci with microscopy can introduce experimenter bias, so these flux assays are qualitative at best.

Tandem mCherry-GFP-LC3 and Ratiometric Flow Cytometry

A more quantitative assay to measure autophagy couples the tandem mCherry-GFP-LC3 construct with ratiometric flow cytometry³. Images from a recent publication⁴ (Panels a and b) exemplify how this assay can be used to detect small changes in autophagic flux in cultured cells at both baseline and after treatment.

(a) To perform this assay, cultured cells with stable expression of mCherry-GFP-LC3 were treated with an experimental stimulus and then harvested for flow cytometry. The necessary controls include cells treated with Bafilomycin-A1 for 24 hours to completely inhibit autophagic flux, which will be used to set the flow cytometry gates. (b) Cells starved of amino acids and glucose for 4-24 hours serve as a positive control for induced autophagy, and unstained cells are used to eliminate low expressing cells from analysis.

Forward scatter and side scatter are used to gate live and single cells. An angled rectangular "autophagy" gate can then be set based on the Bafilomycin-A1-treated cells, such that 5-10% of cells score positive in this condition, and 50-80% of the cells shift left into this gate after starvation. Flow cytometry software like Flow Jo can run a ratiometric calculation as a derived parameter, where autophagic flux is the mCherry signal divided by the GFP signal. This assay is simple and quick with a 1-2 hr process time after treatment and is the most quantitative way to measure autophagic flux.