VAMP-seq Tips

VAMP-seq is a valuable tool for identifying loss-of-function variants for a protein when you don’t have a specific assay for characterizing that protein’s activity. This is because almost all proteins can be “broken” by mutations that cause the protein to mis-fold, mis-traffick, or any other perturbations that may cause the protein to disappear from its normal cellular compartment. In the case of PTEN, we identified 1,138 variants that were lower than WT abundance. Notably, ~ 60 % of variants characterized as pathogenic in people were loss-of-abundance variants, confirming the importance of this property.

VAMP-seq uses a genetic fusion between your protein of interest and a fluorescent moeity (eg. EGFP) to assay its steady-state abundance. Fusions with unstable variants of your protein results in cells expressing unstable fusion proteins that don’t really fluoresce. In contrast, fusions with stable variants (such the WT protein) may fluoresce brightly. Unlike western blots which are ensemble measurements, this assay can be performed at a single-cell level. Correspondingly, this single-cell fluorescence readout makes it possible to test a large library of variants in parallel, in a multiplexed format.

Two VAMP-seq orientation discussed here

Still, when thinking about using VAMP-seq, you must first consider its limitations. While strong loss-of-abundance variants will be non-functional, variants of intermediate abundance may still be sufficiently functional in many contexts. Furthermore, while low abundance correlates with inactivity (and pathogenicity, in many clinical genetics contexts), WT-like abundance in no way indicates activity (or benignity when observed in people). For example, active site mutants often destroy protein function while having little to no effect on protein folding and abundance.

There are also proteins that are inherently incompatible with VAMP-seq. Secreted proteins won’t work because you lose the single-cell, genotype-phenotype link needed for the single-cell assay to work. Marginally stable or intrinsically disordered proteins likely won’t work due to a lack of destabilizing effect. Obligate heterodimers won’t work, though you may be able to get around it by overexpressing the protein partner, such as what I did with MLH1 for assessing PMS2 (See Fig 6). Proteins that cannot be tagged are problematic; this likely includes proteins that normally exist in crowded complexes, or that have key trafficking motifs on their termini. Proteins that are toxic to cells when overexpressed also poses problems, though one of my new landing pad platforms may help with that.

If your protein of interest passes those criteria, the rest is empirically confirming that there is enough signal over background to run the assay. A good literature search is a great place to start. You should look for 1) evidence that an N- or C-terminal tag works well (both good expression and normal protein activity), and 2) known destabilized variants that could serve as controls when performing the preliminary experiments. Ideally, there will be a clear difference in fluorescence distributions that are separable by thresholds used in FACS sorting, and that these differences are physiologically meaningful (as far as we know). Comparing /correlating the results of western blotting with the fluorescence distributions of EGFP-tagged protein is helpful (see below for PTEN). If the initial EGFP distributions between WT and the destabilized variants don’t seem super crisp, see what it looks like when you take the EGFP:mCherry ratio. As you can tell in the below figure, the EGFP:mCherry ratio is quite handy for increasing the precision of each distribution, as it divides out much of the heterogeneity in transcription / translation between cells.

Western and flow results

Regardless, I recommend to most people that they clone both N- and C- terminal fusions, and minimally look at the MFI values of the cells expressing each fusion, as low MFI will likely mean low dynamic range of the assay (from too little signal over background). Ideally, both WT and controls will be tested in both contexts. If the signal is relatively high but there’s concern that the large GFP fusion is causing problems, you could try 1-10/11 split EGFP. This only requires fusion with the ~ 15aa beta-strand 11 of EGFP (and separately co-expressing the larger fragment in the cells), though it’s not completely free of steric hindrance as it requires the spontaneous complex formation of the two subunits for fluorescence. This format also worked with PTEN (See SFig 1d), though I noticed a ~ 10-fold hit in overall fluorescence using the splitGFP format.

Fluorescence levels from different formats

Once it passes all those tests, then follow the steps in the Nature Genetics paper. Good luck!