When doing the Green FPs in HEK 293T cells experiment, we noticed how the same fluorescent protein, EGFP, could have vastly different brightness depending on the construct we were using.

Put another way, cytoplasmic EGFP gave us really high green fluorescence intensity, but a different construct wherein that same EGFP sequence was preceded by a signal sequence (thus causing it to become a secreted / extracellular protein), and succeeded by a transmembrane domain (thus causing the extracellular EGFP molecule to be anchored to the plasma membrane) gave us cells that were roughly 30 to 100-fold less bright. The amount of fluorescence of the transmembrane version was further susceptible to other sequence considerations; for example, addition of a his tag right after the signal sequence (such that the his tag is the most distal sequence on the protein, with the tag flapping around as part of a flexible N-terminus), resulted in a ~ 3-fold reduction in fluorescence as compared to an untagged version. My guess is that this repetitive, pseudo-charged region was interfering with efficient translation into the rough ER, but who knows.

Still, how do I explain this result? Well, I have no evidence-supported answer, but my guess is that it’s about translation process bandwidth and overall real-estate. I’m guessing that cytoplasmic translation and accumulation has pretty high bandwidth, where there are plenty of ribosomes to translate cytoplasmic protein, and there’s plenty of space to accommodate them. In contrast, I’m assuming there are comparatively fewer ribosomes capable of translating transmembrane proteins at the rough ER, and that the overall real-estate on cell-associated membranes (particularly in the vesicular pathway leading up to and including the plasma membrane) is also less (while an imperfect approximation, I’m thinking of it kind of like a difference in surface area or volume of a sphere, type of thing). Although who knows; maybe that’s all incorrect, and it’s more about the signal sequence (possibly from CD8?) and the transmembrane domaine sequence (seemingly from PDGFR-beta) that I used.

Edit 1: Although again, I need to keep reminding myself. Since these are cell surface proteins on adherent cells, some of the reduced signal with the transmembrane protein may be due to some proportion of proteins getting cleaved off the cells during routine trypsinization. I’ve talked to Olivia about trying a side-by-side experiment of resuspending the cells with trypsinization or Versene with gentle agitation. Stay tuned to see if I need to update the above plot or not!

At one point, I was a doe-eyed postdoc reading about new fluorescent proteins (FPs) with improved brightness and thinking it was potentially important to incorporate new FPs into my constructs for cell culture work. I have since come to realize that the intrinsic gains to fluorescence published in those papers do not necessarily translate to brightness in my experiments. The reason are probably multifactorial, including:
1. Commonly accessible equipment is generally optimized for EGFP (or similar FPs), so newer FPs with slightly different excitation and emission spectra may not be captured well with existing microscopes or flow cytometers.
2. The FP brightnesses are typically assess in-vitro, and there may be other factors in eukaryotic cell cytoplasms that may affect the FP brightness (eg. FP half-life).

Well, I’ve still ordered a handful of different green fluorescent proteins anyway, and figured it was worth doing a side-by-side comparison in the transgenic system we use in the lab. This was all done with the HEK293T G542Ac3 (LLP-Tet-Bxb1attP_Int-BFP_IRES-iCasp9-BlastR) cells, which were stably recombined with a single copy of each fluorescent protein at a common genomic locus. The construct organization was: Bxb1attB_[Green FP]_IRES-mCherry-2A-PuroR. Olivia did these recombinations, selected the cells, and ran the cells on the ThermoFisher Attune Flow cytometer, with Sarah’s help. This is what the results look like:

Some interpretations:
1. Rather minimal (~3-fold) difference between mGreenLantern and UnaG. I suppose if we were ever in a situation where we needed every unit of green brightness possible, that we would go with mGreenLantern. That said, UnaG has some benefits; namely, it’s 58% the size of EGFP/mGreenLantern/mNeonGreen, and it lacks the VERY ANNOYING identical sequences at the N- and C-termini (MVSKG … DELYK) which makes molecular cloning a potential pain.
2. Conceptually, I like the idea of fuGFP, but that 20-fold diminished green fluorescence compared to EGFP is potentially problematic. Who knows, maybe I’ll turn this into a target of directed evolution at some point…?

Obviously we make a lot of recombinant DNA constructs and create a bunch of different assays to try to understand biology. Well, at some point I became curious if I could come up with a protease cleavage assay for measuring how various peptide sequences could be substrates for vesicular proteases inside cells. I figured we’d start to test this by looking at furin cleavage, since 1) furin is quite ubiquitous, 2) furin cleavage is relevant for multiple disease-relevant membrane proteins, like many viral entry glycoproteins, and 3) including SARS-CoV-2 spike, where the furin cleavage site in that protein is thought to contribute to its dynamics of spread and pathogenicity during infection.

Well, the first construct that may have worked is a pretty simple one, where I have EGFP targeted to extracellular space using an N-terminal signal peptide, but retained on the cell surface by having it C-terminally fused to a transmembrane domain. Anh has actually been using this construct as part of his undergraduate research project. Well, to turn this construct into one that can study Furin cleavage, I modified it to encode a R-R-A-R peptide between EGFP and the transmembrane domain. Thus, if Furin cleaves this construct, the EGFP protein is now no longer tethered to the cell, and presumably escapes into the media once it reaches the cell surface.

So this is an N=1 experiment so far, so it’s not the most conclusive. That said, there does seem to be decent separation between the construct with and without the furin cleavage site, where cells with the construct with the EGFP that potentially gets released had ~ 10-fold less fluorescence than the construct that could get released. I suppose if this reproduces, it could be worth trying to turn this into a library-based experiment for studying protease cleavage within intracellular compartments.

… Though now that I’m thinking about it further, I’ll definitely need to ask how these cells were prepped. If they were prepped with trypsin (which is likely), this was likely more of a trypsin cutting assay rather than a furin one.

Edit 10/27/22:
Well, so to avoid the whole trypsin complicated, recombined the above constructs in suspension HEK cells that are floating and don’t ever need to be detached off the plate for flow. Here’s what that plot looks like.

So ya, we’re still seeing the ~ 10-fold effect there, so it looks like that’s the real dynamic range of the assay for measuring furin protease cleavage.

I had a product that could have benefitted from using split mCherry to serve an AND function. Put the split mCherry in my usual mCherry spot in the recombination vector (2A’d with Puromycin, also), and I couldn’t see any visible fluorescence when both the small and big fragments were in the same cell. After some time, I saw a paper using split super-folder mCherry fused with the Spycatcher system, so I used that and that seemed to allow us to now see a shift in fluorescence off the background. I found another paper using an improved split super-folder mCherry (sfmCherry3C), and tested that, which seemed to work slightly better.

I think synthetic biology is really cool, and I like playing around with recombinant DNA elements so I can see how well they work in my own hands. If they work OK, then I just let that knowledge stew in the back of my brain until I can eventually figure out a use for it. Reading this paper by Martin Fussenegger made me realize just how easy it is to make cultured cells express melanin. Here was my first foray in creating melanin in HEK cells by overexpressing tyrosinase/

Doesn’t quite work well enough to use as a general reporter (it’s really hard to tell in a cell monolayer, and only becomes noticeable as colonies of cells or in a pellet, like above), but still kind of fun to see. Let’s see if I find an eventual use for this in some future work.