HaloTag pulse-chase labeling for live imaging

Hello all,

I have very recently incorporated a HaloTag strategy for pulse-chase labeling in live cells into my project. I have checked a couple of protocols for live cell imaging using HaloTag ligands but the dilution (final concentration rarely mentioned), vehicle, duration of the pulse, washes, etc. vary widely. I guess they are optimized for different applications but I am quite confused and as a novice in the technology I don’t want to waste reagent (and cells… and time…). Does anybody have any guidelines and tricks that work?

I have HaloTag ligand conjugated to Janelia Fluor dyes as well as AlexaFluor660.

Thanks in Advance!

One crucial control that might help you narrow down dilution and wash conditions is adding ligand to cells that don’t contain a HaloTag. If you see nonspecific binding under your labeling conditions, increase washes or decrease ligand concentration, and if all else fails try a different ligand. I don’t have first-hand experience with Halo ligands, but I’ve seen significant nonspecific binding with SNAP ligands. See this paper for more examples.

Once you have a handle on nonspecific binding, you’ll have some constraints on the conditions to test in your pulse-chase experiment.


Thank you, @Anna! I will definitely do this control :smiley:

My experiences are most with Halo-JF dyes + mammalian cultured cells (e.g., HeLa, Huh7, and SUM159), either with the transient expression system (under CMV promoter for strong expression or TK promoter for weak expression) or endogenous tagging with CRISPR genome engineering. Based on the titration test, I realized that we could use a very low amount of Halo dyes for efficient staining. Here is my protocol (for 35 mm glass-bottom imaging dish):

-Make a 0.5 mM stock solution in DMSO, store at -20C

  1. Add 0.1-0.4 microliter of the stock solution to cells with 2 ml of pre-warmed complete media (1:5000 dilution, final Halo-dye concentration= 25-100 nM)
  2. Incubate in 37 C 5% CO2 incubator for 1 hour
  3. Wash the cell with 2 ml of the pre-warmed complete media for three times
  4. Add 2 ml of the pre-warmed complete media and incubate cells at least 1 hour in 37C 5% CO2 incubator before imaging


  1. Depending on cell type, membrane lipids have a different affinity to Halo dyes (= non-specific background). So, as Anna recommended, do “no transfection + dye incubation” control.
  2. Once the expression of Halo-fused protein is very low (such as for endogenous tagging), the efficiency of Halo staining goes down. We suspect that this is due to a lower chance of Halo-protein meets Halo-dyes. To boost a staining efficiency, you may increase incubation time of Halo-dye with your cells (like overnight)

Thank you so much for your help, @jeeyun! :smiley: This is really useful. I tried to stain last week for the first time and I didn’t get to see much. I’ll try your final concentrations. I though it didn’t work because I am leveraging ultra weak very crippled promoters to get expression levels as similar as possible to the endogenous level… but if you have tried it with endogenous tagging, it should definitely work.

I saw a tendency that protein with lower expression has lower efficiency of Halo staining. You may try overnight staining (with 100 nM concentration) to see whether you can overcome low chance of interaction between HaloTag (enzyme, haloalkane dehalogenase) and Halodye (ligand). Higher concentration + 1hr staining didn’t work for me with a protein of the low expression. Good luck :slight_smile:

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Hi Irene,

the labeling that you will obtain is influenced by several aspects, including the membrane permeability of the dyes you are using (i.e. how long does it take to them to enter the cells?), efficiency of the reaction, fluorogenic properties, accessibility of the tag, etc.
In my experience, good Halo-ligands for live imaging (JFs-Halo, SiR-Halo, 580CP-Halo, 610CP-Halo for example) stain efficiently endogenously tagged proteins in 15-30min of incubation already (1uM in HDMEM) and it never failed. In pulse chase experiments, I used 1uM for 1h, since I wanted to stain really everything, at 5h intervals.
In the end, everything has to be tuned on your experiment, your protein, and on your question: do you want short pulse chases or long pulse chase? How do you want to visualize the protein? How abundant is it and where is it localized (cytoplasmic targets are easier to reach, for instance)?
If you have never tested it before, might help starting with something “easy” and that should work (e.g. overexpression) and then moving to nasty, low copy targets.
Good luck!