Troubleshooting Tip 5: Fluorescently-tagged Protein fluorophore bleaches too rapidly





These micrographs show epithelial tissue labeled with a plasma membrane marker fused to a fluorescent protein (FP). As one can see the flurophore bleaches very rapidly during image acquisition. Here we discuss issues to consider in order to minimize photobleaching of the fluorophore during image acquisition

Possible Causes and Solutions

You need to find a trade-off between the desired spatial resolution, the signal-to-noise ratio, and the speed of image acquisition based on the experimental requirements. Minimize acquisition photobleaching by adjusting one or more of the following parameters:

  • Decrease the excitation light intensity and increase the gain of the detector.
  • Zoom out: the rate of photobleaching is proportional to the square of the zoom factor.
  • Reduce the frame size (number of pixels per frame) (e.g. 512x512 instead of 1024x1024).
  • Reduce the exposure time / increase the scan speed.
  • Reduce signal (line or frame) averaging.
  • Scan only the region of interest (ROI) and not surrounding regions without zooming in ("edit ROI" function in some commercial setups).
  • Minimize the time that the sample or the ROI is exposed to light before the actual acquisition (e.g. when looking for the field of interest using epifluorescence).
  • If the FP still bleaches rapidly consider choosing a fluorophore that is more photostable. Use tables that list the optical properties of FPs to compare the photostability of the FPs (expressed as the time to bleach to 50% emission intensity, typically normalized to the one of EGFP).

    The Fluorescently-tagged Version of My Protein is Very Dim to Work With. What Did I Do Wrong?





    These micrographs show tissue underoing mitosis labeled with a microtubule marker fused to two different fluorescent proteins (FP). As one can see the flurophore on the left is much brighter than the one on the right (imaging conditions and expression levels are all identical between the two cases). Here we discuss how to optimize working with dim samples.

    Possible Causes and Solutions

    When the FP is inherently dim or the expression levels are low one needs to increase the signal-to-noise ratio with minimal acquisition photobleaching. Thus one needs to adjust all the parameters discussed earlier (see Troubleshooting Tip above). Again, one needs to to find a trade-off between the spatial and temporal resolution and the signal-to-noise ratio. Additional issues to consider include the following:

  • Open the pinhole: opening the pinhole to 1.5 to 2 Airy units will significantly improve signal intensity compared to a pinhole of 1 Airy.
  • Work at lower magnification and use high NA lenses for maximal signal collection: The brightness of the acquired image is proportional to NA4/magnification2 (a 40× lens will yield brighter images than 60× or 100× lenses of the same NA with the same resolving power).
  • Use an excitation wavelength that matches the excitation peak of the FP.
  • Use a sensitive detector (e.g. EMCCD cameras).
  • If the signal is too low to work with consider choosing a brighter FP. Use tables that list the optical properties of FPs to compare the brightness of the FPs (expressed as the product of the molar extinction coefficient and the quantum yield time, typically normalized to the one of EGFP).
  • Check the environment/topology of the fluorophore. For example, the fluorescence is quenched in acidic compartments such as the lumen of lysosomes. Use FPs that are not sensitive to acidic pH (pKa < 5.0).
  • Fuse two copies of the FP (tandem dimers) to your protein (depending on whether the protein tolerates such fusions).
  • More information on fluorescent proteins and their properties can be found in Current Protocols in Cell Biology, Unit 21.5 and Unit 21.6. Information on how to design fluorescent protein fusions and on how to optimize imaging parameters can be found in Current Protocols in Cell Biology, Unit 4.5, Unit 4.18 and Unit 21.4.

    Visit for tools, calculators, apps, videos, and information on all Current Protocols methods.

    Contributed by: Manos Mavrakis.