Background suppression in fluorescence nanoscopy with stimulated emission double depletion

Journal name:
Nature Photonics
Year published:
Published online


Stimulated emission depletion (STED) fluorescence nanoscopy is a powerful super-resolution imaging technique based on the confinement of fluorescence emission to the central subregion of an observation volume through de-excitation of fluorophores in the periphery via stimulated emission. Here, we introduce stimulated emission double depletion (STEDD) as a method to selectively remove artificial background intensity. In this approach, a first, conventional STED pulse is followed by a second, delayed Gaussian STED pulse that specifically depletes the central region, thus leaving only background. Thanks to time-resolved detection we can remove this background intensity voxel by voxel by taking the weighted difference of photons collected before and after the second STED pulse. STEDD thus yields background-suppressed super-resolved images as well as STED-based fluorescence correlation spectroscopy data. Furthermore, the proposed method is also beneficial when considering lower-power, less redshifted depletion pulses.

At a glance


  1. Scheme of STEDD microscopy.
    Figure 1: Scheme of STEDD microscopy.

    a, Sketch of the microscope, including the sequence of excitation and depletion pulses. b, Detailed temporal sequence of fluorescence excitation. Shortly after the excitation pulse, the first STED1 pulse (intensity profile visualized in the x–z plane) depletes the majority of excited fluorophores except for those near the centre. A fraction of fluorophores in peripheral regions of the observation volume still escape depletion or are re-excited by the STED beam. The second, weaker STED2 pulse (intensity profile also visualized in the x–z plane) depletes excited fluorophores near the centre but leaves those in the periphery unaffected.

  2. STEDD imaging of fluorescent beads.
    Figure 2: STEDD imaging of fluorescent beads.

    a, Images of 40 nm polystyrene beads in confocal, conventional STED and STEDD microscopy, acquired by applying the excitation pulse only, adding the conventional STED pulse and adding both STED pulses, respectively. Colour bar, counts per pixel; scale bar, 1 µm. b, Close-ups of regions marked by frames in a. c, Intensity profiles (open circles) along the dashed lines in a. Solid lines are Gaussian fits that yield FWHM values of 306 ± 12 nm and 62 ± 2 nm for confocal and STEDD microscopy, respectively. d, Bead imaged by using STED (top row) and STEDD (bottom row) microscopy with different depletion beam wavelengths. Confocal images are also shown for comparison. e, FWHM values of intensity profiles of beads imaged using STEDD microscopy, plotted as a function of the STED beam wavelength.

  3. 3D STEDD imaging of a fixed HeLa cell.
    Figure 3: 3D STEDD imaging of a fixed HeLa cell.

    a,b, Conventional 3D STED image (a) and 3D STEDD image (b) of a 5 × 5 × 1.7 µm3 intracellular region, with the microtubular network immunolabelled with Atto647N. A total of 46 frames were volume-rendered (Imaris, Bitplane) after scaling the raw data to the same intensities. Only pixels with intensities in the range 15–100% are shown, because lowering the threshold would fill the entire structure in a with background. Colour bar, 0–1.7 µm (blue, six frames; all others, eight frames per colour). Scale bar, 1 µm.

  4. STEDD imaging of microtubules in live COS-7 cells.
    Figure 4: STEDD imaging of microtubules in live COS-7 cells.

    a, Combined confocal and STEDD image of a COS-7 cell expressing the mGarnet–RITA fusion protein as a microtubule marker. Colour bar, counts per pixel; scale bar, 1 µm. b, Close-ups of the region marked in a taken in confocal, STED and STEDD modes. c, Intensity profiles along the white dashed line in a, yielding FWHM values of 297 ± 13 nm and 74 ± 4 nm for confocal and STEDD microscopy, respectively.

  5. STED–FCS and STEDD–FCS autocorrelation functions of dye diffusion.
    Figure 5: STED–FCS and STEDD–FCS autocorrelation functions of dye diffusion.

    Data were taken on Atto655 (200 nM) dissolved in a glycerol-phosphate buffered saline (PBS) mixture (72/28, wt/wt), taken using a first-order (left column) and second-order (right column) vortex phase mask and a cylinder phase mask. a,b, STED–FCS autocorrelation curves, with STED powers included with corresponding colours. c,d, STEDD–FCS autocorrelation curves. Dashed lines: autocorrelation curves with STED beam off (confocal FCS), rescaled for better comparison. e,f, Correlation time τD, plotted as a function of r02, the squared lateral dimension of the observation volume. Diffusion coefficients, D =19.1 ± 0.8 µm2 s−1 and 19.8 ± 0.5 µm2 s−1 for the first-order (e) and second-order (f) vortex data, respectively, were calculated from the slopes of linear functions (lines) fitted to the data (squares). These results agree with the reference value, D = 17 ± 3.4 µm2 s−1, obtained by rescaling the diffusion coefficient of Atto655 in pure water28, D = 425 ± 30 µm2 s−1, by a factor of 25 to account for the viscosity change between glycerol-buffer and water.

  6. 3D STEDD–FCS on annexin A5-mGarnet diffusing within a living HeLa cell.
    Figure 6: 3D STEDD–FCS on annexin A5-mGarnet diffusing within a living HeLa cell.

    a, Overview image of the cell. Squares indicate the locations at which data were collected. Scale bar, 10 µm. b, Autocorrelation data (open circles) obtained by confocal FCS, STEDD–FCS with first-order vortex phase mask and STEDD–FCS with second-order phase mask (averages of two FCS measurements of 400 s each). Solid lines, fits of autocorrelation curves; dashed lines, fit of the confocal FCS autocorrelation curve (shown in black), scaled to the amplitudes of the STEDD–FCS autocorrelation curves. We fitted the three curves globally by using Supplementary equations (2) and (3), taking the ratios between r0 and z0 from a calibration measurement (Supplementary Fig. 10) and sharing the saturation intensity Isat as a fit parameter. The resulting values for the diffusion coefficient of the fusion protein in the cytoplasm are D = 17.6 ± 0.8 µm2 s−1, 18.8 ± 1.0 µm2 s−1 and 17.5 ± 0.6 µm2 s−1, for confocal mode and 3D STEDD with first-order and second-order vortex phase masks, respectively. c, Histogram of diffusion coefficients obtained from 40 STEDD–FCS measurements on eight cells using the second-order vortex phase mask.


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Author information

  1. These authors contributed equally to this work

    • Peng Gao &
    • Benedikt Prunsche


  1. Institute of Nanotechnology, Karlsruhe Institute of Technology, 76344 Eggenstein-Leopoldshafen, Germany

    • Peng Gao,
    • Lu Zhou &
    • G. Ulrich Nienhaus
  2. Institute of Applied Physics, Karlsruhe Institute of Technology, 76128 Karlsruhe, Germany

    • Peng Gao,
    • Benedikt Prunsche,
    • Lu Zhou,
    • Karin Nienhaus &
    • G. Ulrich Nienhaus
  3. Institute of Toxicology and Genetics, Karlsruhe Institute of Technology, 76344 Eggenstein-Leopoldshafen, Germany

    • G. Ulrich Nienhaus
  4. Department of Physics, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801, USA

    • G. Ulrich Nienhaus


G.U.N. supervised the project in both its conception and execution. P.G. and B.P. implemented the STEDD technique on our STED microscope, performed measurements and analysed data. L.Z. provided cell samples. G.U.N. and K.N. wrote the manuscript with input from all other authors.

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The authors declare no competing financial interests.

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