Fluorescence Microscopy


Fluorescence microscopy is an essential technique that allows scientists to visualise molecules (proteins, nucleic acids, ions, metabolites, carbohydrates and lipids), large structures and whole cells in fixed and living specimens as well as single molecules, assemblies and enzymes in vitro. Using multiple different imaging modalities, scientists have adapted fluorescence microscopy to advance our knowledge in all areas of biology and across length scales that range from tens of millimetres to a few nanometres.

Key Concepts

  • Fluorescence is an intrinsic property of some molecules and proteins that causes them to absorb and then emit light at given wavelengths.
  • Fluorescence imaging systems are built to excite molecules and then collect the emitted photons.
  • There are many different imaging techniques that can be used for interrogating different types of specimens and obtaining different kinds of data.
  • Localisation of molecules is only one of many readouts that scientists can obtain from fluorescence microscopy.
  • Imaging resolution has been limited to a diffraction limit based on the nature of our ability to collect fluorescence light through the objective, but new techniques have allowed for imaging to almost arbitrary precision.

Keywords: fluorescence microscopy; CCD camera; confocal laser scanning; spectral analysis; optical filters; living cells

Figure 1. (a) Excitation and emission spectra for FITC are shown. The difference between the excitation and emission spectra is the Stokes shift (image courtesy of Chroma Technology Inc.). (b) A Jablonski diagram shows that an electron interacting with an incoming photon of light of the appropriate wavelength (0) and being excited into a higher energy state (S1Vib). Some of the energy is lost by vibrational radiation to reach S1. The electron then returns to a near ground state by emitting a photon of light of a longer wavelength (1). (c) The light path of an epifluorescence microscope – white light is directed into an excitation filter that allows one or more selected bands of light to pass; the excitation light is then reflected off a dichroic mirror and focused by the objective into the sample; emitted photons and reflected photons are collected by the objective, hit the dichroic and the emitted photons are passed through to the detector while the excitation photons are blocked. (d) An example of a triple bandpass excitation filter, dichroic beamsplitter and emission filter set that can be used in a filter cube, coupled with separate emission and excitation filters to perform three‐colour fluorescence images. Image courtesy of Chroma Technology Inc.
Figure 2. (a) In a primary immunofluorescence assay, an antibody that is directly conjugated to a fluorophore recognises its antigen, and its position is directly detected by fluorescence microscopy. In indirect immunofluorescence, the fluorophore is attached to an antibody that detects the primary antibody. (b) An example of a fluorescence image that combines indirect immunofluorescence (a mouse antibody that detects tubulin and a rhodamine‐labelled anti‐mouse secondary that detects the primary antibody), a GFP‐tagged protein and DAPI staining of DNA to label chromosomes (scale bar, 5 µm). (c) An example of a confocal light path – light from lasers is fed into an acoustooptic tunable filter (AOTF), which selects the excitation wavelength, and then focused through a lens to a pinhole which blocks nonfocused photons; the lasers are then reflected off a dichroic and focused through an objective lens; emitted light from the focal plane is then refocused back through the objective, passes through the dichroic and a pinhole to reach the detector. Light from above and below the focal plane are blocked by the pinhole. (d) Two‐photon excitation of a fluorophore produces the same excitation as single‐photon excitation.
Figure 3. (a) A voxel contains many fluorescent molecules that are diffusing in and out of the observable volume. The rate of fluorescence counts for rhodamine B over 100 s (upper) and the autocorrelation (G(τ)) of counts over time (τ(s)) (lower) are shown. (b) In a FRAP experiment, a region is photobleached and the fluorescence recovery is measured to generate curves like what is shown in (c). (c) Before photobleaching, the fluorescence is 100%, and the immediate recovery of fluorescence over the first 20 s shows the reassociation of proteins with the bleached structure. (d) An example of a FRET biosensor that detects phosphorylation by a particular kinase. When the substrate is not phosphorylated, FRET does not happen. Upon phosphorylation, the phospho‐tyrosine binding motif will fold back and lead to high‐efficiency FRET.
Figure 4. (a) A structured illumination image of a Drosophila S2 cell expressing myosin regulatory light chain‐GFP before and after processing. After processing, the two ends of the bipolar thick filaments (BTFs) and tightly packed BTFs can be resolved. (b) An example of a PALM experiment. The wide‐field image is shown and example frames showing the stochastic activation of diffuse fluorophores, each of which is fit to a very precise spot (shown by the cross). At the end, the reconstructed PALM image is built of the thousands of frames and all of the localised spots to build structures with much higher resolution. (c) An example of a stimulated emission depletion (STED) setup. In the standard confocal modality, the laser spot is swept through the field, but in STED, a donut‐shaped depletion beam surrounds the diffraction spot and effectively reduces the size of the spot and breaks the Abbe resolution limit. (d) A Jablonski diagram shows the principle of STED. In standard fluorescence, the excited electron moves from S1 to S0Vib and emits a photon of a wavelength (1) that can pass through an emission filter. In STED, the longer wavelength (2) depletion beam drives the electron to a higher ground state to stimulate the emission of a photon of the same wavelength (2) that can be blocked by the emission filter.


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Further Reading

Enterina JR , Wu L and Campbell RE (2015) Emerging fluorescent protein technologies. Current Opinion in Chemical Biology 27: 10–17.

Ettinger A and Wittmann T (2014) Fluorescence live cell imaging. Methods in Cell Biology 123: 77–94.

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Nienhaus K and Nienhaus GU (2015) Where do we stand with super‐resolution optical microscopy? Journal of Molecular Biology, pii: S0022‐2836(15)00711‐1.

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Griffis, Eric R(Mar 2016) Fluorescence Microscopy. In: eLS. John Wiley & Sons Ltd, Chichester. http://www.els.net [doi: 10.1002/9780470015902.a0005780.pub2]