Fluorescence microscopy is an ideal technique for examining fixed and living specimen. Using the phenomenon that certain materials emit energy detectable as visible light when excited with the light of a specific wavelength, fluorescence imaging allows for the selective and specific detection of molecules at small concentrations with a good signal-to-noise ratio. Furthermore, in combination with confocal microscopy the detection of photons is restricted to photons that originate from the focal plane, neglecting any signal arising from out-of-focus regions.
In short, the basic principle of fluorescence entails the following (Figure 1): A photon with the energy hνex, is supplied by an external source, usually a laser of well-defined wavelength, and is absorbed by a fluorescent molecule. The absorption raises its energy level to an excited, unstable electronic singlet state (S1') as depicted by the green arrow. This excited state is rather instable and thus has a very short lifetime. The excited fluorescent molecule relaxes towards the lowest vibrational energy level within the electronic excited state (S1), loosing the energy in the form of heat (yellow arrow). Fluorescence emission originates from the drop from the excited state S1 to the ground state S0 (red arrow). The energy of the photon that is emitted in this process, hνem, is exactly the difference between S1 and S0. For most fluorophores used in biological applications the light emitted is at a characteristic wavelength that is determined by the difference in energy between S1 and S0.
Figure 1: The basic principle of fluorenscence is illustrated in a Jablonski diagram. Green arrow: excitation from ground state S0 to the excited state S1' by absorption of a photon with energy hνex. Yellow arrow: relaxation from S1' to the lowest vibrational energy level within the electronic excited state S1. Red arrow: emission of a photon with energy hνem, bringing the fluorescent molecule back to ground state S0.
In our applications a confocal microscope is usually used to measure the intensity of the emitted fluorescence signal and create a digital image. In contrast to widefield microscopy confocal microscopy has an increased optical resolution and contrast. It uses point illumination and a spatial pinhole in front of the detector to restrict passage of light that comes from the plane of focus (Figure 2). Out-of-focus light from specimen that are thicker than the focal plane is thereby eliminated. The thickness of the focal plane is largely determined by the emission wavelength and the numerical aperture of the object lens. In confocal microscopy only one point in the sample is illuminated at a time. In order to take 2D or even 3D images one must therefore scan over the specimen. Post processing of images taken by confocal microscopy makes it possible to depict and quantitate the obtained signals.
Figure 2: Upper panel: The microscopy apparatus of a confocal laser-scanning microscope. Lower panel: Schematic Representation of the function of the pinhole in confocal microscopy. A schematic presentation of the function of a pinhole. Light from a point source illuminates a single point of the object and that, in turn, is imaged on a small pinhole (A), making this point confocal. A pinhole conjugate to the focal point passes all light from the focal point and very little from the out-of-focus points, as show in the images (B) and (C) were an illumination from respectively below and above the focal plane are shown. Light from illumination of the left or of the right of the focal volume is represented in (D) and (E). (PhD Thesis, Dr. Gerrit Heuvelman)
Since a cell's endogenous molecules usually do not fluoresce themselves, the fluorescent marker has to be introduced. We use a variety of methods to effectively label different compartments and molecules of cells. Firstly, we use fluorescent dyes that are directly taken up by the cell. For example, we use DAPI (4',6-diamidino-2-phenylindole) that strongly binds to A-T rich regions of DNA and therefore can be used to visualize the DNA content of cells (Figure 3, upper left panel). Secondly, we make use of immunofluorescence, a technique used to very specifically stain one target molecule, usually a protein. Fixed cells are incubated with a primary antibody raised against the protein of interest. This antibody is either labeled with a fluorophore directly or a secondary, labeled antibody is applied to amplify the fluorescent signal (Figure 3, upper right panel). In a similar manner we can detect specific sequences of DNA or RNA, with probes that hybridize to the sequence of interest with a method called FISH (fluorescence in situ hybridization). Similar to immunofluorescence, the probe is either labeled with a fluorophore directly or a secondary step is needed to stain the probe and detect the sequence of interest (Figure 3, bottom panel). Thirdly, we transfect cells with exogenous constructs expressing the protein of interest tagged with a fluorescent marker, e.g. GFP (green fluorescent protein). This method is heavily used to determine the location of the fusion protein in living cells over time, to determine its mobility and its interaction with other cellular components in the cell.
Figure 3: Upper left: DAPI bound to DNA (Cheatham et al., 2004). Upper right: Direct and indirect labeling of a protein of interest with a primary antibody that is labeled with a fluorophore or an additional secondary antibody that carries the fluorophore (http://www.mgormerod.com/_wp_generated/wp41bc17d4.png). Bottom: Basic principle of FISH: a labeled probe that is complementary to the sequence of interest hybridizes with the sequence of interest. (http://www.mydbio.com/eWebEditor/uploadfile/20111222133307707.jpg).
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