Microscopy Techniques

The development of fluorescent labels revolutionized biological imaging by giving researchers a highly selective method for labeling and imaging biological structures. 

A fluorescent label or dye, generally termed a fluorochrome, is a molecule that absorbs a photon of light of one wavelength (e.g. blue) before emitting a photon of light (fluorescence) of a longer wavelength (e.g. green).  The wavelengths of light absorbed and emitted are specific for each fluorochrome.  Utilizing fluorochromes with different emission profiles allows multiple structures to be labelled in the same sample, which can then be used to determine if fluorochromes are located within the same area (i.e. colocalization).  A large number of different fluorochromes exist which can be used to selectively label just about any cell or tissue structure, thus providing an easy way for researchers to study many different biological processes.

AFM is an imaging technique that can provide subnanometer resolution, more than 1000 times better than the optical diffraction limit.  To achieve this high resolving power, instead of utilizing a stream of photons to image specimens, AFM utilizes a specialized mechanical AFM probe (termed cantilever) with a sharp tip to “feel” the surface of the sample being analyzed.  The cantilever tip is only a few nanometers in diameter and deflects when brought into close proximity to the surface of a sample.   This deflection is then measured and translated into structural information.  Scanning of the cantilever across the surface of a sample can then be used to assemble a picture.

Confocal microscopy is one of the most popular techniques for imaging fluorochromes in biological specimens. 

The popularity of confocal microscopy arises from its ability to collect high-resolution in-focused images by removing out of focus light during image collection.  To accomplish this, confocal specimens are illuminated by a focused point of laser light that is scanned across the specimen in the horizontal plane.  Emitted fluorescence from the sample is collected by the microscope objective and passed through a pinhole aperture designed to remove out of focus light originating from above or below the focal plane.  An image is serially built up as the focal point is scanned across the sample and projected onto a photodetection device (e.g. photomultiplier tube).

Multiphoton fluorescence microscopy is another laser based technique for visualizing fluorescently labelled specimens.

In contrast to confocal microscopy, where a fluorochrome is excited using short wavelength light to stimulate the emission of longer wavelength light (i.e. fluorescence), multiphoton microscopy utilizes pulsed long-wavelength light to stimulate the fluorochrome to emit its fluorescence signal.  As a fluorochrome needs to simultaneously absorb the energy from two long-wavelength photons to excite its fluorescence emission, fluorescence is effectively restricted to the focal point of the objective, thus inhibiting out of focus fluorescence without the need of a pinhole aperture.  The focal point is then scanned across the specimen and a fluorescent image is built up using a photodetection device (e.g. photomultiplier tube).

As multiphoton microscopes utilizes longer wavelength excitation light that is relativity low energy, they are well suited for imaging live cells as they cause less phototoxicity than confocal microscopes using shorter-wavelength lasers.  In addition, multiphoton microscopy gives the best sample penetration and is especially good for imaging thick specimens (up to ~ 1 mm).

TIRF microscopy is a technique for visualizing fluorochromes that are immediately adjacent (<200 nm) to a glass coverslip, and is mainly used to image molecular events in cellular surfaces such as cell adhesion, binding of cells by hormones, secretion of neurotransmitters, and membrane dynamics.

To restrict emission fluorescence within such a narrow optical plane, TIRF microscopes direct the excitation laser at an oblique angle at the glass-water interface between specimen and coverslip.  The different refractive indexes of glass and water cause the incident light to be “totally internally reflected” which generates a highly restricted electromagnetic field (or evanescent wave) within the specimen.  This evanescent wave is identical in frequency to the excitation laser (thus exciting fluorochrome fluorescence normally) but decays exponentially in intensity with distance from the glass coverslip, only extending up to ~200 nm into the specimen.

The LSFM technique utilises two microscope objectives arranged perpendicularly.  The first objective focuses a laser beam into a flat light sheet that passes through, and is restricted to, the focal plane of the second objective.  As the light sheet restricts fluorochrome excitation to within the focal plane of the observation objective background signal and thus image contrast is significantly improved, making LSFM images comparable to those collected using confocal microscopy.

Restricting excitation illumination to a narrow light sheet also greatly reduces photobleaching and phototoxic damage when imaging living samples, allowing the collection of far more scans per specimen.  This method is used in cell biology and for microscopy of whole living creatures, such as embryos, or large whole-mount stained tissues/organs.

Live cell imaging is a term given to the observation and recording of living cells grown in culture.

Live cell imaging can be used in conjunction with a wide variety of different microscopy techniques (eg. bright-field, confocal, TIRF), but requires the use of a microscope with specialized incubation chamber for maintaining cultured cells at physiological temperatures and atmospheric conditions.  Cultured cells are placed in the incubation chamber and imaged using time-lapse microscopy for as long as required (minutes-days).

Live cell imaging is a useful technique for imaging a wide variety of different dynamic cell processes including cell division, cell migration, actin/microtubule activity, cytoplasmic trafficking, etc.

Intravital microscopy is a technique used to observe biological processes in living animals (in vivo).  Intravital microscopy can be coupled with a wide variety of different imaging techniques including: wide-field, epifluorescence, confocal, multiphoton, etc.  But instead of imaging discrete biological samples, whole anaesthetised animals are placed on the microscope stage.  Surgical techniques are used to isolate and position the organ or tissue of interest in front of the microscope objective.  Intravital microscopy in a useful technique for imaging biological processes that cannot be easily modelled outside of an organism.

FRAP is an optical technique used to quantify the lateral transport/diffusion of fluorescently labelled molecules.

To accomplish this, the fluorochromes within a subsection of the sample are photobleached using a short pulse of high intensity illumination.  The recovery of fluoresce within the dark photobleached area is subsequently imaged over time as fluorochromes move back into the photobleached area, either via simple Brownian motion or active transport.  This technique is very useful in biological studies of cell membrane diffusion, protein binding, and protein trafficking. 

FRET is an optical technique used to detect molecular interactions and requires specific donor and acceptor fluorochrome pairs.

Excitation of the donor fluorochrome will, instead of emitting fluorescence signal itself, stimulate emission of fluorescence from the nearby acceptor fluorochrome.  The efficiency of this energy transfer between donor and acceptor fluorochromes is inversely proportional to the sixth power of the distance between them.  Thus, for FRET to work, both donor and acceptor fluorochromes need to be in close proximity.

Measurements of FRET efficiency can be used to determine if two fluorochromes are within a certain distance of each other making it a useful tool for quantifying protein-protein interactions, protein–DNA interactions, protein conformational changes, and to detect the location and interactions of genes and cellular structures including intergrins and membrane proteins.

Each fluorescent dye displays a characteristic fluorescence lifetime, namely the length of time following excitation that the dye will remain in its excited state emitting fluorescence.  This fluorescence lifetime is a quantitative signature that can be used to distinguish between different fluorescent dyes, even those emitting the same fluorescent colour.  Thus the lifetime of the fluorescent signal, rather than its intensity, is used to create the image in FLIM, which can be used in conjunction with other microscopy techniques such as confocal or multiphoton microscopy.

A range of biophysical phenomena affect fluorescence lifetime including: ion intensity, hydrophobic properties, oxygen concentration, molecular binding, and molecular interactions.  Therefore the applications of FLIM are many: from ion imaging and oxygen imaging to studying cell function and cell disease.

Fluorescence correlating spectroscopy allows for determining diffusion coefficients and concentrations of fluorescently labelled molecules at nanomolar concentrations both in vitro and in live cells. It is mostly useful for indirect studies of molecular activity in plasma membrane, in cytosol and in nucleus via following relative changes in diffusive behaviour and/or concentration. The method is based on single molecule sensitive confocal detection combined with correlation analysis of the highly fluctuating intensity time traces.

Fluorescence cross-correlation spectroscopy is designed for quantification of strong interactions/binding between two mobile entities both in vivo and vitro. The presence of doubly labelled entities is manifested via synchronised intensity fluctuations for both fluorescent markers – a positive cross-correlation amplitude.

STED is a super resolution fluorescent microscope technique designed to enhance image resolution by bypassing the diffraction limit faced by traditional microscopy.

To accomplish this STED microscopes typically utilize two lasers, a standard fluorochrome excitation laser, and a high-energy depletion laser.  The depletion laser is used to selectively deactivate fluorochromes in a donut shaped area leaving the centre focal spot active to emit fluorescence following excitation.  This centre focal spot can be adjusted to provide resolutions typically between 50 and 80 nm thus allowing biologists to visualize structures and dynamic processes in cells at, or near, the molecular level.

PALM is a probe-based super resolution technique utilizing widefield fluorescence microscopy and photoactivatable fluorescence proteins to bypass the diffraction limit faced by traditional microscopy.  To accomplish this, PALM microscopes require specimens to be labelled with photoactivatable fluorescent proteins that can have their fluorescence switched on using a photoactivation laser.  When using PALM to image a specimen, only a small fraction of these photoactivatable proteins (< 1%) are activated to fluoresce at any one time.  The activation of only a small subset of fluorescent probes allows them to be located within the image with nanometer-level precision.  Once imaged, the photoactivated proteins are then photobleached and a new subset of photoactivatable proteins are activated and imaged.  The process is repeated thousands of times to generate a composite image with single-molecule resolution.

STORM is a super resolution imaging technique similar to PALM, but instead of utilizing photoactivatable proteins and photobleaching, STORM utilizes organic fluorophores that can be driven between active-ON and an inactive-OFF states.  Specimens are immunolabelled with STORM compactable probes and a photoactivation laser is used to activate a small subset (< 1%) to fluoresce.  The activation of only a small subset of fluorescent probes allows them to be located within the image with nanometer-level precision.  Once imaged, the probes are then deactivated and a new subset of probes are activated and imaged.  The process is repeated thousands of times to generate a composite image with single-molecule resolution.

SIM is another super resolution imaging technique, and displays approximately twice the resolving power of traditional diffraction-limited microscopy techniques.  SIM utilizes a widefield technique in which a grid pattern is superimposed on the specimen during image capture; sets of images are captured while the grid pattern is rotated in a series of steps.  Interference between the illumination pattern and sample image is then used to extract image information that would otherwise be unobservable. SIM image sets are processed using specialized computer algorithms to extract the super resolution image.

Fluorescence Polarization is a technique used to assay the kinetics of fluorescent probes and their targets.

When a stationary fluorochrome is excited to fluoresce using plane-polarized light, its emitted fluorescence is also polarized.  However, if the fluorochrome rotates out of alignment between being excited and fluorescing (e.g. via randomized Brownian motion) the polarization of this emitted fluorescence is lost.  Thus factors that influence fluorochrome movement can easily be assessed by assaying fluorescence polarization (e.g. small unbound fluorochromes are more likely to tumble out of alignment causing them to emit less polarized fluorescence than fluorochromes bound to large slowly moving molecules).  This makes Fluorescence Polarization a useful technique for studying a wide variety of biological processes including: Receptor/ligand binding, Protein/peptide interactions, DNA/protein interactions, Tyrosine Kinase Assays, and Competitive Immunoassays.

SHG is an optical scattering effect caused by some molecules with noncentrosymmetric arrangements.  These molecules, when illuminated, can combine two photons into a single photon with half the wavelength but twice the frequency of the original photons.  Microscopes setup for SHG imaging generally contain a short-pulse excitation laser and appropriate filter sets for separating SHG emitted light from the excitation light.  Molecules that can be easily visualized using SHG include collagen, microtubules, and muscle myosin.

When excited, fluorochromes tend to emit a wide spectral band of fluorescence.  Fluorescent signal bleed-through occurs when different fluorescent emission bands overlap in the same sample.  Spectral Imaging is an imaging technique that was developed so that overlapping fluorescent emission spectra could be individually resolved in specimens containing two or more fluorescent labels.

To accomplish this, specimens are scanned using a microscope with a specialized spectral detector unit to generate a three-dimensional dataset of the sample acquired at different emission wavelengths (termed a lambda stack).  Software-based linear unmixing algorithms can then be used to unmix the lambda stack into its constituent fluorescent signals (eg GFP, YFP, DsRed, etc).