Advanced Light Microscopy

🖥 Image Analysis Workstation Pro

Laser scanning confocal microscopy (LSCM/CLSM)

Confocal laser scanning microscopy (CLSM) or laser scanning confocal [LSCM]), often colloquially referred to simply as “confocal”, is a technique for obtaining high resolution optical images with depth selectivity. The key feature of confocal microscopy is its ability to acquire in-focus images from selected depths, a process known as optical sectioning. Images are acquired point-by-point and reconstructed with a computer. This allows three-dimensional reconstructions of topologically complex objects. However, CLSM is significantly slower than widefield or spinning-disk confocal microscopy, because images are acquired pixel-by-pixel and the laser power is concentrated on small diffraction limited spots which increases the phototoxicity load on the samples. CLSM typically requires high-laser power excitation, which can lead to phototoxicity and limited observation of live cells.

Spinning disk confocal microscopy (SDCM)

Spinning disk confocal microscopy is one of the solutions for routine and high-performance fluorescence live-cell imaging applications. SDCM uses a series of moving pinholes on a disc to scan spots of light, in combination with a high-sensitivity camera to acquire instantaneous optical slices. Since a series of pinholes scans an area in parallel, each pinhole is allowed to hover over a specific area for a longer amount of time than on laser scanning confocals (CLSM) thereby reducing the excitation energy needed to illuminate a sample . Furthermore, cameras (often EMCCDs or sCMOS), typically have quantum efficiencies 2-3x higher than PMTs, so much less laser excitation energy is necessary and that reduces photo-toxicity and photo-bleaching of a sample, making it the preferred system for imaging live cells or organisms when optical slicing is necessary. However, optical sectioning in depth is more limited than in CLSM. This technology can be improved with image scanning mic/pixel reassignment to allow non-diffraction limited images which provide resolutions 100-200nm.

Total internal reflection fluorescence microscopy (TIRF)

Total internal reflection fluorescence microscopy (TIRF) is a microscopy technique with which a thin region of the cell, usually less than 200nm can be observed. A TIRF microscope uses an evanescent wave to selectively illuminate and excite fluorophores in a restricted region of the specimen immediately adjacent to the glass-water interface. The evanescent wave is generated only when the incident light is totally internally reflected at the glass-water interface. The evanescent electromagnetic field decays exponentially from the interface, and thus penetrates to a depth of only approximately 100 nm into the sample medium. Thus the TIRF microscope enables a selective visualization of surface regions such as the basal plasma membrane (which are about 7.5 nm thick) of cells. This technique is often used also for observing molecular dynamics in vitro, or study the details of cell locomotion or adhesion to substrata.

Two-photon microscopy (2P)

Multiphoton microscopy - most commonly in the form of two-photon microscopy - is a fluorescence imaging technique that allows observation of living tissue up to about one millimeter in depth. It uses pulsed red-shifted excitation laser light, which can also excite visible fluorescent dyes. However, for each excitation, two (or three) photons of low energy infrared light are absorbed “simultaneously” to provide the required energy for electrons in the fluorophore to reach the excited state. Using infrared light minimizes scattering in the tissue and allows imaging deeper than is possible with common confocal microscopes. Due to the localized multiphoton absorption effect, the background signal is strongly suppressed and a pinhole is not required in front of the detector (PMTs). Both effects lead to an increased penetration depth for these microscopes. Two-photon microscopy is often used to image intravitally, and it may facilitate imaging of unlabelled tissues, using autofluorescence or SHG (second harmonic generation).

Nanoscopy

Single Molecule Localisation microscopy (SMLM)

There are a variety of widely used implementations of single molecule localization microscopy. The unifying principle behind them is that a large series of individual images is acquired, in each of which only a small subset of the fluorescent labels in the sample are fluorescing. This sparse nature of the signal allows the computational calculation of the exact position of each of the emitting fluorophores. By combining the large number of images with precise locations, an image with very high resolution can be formed. Below are descriptions of some of the common SMLM techniques

The fundamental principle behind stochastic optical reconstruction microscopy (STORM) is that the activated state of a photo-switchable molecule must lead to the consecutive emission of sufficient photons to enable precise localization before it enters a dark state or becomes deactivated by photobleaching. Additionally, the sparsely activated fluorescent molecules must be separated by a distance that exceeds the Abbe diffraction limit (in effect, greater than approximately 250 nanometers) to enable the parallel recording of many individual emitters, each having a distinct set of coordinates in the lateral image plane.

Photo activated localization microscopy (PALM) is a widefield fluorescence microscopy imaging methods that allow obtaining images with a resolution beyond the diffraction limit. PALM is based on collecting a large number of images each containing just a few active isolated fluorophores. The imaging sequence allows for the many emission cycles necessary to stochastically activate each fluorophore from a non-emissive state to a bright state, and back to a bleached state. During each cycle, the density of activated molecules is kept low enough that the molecular images of individual fluorophores do not typically overlap.

Single Particle Tracking (SPT)

Single-Particle Tracking involves tracking the motion of individual particles over time to obtain information on their dynamic behavior. These particles can range from molecules and nanoparticles to cellular components like proteins and vesicles. The tracking process typically begins by labeling the particle of interest with a fluorescent marker making it detectable under a microscope.

Mesoscopic imaging

Light-sheet mesoscopic imaging (SPIM)

Light sheet microscopy is a mesoscopic imaging technology that combines optical sectioning with multiple-view imaging to observe tissues and living organisms with impressive resolution. This method is often also referred to as single plane illumination microscopy (SPIM) and many different implementations are available.

Three-dimensional imaging in light-sheet-based microscopy is performed by moving the specimen through the light sheet in small steps and recording a two-dimensional image at each step. Alternatively the light sheet can be moved through the specimen. In multiple-view imaging, the same volume inside the specimen or even the entire specimen is recorded along several angles. The resulting multiple-view information can be combined into a single image stack by data post-processing using a fusion algorithm.

🔬 Light Sheet Microscope

Functional imaging and specialised methodologies

Fluorescence Resonance Energy Transfer (FRET)

Fluorescence resonance energy transfer (FRET) is a mechanism describing energy transfer between two light-sensitive molecules (chromophores). A donor chromophore, initially in its electronic excited state, may transfer energy to an acceptor chromophore through non-radiative dipole-dipole coupling. The efficiency of this energy transfer is inversely proportional to the sixth power of the distance between donor and acceptor, making FRET extremely sensitive to small changes in distance and therefore an excellent reporter on molecule proximity and interaction.

Fluorescence Lifetime Imaging (FLIM)

Fluorescence-lifetime imaging microscopy (FLIM) is an imaging technique for producing an image based on differences in the fluorescence-lifetime rather than its intensity. By quantifying variations in the exponential decay rate of the fluorescence from a fluorescent sample (fluorescence-lifetime) it is possible to report on molecule proximity, pH changes and even polarity. It can be used as an imaging technique in confocal microscopy, two-photon excitation microscopy and multiphoton tomography. Since the fluorescence-lifetime is insensitive to changes in fluorophore intensity or concentration, it is the most quantitatively precise technique to report on fluoresce resonance energy transfer (FRET).

Fluorescence (cross)-correlation spectroscopy (FCS/FCCS)

Fluorescence correlation spectroscopy (FCS) is a correlation analysis of fluctuations in the fluorescence intensity. The analysis provides information on physical parameters of the fluorescent particles (molecules) in solution, such as concentration, average fluorescence intensity and diffusion speed. By following changes on these parameters it is possible to study binding events of the molecules or even conformational changes on them.

Fluorescence cross-correlation spectroscopy (FCCS) extends the FCS procedure in that it looks at the correlation between different colors (cross-correlation) rather than just the same color (auto-correlation). In other words, coincident green and red intensity fluctuations correlate if green and red labeled particles are moving together. As a result, FCCS provides a highly sensitive measurement of molecular interactions independent of diffusion rate.

Single Molecule FRET

Single molecule FRET allows the measurement of distances and distance changes between two fluorescent molecules in a biomolecular complexes, down to the Angstrom and the microsecond timescales. smFRET measurement can be performed on diffusing molecules, in a confocal geometry, to measure distances and fast conformational changes or on immobilized molecules to measure association dissociation kinetics and conformational changes in the seconds timescales.

Microdissection

Microdissection is an established method for a large number of applications, mainly in molecular biology, particularly nucleic acid research, neurosciences, developmental biology, cancer research, forensics, proteomics, plant research, for cutting cell cultures and for single cell isolation.

Microdissection offers a precise and contamination-free solution for the isolation and selection of single cells or tissue. The tissue samples can be embedded, sectioned and stained according to conventional methods of preparation. Paraffin sections, frozen sections, smear preparations, chromosome specimens and cell cultures are all suitable for laser microdissection. The area selected for dissection is drawn on the PC screen and automatically separated from the surrounding tissue with a laser beam.

Photomanipulation

Photomanipulation includes a variety of techniques in which lasers (or other light sources) are used to interact with the sample in a targeted fashion. This includes laser ablation to induce cellular injuries and measure physical properties, photoconversion or photobleaching of fluorescent proteins for tracking of protein movements, uncaging of probes or different compounds in the cell etc. Photomanipulation is key in using optogenetics approaches.

Expansion Microscopy

Expansion microscopy (ExM) is a super-resolution approach that achieves sub-diffraction information by physically expanding the specimen. This allows it to seamlessly obtain resolutions of ~50 nm (up to 10 nm) in four colors in any widefield or confocal microscope.

🔬 Nikon Eclipse TE-2000

🔬 Zeiss Axio Observer 7 + Apotome 3