One of the big aims in microscopy is to achieve optical sectioning in order to bring features of interest into focus or, in other words, to achive confocality.
To reach this goal, many different methods have been developed in the past decades. Some of them adapt specific hardware, others achive confocality post-aquisition through highly specialized software. And some again combine both ways.
Here we strive to give a comprehensive overview of what techniques there are available in the field of light microscopy.
In a Confocal Laser Scanning Microscope (CLSM) a focused laser beam is scanned over your sample, such that the image is aquired point by point. Before each point is collected and amplified by the detector (commonly a Photo Multiplier Tube - PMT), the emission light has to go through a spatial, adjustable pinhole which effectively cuts off the out of focus light.
There is NO CAMERA on your confocal!
Or: the virtues of a cameraless microscope system. What is a PhotoMultiplier Tube (PMT) and how does it work: have a look.
Very cool Java tutorial on how a confocal microscope system works and which adjustable parameters lead to what results, including a comprehensive explanation.
Here you find an introduction to the principles and the history of confocal microscopy.
Read, how to optimally prepare your sample so as to get most out of your confocal microscope.
In this 26min video Dr. Thorn discusses the basic principles of confocal microscopy, with specific discussions of the operation of laser scanning and spinning disk confocal microscopes and of their application to biology.
Multiphoton microscopy is another scanning point illumination technique, where a pulsed IR laser is used to achive a much higher penetration depth than with "conventional" lasers and is therefore very well suited for very thick and/or optically dense samples. Due to the 2-photon effect, your fluorophore is only excited at the focal plane, thus reducing photo toxicity and light scattering.
The lower z-resolution due to the higher excitation wavelength is one of the drawbacks, also multicolour imaging proves more challenging.
On this webpage you can find a detailed introduction to multiphoton microscopy, including some interactive java tutorials.
Here you find an explanation about the fundamentals and applications in multiphoton excitation microscopy
This half an hour talk by Kurt Thorn introduces two-photon microscopy which uses intense pulsed lasers to image deep into biological samples.
A fast rotating, synchronized pinhole disc in combination with virtual "field" illumination and camera detection enables confocal imaging at a much higher temporal resolution than conventional Laser Scanning Confocal. Using Spinning Discs, even confocal live cell imaging of moving objects is possible.
Drawbacks are a lower z-resolution and compromises on the confocality for lower magnification/NA objecitves.
Here you can find a detailed introduction to spinning disc microscopy, including the history and a comparison to conventional confocal laser scanning microscopy.
Here, Zeiss Campus explaines the fundamentals of a spinning disc by using a nice interactive tutorial.
This 25min talk by Kurt Thorn introduces confocal microscopy, and discusses optical sectioning, reconstruction of 3D images, and how the laser-scanning confocal microscope and spinning disk confocal microscope work.
By using high angle incident light, an evanescent wave is created in Total Internal Reflection Fluorescence (TIRF) microscopy, which only excites fluorophores in a thin (~200nm) zone very near a solid surface (eg. a cell membrane sitting on a coverslip).
The resulting image displays very low background noise and virtually no out-of-focus fluorescence.
Here you can find a detailed introduction to the theoretical aspects of TIRF microscopy, including a interactive Java tutorial in which you can explore the TIRFM evanescent wave.
This webpage shows another, nicely illustrated, detailed introduction to TIRF microscopy.
In this 43min. video, the pioneer of TIRF Microscopy (Prof.em. Daniel Axelrod) describes what this technique is used for, explains the principles of the evanescent wave, gives many examples of different microscope configurations used in TIRF, and shows how polarized light TIRF can be used to image membrane orientation.
Light sheet microscopy, better known as single plane illumination microscopy (SPIM) is an emerging technique that combines optical sectioning with multiple-view imaging. The optical sectioning is achieved by using a thin light sheet to illuminate the sample perpendicular to the optical axis, the entire field of view being acquired by a camera.
There are various approaches to mount your (big) sample, many of which aim to assure a free rotation of the object. This way, multiple views of the same sample can be imaged and combined through a processing algorithm (eg Fiji Multiview Reconstruction) to create a 3D representation.
Due to the mainly non-invasive mounting techniques live-organism imaging is not only possible, but one of the main targets.
In this half an hour talk Dr. Ernst Stelzer discusses the new technique of light sheet microscopy, also known as selective plane illumination (SPIM):
Here you can find a reference library on top publications about light sheet microscopy.
OpenSPIM is an Open Access platform for building, adapting and enhancing SPIM technology. It is designed to be as accessible as possible, which includes:
Structured illumination combines hardware and software techniques to achieve optical sectioning. A hardware grid pattern (ie. a pattern of dark and light stripes) is focused onto the sample and shifted several times, one image being aquired by camera after each shift. A subtractive software algorithm is then used to reconstruct the in-focus-image.
The Zeiss ApoTome is one example of the implementation of structured illumination to achieve optical sectioning of thick, fluorescently labeled specimen.
Here, Zeiss explains its structured illumination system with the help of an interactive Java tutorial.
Deconvolution is a software based method to achieve optical sectioning. It corrects the systematic error caused by the objective lens in that smaller and smaller object features come through the lens with less and less contrast than they have in reality. This is true for all widefield, confocal, 2 photon, SPIM/LSFM etc modes of fluorescence light microscopy.
Once corrected by deconvolution, the resulting image is a more quantitative estimate of the true distribution of fluorophores in the object, with increased contrast and dynamic range especially for smaller features. Also, photon and detector noises, as well as background and detector offset are suppressed and the image appears "deblurred" (ie. sharper) to the eye.
In this lecture, Prof. David Agard describes the basic principles of various deconvolution techniques and introduces principles important to deconvolution such as the Fourier transform, points spread function and optical transfer function.
Here, you can find a nice theoretical introduction into how deconvolution in optical microscopy works.
This is a very nice, basic (not too mathematical) explanation of deconvolution.
SVI is the developer and distributer of a professional deconvolution software: Huygens. On their webpage they give nice tutorial videos on how deconvolution (in general and with Huygens in specific) works, including some nice examples.
For deconvolution a proper spherical aberration correction is crucial. Here, you can calculate the correct refractive Index for your immersion oil.