Promoting Microscopy Education
Welcome to The North Atlantic Microscopy Society (NAMS)
We are very proud and excited to announce the formation of a new society, The North Atlantic Microscopy Society (NAMS). Geographically centered at Princeton, New Jersey, our organization envisions our coverage to span Southern New York, New Jersey, and Eastern Pennsylvania.
NAMS is born not simply because we noticed a distinct gap in regional cohesion. Above all else, it is because we are passionate about microscopy in all its forms and believe that we are not alone.
Edwin Hubble famously said, “Equipped with his five senses, man explores the universe around him and calls the adventure Science.”
At NAMS, we believe that some of this exploratory instinct has been muted lately by our disciplinary silos. Individually, we have become exceptional in our specialties and do not take moments to appreciate the many discoveries happening across the entire spectrum of science.
Our mission is to bridge these silos through the lens of microscopy. We seek to achieve this mission by promoting microscopy education, stimulating networking among microscopists, and disseminating microscopy knowledge and skills to the public in the region.
Transmission Electron Microscopy (TEM)
The original form of electron microscopy, TEM involves a high-voltage electron beam emitted by a cathode and formed by magnetic lenses. The electron beam that has been partially transmitted through the very thin (and so semitransparent for electrons) specimen carries information about the structure of the specimen.
The spatial variation in this information (the "image") is then magnified by a series of magnetic lenses until it is recorded by hitting a fluorescent screen, photographic plate, or light sensitive sensor such as a CCD (charge-coupled device) camera. The image detected by the CCD may be displayed in real time on a monitor or computer. Transmission electron microscopes produce two-dimensional black and white images.
Scanning Electron Microscope
Unlike the TEM where the electrons in the primary beam are transmitted through the sample, the SEM produces images by detecting secondary electrons, which are emitted from the surface due to excitation by the primary electron beam. In the SEM, the electron beam is scanned across the surface of the sample in a raster pattern with detectors building up an image by mapping the detected signals with beam position.
Cryo-EM begins with vitrification, in which the protein solution is cooled so rapidly that water molecules do not have time to crystallize, forming an amorphous solid that does little or no damage to the sample structure. The sample is then screened for particle concentration, distribution, and orientation. Next, a series of images is acquired, and two-dimensional classes are computationally extracted.
In the final step, the data is processed by reconstruction software, yielding accurate, detailed, 3D models of intricate biological structures at the sub-cellular and molecular scales. These models can reveal interactions that were impossible to visualize previously, a key to scientific results.
Light microscopy provides researchers with the ability to visualize a variety of samples, from monolayers and small organisms, such as developing fly and fish embryos, to very thick sections from the brain and other organ tissues.
The light microscope, so called because it employs visible light to detect small objects, is probably the most well-known and well-used research tool in biology. Yet, many researchers are unaware of the full range of features that are available in light microscopes.
There are many types of light microscopes, suitable for different applications, including, but not limited to:
Widefield microscopy refers to techniques where the whole specimen is exposed to light. This differs from confocal microscopy where you are using a laser to illuminate only a very small part of the sample.
Bright field microscopy:
In bright field microscopy you illuminate the sample from the bottom (or top) with white light, and observe the sample from the top (or bottom). The main advantages of this technique are the simplicity of the microscope system and the simplicity of sample preparation. The main drawback is lack of contrast.
Phase contrast microscopy:
Samples that are relatively transparent by bright field microscopy can often be imaged by phase contrast microscopy. Differences in the refractive index across the sample result in different light phase shifts. Phase contrast microscopy translates these phase shifts into contrast in the final image.
Dark field microscopy:
In dark field microscopy, greater contrast is created in the image by collecting only the light that is scattered by the sample. Because the background does not scatter much light compared to the sample, dark field images have a characteristically dark (almost black) background while the samples themselves appear bright (or vice versa).
Differential interference contrast (DIC):
This technique uses polarized light which the microscopy splits into two orthogonal beams and then sends through the sample. When the two orthogonal beams are recombined, phase shifts which occurred in each individual beam as it traveled through the sample lead to interference, and this interference is translated into contrast in the final image.
Confocal microscopy is an optical imaging technique for increasing optical resolution and contrast of an image by means of using a spatial pinhole to block out-of-focus light in image formation
In confocal imaging a focused laser beam is used to produce a small spot illumination on the specimen. This is in contrast to conventional imaging processes, in which the specimen is bathed in excitatory illumination. This modification removes out of focus contribution from the final image produced by confocal imaging.
In addition, the shallow depth of field in confocal imaging results in a higher resolution image, which is due to the reduced point spread of the function of the system. This is because the function is the product of the illuminating and objective lenses of the system. The resulting resolution is improved by a theoretical factor of 1.4 in comparison with conventional widefield methods.
Light sheet fluorescence microscopy (LSFM) is a fluorescence microscopy technique with good optical sectioning capabilities and high speed. LSFM aso is much less phototoxic than other existing optical sectioning modalities. In contrast to epifluorescence/confocal microscopy only a thin slice of the sample is illuminated perpendicularly to the direction of observation. For illumination, a laser light-sheet is used, i.e. a flattened laser beam which is focused only in one direction or a circular beam scanned in one direction. The good optical sectioning capability reduces the background signal and thus creates images with higher contrast, comparable to confocal microscopy.
There are two basic types of light sheet systems. Some are made for rotating larger samples, millimeters in size, and others image from one direction only. The larger systems are good for larger samples such as fly, fish, and worm embryos. The smaller systems are optimized for smaller samples, such as monolayers, biofilms, and very early embryos.
Super Resolution microscopy can be generically defined as any modality that can resolve two point sources of light beyond the Abbe defined limit.
In general, there are three ways to accomplish this task, including, but not limited to structured illumination (SIM), localization (STORM, PALM, ....), and point-spread function engineering (STED, GSD, ...),
In SIM, a grid pattern is generated through interference of diffraction orders and superimposed on the specimen while capturing images. The grid pattern is shifted or rotated in steps between the capture of each image set. Following processing with various algorithms, high-frequency information can be extracted from the raw data to produce a reconstructed image having a lateral and axial resolution approximately twice that of diffraction-limited instruments.
The Localization technique relies on stochastic activation of fluorescence to intermittently photoswitch molecules to a bright state, which are then imaged and photobleached. Thus, very closely spaced molecules that reside in the same diffraction-limited volume (and would otherwise be spatially indistinguishable are temporally separated. These techniques can typically obtain 20 nm resolution laterally and 50 nm resolution axially.
Point-spread function engineering or illumination-based superresolution, utilizes non-linear optical approaches to reduce the focal spot size. These techniques can acquire down to 10 nm isotropic resolution!
Mass Spectrometry Imaging (MSI)
MSI utilizes a mass spectrometer to visualize the spatial distribution of a variety of molecules of interest including drugs and other small molecules, metabolites, peptides, proteins, and lipids. The advantage of MSI over traditional methods, such as radiochemistry or immunohistochemistry, is its ability to analyze multiple samples at once and its ability to retrospectively analyze data without prior knowledge of the samples.
Several variations of MSI exist including secondary ion mass spectrometry (SIMS), matrix-assisted laser desorption ionization (MALDI imaging), desorption electrospray ionization (DESI), and laser ablation electrospray ionization (LAESI). The choice of MSI methodology largely depends on the sample being investigated and the nature of the molecules of interest.
Secondary ion mass spectrometry (SIMS) imaging is able to image tissues, single cells, and microbes to reveal chemical species at sub-100 nm spatial resolution. Primary ions from an ion source are accelerated towards the sample surface. The impact of the primary ions ejects molecules (mostly neutral), but a small percentage will carry a charge (secondary ions).
The mass-to-charge (m/z) ratio of the secondary ions are measured in a mass analyzer. Imaging is achieved by either stage movement or beam rastering across the area of interest. 3D reconstruction of 2D serial images are obtained by etching the top (imaged) layer by the primary beam followed by imaging of the lower layer.
Matrix assisted laser desorption ionization (MALDI) imaging mass spectrometry is most commonly utilized on thin sections such as tissues. Tissues are prepared first, after which a matrix is evenly coated across the tissue. The matrix must absorb at the laser wavelength and ionize the analyte. Matrix and solvent selection heavily affects data quality and is reliant upon the analyte
class of interest.
MALDI imaging has the advantage of measuring the distribution of a large amount of analyte at one time without destroying the sample. It allows direct, label-free measurement of proteins, peptides, lipids, drugs, and metabolites from tissue. The distribution of the detected compounds can be seen as images which can be integrated with other imaging modalities to study the molecular signatures of tissue compartments.
The previous methods require samples to be placed under high vacuum. Ambient ionization methods have also been utilized.
These include desorption electrospray ionization (DESI and nano-DESI), laser ablation electrospray ionization (LAESI), matrix-assisted laser desorption electrospray ionization (MALDESI), and liquid extraction surface analysis (LESA). In all cases, ions are created under ambient conditions after which the ions are transferred to a mass analyzer.
Intravital imaging is the application of a variety of microscopy techniques to whole living animals. These techniques include confocal, widefield, light sheet and most commonly, nonlinear multiphoton microscopy. In contrast to fixed tissue or in vitro experiments, where cells and tissues from higher organisms are studied statically, or taken out of their physiological context, intravital imaging investigates biological samples in a manner which maintains their connection to, and interaction with, the rest of the organism.
Performing microscopy in live animals presents a number of unique technical challenges that may at first appear daunting. However, advances in microscope designs, and numerous sample preparation protocols have now dramatically reduced barriers to entry for those new to the technique. These advances include optimizations to microscope optics, detector sensitivity, speed, and automation, as well as tissue fixturing techniques, implantable imaging windows for serial imaging, and novel surgical protocols. These advancements have expanded the range and scope of the tissues that can be accessed, covering nearly all of the organ systems, and allowing imaging at multiple spatial and temporal scales ranging from milliseconds to months, and from subcellular to organism wide.