Designs of Electron Microscope | Laboratories | Microbiology

Electron microscopes are of three basic designs – the transmission electron microscope, scanning electron microscope and the scanning transmission electron microscope. The third one is a hybrid of the two, Specimen as the name implies. Scanning and transmission electron microscopes are similar in that each uses a beam of electrons to produce an image. However, the instruments use quite different mechanisms to form the final image.

The effect of electron microscopy on our understanding of cells can only be described as revolutionary. Yet, like light microscopy, electron microscopy has both strengths and weaknesses.

In light microscopy, most specimens can be prepared easily and examined readily, but resolution of cellular ultrastructure is severely limited by the physical characteristics of light. In electron microscopy, resolution is much better, but specimen preparation and instrument operation are often more difficult.

Design # 1. Transmission Electron Microscopy:

Most parts of the transmission electron microscope (TEM) are similar in name and function to their counterparts in the light microscope, although their physical orientation is reversed. We will look briefly at each of the major features.

Vacuum System:

Because electrons cannot travel very far in air, a strong vacuum must be maintained along the entire path of the electron beam. Two types of vacuum pumps work together to create the vacuum in the column of an electron microscope. A standard rotary pump is used to achieve the initial low vacuum when the instrument is first started up. The high vacuum required for operation is achieved by an oil diffusion pump.

The diffusion pump is an oil-filled reservoir in which oil is vapourized by heating. As the oil vapour rises, it traps air molecules and is then condensed by condensing vanes, which are cooled by circulating cold water. The diffusion pump cannot function independently; it requires backup by the rotary pump to remove the trapped air molecules from the system.

On some TEMs, a device called a cold-finger is incorporated into the vacuum system to help establish a high vacuum. The cold-finger is a metal insert in the column of the microscope that is cooled by liquid nitrogen. The cold-finger attracts gases and random contaminating molecules, which then solidify on the cold metal surface. When functioning at their best, most modern electron microscopes maintain a vacuum of about 1 x 10^-4 mm Hg or less.

Electron Gun:

The electron beam in a TEM is generated by an electron gun, an assembly of several components. The cathode, a tungsten filament similar to a light bulb filament, emits electrons from its surface when it is heated. The cathode tip is near a circular opening in a metal housing called the Wehnelt cylinder.

A negative voltage on the cylinder helps control electron emission and shape the beam. At the other end of the cylinder is the anode. The anode is kept at 0 V, while the cathode is maintained at 50-100 kV. This difference in voltage is called the accelerating voltage because it causes the electrons to accelerate as they pass through the cylinder.

Electromagnetic Lenses and Image Formation:

The formation of an image using electron microscopy depends on both the wavelike and the particle like properties of electrons. Just as a glass lens can bend the rays of light that pass through it, the trajectory of an electron can be controlled using electromagnets. Because electrons are charged, they are subject to magnetic forces when they move. This principle can be used to change the direction of an electron beam.

As the electron beam leaves the upper region of the condenser lens system, it enters a series of lenses made of electromagnets. The lens itself is simply a space influenced by an electromagnetic field. The focal length of each lens can be increased or decreased by varying the current applied to its energizing coils. Thus, when several lenses are arranged together, they can control illumination, focus, and magnification.

The condenser lens is the first lens to affect the electron beam. It functions in the same fashion as its counterpart in the light microscope to focus the beam on the specimen. Most electron microscopes actually use a condenser lens system with two lenses to achieve better focus of the electron beam.

The next component, the objective lens, is the most important part of the electron microscope’s sophisticated lens system. The specimen is positioned on the specimen stage within the objective lens. The objective lens, in concert with the intermediate lens and the projector lens, produces a final image on a viewing screen of zinc sulfide that fluoresces when struck by the electron beam.

How is an image formed from the action of these lenses on an electron beam? Recall that the electron beam generated by the cathode passes through the condenser lens system and impinges on the specimen. As the beam strikes the specimen, some electrons are scattered by the sample, whereas others continue in their paths relatively unimpeded.

This scattering of electrons is the result of properties created in the specimen by the preparation procedure. Specimen preparation, in other words, imparts selective electron density to the specimen; that is, some areas become more opaque to electrons than others. Such electron-dense areas of the specimen will appear dark because few electrons pass through, whereas other areas will appear lighter because they permit the passage of more electrons.

The contrasting light, dark, and intermediate areas of the specimen create the image seen on the screen. The fact that the image is formed by differing extends of electron transmission through the specimen is reflected in the term transmission electron microscope.

Photographic System:

In addition to observing the image on a fluorescent screen, an electron microscopist can also record the image photographically as an electron micrograph, which then becomes a permanent photographic record of the specimen.

Most transmission electron microscopes have a camera chamber mounted directly beneath the viewing screen. The camera is little more than a box that allows photographic plates to be moved manually or automatically to the area immediately beneath the viewing screen. To photograph a specimen, the microscopist simply aligns the image on the screen, focuses the image with the objective lens control, adjusts the illumination to a predetermined intensity with the condenser adjustment, and makes the exposure.

The exposure may be made automatically or by lifting the screen with a lever on the microscope console. Once the exposure is made, the plate is advanced out of the exposure position to a container where it is stored until retrieved from the instrument for later development and printing.

Sample Preparation Techniques in Transmission Electron Microscopy:

Specimens for electron microscopy can be prepared in several different ways, depending on the type of microscope and the kind of information the microscopist wants to obtain. In each case, however, the method is complicated, time-consuming, and costly compared to methods used for light microscopy. Moreover, living specimens cannot be examined because of the vacuum to which specimens are exposed in the electron microscope.

i. Specimen Processing: Fixation:

Specimens to be prepared for electron microscopy must first be chemically fixed and stabilized. This primary fixation kills the cells but keep the cellular components much as they were in the living cell. Primary fixatives are usually buffered solutions of aldehydes.

Glutaraldehyde is the most common fixation. Following primary fixation, the specimen is usually treated with 1-2% solution of buffered osmium tetroxide (OsO₄). The osmium tetroxide stains specific components of the cell, making them more electron-dense.

ii. Embedding, Sectioning, and Post-Staining:

The next step in specimen preparation for transmission electron microscopy is to dehydrate the tissue by passing it through a series of alcohol solutions. The specimen is then placed into a fluid such as acetone or propylene oxide to prepare it for embedding in liquefied plastic epoxy resin.

After the plastic has infiltrated the specimen, it is put into a mold and heated in an oven to harden the plastic. Thereafter, the area around the specimen is trimmed to prepare a face that is appropriate for sectioning.

The specimen is then sliced into the ultrathin sections required for examination with the transmission electron microscope. The instrument used for this purpose is an ultra-microtome. The specimen is mounted firmly on the arm of the ultra-microtome, which then advances the specimen in small increments toward a glass or diamond knife. When the block reaches the knife blade, ultrathin sections (about 60-90 nm thick) are cut from the block face.

The sections float from the blade onto a water surface, where they can be picked up on a circular copper specimen grid. The grid consists of a meshwork of very thin copper strips, which support the specimen while still allowing “windows” between adjacent strips through which the specimen can be observed.

Once in place on the grid, the sections are usually stained with solutions containing lead and uranium. This procedure, called post-staining, enhances the contrast of the specimen because the lead and uranium give still greater electron density to specific part of the cell. After post-staining the specimen is ready for viewing or photography with the TEM. Numerous examples of transmission electron micrographs.

iii. Electron Microscopic Autoradiography:

The auto-radiographic techniques of light microscopy can be used for transmission electron microscopy with only minor changes. For the TEM, the specimen containing the radioactively labeled compounds is examined in ultrathin sections on copper specimen grids instead of in thin sections on glass slides.

iv. Immunoelectron Microscopy:

In electron microscopy just as in fluorescence microscopy, special techniques have been devised to generate contrast, thereby making it possible to identify and localize specific molecules and subcellular structures. Heavy metal atoms such as iron and gold are commonly used to generate contrast because they are very electron-dense and are therefore readily visible as opaque dots on an electron micrograph.

An especially important technique is immunoelectron microscopy, which depends on antibody molecules coupled to colloidal gold particles as a means of tagging and visualizing specific antigens. This approach makes it possible to determine the subcellular distribution of specific proteins or protein- containing structures with great precision.

Caution is necessary, however, because certain fixation techniques (such as osmium fixation) interfere with the fidelity of antibody-antigen binding. In such cases, milder sample preparation techniques must be used.

v. Negative Staining:

In contrast to the considerable effort necessary to prepare ultrathin sections, negative staining is one of the simplest techniques used in transmission electron microscopy. It is the preferred method for examining very small objects, such as viruses or isolated organelles.

For negative staining, the copper specimen grid must first be overlaid with an ultrathin plastic film. The specimen is then suspended in a small drop of liquid, applied to the overlay, and allowed to dry in air. After the specimen has dried on the grid, a drop of stain such as uranyl acetate or phosphotungstic acid is applied to the film surface.

The edges of the grid are then blotted in several places with a piece of filter paper to absorb the excess stain. This draws the stain down and around the specimen and its ultra-structural features. When viewed in the TEM, the specimen is seen in negative contrast because the background is dark and heavily stained, whereas the specimen itself is lightly stained. Negative staining is illustrated by the electron micrograph of an algal cell wall.

vi. Shadowing:

The same cell wall visualized by the technique of shadowing. Shadowing involves the deposition of a thin layer of an electron dense metal such as gold or platinum on a biological specimen. The metal-emitting electrode is positioned at an angle to the specimen, such that surfaces facing the electrode become coated with the metal, whereas those facing away from the electrode do not.

Specimens that can be suspended in water-based solutions are especially suitable for shadowing. The specimen is first spread on a clean mica surface and dried. It is then placed in a vacuum evaporator, a bell jar in which a vacuum is created by a system similar to that of an electron microscope.

Also within the evaporator are two electrodes, one consisting of a carbon rod located directly over the specimen and the other consisting of a metal wire positioned at an angle about 10° – 45° relative to the specimen.

After a vacuum is created in the evaporator, current is applied to the metal electrode, causing the metal to evaporate from the electrode and spray over the surface of the specimen.

Because of the angular positioning of the metal electrode, the metal will accumulate as a thin coating on the sides of any surface irregularities that face the electrode, generating a metal replica of the surface. These same irregularities prevent deposition of the metal on the side facing away from the electrode, thus producing contrast resulting from the shadow effect.

The carbon electrode is then fired, coating the specimen from directly overhead with evaporated carbon to give stability and support to the metal replicas) –  The mica support containing the specimen is then removed from the vacuum evaporator and lowered gently onto a water surface, causing the replica to float away from the mica surface.

The replica is transferred into an acid bath, which dissolves away remaining bits of specimen, leaving a clean metal replica of the specimen. The replica can then be returned to the water surface and retrieved on a standard copper grid. Replicas can be viewed in the TEM in the same way as ultrathin sections.

vii. Freeze Fracturing:

Freeze fracturing is a relatively recent technique that has proved very useful to cell biologists. It is especially valuable for studying the ultrastructure of biological membranes. Freeze fracturing involves the cleavage of a frozen specimen under a vacuum, followed by platinum/carbon shadowing to create a replica of the fractured surface, which is often the interior of a membrane.

It takes place in a modified vacuum evaporator with an internal microtome knife for fracturing the frozen specimen and with provision for precise control of the temperature of the specimen stage and the microtome arm and knife.

Specimens are generally fixed prior to freeze fracturing, although some living tissues can be frozen fast enough to keep them in almost lifelike condition. Because cells contain a lot of water, fixed specimens are usually treated with an antifreeze such as glycerol to provide cryoprotection—that is, to reduce the formation of ice crystals during freezing.

The cryoprotected specimen is mounted on a metal specimen support and immersed rapidly in Freon cooled with liquid nitrogen. This procedure also reduces the formation of ice crystals in the cells. With the frozen specimen positioned on the specimen stage in the vacuum evaporator, a high vacuum is established, the stage temperature is adjusted to around -100°C, and the frozen specimen is fractured with a blow from the microtome knife.

A replica of the fractured specimen is made by shadowing with platinum and carbon as described in the previous section, and the replica is then ready to be viewed in the transmission electron microscope.

Newcomers to freeze-fracturing technique often misunderstand what a freeze-fracture replica represents. One might think that the fracture plane should pass through the specimen in a straight line, as is clearly the case when a fixed and embedded sample is sectioned conventionally with an ultra-microtome.

In actuality, however, the fracture line passes through the hydrophobic interior of membranes whenever possible, because this is the line of least resistance through the frozen specimen. As a result, a freeze-fracture replica is largely a view of the interiors of membranes, showing the inside of one or the other of the two monolayers of the membrane.

Freeze-fractured membranes appear as smooth surfaces studded with intramembranous particles (IMPs) that are either randomly distributed in the membrane or organized into ordered complexes. These are thought to be integral membrane proteins that have remained with one lipid monolayer or the other as the fracture plane passes through the interior of the membrane.

The electron micrograph shows the two faces of a plasma membrane as revealed by freeze fracturing. The P face is the interior face of the inner monolayer; it is called the P face because this monolayer is on the protoplasmic side of the membrane. The E face is the interior of the outer monolayer; it is called the E face because this monolayer is on the exterior side of the membrane.

Notice that the P face has far more intramembranous particles than does the E face. In general, most of the particles in the membrane stay with the inner monolayer when the fracture plane passes down the middle of a membrane.

To have a P face and E face appear side by side, the fracture plane must pass through two neighbouring cells, such that cell has its cytoplasm and the inner monolayer of its plasma membrane removed to reveal the E face, while the other cell has the outer monolayer of its plasma membrane and the associated intercellular space removed to reveal the P face. Accordingly, E faces are always separated from P faces of adjacent cells by a “step” that represents the thickness of the intracellular space.

viii. Freeze Etching:

Freeze etching is related to freeze fracturing, but there is a considerable difference between the two techniques. Freeze etching adds a further step to the conventional freeze-fracture procedure that makes the technique even more informative. Following the fracture of the specimen but prior to shadowing, the microtome arm is placed directly over the specimen for a short time (a few seconds to several minutes).

This maneuver causes a small amount of water to evaporate (sublime) from the surface of the specimen to the cold knife surface. Where the fracture has passed through a membrane, etching will cause small areas of the true cell surface around the periphery of the fracture face to stand out in relief against the background.

By using ultra-rapid freezing techniques and a volatile cryoprotectant such as aqueous methanol, which sublimes very readily to a cold surface, the etching period can be extended and a much deeper layer of ice can be removed, exposing large areas of the specimen surface to view. This modification, called deep etching, provides a fascinating new look at cellular structure.

ix. Stereo Electron Microscopy:

Electron microscopists frequently want to visualize their specimens in three dimensions. Techniques such as shadowing, freeze fracturing, and scanning electron microscopy are useful for this purpose. However, they can be further enhanced by stereo electron microscopy.

Specifically, the same specimen is photographed from two different angles to generate a stereo pair of photos that are then fused optically. To do this, a special specimen stage is used that can be titled relative to the electron beam. The specimen is first titled in one direction and photographed, then tilted an equal amount in the opposite direction and photographed again.

The two micrographs are mounted side by side as a stereo pair. When you view a stereo pair through a stereoscopic viewer, your brain uses the two independent images to construct a three dimensional view that gives a striking sense of depth to the structure under investigation.

Design # 2. Scanning Electron Microscopy:

Scanning electron microscopy is a relatively recent development. It is an especially spectacular technique because of the sense of depth it gives to biological structures, thereby allowing surface topography to be studied. As the name implies, a scanning electron microscope (SEM) generates an image by scanning the specimen with a beam of electrons.

The vacuum system and electron source are similar to those found in the transmission electron microscope, although the accelerating voltage is much lower (about 5-30 kV). The significant difference between the two kinds of instruments lies in the way the image is formed. In the SEM, a magnetic lens system focuses the beam of electrons into an intense spot on the surface of the specimen.

The spot is moved back and forth across the specimen by charged plates called beam deflectors located between the condenser lens and the specimen. The beam deflectors attract or repel the beam according to the signals sent to them by the deflector circuitry.

As the electron beams sweeps rapidly over the specimen, molecules in the specimen are excited to high energy levels and emit secondary electrons. These secondary electrons are used to form an image of the specimen surface. They are then captured by a detector that is located immediately above and to one side of the specimen.

The essential component of the detector is a scintillator, which emits photons of light when excited by the electron incident upon it. The photons are used to generate an electronic signal to a video screen. The image then develops point by point, line by line on the screen as the primary electron beam sweeps over the specimen.

Sample Preparation Techniques in Scanning Electron Microscopy:

When preparing a specimen for scanning electron microscopy, the goal is the preserve the structural features of the cell surface and to treat the tissue in a way that minimizes damage by the electron beam. The procedure is actually quite similar to the preparation of ultrathin sections for transmission electron microscopy.

The tissue is fixed in aldehyde, post-fixed in osmium tetroxide, are processed for dehydration through a series of alcohol solutions. The tissue is then placed in a fluid such as Freon and transferred to a heavy metal canister called a critical point bomb, which is use to dry the specimen under conditions of controlled temperature and pressure. This helps keep structure on the surfaces of the tissue in almost the same condition they were in before dehydration.

The dried specimen is then attached to a metal specimen mount with a metallic paste. The mounted specimen is coated with a layer of gold or a mixture of gold and palladium, using a modified form of vacuum evaporation called sputter coating. These procedures allow electrons to pass through the specimen more readily, thereby minimizing heating of the specimen has been mounted and coated, it is ready to be examined in the microscope.

Preparing specimens for electron microscopy is often expensive and time-consuming. However, the high resolution and unique perspective on the structure and function of cells provide worthwhile insights into the biology of cells and tissues.

Other Imaging Methods:

Light and electron microscopy are direct imaging methods in that they use photons or electrons to produce actual images of a specimen. There are other methods of microscopy that are indirect imaging methods.

To understand what is meant by indirect imaging, suppose you are given some object to handle with your eyes closed. You might feel six flat surfaces, twelve edges and eight corners, and if you then draw what you have felt, it would turn out to be a box. This is an example of an indirect imaging procedure.

The indirect imaging methods described here are scanning tunneling microscopy, atomic force microscopy, and X-ray diffraction. Each method has the potential for showing molecular structures at near-atomic resolution, ten times better than the best electron microscope.

Each method also has certain characteristics that limit its application to biological material, but when these techniques can be successfully used, the resulting images provide exciting information that cannot be obtained in any other way.

i. Scanning Tunneling Microscopy:

Although “scanning” is involved in both scanning electron microscopy and scanning tunneling microscopy, the two methods are in fact quite different. The scanning tunneling microscope (STM) does not use an electron beam, but instead depends on a tip made of a conducting material such as platinum-iridium.

The tip is extremely sharp, ideally with its point composed of a single atom. It is under precise control of an electronic circuit that can move it in three dimensions over a surface. The x and y dimensions scan the surface, while the z dimension governs the distance of the tip above the surface.

The basic principle of the STM is electron tunneling. At the quantum-mechanical level, an electron has both wavelike and particles like properties. These properties allow the electron to cross barriers that it cannot penetrate as a particle, but that it can penetrate in the form of a wave. This penetration is called tunneling. As the tip of the STM is moved across a surface, voltages from a few millivolts to several volts are applied.

Under these conditions, if the tip is close enough to the surface and the surface is electrically conductive, electrons will begin to tunnel between tip and surface. The tunneling is highly dependent on the distance, so that even small irregularities in the size range of single atoms will affect the rate of electron conductance.

An important limitation of the STM is that the specimen must be electrically conductive. Therefore, the technique is better studied to producing images of physical surfaces rather than biological specimens, which are often good insulators.

ii. Atomic Force Microscopy:

Atomic force microscopy (AFM) is related to STM, in that it uses a tip of atomic dimensions. It has the important advantage that the specimen does not need to be an electrical conductor. This is because the tip is actually moved over the specimen surface, bumping along individual atoms in the specimen. The minute movements of the tip are followed with an optical system that sense deflection.

One of the most important potential applications of the STM and AFM instruments is the measurement of dynamic changes in the conformation of a functioning biomolecule. Consider, for instance, how exciting it would be to “watch” a single enzyme molecule change its shape as it hydrolyzes ATP to provide the energy needed to transport ions across membranes.

iii. X-Ray Diffraction:

X-ray diffraction does not involve microscopy but instead reconstructs images from the diffraction patterns of X rays passing through a crystalline specimen. This method can be used to deduce molecular structure at the atomic level of resolution. X-ray diffraction is, in fact, the only method presently available to analyze the structure of proteins, nucleic acids, and other biological molecules at this resolution.

A good way to understand X-ray diffraction is to draw an analogy with light. Light has certain properties that are best described as wavelike. Whenever wave phenomena occur in nature, interaction between waves can occur. If waves from two sources come into phase with one another, their total energy is additive (constructive interference), and if they are out of phase, their energy is reduced (destructive interference).

This effect can be seen when light passes through two pinholes in a piece of opaque material and then falls on a white surface. Interference patterns result, with dark regions where light waves are out of phase and bright regions where they are in phase. If the wavelength of the light is known, one can measure the angle a between the original beam and the first diffraction peak and then calculate the distance d between the two holes with this formula –  

d = wavelength/sin α

The same approach can be used to calculate the distance between atoms in crystals of proteins or nucleic acids. Instead of two holes in a sheet of paper, imaging that we have multiple layers of atoms organized in a crystal. And instead of light, which has much too long a wavelength to interact with atoms, we will use a narrow beam of X rays with wavelengths in the range of interatomic distances.

As the X rays pass through the crystal, they reflect off planes of atoms, and the reflected beams come into constructive and destructive interference. If the X-ray beams are allowed to fall onto photographic plates behind the crystal, distinctive diffraction patterns are produced. These patterns can be analyzed mathematically to determine the organization of atomic layers in the original crystal and thus to deduce three-dimensional molecular structure.

The technique of X-ray crystallography was developed in 1912 by Sir William Bragg, who went on to establish the structures of relatively simple mineral crystals. Forty years later, Max Perutz and John Kendrew found ways to apply X-ray diffraction to crystals of hemoglobin and myoglobin, providing our first view of the intricacies of protein structure. Since then, many proteins and other biological molecules have been crystallized and subjected to X-ray diffraction.

Membrane proteins are much more difficult to crystallize than hemoglobin. In 1985, however, Henri Michel and Johannes Deisenhofer overcame this obstacle by crystallizing the proteins of a bacterial photosynthetic reaction center. They then went on to describe the molecular organization of the reaction center at a resolution of 0.3 nm.

Design # 3. Scanning Transmission Electron Microscopy:

A scanning transmission electron microscope (STEM) contains elements of both transmission and scanning electron microscopes. Like an SEM, or TEM uses an electron beam that sweeps over the specimen. But the image is then formed by electrons transmitted through the specimen, as with a TEM.

An STEM is capable of distinguishing specific characteristics of the electrons that are transmitted by the specimen, thus deriving information about the specimen not obtainable with a conventional TEM. However, an STEM is technically sophisticated, requires a very high vacuum, and is much more electronically complex than a TEM or an SEM.

High-Voltage Electron Microscopy:

A high voltage electron microscope (HVEM) is very similar to a transmission electron microscope except that its accelerating voltage is much higher. Whereas a TEM uses accelerating voltages of 50-100 kV, an HVEM uses voltage of about 200-1000 kv. Because of the high voltage and the greatly reduced chromatic aberration that it makes possible, relatively thick specimens can be examined with good resolution. As a result, cellular structure can be studied in sections as thick as 1 µm, about ten times the thickness possible with an ordinary TEM.