TEM followed by SPA has contributed many macromolecular 3D structures, nearing 2,000 entries in the EMDB by mid 2014. In general, for a given macromolecule, a low-resolution 3D structure is first determined from NS data, which may be followed by a higher-resolution cryoEM 3D structure. The high contrast of the molecular boundaries provided by NS, which facilitates the first 3D reconstruction, is later refined by cryoEM with its ability to map the internal structure of the macromolecule in its fully hydrated state, and the possibility to attain much higher resolution. Another advantage of cryoEM is the elimination of dehydration stress that could result in collapse of the macromolecule. In addition, there is the possibility to omit the carbon support, which eliminates possible surface absorption effects and shows the conformation that the macromolecule adopts in solution. Cryo-negative staining, a combination of cryoEM and NS, affords high contrast in a fully hydrated specimen and can also lead to 3D reconstruction using SPA, however it has been less frequently used, with less than 10 EMDB entries, in part because the stain itself may interfere with the integrity of the macromolecule 31. Two other TEM techniques conducive to 3D reconstruction are 2D electron crystallography and electron tomography. 2D electron crystallography requires a planar or tubular crystal; the crystal's electron diffraction is used for 3D reconstruction potentially leading to atomic resolution 32. In electron tomography of vitrified or traditionally fixed specimens, a macromolecule or sub-cellular component is rotated inside the TEM for subsequent tomographic reconstruction 33, with the advantage that singular objects can be reconstructed; however at present this technique has a resolution limit ranging from 40 to 20 Å 34,35. Among all these molecular TEM techniques, SPA of NS and cryoEM data has been the most widely used. The protocol illustrated here is dedicated to obtaining cryoEM images suitable for SPA analysis; nevertheless most of the protocol is also applicable to cryoEM of 2D crystals and tomographic samples.
A successful cryoEM session depends on the combined success of many critical steps; important aspects to keep in mind, explanation of common cryoEM artifacts and how to avoid them are described in the following paragraphs. This section also describes data collection guidelines to obtain high quality 3D reconstructions using the SPA method.
Avoid transition to crystalline ice. A central aspect is that the sample needs to stay in vitreous state from the moment of cryo-plunging throughout TEM observation. Thus after plunging the sample in liquid ethane all subsequent steps are performed at liquid nitrogen (-196 °C) or liquid helium (-269 °C) temperatures. Warming to above -135 °C transforms vitreous water into crystalline water; then macromolecular rearrangements may take place and water crystals dominate the image (Figure 3A); the sample should be discarded. Accidental sample warm-up could happen if cryo-plunging, transfer of the frozen grid between containers (even if protected by a gridbox), and/or cryo holder insertion into the TEM are too slow, or if the handling tweezers were insufficiently pre-cooled. Significant vacuum deterioration in the TEM upon cryo-transfer (the cold sample traps warmer incoming air) may also warm up the sample. Finally, over-irradiation of the sample could also result in transition to crystalline ice.
Minimize ice contamination. Given the small sample volume (3-5 µl) applied to the TEM grid, evaporation could concentrate buffer components (salt, detergents) and thus affect macromolecular integrity including loss of multimeric state. A high relative humidity (RH) microenvironment or chamber circumvents this problem. Alternatively cryo-plunging can be done in a sufficiently ventilated environmental cold room. After cryo-plunging RH should be as low as possible to prevent ice contamination, or condensation of ambient humidity on the cryo-grid, as the cryo-grid itself acts as a cold trap. Ice contamination consists of particles with high contrast and no substructure with sizes ranging between ~5 nm and several microns that interfere with or even totally block the image. Avoid air drafts, talking/breathing towards the cryo-grid during air transfer, and reduce ambient RH. Open liquid nitrogen containers with condensed water (visible as white suspended particles) should also be discarded.
Maximize macromolecular orientations/views. For sample support, a choice to be made is between true holey grids and thin carbon over holey grids. This depends on (i) how the sample spreads over carbon film versus the carbon holes, (ii) available sample concentration, as bare holes may require a concentration 100X higher than carbon support for a similar particle density on the image (from ~0.02 to 2 µM), and (iii) how random is the distribution of macromolecule orientations. Note that glow discharge, which makes the carbon hydrophilic, will have an important effect in all three aspects and that this will be sample-dependent. Regarding the orientation of the macromolecule, while a recurrent view is desirable for the initial structural determination by random conical reconstruction, randomness of macromolecular views is desirable when refining the 3D reconstruction to higher resolutions. In our example, RyR1 interacts with the carbon with a favorite 'fourfold' view (Figure 2D) and requires holey grids for randomness of orientations and higher resolution 36.
Maximize SNR. The best strategy to increase SNR is to reduce the background sources of noise. Excessive ice thickness and (if present) carbon film thickness beyond what is needed to support and embed the protein will add extra noise. Thus ice thickness should be reduced to the minimum necessary by controlling blotting time, pressure, RH, and filter paper quality. For carbon support, a thin (~5 nm) carbon film is layered over the thicker holey carbon film: the thick holey carbon provides mechanical resistance while the sample is imaged over the thin carbon-covered hole. On the other hand, note that extremely thin ice and/or carbon may result in an easily broken support that can turn into a web (Figure 3D). To maximize SNR, any unnecessary buffer components should be avoided. For membrane proteins, the detergent takes a significant toll on contrast (see Figure 2D, right panel). In addition by reducing surface tension the detergent changes the way in which the sample spreads on the support. Some membrane proteins require the presence of lipid in addition to detergent, which further reduces the contrast. To overcome this the sample can be diluted in a buffer with lower detergent concentration and without lipid just before cryo-plunging 37. New alternative detergents 16 and the use of nanodisks 38 to stabilize the transmembrane domain component appear to be successful approaches for imaging membrane proteins by cryoEM.
Strive for high quality images and prepare for the CTF correction. Vitrified specimens require high defocus values to generate sufficient image (phase) contrast where the CTF, the function that relates intensity to the frequency, has several zero transitions, at which point there is no information. While CTF correction is not necessary for low-resolution 3D reconstruction, when aiming for higher resolution, to recover an accurate representation of the 3D object, images with different defoci need to be collected so that computational CTF correction can be done. CTF determination and correction is included in the major SPA software packages 4-7; details can be found elsewhere 39. For optimal CTF correction, use a range of defoci. A good strategy is to alternate amongst 3-4 defocus values. The values depend on the size of the macromolecule (larger sizes require less defocus), the ice/carbon thickness (thinner sample requires less defocus), and the voltage (lower voltage requires less defocus). For 200 kV, a good starting range of values is 2.5, 3, 3.5 and 4 µm defocus. The CTF manifests as alternating rings of high and low intensity (Thon rings 40) in the reciprocal space representation, the power spectrum. Displaying the power spectrum will reveal the potential for resolution; the frequency of the widest visible Thon ring is the closest a priori estimate of the attainable resolution. The power spectrum should be checked periodically, and astigmatic (non circular) Thon rings (Figure 3C) should be corrected with the objective stigmator. Thon rings should be visible at least up to the resolution desired.
A cryoEM dataset obtained using this protocol can be directly processed by SPA in order to generate a 3D reconstruction. Even with an ideal sample, the quality of the 3D reconstruction will depend on the performance and specifications of the TEM equipment, and on the image processing. With the continued improvements in both these fronts, and with special mention to the recently developed direct electron detectors, the possibility to obtain 3D reconstructions at atomic resolution more consistently is closer than it has ever been.
The History of EM
By the middle of the 19th century, microscopists had accepted that it was simply not possible to resolve structures of less than half a micrometre with a light microscope because of the Abbe’s formula, but the development of the cathode ray tube was literally about to change the way they looked at things; by using electrons instead of light! Hertz (1857-94) suggested that cathode rays were a form of wave motion and Weichert, in 1899, found that these rays could be concentrated into a small spot by the use of an axial magnetic field produced by a long solenoid. But it was not until 1926 that Busch showed theoretically that a short solenoid converges a beam of electrons in the same way that glass can converge the light of the sun, that a direct comparison was made between light and electron beams. Busch should probably therefore be known as the father of electron optics.
In 1931 the German engineers Ernst Ruska and Maximillion Knoll succeeded in magnifying and electron image. This was, in retrospect, the moment of the invention of the electron microscope but the first prototype was actually built by Ruska in 1933 and was capable of resolving to 50 nm. Although it was primitive and not really fit for practical use, Ruska was recognised some 50 years later by the award of a Nobel Prize. The first commercially available electron microscope was built in England by Metropolitan Vickers for Imperial College, London, and was called the EM1, though it never surpassed the resolution of a good optical microscope. The early electron microscopes did not excite the optical microscopists because the electron beam, which had a very high current density, was concentrated into a very small area and was very hot and therefore charred any non-metallic specimens that were examined. When it was found that you could successfully examine biological specimens in the electron microscope after treating them with osmium and cutting very thin slices of the sample, the electron microscope began to appear as a viable proposition. At the University of Toronto, in 1938, Eli Franklin Burton and students Cecil Hall, James Hillier and Albert Prebus constructed the first electron microscope in the New World. This was an effective, high-resolution instrument, the design of which eventually led to what was to become known as the RCA (Radio Corporation of America) range of very successful microscopes.
Unfortunately, the outbreak of the Second World War in 1939 held back their further development somewhat, but within 20 years of the end of the war routine commercial electron microscopes were capable of 1 nm resolution.
Types of Electron Microscopes
All electron microscopes use electromagnetic and/or electrostatic lenses to control the path of electrons. Glass lenses, used in light microscopes, have no effect on the electron beam. The basic design of an electromagnetic lens is a solenoid (a coil of wire around the outside of a tube) through which one can pass a current, thereby inducing an electromagnetic field. The electron beam passes through the centre of such solenoids on its way down the column of the electron microscope towards the sample. Electrons are very sensitive to magnetic fields and can therefore be controlled by changing the current through the lenses.
The faster the electrons travel, the shorter their wavelength. The resolving power of a microscope is directly related to the wavelength of the irradiation used to form an image. Reducing wavelength increases resolution. Therefore, the resolution of the microscope is increased if the accelerating voltage of the electron beam is increased. The accelerating voltage of the beam is quoted in kilovolts (kV). It is now possible to purchase a 1,000kV electron microscope, though this is not commonly found.
Although modern electron microscopes can magnify objects up to about two million times, they are still based upon Ruska's prototype and the correlation between wavelength and resolution. The electron microscope is an integral part of many laboratories such as The John Innes Centre. Researchers can use it to examine biological materials (such as microorganisms and cells), a variety of large molecules, medical biopsy samples, metals and crystalline structures, and the characteristics of various surfaces. Nowadays, electron microscopes have many other uses outside research. They can be used as part of a production line, such as in the fabrication of silicon chips, or within forensics laboratories for looking at samples such as gunshot residues. In the arena of fault diagnosis and quality control, they can be used to look for stress lines in engine parts or simply to check the ratio of air to solids in ice cream!
Transmission Electron Microscope (TEM)
The original form of electron microscopy, Transmission 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.
Resolution of the TEM is also limited by spherical and chromatic aberration, but a new generation of aberration correctors has been able to overcome or limit these aberrations. Software correction of spherical aberration has allowed the production of images with sufficient resolution to show carbon atoms in diamond separated by only 0.089 nm and atoms in silicon at 0.078 nm at magnifications of 50 million times. The ability to determine the positions of atoms within materials has made the TEM an indispensable tool for nano-technologies research and development in many fields, including heterogeneous catalysis and the development of semiconductor devices for electronics and photonics. In the life sciences, it is still mainly the specimen preparation which limits the resolution of what we can see in the electron microscope, rather than the microscope itself.
At JIC we have a high voltage (200kV) TEM, which was installed in 2008. We have two digital cameras on it, one is higher resolution than the other, so that the need for developing and printing film has been negated. Our TEM is designed for use with biological samples and is capable of resolving to better than 1nm. It is also capable of 3-D tomography which involves taking a succession of images whilst tilting the specimens through increasing angles, which can then be combined to form a three-dimensional image of the specimen.
Scanning Electron Microscope (SEM)
Unlike the TEM, where the electrons in the primary beam are transmitted through the sample, the Scanning Electron Microscope (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.
|SEM image of a fly's foot taken at JIC in 2006||From "Micrographia", by Robert Hooke, 1665: plate showing the drawing of a fly's foot|
TEM resolution is about an order of magnitude better than the SEM resolution. Our TEM can easily resolve details of 0.2nm. Our two SEMs at JIC are both relatively recent acquisitions and are high-resolution instruments capable of about 2 nm resolution on biological samples. Because the SEM image relies on electron interactions at the surface rather than transmission it is able to image bulk samples and has a much greater depth of view, and so can produce images that are a good representation of the 3D structure of the sample. SEM images are therefore considered to provide us with 3D, topographical information about the sample surface but will still always be only in black and white.
In the SEM, we use much lower accelerating voltages to prevent beam penetration into the sample since what we require is generation of the secondary electrons from the true surface structure of a sample. Therefore, it is common to use low KV, in the range 1-5kV for biological samples, even though our SEMs are capable of up to 30 kV.
At JIC we currently have two SEMs, both with high-resolution capabilities, digital imaging facilities and cryo-systems which enable them to be used for looking at frozen-hydrated specimens.
Materials to be viewed in an electron microscope generally require processing to produce a suitable sample. This is mainly because the whole of the inside of an electron microscope is under high vacuum in order to enable the electron beam to travel in straight lines. The technique required varies depending on the specimen, the analysis required and the type of microscope:
Cryofixation - freezing a specimen rapidly, typically to liquid nitrogen temperatures or below, that the water forms ice. This preserves the specimen in a snapshot of its solution state with the minimal of artefacts. An entire field called cryo-electron microscopy has branched from this technique. With the development of cryo-electron microscopy, it is now possible to observe virtually any biological specimen close to its native state.
Fixation - a general term used to describe the process of preserving a sample at a moment in time and to prevent further deterioration so that it appears as close as possible to what it would be like in the living state, although it is now dead. In chemical fixation for electron microscopy, glutaraldehyde is often used to crosslink protein molecules and osmium tetroxide to preserve lipids.
Dehydration - removing water from the samples. The water is generally replaced with organic solvents such as ethanol or acetone as a stepping stone towards total drying for SEM specimens or infiltration with resin and subsequent embedding for TEM specimens.
Embedding - infiltration of the tissue with wax (for light microscopy) or a resin (for electron microscopy) such as araldite or LR White, which can then be polymerised into a hardened block for subsequent sectioning.
Sectioning - the production of thin slices of the specimen. For light microscopy, the sections can be a few micrometres thick but for electron microscopy they must be very thin so that they are semitransparent to electrons, typically around 90nm. These ultra-thin sections for electron microscopy are cut on an ultramicrotome with a glass or diamond knife. Glass knives can easily be made in the laboratory and are much cheaper than diamond, but they blunt very quickly and therefore need replacing frequently.
Staining - uses heavy metals such as lead and uranium to scatter imaging electrons and thus give contrast between different structures, since many (especially biological) materials are nearly "transparent" to the electron beam. By staining the samples with heavy metals, we add electron density to it which results in there being more interactions between the electrons in the primary beam and those of the sample, which in turn provides us with contrast in the resultant image. In biology, specimens can be stained "en bloc" before embedding and/or later, directly after sectioning, by brief exposure of the sections to solutions of the heavy metal stains.
Freeze-fracture and freeze-etch - a preparation method particularly useful for examining lipid membranes and their incorporated proteins in "face on" view. The fresh tissue or cell suspension is frozen rapidly (cryofixed), then fractured by simply breaking or by using a microtome while maintained at liquid nitrogen temperature. The cold, fractured surface is generally "etched" by increasing the temperature to about -95°C for a few minutes to let some surface ice sublime to reveal microscopic details. For the SEM, the sample is now ready for imaging. For the TEM, it can then be rotary-shadowed with evaporated platinum at low angle (typically about 6°) in a high vacuum evaporator. A second coat of carbon, evaporated perpendicular to the average surface plane is generally performed to improve stability of the replica coating. The specimen is returned to room temperature and pressure, and then the extremely fragile "shadowed" metal replica of the fracture surface is released from the underlying biological material by careful chemical digestion with acids, hypochlorite solution or SDS detergent. The floating replica is thoroughly washed from residual chemicals, carefully picked up on an EM grid, dried then viewed in the TEM.
Sputter Coating - an ultra-thin coating of electrically-conducting material, deposited by low vacuum coating of the sample. This is done to prevent charging of the specimen which would occur because of the accumulation of static electric fields due to the electron irradiation required during imaging. It also increases the amount of secondary electrons that can be detected from the surface of the sample in the SEM and therefore increases the signal to noise ratio. Such coatings include gold, gold/palladium, platinum, chromium etc.
Disadvantages of Electron Microscopy
Electron microscopes are very expensive to buy and maintain. They are dynamic rather than static in their operation: requiring extremely stable high voltage supplies, extremely stable currents to each electromagnetic coil/lens, continuously-pumped high/ultra-high vacuum systems and a cooling water supply circulation through the lenses and pumps. As they are very sensitive to vibration and external magnetic fields, microscopes aimed at achieving high resolutions must be housed in buildings with special services.
A significant amount of training is required in order to operate an electron microscope successfully and electron microscopy is considered a specialised skill.
The samples have to be viewed in a vacuum, as the molecules that make up air would scatter the electrons. This means that the samples need to be specially prepared by sometimes lengthy and difficult techniques to withstand the environment inside an electron microscope. Recent advances have allowed some hydrated samples to be imaged using an environmental scanning electron microscope, but the applications for this type of imaging are still limited.
It must be emphasised from the outset that every electron micrograph is, in a sense, an artefact. Changes in the ultra-structure are inevitable during all the steps of processing that samples must undergo: material is extracted, dimensions are changed and molecular rearrangement occurs. The best thing we can do is to keep these changes to a minimum by understanding the processes involved so that we make informed choices of the best preparative procedures to use for each sample. Artefacts present themselves in many ways: there could be loss of continuity in the membranes, distortion or disorganisation of organelles, empty spaces in the cytoplasm of cells or sharp bends or curves in filamentous structures that are usually straight, such as microtubules and so on. With experience, microscopists learn to recognise the difference between an artefact of preparation and true structure, mainly by looking at the same or similar specimens prepared in the same or a different way.
Scanning electron microscopes usually image conductive or semi-conductive materials best. Non-conductive materials can be imaged, either by an environmental scanning electron microscope or more usually by coating the sample with a conductive layer of metal. A common preparation technique is to coat the sample with a layer of conductive material, a few nanometers thick, such as 10nm of gold, from a sputtering machine. This process does, however, have the potential to disturb delicate samples and cover some detail. When using chemical fixation and dehydration as part of the sample preparation, there is often much shrinkage and collapse of delicate structures and so, especially for our interests at JIC in botanical specimens which are highly hydrated, we tend to use the cryo-fixation technique which is far less prone to artefacts.
For the TEM, samples are generally prepared by exposure to many nasty chemicals, in order to give good ultra-structural detail which may result in artefacts purely as a result of preparation. This gives the problem of distinguishing artefacts from genuine structures within the specimen, particularly in biological samples. Scientists maintain that the results from various preparation techniques have been compared, and as there is no reason that they should all produce similar artefacts, it is therefore reasonable to believe that electron microscopy features correlate with living cells. In addition, higher resolution work has been directly compared to results from X-ray crystallography, providing independent confirmation of the validity of this technique. Recent work performed on unfixed, vitrified (rapidly frozen, without the use of any chemicals, to form ice without any crystallisation) specimens has also been performed, further confirming the validity of this technique. However, even cryo-fixation techniques are not without their own artefacts of preparation and ice crystal damage, due to the fact that as water freezes it expands, is a common problem when trying to image a large specimen (greater than 200 µm) which cannot be frozen rapidly enough to vitrify the water