The efficiency of implantable electrodes for cardiac pacing depends on the ratio of the conductive surface area to the geometric area of the interface with excitable tissue. New models of heart pacers require reduction of post-pulse polarization, i.e. the potential left on the electrode / tissue interface after a pacemaker pulse. Increasing the conductive surface area is an effective method to this end. Microscopy provides an important tool in elucidating the role of surface structure in electrode performance.

Three different surface textures were characterized on a 90% platinum(Pt)/10% iridium (Ir) polished electrode substrate of roughly 5 mm2 geometric surface area. These consisted of the polished substrate itself, a thin film of textured platinum in the 1-3 micron size range, and a sub-micron platinum black coating. Sample surface effects were characterized via scanning electron microscopy (SEM), in-vitro electrical impedance spectrum analysis, and polarization after-potential measurements.

RecA has been the most intensively studied enzyme in homologous genetic recombination. Actin is one of the most abundant and conserved eukaryotic proteins, and is central to muscle contraction, cell motility, and the control of cell shape. Both form helical filaments, and we have been studying both for many years, primarily using negatively stained specimens. We and others have used cryo-EM of frozen-hydrated RecA and F-actin samples, but the resolution obtained by this method is actually worse than that obtained from negatively stained specimens. The criterion for resolution that we have used is the match with an atomic model for F-actin, since this is a much better standard than statistical measures such as Fourier ring correlations or phase residuals, which can only measure the internal consistency of the data.

F-actin is a flexible structure, both with respect to bending and torsional motions of subunits. The RecA filament is 10 times more flexible than F-actin. The bending of RecA and F-actin require that images be straightened, and the ability of a user to fit a spline function to the filament axis limits the precision of the method. The method is only valid for small perturbations from a rigid rod, and thus will introduce artifacts for real filaments. In order to straighten filament images obtained in ice, one needs sufficient contrast (obtained by a large defocus) to be able to apply such algorithms. The torsional motions of the actin subunits, which may be as large as 5-6° per subunit, limit the resolution that can be obtained as one averages over long lengths.

The identification of tubulin as the major protein component of microtubules (1) occurred at about the same time, a little over thirty years ago, as the first demonstration of the ability to obtain 3- dimensional data from electron micrographs. DeRosier and Klug's paper on image reconstruction of the bacteriophage tail (2) marks what many consider the beginning of electron crystallography - determining protein structures by electron microscopy. Microtubules have long been the target of active study by light and electron microscopy, and even before the discovery of tubulin played a significant role in driving technical developments in EM. The demonstration that tubulin could form two-dimensional crystalline sheets (3) seemed to pave the way for high resolution .structural studies. The discovery of the sheet polymer of tubulin came shortly after Unwin and Henderson's first publications on bacteriorhodopsin showing secondary structure resolved by EM (4). Application of their methods to tubulin in the late 1970's resolved low resolution features of the protein, but technical problems prevented achieving much more than showing the molecular envelope. Many developments in both tubulin studies and electron crystallography followed this work over the next 15 years. As techniques were refined for purifying, labeling and visualizing tubulin and microtubules, the dynamic properties of microtubules and their role in the cell cycle were characterized, and some hints about the mechanism of polymerization and disassembly were derived from biophysical and microscopic investigations.

We have reconstructed the three-dimensional quaternary structure of the complete 480 kDa insulin receptor (IR), complexed with NanoGold-labelled insulin, via sets of electron micrographs obtained by low-dose low-temperature dark field scanning transmission electron microscopy (STEM).

Insulin binding to IR in mammalian cell membranes is essential for its manifold effects such as glucose homeostasis, increased protein synthesis, growth, and development. IR belongs to the superfamily of transmembrane receptor tyrosine kinases that include the monomeric epidermal growth factor receptor (EGFR) and platelet-derived growth factor receptor (PDGFR). In contrast, IR and its homologues IGF-1R (insulin-like growth factor 1 receptor) and IRR (insulin receptorrelated receptor) are sub-types of this family that are intrinsic disulfide-linked dimers of two αβ heterodimers. Monomeric receptor TKs are inactive, but are activated by ligand-induced dimerization that results in autophosphorylation. IR-like TKs are also inactive even though they are already dimeric, and are activated by ligand binding without further oligomerization. Insulin binding to the extracellular domain of IR results in autophosphorylation of specific tyrosines to initiate an intracellular signal transduction cascade. However, because the quaternary structure of IR is not known, the structural basis for the mechanism of IR activation by extracellular insulin binding has not been elucidated.

The insulin receptor was purified from human placenta. Bovine insulin was derivatized with NanoGold at the B chain Phel, a location not directly involved in receptor binding. Binding of derivatized insulin to the purified receptor was reduced only slightly compared to binding of the native insulin.

Recent improvements in digital image acquisition, and automated TEM, have lowered the electron dose requirement for collecting a tomographic tilt series to the point where cryo-electron tomography is now feasible. Theoretical calculations indicate that resolution values of 2-5 nm can be obtained for fully hydrated specimens 100-500 nm thick. This resolution is sufficient for identifying individual macromolecules, and macromolecular assemblies, within-organelles and small cells. However, initial applications to cells and organelles fell short of theoretical resolution limits, indicating further development is necessary before cryo-electron tomography reaches its full potential for characterizing macromolecules in situ. Axonemes are ideal test specimens for cryo-tomography because of their appropriate size (diameter of 250 nm), ease of isolation, and unique construction from regular arrays of well-characterized components. The latter include microtubules, radial spokes, central sheath components, outer dynein arm, inner dynein arms, and the microtubule subunit lattice. These components are convenient benchmarks for evaluating the actual detection limit of tomographic reconstructions within a resolution range of 3-30 nm. Furthermore, axoneme components are sufficiently well ordered that computed diffraction patterns contain layer-lines that are useful for judging the quality of individual 2D images.

The past decade has seen major advances in the analytical capability and utility of both fixed beam and scanning beam electron microscopes. In particular, scanning transmission electron microscopy (STEM) and energy-filtering transmission microscopy (EFTEM) have benefited from the development of devices and techniques—including improved electron optics, sensitive solid-state detectors and new software for imaging and electron energy loss spectroscopy (EELS)—that optimize detection of weak spectroscopic signals arising from biological specimens while minimizing specimen damage. Here we discuss and illustrate some of these advances, especially in the context of structural imaging, detection limits and mapping techniques for the biologically important elements phosphorus and calcium. Analytical microscopy of biological tissues is absolutely dependent on cryotechniques. It is generally agreed that rapid freezing and subsequent low-temperature processing, e.g., cryosectioning or direct cryotransfer of frozen-hydrated specimens, is the most reliable way to preserve the native distribution and organization of biological structures. Equally important, however, as an adjunct to spectroscopic analysis is the use of established low-temperature, low-dose techniques for recording optimized images. By limiting beam exposure, low-dose methods greatly improve the quality of images from fragile, freeze-dried preparations. In this case, the quality and information content of, e.g., cryosections are virtually as good as conventional preparations (Fig 1).

Cryo-electron tomography can be used to obtain highly-detailed 3-D reconstructions of micrometer-sized biological structures which are unfixed, unstained and embedded in vitreous ice. Important practical considerations for acquiring the requisite tilt-series images are the choice of electron dose and defocus value, and the effects of contamination and ice thickness.

It is essential to reduce the total dose enough to prevent excessive ice radiolysis and damage to the specimen itself. To reduce the dose, a sensitive cooled CCD camera and microscope automation system are all but required. Because the specimens are thick, it is advantageous to use a high accelerating voltage, and/or energy filtering to reduce inelastic scattering. At high accelerating voltage, it is important to keep the dose high enough so that the signal is above the “scintillator noise” generated by the interaction of the high-energy electrons with the phosphor scintillator of the CCD camera. Initial test reconstructions indicate that scintillator noise, unlike electron shot noise, cannot be mitigated using dose fractionation. Fortunately, we find that large objects embedded in a thick (ca. 1 μm) vitrified ice layer are more resistant to beam damage (as well as charging) compared to smaller objects embedded in ice 100-200nm thick.

Electron microscopic tomography is providing important new insights about the internal structure of the mitochondrion. In particular, the infoldings of the mitochondrial inner membrane (cristae), which are usually rendered as lamelliform baffles, are revealed to have considerable tubular nature. Rather than opening wide to the peripheral compartment (between the inner and outer membranes), the cristae connect to the outside and to each other through narrow (20-30 nm) tubular segments, which can be hundreds of nanometers long. This suggests that diffusion of ions, metabolites and proteins between the intracristal and intermembrane spaces may be restricted.

The earlier tomographic reconstructions were done on conventionally prepared, plastic-embedded specimens, which raises the usual concerns about structural preservation. More recently, we have undertaken tomography of isolated rat-liver mitochondria that have been embedded in vitreous ice (by plunge-freezing in iso-osmotic buffer without chemical fixatives or stains). These frozen hydrated specimens are imaged with a JEOL 4000FX equipped with a Gatan cryo-transfer holder and a Tietz automated data collection system, with a Ik × Ik CCD. For 3D reconstructions, images were recorded at a dose of 5 e−Å2 at 2° increments over the range +/− 60°.

When resins are ultrathin sectioned they are compressed in the sectioning direction. The amount of compression depends very much on the sectioning angle on one hand, and on the properties of the resin on the other hand. Shortening of the sections in the sectioning direction is in the range of 5 to 35%. Even more compression occurs during cryosectioning of vitreous samples at very low temperatures (-170°C); the range was measured to be 30 to 60% (1).

To reduce these compression phenomena, there is basically only one approach possible. The sectioning angle has to be reduced as much as possible. In the past it was shown that by reducing the knife angle from 45° to 35°, a reduction of compression by appoximately one third was achieved (2). The problem with diamond knives is, that the smaller the knife angle, the more susceptible the cutting edge becomes. In practice angles smaller than 35° are not used because of this limitation.

Field emission scanning electron microscopy (FESEM) provides a direct view of a biological sample with high spacial resolution. Combining FESEM with cryo-techniques, macromolecular structures have been obtained from single SEM image successfully (Hermann & Müller, 1992; Chen et al., 1995, 1997). This protocol is now applied to study the extracellular matrix of the diatom A. longipes.

Stalk, which is originated from the terminal region of diatom cells, consists of 3 regions: a surfaceadhered pad, a shaft that separates the cell from the substratum, and a ring-like collar (Fig. 1). Investigating the structure and function of the stalk will help to understand the adhesion and interaction of diatom with the external environment. The extracellular matrix of diatom is highly hydrated, it is difficult to preserve them for EM. It will be easily extracted and collapse during specimen preparation (e.g. fixation, dehydration). In order to preserver the stalk structures close to their native condition and observer them with high fidelity, cryo-preparation techniques are implemented. Cryo-technique is widely used in the plant biology. However, to our knowledge, it has not been used in the diatom research.

The cells of protein-storing plant tissues, including the aleurone of monocots and the cotyledons of dicots, have been widely used to study biological phenomena. The cells of protein-storing tissues share many characteristics including the presence of protein storage vacuoles (PSV) which act as reservoirs for the proteins, lipids and minerals required for germination. Many proteins synthesized in these tissues during germination are transported to the PSV. As a result, protein-storing tissues have become a model system for the study of protein transport.

Research in our laboratory focuses on the protein-storing barley aleurone layer. The aleurone layer is notoriously difficult to prepare for transmission electron microscopy (TEM). Aleurone cells are surrounded by thick cell walls that are difficult to penetrate with fixatives and resins. To increase penetration of chemicals, most of our studies are performed on protoplasts, aleurone cells that have had their walls enzymatically removed. Protoplasts, however, are sensitive to the osmolality of the fixation medium and solutions must be closely monitored during specimen preparation. The development of cells in the aleurone layer adds to the difficulty of preparation for TEM.

A structure preservation of biological and organic samples, close to native state, can only be reached by cryo immobilization techniques. Cryo immobilization allows not only to preserve the high structural integrity but also to arrest dynamic processes in the μs- ms range.

After freezing the sample and preparing the surface of interest, it is important to prevent the sample from ice crystal damage, removal of structural water, condensation of water or other contaminants until imaging. Therefore, ideally the samples are kept below the recrystallisation temperature of water (< 147K) during the transfer from the preparation environment into the imaging chamber.

For the transfer of frozen samples several concepts have been followed in the,past: a) the specimen after manipulation/preparation were submersed in liquid nitrogen and transferred to the cold stage of the microscope or b) a preparation chamber was permanently attached to the microscope column allowing the direct transfers between the preparation chamber and the cold stage in the microscope. These concepts allow either a high grade of flexibility combined with a high risk of contamination or to prevent contamination but combined with inflexibility. In addition the later also does not allow using the microscope during the specimen preparation procedure, nor transferring the specimen to an other imaging device.

It should be no secret by now that ultrarapid freezing is a superior method of specimen preparation for many biological EM projects, and that high pressure freezing is the most versatile of the freezing methods. While cryopreservation is not necessary for all EM studies, it is the method of choice for high resolution work and where “fixation artifacts”, such as distorted membranes, or extraction of the cytosol is a problem. It is true that the machines are expensive, and not always immediately available, but for some questions it is the appropriate technology to use. In the U.S., there are machines available for general use in the far West (Berkeley, CA), the upper Midwest (Madision, WI and Minneapolis, MN), and in the Northeast (Albany, NY). For locations of any of the other 8 machines around the country, interested users can call Technotrade, International at (603) 622-5011.

Current Status of High Pressure Freezing: A decade ago, Studer et al. wrote an article entitled: “High Pressure Freezing Comes of Age” that illustrated how high pressure freezing (HPF) had become a proven technology, useful for preserving ultrastructure with unmatched fidelity in cell types that had previously been difficult to fix well for electron microscopy (EM).

Sample preparation techniques for electron-microscopy (EM) dictate our perception of the microworld: any structural information which is lost or distorted during preparation can not be regenerated later and might lead to wrong interpretation of the observed micrograph.

Cryofixation based procedures for specimen preparation can avoid most of the structural alterations associated with standard techniques based on chemical fixation. The ultrastructure can be represented in a near “native state” thanks to the high time resolution for dynamic cellular events (1).

High pressure freezing (2) permits to cryoimmobilize biological samples up to approx. 200μm thick, in contrast to rapid freezing procedures at ambient pressure that are useful to cryoimmobilize samples up to 10-20 μm thick. The actual samplethickness that can be adequately frozen (=without visible damage due to ice crystal formation in freeze-substituted or freeze-fractured specimens) depends on the concentration of naturally occuring substances that exhibit cryoprotective activities.

Cells are information driven systems. Cellular information is stored in certain molecules, at certain places, in a certain concentration, at a particular time and under given physiological conditions. The goal of biological electron microscopy is to provide this information network, to correlate the cellular ultrastructure and its function. In this sense, it is essential to combine the high resolution of our electron optical instruments with a high information density of the biological system. Most of the structural information is lost in the course of the different preparation steps prior to electron microscopy. For this reason it is necessary that all preparation steps such as: 1. sampling: e.g. excision of tissues, 2. cryoimmobilisation, 3. follow-up procedures: e.g. freeze-fracturing, freeze-substitution and embedding, should have identical high quality levels preventing or minimizing the loss of structural information. To this aim our methodological activities focus on the development of special micro-techniques for the sampling of: a) native tissues, with an automatic fine-needle biopsy technique (1), of b) suspensions, with a special cellulose capillary technique (2), of c) cell monolayer, with a thin film cultivation technique (3) and the application/perfection of cryotechniques (high-pressure freezing (HPF) and freeze-substitution). In particular, the high-pressure freezer (HPM 010, Bal-Tec) has proven to be a highly useful tool for successful cryoimmobilization of almost any kinds of cells and tissues, bulk specimens (< 200 μm in thickness) being included. This freezing technique does not require any cryoprotection, and if combined with our micro-techniques the risk of inducing artefacts as a result of specimen preparation prior to freezing is minimized.

To date, high pressure freezing is the only reliable cryofixation method for vitrifying bulk biological samples up to 150μm in thickness (1). This proves the enormous potential of the method. Nevertheless, the procedure is not well accepted by the scientific community. It seems that people not completely convinced by the method have reservations to use high pressure freezing machines. The main reasons seem to be that the commercially available machines are noisy, large, very heavy, expensive and consume a lot of liquid nitrogen. Last but not least, a relatively large volume of cryogen is pressurized to 2000bar representing a potential risk of accident. Considering high pressure freezing to be a very potent method, it was tried to establish a new machine avoiding the above mentioned disadvantages.

The first functional high pressure freezing apparatus was freezing fine metal rods with a boring of 0.2mm in diameter (2). This boring was filled with suspensions, the rod was connected to a pressure generator thus pressurizing the suspension, and simultaneously the metal rod was frozen from the outside by a low pressure jet of liquid nitrogen. The big disadvantage of the method was that only freeze-fracture experiments could be performed at the time because the sample could not be removed from the boring. As a consequence, this approach was never commercialised. In the mean time sample preparation has improved. Cellulose capillaries can be used to freeze suspensions and tissue samples (3,4). To freeze these capillaries in the new machine, they are introduced in a sample holder which consists of a copper tube with an inner diameter of 0.3mm and a length of 16mm. This tube fits into a concentrical stainless steel tube which stabilises the copper rod and helps to fix the sample in the cryofixation device.

In the interactions of Candida albicans with host cells, the cell wall of the yeast may play important roles in the adhesion of yeast cells to tissues. The outer cell wall of yeast (e.g. Saccharomyces cerevisiae, C. albicans) has been shown to consist of a dense network of radially projecting fibrils composed of mannoproteins that are known as fimbriae and which previously have required cryopreservation either by jet propane freezing or by plunge freezeing for their visualization. High pressure freezing provides an advantage over jet or plunge freezing in terms of the higher consistancey in the quality of freezing, and the minimization of formation of ice I with this method. Hohenberg et al reported a method utilizing cellulose capillary tubes to cryoimmobilize suspensions of microoganisms by high pressure freezing (HPF) and freeze substitution (FS), and herein, we describe an adaptation of this method by culturing microorganisms within the tubing to increase cell density prior to high pressure freezing and freeze substution.

In vertebrate somatic cells, sister kinetochores attach chromosomes to the mitotic spindle by capturing microtubules (Mts) emanating from opposite spindle poles. Kinetochores are also required for chromosome alignment at the spindle equator, stabilizing kinetochore Mts, cell cycle control of anaphase onset, and poleward migration of sister chromatids during anaphase. Thus kinetochores play a critical role in the distribution of genetic information to daughter cells during cell division. Understanding the molecular mechanisms behind kinetochore function requires knowing how the molecular components are arranged relative to each other, and relative to the kinetochore Mts. Currently, however, our knowledge of kinetochore composition is incomplete, and until recently our knowledge of kinetochore ultrastructure was limited to the familiar trilamellar model derived from conventionally fixed and dehydrated specimens(Figures 1a&b).

Highly fibrous structures, such as the kinetochore, are particularly vulnerable to distortions caused by the chemical fixation and dehydration methods used in conventional specimen preparations. Such distortions can be largely avoided by using high pressure freezing and freeze substitution (HPF/FS).

As the defining structure in eukaryote cells, the nuclear envelope is completely dismantled and reformed within an hour or so at each cell division (open mitosis). In yeast and some insect tissues, closed mitosis occurs, in which the nuclear envelope is maintained largely intact throughout chromosome separation. Use of cell free systems has allows us access to the mechanisms of cell division and NE dynamics in vitro by FEISEM (Field Emission In Lens Scanning Electron Microscopy. We have used demembranated Xenopus sperm heads as a source of DNA, which is incubated in an extract of Xenopus egg cytoplasm, where it becomes assembled into a normal nucleus with functional nuclear envelope (1-4). DNA replication proceeds under normal cell cycle controls, followed by an in vitro mitosis in suitable conditions. The cytoplasmic extract can be separated into membrane and soluble fractions that can be supplemented with, or depleted of, specific proteins. Inhibitors and other effectors can be .added to modulate both assembly and transport (5). Using the lectin WGA we have depleted Xenopus cytoplasmic extract of the major nucleoporins, CAN, Nup 98 and p62 and their associated proteins, whose removal effectively inhibits three aspects of nuclear formation, namely NPC formation, nuclear growth, and the reorganisation of the DNA in the depleted nuclei. Adding back these eluted nucleoporins restores normality with respect to nuclear growth, DNA reorganisation and NPC assembly. Current work involves purification of complexes containing these proteins by HPLC to allow add back of the complexes singly and in combination, to characterise their individual roles in NPC assembly, structure and transport (Figures 1,2,3).

High-pressure facilitates freezing of biological specimens: pressurizing to 2000 bar before cooling retards growth and reduces size of ice crystals in the specimen. Although the lung is 80% air and is generally difficult to freeze, we report here good cellular ultrastructure in high-pressure frozen lung when: 1) the lung is allowed to collapse (empty of air) prior to sampling, or 2) air-filled lung is allowed to compress during pressurization in the high-pressure freezer prior to freezing or 3) airspaces are filled with relatively incompressible fluorocarbon liquid.

Small Balb mice (28-30g) were anaesthetized and ventilated via endotracheal tube. The chest was opened and a 2mm diameter disk of lung at the thin anterior margin of the right lung was clamped between two aluminum tophats (200μm wells), sealing in air. In other experiments, the lung was allowed to collapse to 0 cm water transpulmonary pressure prior to clamping. In the third series of experiments, air was gradually replaced with fluorocarbon liquid (FC5312, 3M, Minneapolis MN) and the lung inflated with liquid to functional residual capacity. The top hat assembly containing the sealed lung was transferred to the high-pressure freezer (HPM 010, Bal-Tec, Balzers, Lichtenstein) where they were pressurized and frozen.