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Abstract

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED

Scanning Electron Microscope (SEM) is a powerful research tool, but since it requires high vacuum conditions, the wet materials and biological samples must undergo a complex preparation that limits the application of SEM on this kind of specimen and often causes the introduction of artifacts. The introduction of Environmental Scanning Electron Microscope (ESEM), working in gaseous atmosphere, represented a new perspective in biological research. Despite the fact that many biological applications have demonstrated the convenience of ESEM, the full potentialities of this technology are still under investigation. In this review, the exploration of the recent literature data confronted with the first results obtained in our experimental work suggest that ESEM represents an important extension of conventional scanning microscopy. © 2005 Wiley-Liss, Inc.

The aim of this study is to depict the current state of the art in the application of Environmental Scanning Electron Microscope (ESEM) in bio-medical research. Since its introduction, ESEM has represented a very attractive innovation for many researchers because it allows imagery of biological as well as material samples, in wet mode, with no need for the conventional Scanning Electron Microscope (SEM) preparation.

Our laboratory is involved in biomaterials and tissue-engineering research, a field in which the use and importance of ESEM have increased progressively during recent years (Manero et al., 2003; Stokes, 2003; McKinlay et al., 2004). The possibility of observing wet samples, as well as the fact that sample preparation is minimal, has resulted in shorter time scales and lower costs in microscopy. This was very important for us when dealing with samples from biopsies or prosthetic explants (Fig. 1, parts 1–4). Minimal sample manipulation has also reduced the possibility of introducing artifacts. We obtained good results studying cell adhesion and substrate-induced morphological features on PET, on fluorinated (C2F4) flat and nanostructured PET.

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Figure 1. ESEM micrograph of polymers and surgical explants. Part 1: PVA dehydratation process (4.90 Torr, 5°C, 12.5 kV). Part 2: Poly(ethylenetherephtalate) prosthesis explant (Marlex) (3.90 Torr, 5°C, 12.5 kV). Part 3: Parathyroid external adipose cells (4.20 Torr, 5°C, 10 kV). Part 4: Thyroide section showing phollicular structure (4.20 Torr, 5°C, 10 kV).

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Previously, we employed only the SEM technique to evaluate the cytomorphology and cytotoxicity of primary human fibroblasts and polyelectrolyte complexes (Rosso et al., 2004). Currently, our preliminary results also include a direct comparison of the morphology of 3t3 Swiss albino mouse fibroblasts as imaged with SEM and with ESEM. As the former uses higher magnification and more exact contours, the latter respects the three-dimensional structure of the cell and prevents the loss of the finer surface features. Based on our experience, the extra-flattened cell structure and extremely rough or smooth surfaces, even in the advanced stages of adhesion, are to be considered desiccation artifacts (Fig. 2, parts 5–8). This can be confirmed by their appearance in ESEM under defective pressure-temperature conditions or beyond the acceptable observation time. In contrast with this, the real potential of this instrument is still under discussion and in part unclear. Thus, we felt a comprehensive review was warranted.

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Figure 2. Cell morphology evaluation by SEM/ESEM. Part 5: SEM image of 3T3 Swiss Albino Mouse fibroblasts (20 kV). Parts 6–8: ESEM images of 3T3 Swiss Albino Mouse fibroblasts adhesion and proliferation on biomaterials (4.60 Torr, 5°C, 7 kV).

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Before comparing the two techniques, it will be useful to give a brief overview of SEM sample preparation and then of ESEM working principles.

BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED

Using SEM to visualize cells and biological tissues requires to apply several rigorous processing steps to dehydrate the sample and prevent it from charging when exposed to the imaging electron beam (Bozzola and Russel, 1998; Hayat, 2000).

As is well known, fixation is a technique that quickly arrests biological activity and stabilizes cellular components with minimal distortion of conformational and spatial relationships between cellular constituents. Normally, the first principle to consider before deciding on a fixation treatment is the difference between killing and fixing. Generally the term fixing includes both processes, which may occur at about the same time, but they are not one and the same process. Cells and tissues immersed in typical aldehyde fixatives can exhibit physiological and physical changes before cellular death occurs. In many cases, it may be necessary to kill cells with toxic fumes (osmium tetroxide) before immersing them in aldehyde fixatives to prevent structural change.

The most common reason for poor fixation is large specimen size. To minimize auto-lytic changes, the slice, ribbon, or 0.5/1 mm cube of tissue should be placed in the fixative promptly (Slayter and Slayter, 1992). Chemical fixation for electron microscopy prepares cells for the preservation of damage due to subsequent washing with aqueous solvents, dehydration with organic solvents such as ethanol or acetone, and imaging with high-energy electron beams in an electron microscope.

Materials prepared for scanning electron microscopy are not embedded with resins but are subjected to high pressure while immersed in solvents during critical point drying (CPD). An ideal fixative would transform the viscous colloidal protoplasm of a cell into cross-linked and stabilized cellular components. The spatial relationship between all organelles and cellular structures would not be altered, the cellular components would not be solubilized, and the biological activity of complex proteins such as antigens and enzymes would remain undiminished (Dykstra, 1993).

Good penetration speed, to a large extent, determines the success of a fixation procedure. Small fixative molecules (formaldehyde) penetrate more rapidly than larger ones (glutaraldehyde), but the larger molecules possess greater numbers of reactive sites and can thus cross-link and stabilize cellular components more thoroughly. The buffering system is also used typically to stabilize the pH of the tissue somewhere near physiological levels during fixation and to maintain near isotonic conditions. The nature of the specimen and the aim of the observation dictate the type of buffer and the pH that must be maintained during processing.

For structural studies, most cells fix well within a pH range from 7.0 to 7.4. Certain highly hydrated tissues fix better at a more alkaline pH (i.e., 8.0–8.4), whereas plant cells, nuclear material, and the fibrils of mitotic spindles fix better at a more acid pH (6.0–6.8) (Hayat, 1981). The fixation of biological samples for scanning electron microscopy (SEM) involves all the same principles for preserving specimen structural integrity for transmission electron microscopy (TEM), but often osmium tetroxide post fixation can be omitted, although samples that have charging problems leading to image distortion can frequently benefit from osmication.

Subsequent dehydration in an ethanol or acetone series followed by CPD, freeze-drying, or drying with one of the chemical techniques, produces a sample that can be introduced into the high vacuum system of the SEM. The CPD technique is used for samples of soft tissue that are hydrated. If the sample is already dry, CPD is often inappropriate. In such cases, the dried sample can be immediately mounted on SEM stubs, sputter coated, and examined with the SEM. A hydrated biological sample should be fixed and dehydrated as described for routine SEM preparation. CPD is based on the concept that at a certain temperature and pressure, the vapor and liquid phases of carbon dioxide become indistinguishable. Liquid CO2 from a siphon-tube tank is introduced into the chamber and used to replace the 100% ethanol in the specimen. After the ethanol has been totally replaced by the CO2, the CPD chamber is raised above the critical point. The temperature is kept above the critical point while the gaseous CO2 is vented from the chamber. The process is finished when the CPD is returned to atmospheric pressure. Form, dimension, and measure of the sample can change during CPD treatment, constituting the correct evaluation of the SEM observation study. After CPD, the specimen should be totally dry and is ready to be introduced into the vacuum system of the sputter coater and SEM (Dykstra, 1993; Hayat, 2000).

Sputter coating is a technique for deposing a metal coating on specimen surfaces to be examined with an SEM. The dried sample is attached to support stubs with a variety of materials (colloidal silver, colloidal carbon, double sided tape, or conductive carbon tape, among others) prior to coating with precious metals such as gold-palladium to ensure the electrical conductivity of the specimen surface. Less specimen heating is developed during sputter coating than with vacuum evaporation. In modern sputter coating, the temperature rise during evaporation increases to even less than 10°C and in many cases is even stronger for a very sensitive sample that is free of heat damage. Thus, even using a more simple procedure for SEM preparation, the sample requires a few hours before reaching the electron microscopy phase.

LIMITATIONS OF CONVENTIONAL SEM (CSEM)

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED

During the last 40 years, scanning electron microscopy (SEM) has increasingly represented a very important structural characterization for material scientists. Scanning the material surface with electron beam and detecting the emitted secondary or backscattered electrons, allows microscopists to see down to resolutions of 10 nm or less, giving them intricate details of the material's structure. However, the requirements of SEM, such as a high vacuum and the need for a coating film if an insulator is being analyzed, mean that certain types of materials have always proven difficult or impossible to image frankly. For example, the coating can obscure the fine surface detail on some specimens, although SEMs equipped with field emission guns have made such samples easier to image.

Another difficulty arises with wet and damp samples such as paints, inks, emulsions, and biological tissue: these materials prove particularly challenging for SEM. The high vacuum requirements in the chamber mean that long specimen preparation techniques are required to remove or fix the water before imaging, raising the risk of introducing artifacts, destroying the finer features, and leaching out the ions that could otherwise be analyzed by microanalysis (McKinlay et al., 2004).

INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED

These problems can now be overcome, thanks to the new environmental scanning electron microscope (ESEM) that permits the imaging of wet systems with no prior specimen preparation. Since the introduction of this instrument, many attempts have been made to perform SEM imaging of samples in a gaseous environment.

The modern environmental scanning electron microscopy (ESEM) was developed about 15 years ago, only after the introduction of the gaseous detector device (GDD) (Danilatos, 1988, 1990). The advantage to using the ESEM operating in wet mode is that it is not necessary to make non-conductive samples conductive. Material samples do not need to be desiccated and coated with gold–palladium, for example, and thus their original characteristics can be preserved for further testing or manipulation. The development of this instrument means that whole new classes of materials, previously undreamed of, can be imaged in their natural state. But the potential of ESEM is even greater than this. Because the sample environment can be dynamically altered, hydration and dehydration processes can be followed as they happen in the sample chamber.

In the ESEM instrument, a series of pressure-limiting apertures (PLAs) are placed down the column, across each of which a pressure differential is maintained. The PLAs are simply discs with small holes bored through the center. The principle at work here is that if there is a small enough pinhole between two different vacuum levels, and the difference between levels of vacuum is not that great, then the vacuum will not ‘diffuse’ from one level to another through the pinhole. So we can have a very good vacuum at the electron gun, at the top of the column where we need a very good vacuum, and in the specimen chamber at the mid-portion of the column we can have a relatively poor vacuum, without endangering the electron gun. Various gases as nitrous oxide, carbon dioxide, helium, argon, nitrogen, and water vapor can be introduced into the specimen chamber via a separate dedicated vacuum pump that can control the chamber pressure with great accuracy. Water vapor is the most common gas used in ESEM both for its amplifying efficiency (Fletcher et al., 1997) and useful thermodynamic properties. Many authors provided a detailed description of the cascade effect (Danilatos, 1988, 1990; Durkin and Shah, 1993) and its relations with image intensity and resolution (Thiel et al., 1997; Stokes et al., 2000).

The primary electron beam is very energetic, and it penetrates the water vapor with little apparent scatter, scanning across the surface of the sample. Secondary electrons are released from the surface of the sample, as they are in normal SEM, but they encounter water vapor molecules once they exit the surface. The water vapor molecules, when they are struck by the secondary electrons, produce secondary electrons themselves, which in turn produce secondary electrons from adjacent water vapor molecules. Thus the water vapor functions as a cascade amplifier, amplifying the original secondary electron signal from the sample. At the same time, the positive ions resulting from this cascade process are an essential feature in ESEM: ions drift towards the specimen surface and thereby help to compensate for negative charge build-up, hence insulating samples can be imaged without the need for a conductive coating.

Having a vapor in the sample chamber, the conventional detector used to pick up the secondary electrons emitted from the sample surface in standard SEM cannot be used for ESEM. The amplified secondary electron signal is collected by an on-axis GSED (gaseous secondary electrons detector), with its 300–550 V local positive charge that forms a fitted seal over the pole piece insert. The hole in the center of the GSED functions as the final aperture through which the primary electron beam passes, and its bore size determines how poor the vacuum can be in the specimen chamber. If the GSED has a 500-micron aperture in it, we can increase the pressure in the chamber to as high as 10 Torr; if the GSED has a 1-mm aperture in it, we can take the pressure in the chamber only as high as 5 Torr. And if we use the large-field detector (LFD) version of the GSED, we do not actually fit it over the pole piece insert, so the wet bullet itself provides the final aperture, and we can take the pressure in the chamber only as high as 1 Torr. Scattering of the electron beam between the gun and the sample is an obvious concern, given that there are many atoms/molecules of vapor in the ESEM chambers. The gap between the final PLA and the sample is kept reasonably small, 10 mm, and the pressure is kept quite modest, less than 10 Torr. Since vapor is tolerated in the sample chamber, ESEM makes it possible to carry out ‘wet imaging’ of samples. In order to view a wet sample, such as a colloidal dispersion, the atmosphere in the sample chamber must be carefully controlled at all stages. After the sample is placed in the chamber, air at atmospheric pressure must be replaced by water vapor at a few Torr. It is important to carry out the pump-down carefully so that premature dehydration of the dispersion does not occur. If done correctly, no accidental aggregation of the particles occurs, and a truly dispersed state will be imaged.

During imaging it is also important to ensure that neither water evaporation nor condensation occurs. This is achieved by using an atmosphere at the saturated vapor pressure (SVP) of water. However, this raises an additional difficulty in imaging samples at room temperature, because the SVP of water is relatively high at this temperature compared to the modest pressures of a few Torr that are acceptable in the sample chamber. The trick is to use a Peltier stage to drop the temperature down to 3–5°C so that SVP is easily maintained without undue loss of image quality and resolution.

ESEM BIOLOGICAL APPLICATIONS

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED

Given that SEM has been an instrument dedicated to material science since its birth, one of the main features of ESEM offers the opportunity to work on biological samples without complex and artefact generating manipulations. While few published works offer a direct comparison between these two different tools, many others offer a clear evaluation of the ESEM potential in biomedical research. One of the earlier works that reported the advantages of ESEM came from the study conducted by Collins et al. (1993) on microorganisms.

Biomaterials and tissue-engineering research is probably the field in which ESEM has found its fullest application, because it enables investigation of both cell and material surface morphology in hydrated conditions. In addition, performing observations with minimal specimen preparation is very attractive for studying interactions between mammalian cells and biomaterials that are developed for tissue engineering (Baguneid et al., 2004; Motta et al., 2004).

Rizzi et al. (2001) studied biodegradable polymer/hydroxyapatite (HA) composites as possible bone graft substitutes. Thin films of polymer/HA composites were produced using poly (ε-caprolactone) (PCL) and poly (L-lactic acid) (PLA), and the initial attachment of primary human osteoblasts (HOBs) was assessed to investigate the biocompatibility of the materials. Using ESEM, the authors investigated surface morphology of the polymer and cell morphology on the surfaces of samples after 90 min, 4, and 24 h of cell culture. Using Alamar blue and DNA assays, cell activity and viability were assessed after 24 h of cell culture. In the first 4 h of culture, the cells were spread to a higher degree on exposed HA regions of the composites and on PLA than they were on PCL. After 24 h, the cells were spread equally on all the samples. Based on cell morphology and activity assays, they concluded that a polymer surface exhibiting “point exposure” of HA appeared to provide a novel and favorable substrate for primary cell attachment.

In many studies, especially in the earlier years, the procedures followed for sample preparation, even if ESEM was used, involved fixation and alcohol series dehydration.

Schmidt et al. (1997) studied the stimulating effect of a conducting polymer PP (oxidized polypyrrole) on neurite outgrowth. Using optical and electron microscopy, the authors demonstrated a conspicuous neurite length increase compared with other polymers such as PLA, poly (L-lactic acid), and PLGA, poly (lactic acid-co-glycolic acid), and concluded that PP is a suitable material for in vitro nerve cell culture and in vivo implantation. ESEM was used on fixed, alcohol-dehydrated and overnight dried cells but no comparison with wet-mode images of the same fixed or unfixed cells was made to exclude morphological changes caused by the preparation.

In this respect, the work of Stokes et al. (2003) is very interesting. They imaged HOB-like cells using ESEM. The cells were hydrated, unfrozen, and uncoated. Specimens were cooled to 3°C and imaged in water vapor, with partial pressures varying from saturated conditions to a humidity of approximately 50%, relative to pure water. The ESEM images show the presence of cell nuclei, nucleoli, and cytoplasmic membranes. Comparisons between chemically fixed and unfixed specimens show that cell morphologies are similar in both cases. These results are compared with a fixed, dried, carbon-coated specimen. Thermodynamic and kinetic arguments are used to show that humidity significantly lower than 100% corresponds to metastable states suitable for stabilizing hydrated biological tissues and cells. This argument has been extensively discussed by Stokes (2003) in a brilliant review work about the physical principles involved in ESEM technology. Real aqueous phases, such as those found in the interiors of mammalian cells, are neither dilute nor ideal: macromolecules such as proteins and polysaccharides interact strongly with water molecules (Ellis, 2001). Tai and Tang (2001) empirically noted that, in biological samples, lowering of the equilibrium vapor pressure at humidities of ca. 90% occurs. Additional lowering of the chamber pressure (20–25%) could be introduced due to kinetic properties of the physiological solutions, thus specimens tolerate slowly dehydrating conditions in a finite time (Stokes, 2003). Our experimental results confirm this statement, as we achieved the best conditions between 60% relative humidity, for bulk samples such as surgical explants (see Fig. 1, parts 2–4), and at 70% r.h. for cell monolayers (see Fig. 2, parts 6–8).

On the other hand, some other researchers use ESEM but adopt a preparation protocol similar to that used for conventional SEM, comprising even CPD but not sputter coating with metals. For example, in immuno-gold staining experiments used to locate membrane receptors and to characterize cell types, metal coating represents an obstacle, making it advantageous to employ ESEM even if partially exploiting the instrument potentialities. This approach was used for phenotypical characterization of the cells that are responsible for regression of tunica vasculosa lentis (TVL), a transient vascular network surrounding the developing lens in pre-natal humans and in post-natal rodents. In fact, Djano et al. (1999) and McMenamin et al. (2002) utilized a novel combination of silver-enhanced immuno-gold staining and ESEM. The cells surrounding the developing lens that are postulated to play a role in regression of the TVL have the morphologic and immuno-phenotypic characteristics (ED1+, ED2+, MHC class II, and CD11b/18) of resident tissue macrophages similar to those previously identified in the adult rodent uveal tract and the vitreous (hyalocytes). This phenotype differs from that of dendritic cells and microglia; however, it is postulated that lens-associated macrophages are ideally located to act as a source of retinal microglia after completion of their role in TVL regression.

Secondary electron (SE, generated by inelastic collisions) imaging is the main method for examining structural characteristics of specimens, while back-scattered electrons (BSE, generated by elastic collisions) imaging offers less resolution power but reveals compositional differences. Because of the non-conductive nature of biological tissues, ESEM imaging of colloidal gold stained tissue offers a very strong contrast with SE-BSE component mixed signal, thus making it very simple to identify the immunopositive cells in the microarchitecture of the tissue.

Previously published works on other tissues employed the colloidal-gold staining technique in conjunction with conventional SEM (Goode and Maugel, 1987; de Harven et al., 1990). In high vacuum microscopy with chromium-coated cells, SE mode failed to identify gold labels while the BSE signal, because of the great difference of atomic number between chromium and gold, produced a contrast sufficient to correlate the distribution of gold particles to the morphological details of the cell surface (Herter et al., 1993). Coating with carbon offers less contrast than chromium, so it maximizes gold evidence but hides cell surface details. When metal coating is absent, the sharpness of the images is slightly different, but nonetheless it is clear that ESEM offers great advantages when using this kind of technique. In other published studies, condrocites and epithelial cells have been used to visualize and characterize the hyaluronian coat on the cell surface. Conventional SEM studies failed to reveal cell-associated material outside the plasma membrane. However, ESEM revealed a 4.5-μm thick halo, removed after hyaluronidase treatment, which can likely be removed by CPD. Furthermore, uranyl-acetate staining favors visualization of the gel borders, because whilst osmium tetroxide is well incorporated in the plasma membrane, heavy uranyl ions Uo22+ bind to the negatively charged hyaluronian. The main conclusion drawn here is that this gel coating is responsible for the first phase of the cell adhesion (Cohen et al., 2003). In some cases, some features considered normal for certain biological tissues were discovered by ESEM imaging to be histological artifacts. For example, Suso et al. (2003) observed that articular cartilage had aspects similar to those described previously with CSEM, except for the absence of splits and fractures that were then considered to be artifacts derived from the preparation procedures.

LIMITATIONS OF ESEM

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED

Besides the great number of biomedical samples that have been analyzed using ESEM, many authors have reported their difficulties in dealing with wet samples. Mestres et al. (2003) pointed out the frequent occurrence of contrast effect that impedes the ideal imaging of cells and tissues. The condensed water layer on the sample can cause low contrast and limited visualization of small surface details, such as microvilli. In addition, especially in bulk tissue samples, alternate dark and light areas, corresponding to different charge status, can confuse the sample topography. In many cases, increasing the tilt angle results in a much better image definition, resolving the first problem, but at the same time increases the second one, so that it becomes difficult to take images at low magnification.

Obviously tilt is a very critical condition when working with wet samples, because of liquid displacement. Similar conclusions were drawn by Martinez-Alvarez et al. (2000) in their work on palatal fusion in mouse embryonic development. The authors observed the presence of bulging epithelial cells on the medial edge epithelium prior to palatal fusion. These cells had also been observed in classic histological sections but never in past SEM studies, probably because the dehydration caused them to collapse. On the other hand, cell filopodia and lamellipodia previously seen with SEM were no longer distinguishable. Opposite conclusions were drawn by McKinlay et al. (2004), when they compared HOB morphology during the adhesion process on oxidized titanium surfaces as obtained by conventional SEM and environmental SEM. They found that in wet samples, there were more filopodia in both cell-cell and cell-substrate contact compared to dehydrated samples. The majority of these was of the cell-cell type, while the few on the surface appeared to be very thin.

A first explanation for these contrasting results could be in the nature of the sample, as filopodia in a bulk tissue sample are all cell-cell and thus suffer less severe consequences when they are dehydrated. On the other hand, filopodia on a material surface will be lost during processing for conventional SEM, if they are not strongly adhering. Second, as we can testify, ESEM cannot produce an image through a significant thickness of water, and the ability to view this kind of cell structures in wet mode is strictly connected to the water content: if the water film is not removed before the observation, by whatever means, it will surely mask small filopodia and lamellipodia.

It is noteworthy that in most of the works mentioned, the complications potentially arising from the presence of water in the sample were simplified by submitting the samples to slight fixation and dehydration using the alcohol series, avoiding only CPD and metal coating. This means that observation of biological material in a living state is a task that has not yet been fully achieved. In this respect, it is important to consider data suggesting that merely fixing the living cells does not significantly affect their morphology (Stokes et al., 2003). It is important to remember that even a slight fixing affects the cell structure, introducing relevant changes to its internal ultra-structure; but those changes are found to be determinant if the cell is observed with TEM, while they are revealed in a minimum way by scanning analysis with SEM/ESEM. Therefore, fixing can be accepted as long as it is likely to increase the cells resistance under observation conditions. Even when high relative humidity and low acceleration voltages are used, these conditions still represent stress for the biological material, hence it is generally accepted that a single ESEM sample cannot withstand a time span longer than half an hour. For the same reason, it is possible to view a sample in ESEM only once, in contrast with multiple observations of samples in SEM. A certain limitation is the field of view due to size of the PLA inserted on the pole piece. Since we use the 500-μm PLA, the lowest magnification allowed is about 200×, thereby prohibiting the complete visualization of the sample's different parts. Under these conditions, it may often cost precious time to understand the general sample morphology and to identify the regions of interest. A possible approach to this problem could be to increase the working distance up to 15–18 mm, but it is clear that if the gap between the sample and the detector is too large, beam scattering increases and the signal becomes too weak. Successful imaging in these conditions should require less gas pressure inside the chamber or higher accelerating voltage applied; both being very difficult when working with wet and uncoated samples. Taylor and Wight (1996) reported a new method for low magnification in ESEM capable of reaching about 30×, significantly better than the original 200× imaging limit. This was achieved by using an additional aperture to limit the pressure at a point where it will not block the electron beam, and a larger open plate for the combination final aperture/secondary electron signal collection surface that also does not block the electron beam significantly. In this sense, important technical innovations, such as the reverse flow PLA and the radio frequency gas detection device, have been introduced but not yet incorporated in the commercial ESEM (Danilatos, 2000a,b).

ESEM has offered significant advantages over the last few years in biological and medical research; nonetheless it seems quite clear that the methods used in this application are far from being standardized, as experimentation is still on going. This is perhaps especially true of its application to studying wet systems. In part, it is true that the optimal working conditions are always sample specific and thus will probably continue to be determined empirically. On the other hand, ESEM still suffers from some suspicion by the researchers community that can be attributed to an early failure to appreciate the careful control of chamber conditions required at all times. With improving awareness of this and other drawbacks, ESEM should take on a key role in an increasing number of materials and cell biology laboratories, expanding its range of applications in industries and production processes.

Furthermore, based both on the published data available and on our laboratory experience, we believe that at the present moment, ESEM could not definitively replace SEM. More likely they appear to be complementary, as the former provides a real image of the sample, while the latter is still vital in resolving the finer structures in detail.

LITERATURE CITED

  1. Top of page
  2. Abstract
  3. BIOLOGICAL SAMPLE FOR SCANNING ELECTRON MICROSCOPY (SEM)
  4. LIMITATIONS OF CONVENTIONAL SEM (CSEM)
  5. INTRODUCING THE USE OF ENVIRONMENTAL SCANNING ELECTRON MICROSCOPE (ESEM)
  6. ESEM BIOLOGICAL APPLICATIONS
  7. LIMITATIONS OF ESEM
  8. LITERATURE CITED
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