TITLE: No Title
AUTHORS: 

[DESCRIPTION]
This protocol is from:<br /><br />Hans-W. Ackermann and Mikal Heldal. 2010. Chapter 18: Basic electron microscopy of aquatic viruses. Manual of Aquatic Viral Ecology. Waco, TX:American Society of Limnology and Oceanography. doi:10.4319/mave.2010.978-0-9845591-0-7<br /><br /> Please see the <a href="http://www.aslo.org/books/mave/MAVE_182.pdf" target="_blank">full manuscript</a> for additional details.

[GUIDELINES]
<em><strong>General<br /><br /></strong></em>Virological electron microscopy has essentially descriptive purposes, namely the identification and characterization of free viruses. Of the many biological applications of electron microscopy (EM), only transmission electron microscopy (TEM) is of significant importance in virology. Its other applications, e.g., scanning electron microscopy (SEM), cryo-EM with or without three-dimensional image reconstruction, immuno-EM, sectioning of infected cells, shadowing of individual particles, or enzymatic virus digestion on grids, have limited applications and are sometimes practiced in a few specialized laboratories only. In aquatic virology, sectioning is still important because it permits identification of intracellular assembly sites of novel viruses and the detection, in mixed host systems, of which host cell produces viruses. An excellent overview of sectioning techniques may be found in a recent book (Kuo 2007). The direct harvesting of particles onto grids for total viruslike counts was and is of great importance for the understanding of viral activity in aquatic ecosystems (Bergh et al. 1989).<br /><br />Negative staining was introduced in 1959 (Brenner and Horne) and revolutionized virology in generating virus images of unprecedented clarity. It is indispensable for structural studies and virus identification, but is also used for particle counts (below). The principle of negative staining is to mix the particles to be examined with an electron-dense solution of a metal salt of high molecular weight and small molecular size. On an electron microscopic grid, the particles then appear white or gray on a dark background. Many variant procedures have been devised. A detailed description of stains and procedures may be found elsewhere (Ackermann and Dubow 1987; Hayat and Miller 1990; Kay 1965).<br /><br /><em><strong>Specific Techniques</strong></em><br /><br /><strong>a.)</strong> <strong>Purification</strong><br /><br />For high-resolution images, samples should be sterile and free from bacteria and other large particles. This is achieved by filtration through membrane filters of 0.22–0.45 µm pore size. Except for particle counts on EM grids (below), purification is a must because proteins and slime interfere with staining and resolution and their amounts must be reduced to acceptable levels. The most convenient procedure is washing in a buffer, e.g., 0.1 M ammonium acetate (pH 7), in an ultracentrifuge with a fixed-angle rotor (25,000g or 25,000 rpm for 1 h). Centrifugation in swinging-bucket rotors requires higher g forces and centrifugation times (e.g., 70,000–80,000g for 60–90 min). Clearly, fixed-angle rotors are advantageous as they permit the use of relatively inexpensive medium-size centrifuges. Phosphate buffer is acceptable, but physiological saline is not because it leads to the formation of salt crystals on grids. Alternatively, viruses may be purified by CsCl or sucrose gradient centrifugation followed by dialysis.<br /><br /><strong>b.) Particle counts on EM grids</strong><br /><br />Viruses in lysates are counted by TEM after depositing or vaporizing a fixed volume of virus suspension on the grid (Figs. 1 and 2). Viruslike particles (VLPs) in water can be counted by TEM, flow cytometry, or epifluorescence microscopy (EFM). The direct harvesting of particles on TEM grids ensures that bacteria, algae, and other particles can be observed simultaneously. These preparations can also be used for estimating bacterial size distribution and the frequency of infected cells and dividing bacteria, and to visualize lysed bacteria.<br /><br />The technique requires an ultracentrifuge with a swingingbucket rotor and centrifugation tubes (Polyallomer®) with flat bottoms, created by introducing Epoxy resin into a conical tube. The resin is left to harden at 60°C overnight, and an electron microscopic grid is deposited on the flat resin surface. Samples are centrifuged at 80,000g for 90 min (Borsheim et al. 1990). Tubes fitting Beckman SW-41 rotors allow particle harvesting from a 60-mm water column. Use cellulose nitrate filters fitting the diameter of the tubes, add a small piece of double-sided tape, and position the grids at the edge of the tape in the middle of the tube. Fill the tubes with sample and then add the filter paper with the grids attached to avoid air bubbles being trapped under the grids. The tubes should always be filled up to 2–3 mm from the top. If particle density is high (10<sup>7</sup>–10<sup>8</sup> cells mL<sup>–1</sup>), the sample volume can be reduced. This can be done by reducing the height of tubes. The calculation of total virus contents is described by Bratbak and Heldal (1993).<br /><br />Montanié et al. (2002) presented a different approach to total virus counts by TEM. A purification step is followed by pelleting viruses from the supernatant by means of an ultracentrifuge. The particles are then transferred by capillarity to grids with carbon–collodium membranes. The recovery of viruses is reported to be 71% to 79%. The filtration technique uses membrane filters of 0.2 µm pore size for virus concentration. Virus particles on the filter are then resuspended in water and the suspension is examined in the EM (Torrella and Morita 1979). Again, this technique is inexact because particles smaller than 0.2 µm may be lost and also because of uncertainties introduced by resuspension. The technique of Ewert and Paynter (1980) involves sedimentation of viruses by means of a special rotor onto agar blocks and the preparation of pseudoreplicas, which are then stained and examined in the EM. The technique is quite complicated and has, to our knowledge, not been used by anybody else. We have no personal experience with it.<br /><br /><strong>c.) Special Applications</strong><br /><br />TEM has been used to visualize viruses within infected, unstained cells (Fig. 1). The aim is to determine burst sizes, lytic cycle lengths, percentages of bacteria in a stage of lytic infection, and mortality rates. The subject has been aptly reviewed by Weinbauer (2004).<br /><br /><strong>d.) Stains</strong><br /><br />The stains used are tungstates (potassium, sodium, lithium, silicotungstates), uranyl salts (acetate, formate, magnesium acetate, nitrate, oxalate), molybdic acid, and ammonium or vanadium molybdates. The most common stains are aqueous solutions of sodium or potassium phosphotungstate (PT; usually 2% and pH 7.2) and uranyl acetate (UA; 1%–2%, pH 4–4.5). Stains are prepared by dissolving phosphotungstic acid or uranyl acetate in distilled water and adjusting the pH with KOH or NaOH. They can be kept in stoppered bottles at 4°C for 2 years. Uranyl salts have a strong affinity to doublestranded DNA (Huxley and Zubay 1961), are toxic and radioactive, and should be handled with care. PT produces negative staining only. UA produces negative staining and, because of its affinity to dsDNA, will stain positively in black any particles with sufficient dsDNA content, for example, phage heads or herpesvirus capsids.<br /><br /><strong>e.) Staining</strong><br /><br />The required supplies include pointed pipettes and tweezers, strips of filter paper, and common electron microscopic grids (Athene-type, copper or steel, 100–400 mesh or square holes). Grids must carry a Formvar or collodium film stabilized by a carbon layer. They may be prepared by the electron microscopist or purchased ready-made. After 1 month, grids tend to be hydrophobic and reject particles. Grids can be made hydrophilic again by glow discharge in a carbon evaporator or by rinsing with a wetting agent (Alcian blue, bacitracin, poly-L-lysine, serum albumin, sucrose) (Gentile and Gelderblom 2005; Gregory and Pirie 1973). One of the authors (H.-W. Ackermann) sometimes uses UA or PT supplemented with 2 drops/mL of a 0.5% bacitracin solution.<br /><br />The staining procedure is extremely simple and nearinstantaneous. A drop of virus suspension is deposited on a grid, left 30 s for adsorption of viruses, and then a drop of stain is added, and the liquid is withdrawn with filter paper. Positive staining takes a little longer, but is complete after 1 min. Preparations prepared for TEM by centrifugation are stained with 1%–2% UA in water for 2–3 min, gently washed in distilled water, and air-dried. This will give a positive staining which facilitates total virus counts and an estimation of the number of infected cells and burst sizes (see Weinbauer 2004 and references therein). Positively stained viruses appear as electron-dense and deep black. The presence of impurities may create a high background noise.<br /><br /><strong>f.) Electron microscope and Imaging</strong><br /><br />Transmission electron microscopes (TEMs) are operated at 60–80 kV. To limit contamination by hydrocarbon molecules present in the column and hence loss of resolution, the object stage may be cooled with liquid nitrogen. This is not necessary for experienced users. Three types of instruments are presently in use.<br /><br />    1. Conventional TEMs - Lens columns, astigmatism, focusing, and magnification are adjusted by the user. Images are recorded on films or plates and are as much a product of the darkroom as of the EM. They require a darkroom equipped with a enlarger of professional quality. Mercury lamps and exhausts are not necessary, but small table-top enlargers as used by amateur photographers are inadequate. Supplies include fine-grain films, high-speed developers, polycontrast paper, and graded gelatin filters for enhancing contrast. Image contrast generally depends on developers, and papers and can be dramatically improved by filters. It is unfortunate that some photographical supplies, especially papers, have become difficult to find or have even disappeared from the market following the introduction of digital cameras.<br /><br />    2. Automated TEMs- Images are recorded as above on film or plates, but focusing is automated, columns are largely selfcontained, and resolution and magnification can no longer (or only with great difficulty) be adjusted by the electron microscopist. Their operation has thus been considerably simplified, but the operator has little control over the instrument and depends largely on the installer for alignment, resolution, and magnification control. Most of these instruments are now equipped with digital cameras and are commonly called “digital electron microscopes.”<br /><br />    3. Digital (charged-couple devide [CCD]) Cameras- The cameras are not made by EM manufacturers and must be purchased separately. Images are recorded electronically and are easy to store, retrieve, copy, and exchange. A darkroom is no longer necessary, but images must be printed on special paper and by means of special printers.<br /><br />If no TEM is available, it is possible to use a scanning electron microscope (SEM) equipped with a field emission gun (FESEM) and scanning–transmission electron microscopy (STEM) detector. Most of these instruments can be run in STEM mode and used as ordinary TEMs. Preparation of samples and staining are as described for TEM. FESEM instruments are normally operated at 10–40 kV, but for magnifications up to ×150,000, reasonably good total counts of viruslike particles are obtained. One example is the JEOL instrument JSM-7400F (cold emitter), with which unfixed and unstained marine viruses were observed in STEM mode at a magnification of ×100,000 (M. Heldal, unpubl. data). <br /><br />The other advantage of FESEMs is the possibility of obtaining high-resolution images in scanning mode, even at low accelerating voltages (1.5–2.5 kV). In such cases, the evaporation of gold/palladium on the material may give a coarse metal surface, and evaporation of iridium should be preferred. As an example, a Zeiss SUPRA 55 VP instrument (FESEM) equipped with an in-lens detector can be run at enlargements of up to ×500,000. Altogether, the new-generation SEMs (FESEMs) may be a good choice for providing good resolution microscopy for both TEM and SEM preparations. A drawback is the time-consuming setup of the STEM detector.<br /><br /><strong>g.) Magnification Control</strong><br /><br />High magnification (×250,000 or more) is controlled by means of grids with beef liver catalase crystals (Luftig 1967) or T4 bacteriophage tails. Diffraction grating replicas are suitable for low magnification only (below ×30,000). Calibration grids can be prepared in advance and kept for at least 1 month. Catalase crystals have parallel lines with 8.8-nm periodicity, can be used after PT and UA staining, and are commercially available. At a magnification of ×300,000, 20 lines correspond to 5.2 cm. T4 phage tails are 114 nm long, which corresponds to 34 mm at the same magnification. In conventional electron microscopes, magnification should be monitored often because of possible fluctuations in the local electricity supply. This is easily done by taking a picture of a calibration grid every 20–50 photographs and adjusting the darkroom enlarger. Digital electron microscopes and cameras are calibrated by the supplier or installer. This is a difficult procedure; it is doubtful whether these adjustments are all done by competent operators and it is so far unclear how stable digital cameras are over long periods.<br /><br /><em><strong>Assessment</strong></em><br /><br /><strong>a.) Purification</strong><br /><br />Centrifugation does not eliminate bacterial debris (cell wall fragments, flagella, pili, capsule material), but this is not a real problem as these particles do not interfere with resolution and it is always possible to find areas without them. In addition, almost all centrifuged preparations contain phage debris (e.g., isolated heads or tails) and abnormal or defective particles (polyheads or polytails). Using a fixedangle rotor, a force of 25,000g is amply sufficient for tailed phages, the largest of which sediment at 6000g for 1 h. It is just sufficient for very small viruses, e.g., microviruses or filamentous phages, which require 30,000g for 1 h for complete sedimentation (Ackermann and Dubow 1987). The technique described above is akin to centrifugation in the Beckman Coulter Airfuge which, however, uses extremely small volumes (180–240 µL) and is thus unsuitable for environmental studies. Density gradient purification is not mandatory for environmental studies. Such preparations can be very rich and pure, but some are full of crystals and impurities. The technique critically depends on adequate dialysis, demands large amounts of material, and can be recommended only if the performing scientist is sure of his or her skills.<br /><br /><strong>b.) Particle counts &amp; Special Applications</strong><br /><br />Counting of viruses in lysates can be difficult because viruses often do not disperse well or adhere poorly to the grid. For example, phages may aggregate with each other or cellular debris and even separate into areas of intact and damaged particles. Both problems can be overcome by washing grids with a solution of poly-L-lysine (Müller et al. 1980). Visualizing viruses within unstained infected bacteria is problematic because of bacterial inclusion bodies, notably carboxysomes (Shively et al. 1973) and polyhydroxybutyrate (Tian et al. 2005). Carboxysomes, which occur among others in the important cyanobacterium <em>Synechococcus</em>, resemble phycodnaviruses in diameter and hexagonal shape, but are recognizable by their variable size and distinctive capsomerlike structures.<br /><br /><strong>c.) Stains &amp; Staining</strong><br /><br />Phosphotungstate (Table 1) is a neutral stain and produces negative staining only. Artifacts are few and results are predictable and constant. Full phage heads often appear rounded in PT; empty heads appear as membranes or are filled by stain and then resemble black vesicles. Ammonium molybdate produces the same type of staining as PT and has no advantage over it.<br /><br />UA is an acid stain and causes both negative and positive staining. It is unpredictable and there is no way to produce positive or negative staining consistently and reliably. Both types of staining are usually present on the same grid and even on adjacent parts of the same grid hole (Fig. 3). However, proteinic impurities seem to facilitate negative staining. Negatively stained phage heads are white or grayish when full, but appear as thick membranes when empty. Positively stained viruses (or phage heads) are deep black, indicating the presence of dsDNA. They can easily be detected at low magnification, which is advantageous in environmental studies (see Borsheim et al. 1990). Positively stained phage heads are invariably shrunken by 10% to 15% (Figs. 4 and 5), and their dimensions are useless for virus identification and description. UA produces positive staining with large dsDNA-containing particles and not with ssRNA-containing phages, filamentous phages, or dsDNA viruses with low DNA content such as marine phage PM2. Consequently, these viruses cannot be detected by positive staining. In addition, UA causes swelling of purely proteinic structures such as phage tails and empty capsids. UA produces excellent contrast and is particularly suitable for resolution of tail striations. Negatively stained heads are more angular and presumably better preserved than in PT. UA spreads more evenly than PT, but tends to crystallize on grids.<br /><br />Both PT and UA form glassy films around particles and act as fixatives. PT-stained preparations are stable for a month without loss of detail. UA-stained grids keep easily for 10 years in grid boxes at room temperature. One of the authors (H.-W. Ackermann) has several grids that are 30 years old and still observable. Because PT and UA are complementary, investigators should always use both of them. Fixation, for example with 2.5% glutaraldehyde, may be necessary for marine phages that cannot be processed immediately and have to travel weeks before examination (Cochlan et al. 1993). The stability of glutaraldehyde- or formaldehyde-fixed material seems to be highly variable. Bacteria (Gundersen et al. 1996) and some viruses have a variable loss rate in fixed samples, whereas the formaldehyde-fixed Emiliania huxleyi virus showed reasonably high stability for 2–3 years under storage in the dark (Bratbak et al. 1993). As a rule, we recommend harvesting and preparation on grids of fresh material without any fixation.<br /><br /><strong>d.) Electron microscopes and imaging</strong><br /><br />The electron-optical resolution of EMs has not improved since the 1960s; all technical improvements made since concern ease of operation, automated focusing, digitalization, and gadgetry. The introduction of digital cameras (and hence digital electron microscopes) has not improved image quality, speed or ease of manipulations, or image recording. On the contrary, it has generated a flurry of gray, contrastless pictures of poor resolution which may irreverently be called “gray pulp.” Even company prospects show similar gray pictures. Neither prospects nor instruction manuals provide guidance for contrast improvement. It appears that the various EM and camera companies marketed their products in haste and without knowing how to operate them. The relative failure of digital electron microscopes is largely due to lack of information. Fortunately, image contrast can be greatly improved by manipulation of the camera software, particularly of the histogram governing luminescence (gamma), brightness, and intensity values (Tiekotter and Ackermann 2009). These parameters can be adjusted separately for selected areas, and excellent, high-contrast pictures can be so obtained. A major drawback of CCD cameras is the high price of cameras ($50,000 to $300,000) and printers ($5,000–10,000). This compares poorly with $2,000 for a Philips EM300 conventional camera and $10 for photographic trays and puts digital EM out of reach of most laboratories and institutes.<br /><br /><strong>e.) Magnification control</strong><br /><br />The specifications of manufacturers cannot be trusted, as they depend on lens setting and local electricity supply. Conventional and digital EMs suffer from the same problem and both must be controlled by means of test specimens. Unfortunately, both catalase crystals and T4 tails have their problems. Catalase crystals may be deformed and it can be difficult to find enough parallel lines of subunits for accurate measurements. In addition, the crystals are soluble in water and tend to dissolve into subunits. T4 tails stained with UA (but not PT) may be stretched beyond their normal length. On the other hand, T4 phages are easy to prepare in a microbiological laboratory and much easier to use than catalase crystals. It remains to see how stable and trustworthy digital cameras are over long periods of operation. Unfortunately, to our knowledge, the adjustment of digital EMs and cameras is lengthy and difficult and may take 2–3 h. It is essentially out of reach for investigators, who depend for adjustments on manufacturers and their technicians.<br /><br /><strong>f.) Measurements</strong><br /><br />Virus particles on conventional photographic images must be measured after enlargement on prints and never on films, plates, or screens. The reason is that measurements of small particles at low magnification are difficult per se and cannot be accurately extrapolated to high magnification. Digitized particles can be measured directly on the monitor screen. Long flexible phage tails may be measured with special flexible rulers. Investigators should be aware of artifacts, should not measure positively stained particles, and should measure at least 10 particles to obtain statistically meaningful data at sufficiently high magnifications (×250,000 or higher) to minimize measurement errors. Isometric viruses or phage heads should be measured between opposite apices. The literature sometimes includes mentions of phage heads of 30 nm in diameter. This puts tailed phages into the vicinity of polioviruses and is clearly an error. The smallest tailed phages known, podoviruses of the φ29 type (Fauquet et al. 2005), have head diameters of 40 nm. All others have heads of 50–60 nm in diameter or larger.<br /><br /><strong>g.) Virus detection and identification</strong><br /><br />Preparations enriched by incubation with host bacteria may reveal a wide variety of phages or phage-like particles (Rachel et al. 2002). Virus detection is both simple and difficult. It is easy because tailed phages, the mainstay of water samples, are instantly recognizable and differentiated in the EM. They belong to three clear-cut families, <em>Myoviridae</em>, <em>Siphoviridae</em>, and <em>Podoviridae</em> (Ackermann 2007; Ackermann and DuBow 1987; Fauquet et al. 2005). Members of the <em>Myoviridae</em> family are characterized by contractile tails like phage T4. Tails are generally thick, rigid, and constituted by a tail tube and a sheath separated from the head by a constriction or “neck.” Tail sheaths may be contracted. <em>Siphoviridae</em> have long, noncontractile, often very flexible tails like phage lambda. <em>Podoviridae</em>, similar to coliphage T7, have short tails of about 10 nm in length. Positively stained podoviruses may appear as headless tails and be difficult or impossible to identify. All tailed phage families have members with isometric or elongated heads. Empty capsids and headless tails will not elicit positive staining and remain invisible.<br /><br />The detection of isometric, filamentous, or pleomorphic viruses is intrinsically difficult. These viruses seem to be rare in nature, constituting only 4% of prokaryote virus isolates (Ackermann 2007). Further, ssDNA, ssRNA, and small dsRNA viruses do not stain positively. In crude, nonenriched samples, they can be diagnosed (a) after negative staining only and if they (b) occur in large numbers of identical particles, (c) present regular structures such as capsomers, and/or (d) have dimensions corresponding to those of known viruses. The identification of enveloped viruses without inner capsids, such as paramyxoviruses, is difficult or even impossible. Indeed, no pleomorphic virus has ever been identified by simple electron microscopic examination of environmental samples. Even the visualization of spikes on membranous particles is no help, as they also occur on mammalian cell membranes. The detection of filamentous phages is particularly difficult because these viruses are thin, break easily, and may be confounded with pili, flagella, or slime filaments. Algal viruses are isometric or filamentous (Van Etten et al. 1991). Their presence may be suspected if particles are isometric, at least 100 nm in diameter (Fauquet at al. 2005), and show capsomers and transverse edges. Filamentous algal viruses are known in the brown alga, <em>Chara corallina</em> (Skotnicki et al. 1976), but should be diagnosed only if they show transverse striations. Pitfalls are many; for example, one of the authors (H.W. Ackermann) once misidentified the mastigoneme filaments of <em>Ochromonas</em> algae (Bouck 1971) as possible filamentous viruses. In conclusion, isometric, filamentous, or pleomorphic viruses should be diagnosed only after their concentration in a density gradient or their propagation as pure cultures.<br /><br />For a definitive diagnosis and identification with known viruses, high-magnification (×250,000 to ×300,000) images and exact dimensions of negatively stained viruses are required. Images of positively stained viruses are unsuitable for this purpose and their dimensions are always inexact (see above, 3c; Figs. 4, 5).<br /><br /><strong>h.) Artifacts</strong><br /><br />Damaged particles and bacterial debris are frequent in phage lysates and environmental samples. For example, tailed phages are quite frequently decapitated, simulating tailless viruses or short-tailed phages. The frequency of tail loss depends on the phage and, possibly, storage conditions. Some phages are stable over years and some, such as coliphage Mu, fall into pieces within weeks. The frequency of damaged particles increases with repeated freezing and thawing. This should be considered if samples are taken during a maritime expedition, frozen, and processed weeks later. Bacterial debris include round particles derived from cell walls or cytoplasmic membranes, pili, flagella, and capsule-related mucus. Round particles (Fig. 6), often of the size of normal phage heads and thick-shelled, are particularly frequent in lysates of <em>Pseudomonas</em> phages, but also occur in those of <em>Vibrio</em> and <em>Synechococcus</em>. They may be misdiagnosed by inexperienced observers as enveloped viruses. We have even seen round cell debris lodged within other spherical cellular vesicles, simulating enveloped viruses (Fig. 6c) or <em>Tectiviridae</em>. All these debris are likely to adsorb DNA. Unless treated with DNase, they are bound to mimic DNA phages in epifluorescence microscopy. Slime, produced for example by xanthan-producing <em>Xanthomonas</em> bacteria, is likely to occur in freshwater. When centrifuged, slime blobs may become elongated and then masquerade as filamentous viruses (Fig. 7) and even tailed phages. Short flagella fragments often resemble contractile phage tails, and pili are almost impossible to distinguish from filamentous viruses of the <em>Inoviridae</em> family. Further, salts may obscure phage structures (Fig. 8), and UA sometimes simulates envelopes by producing haloes around positively stained viruses (Fig. 9). In conclusion, observers must be wary and constantly on the outlook for artifacts (Table 2).<br /><br /><em><strong>Figures &amp; Tables</strong></em><br /><br /><strong>Table 1: Comparison of PT and UA</strong><br /><br />
<table>
<tbody>
<tr>
<td><strong>Parameter</strong></td>
<td><strong>PT</strong></td>
<td><strong>UA</strong></td>
</tr>
<tr>
<td>pH</td>
<td>Neutral</td>
<td>Acidic</td>
</tr>
<tr>
<td>Crystalizes on gris</td>
<td>Generally not</td>
<td>Yes, often in blades</td>
</tr>
<tr>
<td>Acts as a fixative</td>
<td>No</td>
<td>Yes</td>
</tr>
<tr>
<td>Type of staining</td>
<td>Negative</td>
<td>Negative or positive</td>
</tr>
<tr>
<td>Grid life</td>
<td>1 month</td>
<td>10 years or more</td>
</tr>
<tr>
<td>Contrast</td>
<td>Poor to good</td>
<td>Poor to excellent</td>
</tr>
<tr>
<td>Negatively stained capsids</td>
<td>Often rounded</td>
<td>Angular</td>
</tr>
<tr>
<td>Positively stained capsids</td>
<td> </td>
<td>Black, shrunken</td>
</tr>
<tr>
<td>Protein structures</td>
<td> </td>
<td>Swollen</td>
</tr>
</tbody>
</table>
<br /><strong><strong>Table 2: Frequently Asked Questions<br /><br /></strong></strong>
<table>
<tbody>
<tr>
<td>Can I use grids without a carbon layer?</td>
<td>No. The support film is unstable.</td>
</tr>
<tr>
<td>What is better, PT or UA?</td>
<td>Neither. They are different</td>
</tr>
<tr>
<td>What is negative staining for?</td>
<td>Virus descriptions</td>
</tr>
<tr>
<td>What is positive staining for?</td>
<td>Virus counts on grids</td>
</tr>
<tr>
<td>Which viruses can be positively stained?</td>
<td>Those with high dsDNA contents.</td>
</tr>
<tr>
<td>What happens to the others?</td>
<td>They are stained negatively or not at all.</td>
</tr>
<tr>
<td>Aspect of empty phage heads in PT?</td>
<td>Membranes or black stain-filled vesicles.</td>
</tr>
<tr>
<td>Aspect of empty phage heads in UA?</td>
<td>Membranes or shadows.</td>
</tr>
<tr>
<td>How to improve virus adhesion to grids?</td>
<td>By glow discharge or wetting agents.</td>
</tr>
<tr>
<td>Why should samples be washed?</td>
<td>To remove proteins, sugars, and salts.</td>
</tr>
<tr>
<td>Shall I examine unwashed lysates?</td>
<td>Never.</td>
</tr>
<tr>
<td>Is fixation necessary?</td>
<td>Generally not</td>
</tr>
<tr>
<td>What is the average diameter of phage heads?</td>
<td>55 to 60 nm.</td>
</tr>
<tr>
<td>Do phage heads of 30 nm in diameter exist?</td>
<td>Probably not.</td>
</tr>
<tr>
<td>When to suspect large algal viruses (<em>Phycodnaviridae</em>)?</td>
<td>In icosahedral, nonenveloped, tailless particles of more than 120 nm in diameter</td>
</tr>
<tr>
<td>How to correct faulty magnification in digital EMs?</td>
<td>By calculation after photographing and measuring phage T4 tails.</td>
</tr>
</tbody>
</table>
<br /><strong>Figures 1 &amp; 2: </strong>Particles harvested by centrifugation onto electron microscopic grids. Samples from Raunefjord near Bergen were sedimented onto 400-mesh Ni grids carrying carbon-coated Formvar films, using a Beckman Coulter ultracentrifuge, an SW-41 rotor, and Polyallomer tubes with molded flat bottoms. Grids were stained for 1 min with 2% UA, air-dried, rinsed in distilled water, and examined in a JEOL TEM 100.<br /><br /><img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5jc2w6.jpg" alt="" width="297" height="233" data-src="https://s3.amazonaws.com/pr-journal/5jc2w6.jpg" data-ofn="figure 1.jpg" />                                            <img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5nc2w6.jpg" alt="" width="304" height="233" data-src="https://s3.amazonaws.com/pr-journal/5nc2w6.jpg" data-ofn="figure 2.jpg" /><br /><br />Fig. 1. Marine bacterium with mature phages before lysis                                    Fig. 2. Viral cluster outside a lysed cell.      <br />and free phages in the vicinity. Note the size variation in <br />viruslike particles. The squares and crownlike structures are <br />scales of algae, e.g., <em>Pyramimonas orientalis</em> (Chlorophyta). <br /><br /><strong>Figures 3–8:</strong> Various bacteriophages and artifacts. Philips EM 300 electron microscope; uranyl acetate (2%, UA) or phosphotungstate (2%, PT) except Fig. 5b, which is a section. Bars indicate 100 nm.<br /><br /><img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5pc2w6.jpg" alt="" width="301" height="188" data-src="https://s3.amazonaws.com/pr-journal/5pc2w6.jpg" data-ofn="figure 3.jpg" />                                                  <img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5sc2w6.jpg" alt="" width="302" height="200" data-src="https://s3.amazonaws.com/pr-journal/5sc2w6.jpg" data-ofn="figure 4.jpg" /><br /><br />Fig. 3. Negatively and positively (center) stained Salmonella                               Fig. 4. Negatively and positively (right) stained T4-like Vibrio <br />phages within the same field; UA, ×92,400.                                                       parahaemolyticus phage; UA, ×297,000.<br /><br /><br />
<div style="text-align: center;"><img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5tc2w6.jpg" alt="" width="343" height="208" data-src="https://s3.amazonaws.com/pr-journal/5tc2w6.jpg" data-ofn="figure 5.jpg" />                   </div>
<br />Fig. 5. Negatively and positively (right) stained phage 71A-6 of <em>Vibrio vulnificus</em>; UA, ×297,000. The phage is a podovirus with a very long capsid and a barely visible short tail.    <br />
<div><img id="s-mce-img" class="s-mce-img" style="display: block; margin-left: auto; margin-right: auto;" src="https://s3.amazonaws.com/pr-journal/5vc2w6.jpg" alt="" data-src="https://s3.amazonaws.com/pr-journal/5vc2w6.jpg" data-ofn="figure 6.jpg" /><br />Fig. 6. (a), Cell wall and cytoplasma membrane debris of lysed <em>Pseudomonas aeruginosa</em> cells; UA, ×92,400. (b), Vesicles budding from the cell membrane of a phage-infected Salmonella bacterium; thin section, ×59,400. (c), False herpesvirus simulated by cell wall debris surrounding an unknown particle; Vibrio phage lysate, UA, ×297,000.<br /><br /><br /><img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5wc2w6.jpg" alt="" width="300" height="184" data-src="https://s3.amazonaws.com/pr-journal/5wc2w6.jpg" data-ofn="figure 7.jpg" />                                                   <img id="s-mce-img" class="s-mce-img" src="https://s3.amazonaws.com/pr-journal/5yc2w6.jpg" alt="" width="158" height="203" data-src="https://s3.amazonaws.com/pr-journal/5yc2w6.jpg" data-ofn="figure 8.jpg" /><br /><br />Fig. 7. Slime filament and various bacterial debris.                                      Fig. 8. Phage T4 with a NaCl crystal, UA, ×297,000. <br /><em>Bacillus cereus</em> phage BL1, PT, ×148.500.<br /><br /></div>
<div><img id="s-mce-img" class="s-mce-img" style="display: block; margin-left: auto; margin-right: auto;" src="https://s3.amazonaws.com/pr-journal/5zc2w6.jpg" alt="" width="241" height="319" data-src="https://s3.amazonaws.com/pr-journal/5zc2w6.jpg" data-ofn="figure 9.jpg" /><br /><br />Fig. 9. False envelope in a phage of <em>Rhizobium</em> sp.; UA positive staining, ×297,000. Haloes of this type are sometimes seen after UA-positive staining. They expand with prolonged irradiation. Their nature is unclear.<br /><br /><em><strong>References<br /><br /></strong></em>Ackermann, H.-W. and M. S. DuBow. 1987. Description and identification of new phages, p. 103-142. In Viruses of prokaryotes. CRC Press. V. I, General properties of bacteriophages.<br /><br />Bergh, O., K. Y. Borsheim, G. Bratbak, and M. Heldal. 1989. High abundance of viruses found in aquatic environments.<br />Nature 340:467–468.<br /><br />Borsheim, K. Y., G. Bratbak, and M. Heldal. 1990. Enumeration and biomass estimation of planktonic bacteria and viruses by transmission electron microscopy. Appl. Environ. Microbiol. 56:352-356.<br /><br />Bratbak, G., J. K. Egge, and M. Heldal. 1993. Viral mortality of the marine alga Emiliania huxleyi (Haptophyceae) and termination of algal blooms. Mar. Ecol. Prog. Ser. 93:39-48.<br /><br />Brenner, S., and R. W. Horne. 1959. A negative staining method for high resolution electron microscopy of viruses.<br />Biochim. Biophys. Acta 34:103-110.<br /><br />Cochlan, W. P., J. Wikner, G. F. Steward, D. C. Smith, and F. Azam. 1993. Spatial distribution of viruses, bacteria and chlorophyll a in neritic, oceanic and estuarine environments. Mar. Ecol. Progr. Ser. 92:77-87.<br /><br />Ewert, D. L., and M. J. B. Paynter. 1980. Technique for determining total bacterial virus counts in complex aqueous systems. Appl. Environ. Microbiol. 39:253-260. <br /><br />Fauquet, C. M., M. A. Mayo, J. Maniloff, U. Desselberger, and L. A. Ball [eds.]. 2005. Virus taxonomy: VIIIth report of the International Committee on Taxonomy of Viruses. Elsevier Academic.<br /><br />Gentile, M., and H. R. Gelderblom. 2005. Rapid viral diagnosis: Role of electron microscopy. New Microbiol. 28:1-2.<br /><br />Gregory, D. W., and B. J. S. Pirie. 1973. Wetting agents for biological electron microscopy. I. General considerations and negative staining. J. Microsc. 99:251-255.<br /><br />Hayat, M. A., and S. E. Miller. 1990. Negative staining. McGraw-Hill.<br /><br />Huxley, H. E., and Zubay, G. 1961. Preferential staining of nucleic acid-containing structures for electron microscopy. J. Biophys. Biochem. Cytol. 11:273-296.<br /><br />Kay, D. H. [ed.] 1965. Techniques for electron microscopy, 2nd ed. Blackwell Scientific. See especially pp. 58 and 328.<br /><br />Kuo, J. [ed.] 2007. Electron microscopy: Methods and protocols, 2nd ed. Humana Press. See chaps. 1, 2, and 4–6. Methods in molecular biology 369.<br /><br />Montanié, H., H. J. Hartmann, C. Crottereau, and C. Trichet. 2002. Virus-like particle analysis in a seston-rich coastal pond using transmission electron microscopy. Aquat. Microb. Ecol. 28:105-115.<br /><br />Müller, G., G. Nielsen, and C. L. Baigent. 1980. Electron microscopical particle counting in virology. III. Parameters<br />governing the efficiency of sedimentation techniques and its importance with the application of poly-L-lysine. Arch. Virol. 64:311-318.<br /><br />Shively, J. M., F. S. Ball, and B. W. Kline. 1973. Electron microscopy of the carboxysomes (polyhedral bodies) of Thiobacillus neapolitanus. J. Bacteriol. 116:1405-1411.<br /><br />Skotnicki, A., A. Gibbs, and N. G. Wrigley. 1976. Further studies on Chara corallina virus. Virology 75:457-468.<br /><br />Tian, J., A. J. Sinskey, and J. Stubbe. 2005. Kinetic studies of polybutyrate granule formation in Wautersia eutropha H16 by transmission electron microscopy J. Bacteriol. 187:3814-3824.<br /><br />Tiekotter, K. L., and H.-W. Ackermann. 2009. High-quality virus images obtained by TEM and CCD technology. J. Virol. Methods 159:87-92.<br /><br />Torrella, F., and R. Y. Morita. 1979. Evidence by electron micrographs for a high incidence of bacteriophage particles in the waters of Yaquina bay, Oregon: Ecological and taxonomical implications. Appl. Environ. Microbiol. 37:774-778.<br /><br />Weinbauer, M. G. 2004. Ecology of prokaryotic viruses. FEMS Microbiol. Rev. 28:127-181.<em><strong><br /></strong></em></div>

[STEPS]
?. 
?. 
?. 
?. 
?. 
?. 
?. 
?.