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How to identify an amoeba?   Ecology of amoebae  

Getting started: brief manual to identification of gymnamoebae

Alexey Smirnov1 and Susan Brown2

1Department of Invertebrate Zoology, St.Petersburg State University
2CEH Windermere, UK


Introduction: organisation of the cell and biology of amoebae

1. Light- microscopical morphology

      The cell of an amoeba is capable to sufficient conformations, especially when it is stationary, resting. When a cell starts to move, it change shapes rapidly, and it is hard (if even possible) to describe the form of an amoeba in non-directed movement. However, when the cell starts continuous, directed locomotion it becomes more stable. The shape of such cell still undergoes minor changes, however it keeps general type of organisation until it stops the movement or change the direction. The form of actively, continuously moving cell is called the locomotive form , first recognised by A.A.Schaeffer in 1926 and established by F.C. Page in  70th.

Rhizamoeba saxonica CCAP 1570/2: A - stationary, nearly non-mobile cells ; B - cells in non-directed movement; C - locomotive form. This non-remarkable and rather small amoeba is specially choosen to show radical difference of the locomotive form from others.

      Usually in the frontal area of a cell (especially in fan-shaped amoebae) you may see the transparent area of the cytoplasm - the hyaloplasm, which does not contain any optically visible inclusions. All the rest of the cytoplasm is filled with different granules, crystals and other inclusion and is called the granuloplasm. Usually in the locomotive form the hyaloplasm is situated anteriorly and forms either frontal hyaline area, antero-lateral hyaline crescent or anterior hyaline cap.  However it should be stressed that these terms are partly artificial, as they are normally applied to the cell viewed from the top, and do not take into account the fact that, for example, in Thecamoeba spp. dorsal ridges consist of the hyaloplasm as well.

Typical appearence of the hyaloplasm in amoebae. A - frontal hyaline area in Vannella lata CCAP 1589/3; B: antero-lateral hyaline crescent in Thecamoeba similis CCAP 1583/8; C - anterior hyaline cap in Rhizamoeba flabellata CCAP 1546/2

      Many (but far not all) amoebae produce pseudopodia  - variable projections of the cell, which include both the granuloplasm and the hyaloplasm and participate in the relocation of the main cytoplasmic body of the cell. However, most of medium-sized and small amoebae move "as a whole" - without formation of distinct pseudopodia. Besides that most of amoebae are capable to form subpseudopodia - small hyaline projections of different form that do not take part in the movement of the cell. There function remains basically unclear. Often subpseudopodia are formed from the wide pseudopodium, in its anterior part.

      The posterior end of the locomotive form - uroid - in many amoebae species has characteristic distinct appearance. Various posterior formations are called uroidal structures. They are known for a long time, it was Wallich (in 1838) who first decided that the appearance of the uroid may be taxonomically important in amoebae. Later, in 1974 F.C. Page defined this term and classified uroidal structures into several types.

      Many amoebae have folds or wrinkles on the dorsal surface of the locomotive form. Wrinkles may appear also on the lateral surface of the cell.

A - finger-shaped subpseudopodia in Korotnevella stella CCAP 1547/6; B - pseudopodia in Deuteramoeba mycophaga CCAP 1586/1; C - dorsal hyaline ridges in Thecamoeba similis CCAP 1583/8 D - villous-bulbous uroid in Rhizamoeba flabellata CCAP 1546/2

Typical uroidal structures in amoebae (scheme). A - bulbous; B - morulate; C - fasciculate; D - spineolate; E - villous-bulbous; F - plicate; G,H - adhesive uroidal filaments.

      If you observe an amoeba for enough long time (or vice versa just made fresh preparation) you may see amoebae floating in the water. These amoebae - so-called floating forms (in this case the word "form" means the "cell")  usually have several pseudopodia radiating from the central body mass, however variations are possible. All amoebae are capable to float, and due to the existing of floating forms it is impossible to subdivide amoebae into “benthic” and “planctonic” species in water habitats. In one habitat the same amoebae may be found and isolated both from the plankton and the benthos, but quantitative distribution of species may differ.

Typical floating forms of amoebae. A,B and D,E - with radiating pseudopodia of different type; C - without defined pseudopodia.

      Observations on the locomotive forms of several amoebae species soon will  reveal the fact that amoebae differs clearly in the character of the cytoplasmic flows during locomotion. In some amoebae cytoplasmic flows are steady, continuos, even hardly visible sometimes. In other species, mostly in small monopodial amoebae of so-called limax type, cytoplasmic flows are eruptive, short periods of the formation of leading pseudopodium are followed with short breaks, when an amoeba does not seem to show any activity. This is so-called eruptive movement - characteristic feature of heteroloboseans. It is important that some monopodial lobose amoebae also can show single eruption of the cytoplasm. It is characteristic  for leptomyxids, but they never move by mean of eruptive movement continuosly.

      If you are equipped with the DIC optics or phase contrast you are usually able to see the nucleus of the cell. Inside the optically empty nucleus there is a big dense patch (or many small patches) called endosome  or (if confirmed with EM observations or cytochemistry) - nucleolus. To stress difference we should note that any dense body within the nucleus may be called “endosome”, but only confirmed aggregation of ribonucleoproteins (RNP) may be termed as “nucleolus”. In the literature these terms usually used as synonyms. Position of the nucleolus and number of nucleoli may be different; they determine the type of the nucleus.

Basic types of nuclear structure in amoebae. A - granular nucleus; B - vesicular nucleus; C - nucleus with peripheral nucleoli; D - nucleus with complex nucleolus.

      In most amoebae you may see one or several contractile vacuoles. The last remarkable detail, visible at LM level is crystals and other cytoplasmic inclusions. They are usually very characteristic, however their appearance and shape may depend on the culture conditions and food of the cell. It is believed that in most cases crystals represent an excretes of an amoebae, and most of other visible refractive inclusions are either lipid drops of different size of various endobionts. Mitochondria and other organelles may appear as a dark spots, however they are not identifiable at LM level and require involvement of EM.


2. Electron - microscopical morphology

      Electron microscopy, been applied to amoebae, make a revolution in our views on the organisations of amoebae cell and in approaches to amoebae systematics. Been applied initially to selected objects and then systematically, EM recovered a variety of  structures, which are used now in gymnamoebae systematics.

      Cell coat of amoebae represents highly differentiated glycocalix. It forms variety of structures, and various types of the cell coat in amoebae may be listed in the following order:

Cell coat of some amoebae species. A - amorphous cell coat of Chaos glabrum; B - filamentous cell coat of Polychaos annulatum; C - glycostyles of Vannella; D - thick, multilayered cell coat called "cutuicle" of Mayorella; E - scales of Korotnevella bulla. Scale bar is 100 nm.

      One of the most remarkable structures in the nucleus  is the nucleolus (or numerous nucleoli). Their number and position determine type of the nucleus. According to simplified classification by I.B. Raikov in 1982, we distinguish vesicular nucleus with single central nucleolus and granular nucleus with many small nucleoli. Intermediate types represented by the nuclei with several large peripheral nucleoli, like in Thecamoeba striata or with a very complicative structure of the nucleolus, exemplified with Polychaos fasciculatum, Polychaos annulatum and several more species.

      Another remarkable structure which appears in some nuclei is the internal nuclear lamina . This term mean layers of hexagonal, honeycomb-like structure (like in Amoeba proteus and Thecamoeba striata) or consisting of a fine filaments (like in Saccamoeba limax). Functional role of this layer is unclear; speculation about its mechanical properties does not seem to be reasonable. More probably, honeycomb-like layer fulfil some regulatory functions in the exchange of the material between the nucleus and the cytoplasm.

Nuclei of some amoebae species. A - vesicular nucleus of Saccamoeba limax with fibrous nuclear lamina (arrowed in A and B); C - granular nucleus of Chaos glabrum with honeycomb nuclera lamina (arrowed in C; D - cross-section; E - tangental section of the lamina); F - nucleus of Thecamoeba striata with peripheral nucleoli; G - complex nucleus of Polychaos annulatum. Scale bar 500 nm.

      Mitochondria of amoebae may be of  two principally different types - with tubular cristae and with flattened, discoid  cristae. If to consider the shape of cristae more closely , which was done by L. Seravin in 1992, it becomes clear that it is more correct to say "the cristae of tubular type", as many particular modifications are possible within this type. The same probably is true for discoid cristae, which usually are plate-like, i.e. represent a plate, growing on a fine leg. Type of the mitochondria cristae, together with the character of movement and organisation of the Goldgi complex (see below) differs well schizopirenids, acrasids and other members of the class Heterolobosea from the lobose amoebae, members of the class Lobosea.

      Goldgi complex of an amoebae may be either organised as a dyctiosomes , well-visible in TEM sections (this is a characteristic of the class Lobosea) or to exists as a set of small vesicles, non distinguishable in TEM and discernible only with cytochemical methods (in Heterolobosea). Among the cytoplasmic inclusions crystals are usually washed out during the treatment of the cell (only so-called “places of crystals” may be found), and lipids drops are usually numerous and well-visible. Endobionts present in most of amoebae, and they are well-visible in TEM sections, however it is sometimes difficult to differentiate endobyotic bacteria from engulfed food bacteria.

A - dyctyosome of lobose amoeba Chaos glabrum; B - mitochondria of lobose amoeba Thecamoeba striata with tubular cristae; C - mitochondria of heterolobosean Euhyperamoeba fallax with flattened cristae (arrowed). Scale bar 500 nm.


Basic methods for recovery, study and identification of gymnamoebae

1. Approach

      Amoebae are nearly invisible (with rare, occasional exceptions) in fresh samples, been mostly rather flattened and attached to various particles. In experiments W. Foissner recovered by direct microscopical exemanation only 2% of individuals added to the soil suspension (Foissner, 1987). Thus, amoebae must be isolated first, using a variety of enrichment cultivation methods. The background of enrichment cultivation is to create specific selective conditions for some group of protists in order to give them some advantages (in the grows rate, or by absence of grazing, for example) over other protists. This is the basic, and nearly the only currently possible approach to isolation of amoebae.

2. Media

      Following list of media is based on that compiled by Page (1976, 1983; 1988, 1991). Most of protocols and abbreviations are taken from these sources. This is rather limited, minimal set as we suggest here only the media, which were found to be most useful in practical work. Many of Page’s media are not mentioned here. In our experience they are too reach with nutrients and their use result in rapid fungal growth, especially in initial cultures. The reader is addressed to above cited books for more protocols.

2.1 Saline solutions:

AS (modified Neff’s amoeba saline).
Source: Page (1988):

      Prepare 5 separate stock solutions. To make AS  combine 10 ml of each stock solutions with 950 ml of distilled water

PJ (Prescott and James solution).
Source: Prescott & James (1955), protocol adopted from Page (1988)

      Prepare 3 stock solutions. To make PJ combine 1 ml of each stock solution with 1 l of distilled water. In our opinion this solution can fully replace AS for experimental purpose, thus been much easier in preparation.

      For marine amoebae the nost popular mineral media is the artificial or natural (Millpore filtered) seawater. Marine salt is available from virtually any Zoo shop all over the world. Any kind will work; we cannot reccomend some preferable producer. Do not forget to check any particular portion of the seasalt for presence of protozoa before use. For the same purpose, it is desirable to make negative controls during the cultivation. Do not forget to check the salinity just before the usage of seawater.

2.2 Liquid media

SE (Soil extract with salts).
Source: Page (1988).

      Into a beaker put garden or agricultural soil (preferably this one which seems to undergo no or minor treatment with chemical nutrients) and natural or tap water so that the supernatant water occupies approximately four/fifths of the depth. Autoclave for one hour, than filter. The liquid is a soil extract. Combine with water and stock solutions of salts.

CP (Cerophyl-Prescott infusion).
Source: Page (1988), modified.

      Cerophyl is a cereal derivate, which have been manufactures by Cerophyl Laboratories, Inc., Kansas City, Missouri, USA. The production has been stopped recently, and the new product with the same name has nothing common with old “Cerophyl”. However, many laboratories still have plenty of old Cerophyl in stocks, thus this protocol is listed here. Sigma Chemical supplies a product C7141 Dehydrated Ceral Leaves which probably can substitute Cerophyl.

      Boil 0.5g of Cerophyl in 1l of PJ for 5 minutes, filter and restore the volume with PJ. For marine amoebae use artificial or natural seawater of appropriate salinity; do not forget to check and adjust the salinity of the final infusion

      Resulting media is rather reach with nutrients, thus additional dilution with PJ may require to avoid superfluous bacterial growth.

2.3 Agar media:

NNA (Non-nutrient agar).
Source: Page (1988). Perhaps the most common and oldest medium for amoebae. First was offered in 1922 (Severtzoff, 1922).

      Add 15 g of non-nutrient (important!) agar  to 1 l of AS or PJ solution. Autoclave (0.5 atm; 30 min).

CPA (Cerophyl-Prescott agar).
Source: Page(1988).

      Add 12 g of non-nutrient (important!) agar  to 1 l of CP infusion. Autoclave (o.5 atm, 30 min).

      For marine amoebae use marine water instead of PJ and CP made on marine water, respectively. Note that under the salinity higher then 35 ppt the agar may coagulate in the solution. In this case we can advice to make a freshwater agar of double concentration and to dilute in just after autoclaving with the double-concentrated seawater or cerophyl infusion to approach desirable final salinity.


Sampling, inoculation and examination of initial cultures for faunistic survey

1. Soil samples
      The sample, taking by amoebae-free instruments (sterilisation is not obligate, however preferable) should be diluted with PJ or AS up to the appropriate concentration of soil particles. In case of faunistic survey dilution level should be adopted by the investigator, the primary criteria is that soil particles should leave enough free space for amoebae observation on the agar or plastic surface of the dish after inoculation. We can advice to start from approx. 15g of soil /1l of PJ or AS solution and adopt it during further work.

      We advice following set of media for initial inoculation:

      Each dish should be inoculated with approximately 1-2 ml of diluted sample, it is important to have not only the liquid but several soil particles in each inoculated dish. In dishes with liquid media (both agar and non-agar dishes) the "spot" of particles should be accurate, local, and placed preferably near the edge of the dish, to leave enough bottom space for amoebae observation. In dishes with agar media, 1-2 ml of diluted sample should be dropped near the edge of the dish, than the dish should be placed vertically to allow the drop to flow down, thus forming a path across the dish. Agar cultures should be tightly closed with Parafilm or any similar laboratory film to avoid drying. Incubate cultures under room lighting and temperature. Termostabilised cabinets, with the temperature close to that of initial habitat seems to have no influence on the success of species isolation.

2. Freshwater and marine benthic samples

      Samples should be appropriately diluted to reach relatively low concentration of particles in inoculated dishes. A test of 10 most popular enrichment media (Smirnov, 2003) revealed that few media allowed rather comprehensive species recovery, and that the number of  inoculated dishes is of more importance than the number of media used. It is possible to offer following set of media that allow recovery of most amoebae species:

      Filtered water from original habitat may be replaced with the cerophyl infusion or PJ medium. However, it seems to be logical to suggest that the water of original habitat that has characteristic chemical composition may allow for the creation of more appropriate michoniches. It results in the development of more species than an artificial, chemically-defined media. Inoculate not more than 0.1-0.2 ml of the sample per 100 mm dish. Patches of the inoculated material must leave enough free bottom space to observe amoebae and bacterial and fungal growth in the dish must not suppress the amoebae.

3. Plankton samples

      The only difference with benthic samples is that the number of amoebae in planktonic samples is relatively low (with rare exceptions). No dilution require for such samples, the sample may be directly inoculated onto same media as benthic ones. The water of the sample play the role of liquid media. In certain cases techniques for concentration of samples may allow to reach better species recovery due to appearence of rare species.

4. Examinatiom       All initial cultures should be examined several times, as the succession of amoebae species is considerable and rapid. We advice to examine liquid cultures at 5-6; 10-11; 17-20 and 30 days after inoculations. For agar cultures the last time may be omitted. In case of the time shortage, two examination - at 10-11 and 17-20 days are generally sufficient. Cultures should be examined using both dissection microscope (to recover large amoebe) and inverted phase-contrast microscope with the magnification 200x or (preferable) 400x.

      Agar cultures usually show more rapid amoebae growth, however only small and partly medium-sized species are developed in these cultures. They should be examined without opening, under dissection microscope at the magnification 40x or higher. It is easy to find amoebae, as in contrast with other protists they have strong tendency to migrate on the agar surface outside of the initial path of the inoculate, forming a set of small clamps on the agar surface with the narrow tracks of liquid below them. However, some species prefer to stay within the initial paths. Any portion of the agar, suspicious on amoebae growth must be a subject of attention and subsequent cloning (see below).

5. Cloning techniques

      In terms of the modern systematics of amoebae identification of a species really represent an identification of a strain. Thus, after finding of an amoeba species in initial culture, amoebae should be cloned (in order to avoid errors related with the similarity of many amoebae species) and prepared for further studies.

      A variety of methods is suggested to clone amoebae. However, an experience shows that the level of success (% of obtained clones) is nearly similar in all of them. We suggest to use three basic methods, the reader is refereed to the cited literature for more techniques. The reader always should remember that the success of cloning (the number of resulted cultures) is rather low, and normally does not exceed 10%.

Migration method.
      It is appropriate for amoebae growing on the agar (with or without overlay) and should be applied as following:

      When you observe initial culture of amoebae on the agar without overlay under dissection microscope, you need to find sites where amoebae migrated for sufficient distance from the initial path of the inoculate and are not too abundant (if cells are to abundant or different types are mixed this may mean that you need to increase dilution of the soil sample for initial inoculation). Cut of a small block of agar with one amoeba or one cyst with a scalpel or a needle with flattened end. Resulted block should be transferred to a fresh Petri dish with the same agar media, and the cell should be washed of from the block by addition of a drop of  respective liquid (same as used for initial isolation).

Pipette technique.
      This is intended for amoebae, growing in liquid media. The method is trivial and require experience in pipetting. Make very fine cappillar end of a Pasteur pipette and try to capture one cell (or cyst) from the culture. Resolution of this method is much lower (more mixed cultures as a result), however it nearly the only availiable in practice for cultures growing in 100 mm Petri dishes with grains.

Dilution technique.

      This method based on the critical dilution of amoebae. Wash all amoebae (and other material) from the surface of a dish by a portion of fresh  media, and dilute supernatant  1:100, 1:1000, etc. (should be adjusted from preliminary results). Inoculate new dishes with 1ml of dilution each. If during examination at all times of the check you see only one species in the dish it is suggested to be a clone.

      For all three techniques it is preferable to repeat cloning. Only subclones may be relied on in further investigations. Suggested clones must be monitored for some time (about a month), at different growth time to make sure that no other amoebae (and preferably no other protists of comparable size) are  present. The last is important for further EM studies.


Identification of amoebae

      Identification of amoebae remains one of the most difficult problems, and a lot is written about this. For correct generic and specific identification EM is obligate  in the very most of cases. Another problem is our relatively low level of knowledge about amoebae biodiversity - the chances to find new species in any habitat are very high. For example, detailed faunistic survey of a freshwater lake revealed 32 Gymnamoebia species, of which 15 were found to be new for science (Smirnov & Goodkov 1995). You should always be ready to meet unknown species in your cultures.

      In order to identify an amoeba you need first to decide with the appropriate level of detalisation of your identification. If your are satisfied with the level of a morphotype, you have no need to clone amoebae, observation from initial cultures are sufficient. However, if you are going to follow further in systematical identification you must fulfil all requirements, for example, of Page’s key (or other respective literature) concerning the set of  necessary species data. This may be rather laborious. It is strongly preferable to stop with the reasonably identified morphotype or to identify genus using respective EM and respective literature, rather than to make non-reliable suggestions about generic or specific position of your strain, if you are not sure or unable to fulfil all requirements of systematic identification.

      Identification itself consists of several distinct steps. We will consider them subsequently, and this schedule should be used in real work as it is described. You are welcome to stop either at the first step, if you are going to identify a morphotype only, or to follow them all for systematic identification, using methods as required in respective literature (cited for each morphotype).

      If you are going to deal with systematic identification of amoebae first consult F.C. Page’s keys (1988 - in English, 1991 - in German). They allow you to have a good deal of information and provide you with the basic steps for systematical identification. They may be sufficient for species identification, however when you will decide with the species always check the original description and latest papers dedicated to this species (if availiable). Review the literature, dedicated to your species and relative taxa which was published after 1988. These papers may contain descriptions of new, “post-Page” genera and species which have been found already in many taxa of amoebae. Consult the cheklist of valid amoebae species at this homepage. It is difficult to give further advises, as here you will be already at the expert level of identification, thus follow the recommendation of Page's keys and  related  literature for further work.

Step 1. Locomotive form.

      Locomotive form - the form of an amoeba in continuos, directed movement is a background for any further speculations. If you are working with water-immersion or inverted optics, find locomotive amoeba on the clean area of the bottom, free of sufficient debris of bacteria and detritus (presence of a material on the bottom of the dish may influence locomotive form). If you are working with agar culture, wash amoebae from the agar with a drop of respective media,  place this drop on the object slide and cover with a coverslip. The drop should be of a size that the coverslip does not touch cells. If you are working with  large amoebae, scrape a piece of vax by each corner of a coverslip prior to use it. This will form small “legs” on the coverslip which predicts contact with the cells. It is very important to avoid depressement of amoebae with the coverslip, as this influence sufficiently on the locomotive form and may result in misidentification. It may require some time for amoebae to start movement and adopt locomotive forms, thus it is better to place ready preparations in a wet camera for two-three hours and only than to observe them.

      Choose actively moving cell and note the shape and characteristic details of the locomotive form (uroid, hyaloplasm, ridges, lateral flatness, shape of subpseudipodia, lobs and wrinkles, if  present).  Sketches, videoprints or photographs  of moving amoeba may be highly useful in further work. Measure the locomotive forms using micrometer. Preferably several amoebae should be measured, but make sure that they all belong to the same species! If you are going to identify species you should work with clones, and measure not less than 30 amoebae to have average measurements. Note the nuclear size and structure, shape and size of crystals (if present), typical position of contractile vacuole (if any).

      If amoebae were maintained on the agar without overlay, it may be extremely difficult to observe moving cells. It seems that been cultivated under these conditions for several generations, cells loose partly their locomotive capacities. To avoid this, prior to observe locomotive forms cover the agar with the overlay of respective media and leave  it for two-three days. In the very most of cases this is enough for cells to restore locomotion. In worst case try several passages using agar with overlay.

Step 2. Floating form

      Floating form is very important for species identification. Observation of the floating forms should be done preferably in clonal cultures, unless the size difference of amoebae you are interested in from any other existing in this dish is sufficient. In some cultures with overlay or in liquid you may easily see floating forms at any time under dissection microscope. If not, to observe floating form, shake a  culture (if it has an overlay or is liquid one) carefully and observe under the dissection microscope development of the floating forms. Far not all amoebae form them readily, and you need to see the dynamic  of the shape changes in floating amoebae to make sure that you have seen developed floating forms. For smaller amoebae and amoebae which are maintained on the agar without overlay,  wash of cells from the dish with the drop of respective media, place the drop on the object slide, close with coverslip and observe immediately. Sometimes it is possible to see floating form  and than locomotive forms, subsequently, on the same object slide.

      Note the appearance of the floating form, shape and number (min/max.) of pseudopodia, the material of pseudopodia (hyaloplasm only, or with the granuloplasm), shape of the ends of pseudopodia, their thickness. Note if amoeba has tendencies to form coiled or spiral pseudopodia. Measure the length of pseudopodia comparing it with the size of a central mass of the cytoplasm in radiate floating forms. Some amoebae species has a tendency to gradual modification of the floating form  with the increment of the time of cultivation, thus floatings form of fresh isolates may differ slightly from the floating form of the same species in culture collection.

      There are amoebae species which do not adopt any specific floating forms. They flotate remaining usual, locomotive-like. There are minor of them, and careful observations required to make a conclusion that an amoeba do not have differentiated floating form.

Step 3. Nuclear structure and crystals

      Nucleus and crystals are well visible with oil immersion optics, using  100x objective lense. Amoeba for these observation preferably should be slightly pressed with the coverslip in order to make nucleus and crystals better visible. Note nuclear structure, number and position of nucleoli, shape and size of crystals, count approximate number of crystals. However you should not measure nucleus under these conditions! It should be done in locomotive form.

Step 4. Cysts

      Cyst formation and cyst structure is a very important criteria in amoeba systematics. However, far not all species form cysts in culture. Basically, to observe cysts you need to have pure clonal culture. Cyst may be found after 7-15 days in agar cultures and after longer periods (up to month) in liquid. Some species loose the capacities of encystment after some time of cultivation, some do form cysts only in cultures with overlay. Different conditions should be applied and all cultures should be traced for at least a month prior to conclude about cyst formation. Cysts should be observed with LM under 100x oil immersion, shape, structure and number of cyst walls, presence and disposition of cyst pores, size of cysts must be noted. Cysts should be a subject of EM studies as well as trophozoites.

Step 5. TEM studies

      TEM studies are obligate if you would like to identify an amoeba up to the genus and species, as the microsystematics of gymnamoebae itself is based on EM features.  This is a great disadvantage, of course, but so far we have no other way do distinguish species with sufficient level of confidence. We do not describe here all EM protocols, given that the reader is familiar with these techniques. Only important points are listed.

      To prepare amoebae for EM wash them of from the agar with a drop of media or shake the dish with liquid culture carefully. Concentrate amoebae with gentle centrifugation. We do not advice to embed amoebae in the agar blocks after fixation, as it seems to damage cell coat. If amoebae are adhered well to the agar, it may be easier to cut off small (max. 2x2x1 mm) blocks of agar with adhered amoebae and treat them further in a glass wells. For amoebae which are non-numerous in culture the following approach may be useful (Smirnov & Goodkov, 1994).

      Prepare 40 mm Petri dishes with a layer of polymerised resin. Let them stay opened for 2-3 days after polymerisation. Place several drops containing amoebae on the resin and mark the position of the drops (scrape the resin around the drops with a needle). Leave them in wet camera for 30-60 min in order to allow amoebae adhere to the resin. Fix amoebae and treat them as in the well-glass, in these dishes. The only difference is that you should not use acetone or any other substance which may dilute plastic of the dish during the embedding (we use 100% ethanol to dilute resin at the final steps of embedding). Cover amoebae  in the dish with a thin layer of  resin, preferably from the same initial portion of resin. Blocks should be sectioned so that the border of the layers of resin is either parallel or perpendicular to the knife.

      This method allow to work with very small amoebae and even with single cells, as it is possible to examine embedding even under the microscope (marks of the initial position of drops with amoebae are highly useful here), to ensure that you have really the organism of interest in the embedding and to mark its position in the block exactly. This method often gives better results then these which involves centrifugation; it seems that it is preferable to fix locomotive forms to have better quality.

      Fixation. This is the most important part of TEM, and only appropriately fixed specimens may be considered. Numerous artefacts are possible, especially the elimination or coagulation of filamentous cell coat under unsuitable fixation (Smirnov & Goodkov, 1998; Smirnov, 1999). Three basic protocols may give satisfactory results, depending on species. They far not exhaust the variety of existing methods, and the reader is referred to Page (1983; 1988) and other respective literature for more information . All fixative in following protocols are suggested to be made on 01.M Phosphate  buffer (PBS) Ph 7.4 or 0.05M NaCacodylate buffer Ph 7.0 - there is no noticable difference in the fixation quality in the very most of cases.

Procedure 1. This is the most common protocol

Procedure 2. This procedure appears to be best for marine amoebae

Procedure 3. Appears to preserve well even very fine glycocalix

      The last protocol is most complicative, however sometimes it gives perfect results. For example it was the only which allow to recovery filaments in the cell coat of Polychaos annulatum (Smirnov & Goodkov, 1998), while all other resulted in the collapse of filaments.

      All protocols are followed with the standard dehydration, embedding in any appropriate resin, sectioning, staining of sections with uranil acetate and Reynold’s lead citrate and examination.

      Some amoebae have glycostyles or scales on their surface. This is characteristic for amoebae of fan-shaped, dactylopodial and paramoebian morphotypes. These  structures are poorly visible in TEM, thus chromium shadowing may be simple and highly useful technique to recover them. In fan-shaped amoebae only chromium shadowing can ensure in the absence of glycostyles.

Shadowing protocol adopted by Ken Clarke (CEH Windermere)

      For shadowing collect amoebae from liquid cultures or wash them off from the agar surface. In the very most of cases, cells in cultures are densely covered with adhered bacteria (this is especially true for agar cultures, thus try to wash bacteria off  by active pipetting of the drop. Preferably the drop containing amoebae should be placed in a glass well, diluted with sufficient amount of distilled water (osmotic shock does not seem to be sufficient) and amoebae should be re-collected with pipette. Sufficient losses of cells are possible at this stage. Floating form are preferable for such examination, thus wait until the cell adopt them (if they do). Place a drop with washed amoebae on a formwar-coated grid, and cover the grids with a Petri dish cover with several drops of osmium tetroxide for half an hour. Let  the drops dry on the grids.

      Remove the grids to a vacuum coater for shadow-casting. The ‘grazed lighting’ effects produced by this process will enhance the visibility of fine structure such as glycostyles and scales. Use gold-palladium (medium resolution, high contrast shadows), platinum (high resolution, low contrast shadows) or chromium (recommended for routine use) as a shadow source. Shadow at 30o-40o to the horizontal for the examination of general cell shape, or 20o-30o to the horizontal for viewing surface filaments, scales, glycostyles, etc. Examine each grid by TEM.

      NB. As shadowed cells on the formvar grid-coating may shrink in size when exposed to the electron beam, be sure not to confuse shrinkage artifacts with fine surface structure.

Step 6. Special techniques

      The four above steps  gives enough data for identification of the very most of Gymnamoebia. However, some groups may require special techniques. Identification of Acanthamoeba, for example,  require physiological tests, impregnation of cysts and biochemical tests as described in the respective literature (references within a key). Amoebae of eruptive morphotype can be identified at the morphological level mostly up to genus only, and this identification require enflagellation tests. For most of these amoebae reliable identification of species is possible only with molecular methods, as they are indistinguishable at morphological level. Here you must follow recommendations in the literature, cited for the respective  morphotypes.