Part 1: Introduced the taxonomy and biology of the Class Bdelloidea.

Part 2: The key to the 15 genera of the Class Bdelloidea.




This article forms the third part (Technical Notes) of the

Key to the Genera of the Bdelloid Rotifers

already published. Once written I realized that most descriptions of the micro-habitats, and also the techniques to be used for each of them, could even be applied almost without modification not only to Bdelloidea but to many other groups of micro invertebrates (gastrotrichs, nematodes, rotifers Monogononta, the inhabitants of the meiobenthos, including the kinorhyncha, and with a few modifications that surely the imagination of the amateurs will discover quickly, to the entomostracans and the hydracarina). Consequently I modified the title to make it more accessible to the indexing for the Internet, although I maintain the text without change. My primary objective is still to help stimulate more amateurs' work on bdelloids.

I use here some terms to designate “biocenosis” that probably are not supported by the European or even American bibliography (they were coined in Institutes of Limnology from the southern hemisphere, where the researchers must deal with a much diversified biological world). I believe that it is very useful to separate which most of the time are different and identifiable species groups, which could be otherwise confused if you don’t make a careful selection of the microhabitats. I make a short definition of the used terms.


Leaving alone Zelinkiella, an exclusively marine genus, which lives as a commensal on holothurians, the bdelloid rotifers are found in practically all ecological habitats, although most of the time in freshwaters (or systems that could dry completely, being periodically dampened by the rains). Only 21 species in 8 genera have representatives which have been found in continental waters, salty or brackish. (Fontaneto et al, 2008) and the majority are haloxenic species, that is to say, species that are suspected to have only arrived and survived by chance in those habitats.

In fresh waters they can, and they must, be looked for in zooplankton of the larger water bodies (fishing with a fine mesh of about 70 microns at the most) but there will be found only a few species mixed with the frequent Monogononta. The most fruitful habitats are described next.

The classification of the biocenosis is not capricious. Each one has its own physical, chemical and structural characteristics, which determine which fauna can live there. One must learn how to recognize them.

a) littoral forms: those that swim free near the edges of the water bodies, (sometimes called heleoplankton) or crawl on the littoral plants. Between these must be distinguished those related fundamentally to pleuston,(floating plants, like Salvinia, Pistia or Eicchornia) and those that are periphytic (growing on and around a support) over the bafon,(submerged part of the rooted plants). Or the ones bound to plocon (floating filamentous algae, fixed to the shore, stones, or objects) and to heteroplocon (free floating filamentous algae). The relationships of these faunas, between them and with those free swimming, are debatable, but in most cases they are distinct species.

b) inhabitants of other bioderms (also called biofilms), thin layer of bacteria, micro-algae and micro-invertebrates that occur on permanently or temporarily submerged substrates, like stones, wood, the submerged areas of the floating or emergent plants, submerged roots of trees, walls, wood, etc. (periphyton).

c) those that live between the grains of littoral sands (psammon) or in the thin layer of the superficial benthos (ooze).

d) sapropelic, those that live in ponds and small pools contaminated by organic matter in decomposition (in any one of the previously described situations).

e) those that live in waters that are collected periodically in cavities of trees (tree-holes), or in the “pitchers” (pitfalls at the end of leaves) of the carnivorous plants like Nepenthes, or in the “cup” or “vase” reservoir, that forms in the center of many bromeliads, because of a rosette of overlapping leaves (phytothelmics).

f) those that live in periodically dried mosses or lichens (bryophytes), in and under pine needles and litter (edaphic) or in cavities, be they always humid, or flooded periodically, on rocks, walls, buildings, pavements, etc. (lithothelmic).

g) in addition, there are 3 epizoic genera (Anomopus, Embata, Zelinkiella), commensals on freshwater animals; (one of them, Anomopus in brackish water).

It is becoming increasingly common to call the species that thrive in the e and f situations limnoterrestrial, because evidently they are in terrestrial environments, but thriving only when they are wet or flooded.

Sampling and conservation

All trips in search of qualitative samples of bdelloid rotifers must be specific, directed to a certain habitat (i.e: periphyton, bafon, etc), and very limited in the sampled volume. A single live sample can provide sometimes many species, with few individuals each, which will force a slow and individualized study. Three or four samples would be the ideal. An ample investigation of a selected habitat, must be divided into many small samplings throughout a certain time. This will allow not only a better idea of the specific richness (the number of species present at the site) but to obtain a first sight of the possible successive substitution of the species, driven by physical events such as the seasons of the year, or the state of humidity of the materials. The live samples must be protected from abrupt changes of temperature, they may be best transported in a thermally isolated box.

Of course the sampling methods are as varied as the habitats that must be investigated, and in addition not only provide rotifers but a very varied microfauna, although here I have concentrated on the methods best adapted for bdelloids.

The epizoic rotifers can only be collected by examining the animals that lodge them and washing the open surfaces (which include, by example, the cavities where gills of the crustaceans are sited, or the mantle cavity of mollusks) to gather the agile or fixed organisms that inhabits them. (Vagile: not fixed, moving animals.)

The coastal forms raise a true challenge to the talent of the collector. It is easy to gather, with small plankton nets, the forms that swim free near the shore, or between the plants. But there also exists a varied microfauna that adheres more intensely to the surfaces, or that inhabits more intricate habitats.

For pleuston, plocon and heteroplocon there is no other remedy than to gather portions of the corresponding flora, carefully sliding them with its water and fauna within bags or bottles.

Periphytic fauna. This is most difficult to gather. For the inhabitants of the bioderms which cover the submerged part of diverse rooted plants, stones, wood, etc., one generally resorts to lift off, with care and skill, with the help of a brush or spatula, the material adhered, directing it (with luck) towards the harvesting containers. For rooted emergent plants (e.g. rushes) a somewhat safer method is to cut (whenever you can) the aerial part, inverting a bottle or plastic bag to surround the submerged stem with its adhered microflora and microfauna, and cutting the plant below the mouth of the container, closing this and removing from the water. This even allows advanced researchers a quite exact quantitative sampling.

For plankton the conventional plankton nets with 70 or 45 micron mesh are used. They can be towed behind a boat driven at a low speed. Or the fauna can be fished-out of a “water column”, lowering the net to the bottom and recovering it vertically with a slow and steady ascent.

For the rotifers of mosses, lichens, litter, sands and soil, samples will be taken which must be investigated later at the laboratory.

The sands must be collected with abundant water from the sampling site. Shaking them, to suspend them, in a great volume of water, can help to loosen and suspend the interstitial fauna. Let it rest some seconds so that the sand settles, and pour off the supernatant water. Concentrate it by straining through a fabric of 40 to 70 microns mesh. As many of the psammobionts (animals which live in the interstices of the sand, at the shore of the water bodies) adhere strongly to the sand particles, researchers generally try to anesthetize the microinvertebrates using magnesium sulphate, or bupivacaine, (see “anesthetics”) which facilitate its removal. It must be investigated with care the effectiveness and the form of administration of the anesthetics when collecting bdelloids. The times and suitable concentrations vary according to the species present in the samples.

Some investigators propose to leave the sample alone for two days. The oxygen at the bottom will be consumed and the animals will creep towards the surface. It will be possible to gather them with a pipette at the interphase between the sandy substrate and the water. The best site to search is the angle between the sand and the glass walls of the container. Exploration can be continued over one week or more.

The lichens and mosses must be wetted with distilled water or, better, rain water, and they will be let to rest for two days. They can be squeezed to separate the water from its microfauna, and the liquids will be investigated during several days with the stereoscopic microscope or under the low power objective of the binocular to separate the detected fauna.

The rotifers of ground and litter are collected by treating the samples in the laboratory, using a Baermann funnel. This is a funnel with a sieve applied to its mouth, where the sample is placed. The tip of the funnel is closed with a rubber tube and a Mohr clamp. The funnel is filled with water up to the level of the sample. The vagile animals can cross the mesh of the sieve and accumulate in the rubber tube, from where they will be collected periodically.

Phytothelmic fauna is better studied if the plants are carried to the laboratory, if it is possible. Otherwise, the liquids in the vegetable container must be siphoned to a sampling tube or bottle and must be transported as any other liquid sample.

Lithothelmic habitats will be sampled, absorbing the water filling the cavities, and scraping the sediment, generally rich in algae, and especially in cyanobacteria.



In the following technical discussion I refer sometimes to the late H. Taylor. He was a specialized technician that worked all his life with rotifers, and started his career with the great rotiferologist F.J. Myers. He published a series of technical articles that are now bound in a book. Although aimed at the professional researcher, many of his suggestions are worth trying by the amateur microscopist.

Those individual fauna selected for a later more detailed study, or to be mounted as vouchers”, (duplicate specimens, which are filed for future reference and comparison) will have to be anesthetized, if it is possible.

The best ever known anesthetic for rotifers, including bdelloids, is the “Liqueur de Rousselet”, for which we have two formulations, the original, and a modification due to De Beauchamp. The Italian limnologist Mario F. Canella says (1954) that even traces of this reagent were so effective that they stretched immediately the individuals of Rotaria neptunia.

Even if all the species were not so collaborative, it is worth the trouble to try the Rousselet if its basic ingredient can be obtained: Cocaine Chlorhydrate.

ROUSSELET original (0.006%)

 Cocaine chlorhydrate 2% solution…………………..3 ml

 Methyl alcohol (full strength)…………………………..1 ml

 Distilled water……………………………………………….....6 ml

De BEAUCHAMP, modification (0.05%)

Cocaine chlorhydrate………………………………………..0.5 g

Methyl alcohol (full strength)………………………….5 ml

Distilled water………………………………………………....5 ml

With either of the two formulae it is recommended to add 1 drop of anesthetic to each mililiter of sample, every 5 minutes, until obtaining the narcosis.

In addition to the Rousselet formulae, many other anesthetics have been tried.

Most used now is Bupivacaine (also known as Marcaine).

Bupivacaine (Marcaine) was recommended by H. Taylor as a 0,5% mother solution. It is an anesthetic used by dentists, which I obtained in presentation of 30 milliliter with 150 mg. of active substance. Dilute it and use carefully drop by drop, or infiltrate the diluted solution under the coverslip. It is a proven narcotic with Monogononta, I tried it with bdelloids, but results were successful only a few times. Segers has successfully anesthetized Adineta ricciae with this substance.

1% Magnesium chloride or sulphate, was also recommended by Mario F. Canella (1954). Also use it drop by drop. There is some note of its successful use in some species of bdelloids, but most of his pictures are from Monogononta. An 8% solution is regularly used by meiobenthologists to loosen the microfauna clinging to the sand from marine habitats.

Neosynephrine (Phenylephrine) ophthalmic drops are used by ophthalmologists to dilate the pupil of the eye. They are also sold as nasal sprays. You must take care with these, because the drug is very addictive. Doctors recommend not to use it as a medicine for more than five days! I had tried it a long time ago, and it was very good (as always) with Monogononta. It was deceptive applied to bdelloids.

Homatropine can be obtained in drug stores as a medication for liver diseases, and also as ophthalmic drops. This, and a lot of other drugs as atropine, benzamine hydrochloride, xylocaine (lidocaine), tetracaine, metoprolol, etc. were recommended to be tried. But no one reports today an always successful anesthetic for bdelloids. But all of them merit a trial.

The anesthesia must be judged by the lack of response to the contact stimulation, because cilia do not anesthetize generally, and the rotifers continue moving, even if they are well anesthetized.



Sometime ago the anesthetized rotifers were fixed adding to the containing drop a similar volume of 8% neutral formol, or 6% glutaraldehyde (H. Taylor used it as a 2% solution). Formol is now banned as carcinogenic, but glutaradehyde is not. (See footnote, added March 13th). Although bibliography shows that professionals continue to use both products, even if many laboratories have changed to less toxic fixatives.

GALA 60 (Dioni) was used in Italy by Dra. Ulrique Uehliger at 10% concentration, in planktonic rotifers samples, and they reported a good behavior in samples conserved in the fixative for two months, (of course most of the species were Monogononta). Nevertheless it is highly advisable to discard the liquid of the sample, and wash it with 70% alcohol after no more than a week, and preserve it afterwards in 70% alcohol plus a 5% of glycerin. This has three advantages: it prevents the possible corrosion of organisms by the strong acids, provides a permanent preservation medium, and facilitates the later work and the faunistic counts eliminating the irritating fumes of acetic acid.

As the specimens (fixed or alive) are selected from the sample being searched, they will be withdrawn with a micropipette and accumulated in a small watchglass or any small capsule, to conserve them for future study.

Although it is always advisable to bring a live sample to the laboratory, at least in the beginning of a project, most of the time the samples will be generally fixed in the field. If bdelloids are the objective, the traditional methods of fixation will show only unidentifiable contracted units.

So, I propose to divide the sample in three:

A subsample will be anesthetized with Bupivacaine (or the anesthetic that would be finally selected by experimental trials) until it only shows ciliar activity, and the affected animals do not contract. H. Taylor suggested, as a standard technique, to split the concentrated sample in 5 or 6 subunits of 4 to 5 ml each, and to apply to each one an increasing number of drops of the anesthetic, expecting that one turns to be optimal. He also says that a uniform time (9 min. max.) would be used, to fix the samples. It seems that more time allows for the specimens to contract even if they were already well expanded. At this moment one will add to each subsample, 10 to 20% GALA, with or without any in toto stain. Before, I used 0.2% Rose Bengal, but it is now considered strongly carcinogenic. Try the use of much diluted Allura Red, or Tartrazine (respectively red and yellow food colorants), or, even better, 0.5% to 1% aqueous Eosin.

The second can be treated with CO2 (mixing with the sample some gasified commercial water) or by asphyxia (small flask, concentrated collection, very tight sealing), which, in this material, can rarely produce some anesthetized bdelloids.  If the rotifers do not die stretched by the boiling water, there would be certainly many other invertebrates that do, which could also be very interesting for the microscopist which for this reason could be prone to do a more complete inventory of the sample.

Third unit - The samples gathered by filtration of the sediment detached from pleuston or bafon, or gathered as benthonic ooze, are generally too voluminous for an individualized treatment in the field. In this case the best strategy (apparently designed by Frank Myers) is to place the sample in a container of a volume 6 times greater, located as a safety precaution, within another one 2 to 3 times bigger. Time is allowed, so that individuals stretch out and retake their normal rate of activity, at which moment, 3 to 5 volumes of almost boiling water is poured into the first container. Sediment is allowed to settle for a long time and upper liquid is poured out to the maximum possible. A volume of fixative similar to the sediment volume is added to the sample. As in the second suggested treatment only a few species of bdelloids respond to this treatment, which is normally very useful with Monogononta.

Alternatively, the filtrates or sediments can be placed in a “detachable-neck flask”, with the base painted in opaque black (Dioni). After a time, the geotrophic negative, and phototrophic positive, swimming animals, plus those that suffer with the oxygen rarefaction, will meet in the detachable neck. This one is taken apart, its content is poured in the definitive container, and the fixative is added; or the microfauna is first massively anesthetized before to fix it, or it is treated with almost boiling water or almost boiling fixative.


Methods of study

The sample is first studied with a stereoscopic microscope, or the low power objectives of your microscope. Preferably over a dark background. And the individuals whose morphology must be specially studied, or which must be identified taxonomically, will be separated, using micropipettes with buccal or mechanical control…… (if you try your pipette under the low power, remember that the movements has its directions inverted. If your microscope allows it, use the “Dioni’s poor man stereoscope”. Some investigators consider that pipettes allow the adhesion of the animal to the glass, preventing them to be unloaded to the slides, and suggest to learn to manipulate the rotifers, even live, exclusively with micro-loops, micro-spatulas or micro-needles. They are saved in watchglasses, or suitable capsules of any type, where they are accumulated, and are later transferred to slides in a drop of water, and first studied uncovered, to verify its behavior in an unrestricted medium, style of swimming, etc., or they are covered with a coverslip. As water under the coverslip evaporates the weight of the coverslip can apply pressure on the specimens. If this is not prevented the rotifer could be squeezed. To avoid this

  1.  For voluminous species some support must be included (paper, cellophane, small coverslip parts, hairs, etc).
  2.  A more technical solution, but a not so simple one, is to create a petroleum jelly compressor.

Place a small drop of water, with the specimens to be studied, on one slide. Spread a very thin layer of petroleum jelly on the palm of your hand, and slide over it two opposite edges of a coverslip, to gather a thin line of jelly on them. Invert the coverslip and, carefully, lower it VERTICALLY over the drop, watching to keep it centered. With the aid of two thin and blunt needles, (or even toothpicks), adjust the height of the coverslip, looking at the preparation with the stereoscopic microscope, or with the low power objective of the binocular, until a delicate compression of the rotifers is reached, that will be controlled with great care, to avoid destroying them.

Individualized processing:

a) The selected living individuals, can be treated with some anesthetic, if any one is useful, fixed with GALA, to be stained and mounted in permanent preparations, as described below.

b) The individuals of the species already fixed in the field, which would be needed for further study, must be separated and collected in a small capsule. If they have been massively stained with Tartrazine, or Allura Red or Eosin, they go directly to the glycerin as it is explained below. But, if not, 0.5% Eosin will be added to them letting it act for the necessary time. By means of micropipettes, or working with microloops or microspatulas or even microneedles,  the animals must be worked out of the staining solution, and they will be washed in water, to be mounted in glycerin.

Glycerin will be applied through several graduated steps of 10, 25, 50, 75 and 100%, a few minutes in each one, or they can be placed in a 5% solution that are left to concentrate under a dust cover, over several hours. The goal is to avoid the specimens wrinkling, but to concentrate and mount them in pure glycerin (H.Taylor used glycerin with a “touch” of phenol, as a bactericide and an aid in clearing). Apparently the fragile species that wrinkle during the mounting process can become suitably hardened if they are previously treated with 10% acetic acid.

The mastax is indispensable to identify the species, even if its general structure is very similar throughout the class. A solution of 0.3% of commercial sodium hypochlorite (supermarket cleaner bleach) is recommended by H.Taylor. The commercial solution is normally a 5% solution. Dilute 1 drop of commercial solution with 8 drops of water (9 final drops). This gives a 0,6% solution approx (5%/9 = 0,6%) Add 1 drop to another water drop with the rotifer, the working solution is now 0,3%, and it starts dissolving the cells and leaving only the hard structures. It is better to work with a well slide, and with small drops to prevent losing the tiny structure.

It is also possible to work under the coverslip, adding a 0.5-1% solution drop to the cover margin and absorbing with great care the water at the opposite margin with a thin but long strip of filter paper. It is also possible to use the same technique to replace the water by glycerin which has a better refractive index.



Put your sample in a drop of water on a slide. Ricci and Melone (2000) suggest that, at first, you study your material without applying a coverslip. This is easy with the low power stereoscopic microscopes, but only for the 4x and 10x objectives with the compound microscopes, and with more or less quiet animals, but could give you a good appraisal of the general morphology and activity of the rotifer.

The general morphology will be described from observations with a 10x eyepiece and the objectives 4x and 10x, with complementary details with a 40x, and some details (the trophi as an example) will even be recorded with the 100x Oil Immersion objective, if you have one. The law here is to “document all that is possible, in the best way which will be possible”.

Digital pictures are a rapid recording technique. Now you can resort to one of a wide range of digital cameras, from the “big names” valued at many thousands of dollars, to the most modest webcams. In Europe Philips is the preferred trade mark, replaced by Logitech’s in America. Be cautious and do not acquire for your microscope any camera with less than 1.3 megapixels. Study with care the many articles on this theme published in Micscape. If you can read French, a detailed view of the posts in the Forum Microscopies (see the link in Micscape front page), could be of your interest, long theoretical discussions and detailed camera presentations has been published. You can work without a photomicrography camera, even if one of them can make things more easy for you. But if no camera at all, you can draw, as hundreds of rotiferologists did fruitfully in the past. They study and document with drawings and descriptions almost all the actually known species.

If the rotifer is more or less immobile, take the needed images to later compose a whole body mosaic. In all the cases it is highly useful to make z-arranged stacks, open the pictures in the desired order, and using the Screen Grid function of your image processor, make detailed drawings, even if some of them are only sketchy ones. When you have a clear concept of the relationship of the organs, you can select and arrange the images of the stack to apply CombineZ (in any of its later versions) or to use instead Helicon-Focus.

When studying a new specimen it could be useful to adhere to the following list, (It is not exhaustive, even if in many cases could be excessive).

Write and draw all that you can, you will verify that this is a very useful approach. Don't forget that, with every note, drawing, or picture, you must record the date and, if it could be important, the hour. Record also the microscope, objective and illumination technique, the technique used to record the data, any other instrument involved. Put a scale on the picture or drawing now. Not a general statement about a group of images, but an individual scale. You don’t know when, or what kind of phenomena can hit tomorrow, and separate you from your materials and notebooks. Record all you can do for your own benefit, or to make your laboratory work comprehensible to other scientists. And don’t worry about the word. If you work seriously along this proposal, what you are doing is SCIENTIFIC WORK (yes, let stand the capitals) even if you don’t have the ultimate tools to do the publications, or you don’t work on “The Big Problems” of Physiology, or Evolution, your contribution to taxonomy and zoogeography will be recognized and stimulated. These two areas are those which need more of the push up that the amateurs can give. Many, many years ago, Huxley states that “Taxonomy is in the hearth of Biology”.

General data

First of all register the origin of your sample, describe the habitat in your notebook, record, if you can, latitude and longitude (NO, now you don’t need complicated apparatus, nor even expensive topographical maps. You need to use your FREE “Google Earth”?) In some cases and in many countries you can even include in your files one or more screen captures to record the locality. Record any data you can obtain about the source (this includes of course physical and chemical data if you can) …But, surely you have your computer, your word processor, your Photoprocessor, your digital camera and your Google Earth. But a kit of chemical tests, even for aquarists, is not in the standard equipment of the amateur: search in your wallet if you can afford the expenses, this would be a great addition to your lab. Use your digital camera to file some pictures of the site. They are invaluable aids today, and especially month or years in your future.

External anatomy

            It is necessary to describe their appearance and activities when eating, crawling and swimming.

Body structure - Verify the shape, in dorsal and lateral view, and the relationship between the pseudo-segments. Measure the length of the foot, trunk, neck, and the segments of the head.

Cuticle - Write down the appearance and thickness of the cuticle, and the presence of folds, longitudinal furrows, or reliefs. If they exist, note the number, disposition, form and size, of the cuticle thorns, or any projection, warts, or papillae they have.

Head and corona - Verify the form of the head, its width, and the shape of the “corona”, its size with respect to the head when they are displayed, the order in which the trochas opens, pedicel length, and its mobility.

Superior lip - While the specimen eats, verify in detail the shape of the superior lip, especially in dorsal view.

Antenna - Verify the position, form, and length of the antenna (in lateral view).

Eyes - Verify the presence or absence of eyes, color and position (on the brain or at the end of the rostrum or proboscis, or at any other situation).

 Foot: number and shape of segments, form and details of union with the trunk.

Spurs: Shape and length, insertion on the foot, and width at their base, separation among them, and any mark that they have.

Toes: number, and disposition. Absolute and relative lengths, among them and with the spurs. (It is better to study the bdelloid in a hanging drop, over an o-ring or well slide. It sticks to the coverslip and one can see much better its foot and toes, or any other adhesive system it has).

Adhesive disc: (if present) position, orientation, structure.

The study of the foot could take a long time, and a lot of effort, most of the time it is hidden between algae or detritus. A very fine pipette to isolate some clean specimens is a must.

Internal anatomy

        Position, form and size of the brain

        Form and disposition of the sub-cerebral glands and the retro-cerebral sacs

        Mastax: position, form and size

        Trophi: form and size of each piece. Teeth, dental formula (100x)

        Gastric and salivary glands: aspect, size, position.

Stomach (Intestine): form, size disposition presence or absence of a lumen, (it can be necessary to verify it at 400x in some compressed individuals). Only in the Habrotrochidae there is no lumen

Intestine (rectum): location, size.

Feces: if the opportunity allows their observation. In Habrotrochidae the feces are pelleted, similar to the gastric content, in the rest of the bdelloids they are a loose material

Protonephridia: 40x and 100x; compressed; they could be easily overlooked, look for the flame cells.

Bladder: shape, position, size, relationships. Time for repletion and voiding?

Foot glands: also called cement glands (number and disposition) 10 and 40x with extended foot.

Gonads (germovitellarium): made from two syncitia:

Vitelogen, The big nuclei that are normally seen even at moderate powers.

Ovaries (Small nuclei between the nuclei of the vitellaria. They are only well seen in compressed individuals.)

Oviduct: one, common to both germovitellaria, difficult to see or to identify.

Developed eg:, presence, size (in oviduct).

Uterus (in the viviparous species): mostly indistinguishable.

Position of the opening of the cloacae: Shape of the near segments if they have any interesting characteristic.

Egg (layed) - Form, size, type of covering and any sculpture on the surface.

            If it can be recorded: time to eclosion. You must separate some females into a solution innoculated with bacteria, with very few gross particles, in a capsule protected from evaporation, and record the laying time, the egg structure and the time to eclosion.

As far as is possible all the studied details would have to be photographed.

Probably the specimens that are used for the complete study do not survive the treatment.

If pictures are not completely satisfactory (as rarely they are) one can make drawings based on them, using as a guide the screen grid which can be displayed over the picture in almost all the image processors. Complete your drawings with details from live specimens

Some times ago I published in Micscape an article on the utility of the drawing for the microscopists

If a specialist can appraise most of the pictures, drawings, notes and measurements here advised, (and it would be a lot of good information) they will have very little difficulty to classify the material, even if this could be new to science.



Present situation

Claudia Ricci and Giorgio Melone in their work of 8 years ago (2000) hoped that their paper would wake up the interest of amateurs and professionals on Bdelloidea.

The reasons that prevented a fast concretion of this desire are also explained by the authors: small size, excessive mobility, lack of useful media to slow or anesthetize them, incredible speed for total contraction, difficulty to stop them by compression without badly deforming them, necessity to study the individuals for hours, alive, to be able to decipher their organization.

The books with suitable keys and descriptions are mostly out-of-print editions, and it even influences negatively the fact that outside Europe few students have a good knowledge of German, because the fundamental work on Bdelloids is

Donner. J. 1965. Ordnung Bdelloidea. Bestimmungsbücher zur Bodenfauna Europas, 6, 297 pages.

This last difficulty could be partially resolved if this book is translated to English and or French.

This illustrated Key, which we include today in the Micscape Magazine tries to make available to students, in direct visual form the morphological basis that allows separation of the Bdelloidea genera.

It is a pity that I cannot include a picture of each genus, When I found them I took some drawings from Internet. But even then there are some for which I could not find images (I live far from the scientific libraries, and on the Internet there are not many images).

But it will continue being true that their does not exist a sure medium to anesthetize the Bdelloids. Apart from Rousselet's, any tried technique gives as a most probable result the contraction of the rotifers so fast and complete that it disqualifies any further study, except for the recovery of the trophi by dissolution of soft parts with hypochlorite. In this Class the trophi are not generally a very important specific character, mostly confirming, better than defining, the determination. But the Italian team at the University of Milano is assembling an important library of trophi images captured with the SEM and this could change things in the future.

Nevertheless there is a tool that surely will facilitate the investigation of the species of Rotifera. The professionals of course, but now even many advanced amateurs, have incorporated to their equipment not only the powerful Nomarski DIC microscopes, but also the photography with electronic flash.

Figure 2, of the first part, which we reproduce below by courtesy of its author, Charles Krebs (who with it inaugurated a new approach that surely will attract the efforts of many other amateurs) illustrates the magnificent results that augur this technique. We add other pictures by Krebs and Abel Lear, and others shown below, which confirm that.

The availability of flash, digital photography in hi-res, and the now popular programs for three-dimensional reconstruction (CombineZ5, CombineZM, Combine ZP, and Helicon Focus (a and b methods)), augur an important generalization of the capacity to investigate and to document this difficult group of microinvertebrates.

Amateurs have an important opportunity here to collaborate with the professionals (who have the suitable bibliography, and the scanning electronic microscopes) sending to them high quality descriptions, measurements, and detailed photos of the species they observe. And this can be made on a world-wide scale thanks to the aid of Internet overcoming the geographic barrier.

I hope that the insatiable curiosity of present and future members of the forums of microscopists, will produce an ample harvest of documents to make them available to the specialists.

Nevertheless, it is clear that, due to the exigencies of bibliographical availability, and access to electron microscopes, the specialists in the Universities and Research Institutes, will continue to be the ones who can identify with certainty the species of bdelloids, and while the number of those does not increase substantially, and the photomicrographic documents harvested by amateurs don’t run fluently: “The biogeography of the Bdelloidea, (also in general for all Rotifers – author’s note) will continue to be, really, the biogeography of the students of the group.” (Ricci and Melone, 2002)”

To facilitate the contribution of amateurs to this discipline I add at the end a list of the more active specialists I know. I think that most of them would be glad to receive taxonomic novelties from all over the world.

SENDING SAMPLES - If the amateur does not feel himself able to accomplish the more difficult tasks, he or she can make a huge contribution by sending samples to the specialists. Remember you are working with species that (almost all) can survive after desiccation. Using a piece of laboratory filter paper (or coffee-makers filter paper) you can prepare a sample that could withstand the hazards of even the surface mail, and can be included in any mail envelope. THIS IS SIMPLE AND VALUABLE.

But remember that a researcher is a very busy person. Take care to contact first the specialist of your election to ask for their permission to send your materials, pictures and dehydrated samples.

If after seeing your notes, pictures and drawings, the scientist tells you that your material is of real interest, prepare a desiccated sample to send her or him some specimens.

How to prepare the sample

Concentrate as many as you can of the individuals in a little volume of water. Prepare a Petri dish, or a similar flat dish with a cover, with a piece of filter paper on the bottom. Lightly wet (not flood) the paper with some drops of distilled water. Deposit the sample with the living rotifers in the center, and cover, letting a thin open gap to allow evaporation. Dryness must be attained in 4 or 5 hours. Cut the dry filter paper to recover the area with the sample and put it inside a folded paper. Include this with the letter you send with data. Don’t use plastic for the holding paper or the envelope. There is a good chance that the individuals in your sample can be easily recovered in the destination laboratory.

Even if you don’t result being the happy parent of a new species, your sample could have zoogeographical and ecological interest. If you are doing that you are a very careful amateur, and of course you know that a full series of data must be sent with the sample.


    Jersabek, C.D., H. Segers, and P.J. Morris, . An illustrated online catalog of the Rotifera in the Academy of Natural Sciences of Philadelphia (version 1.0: 2003-April-8). [WWW database] URL

    Diego Fontaneto and Claudia Ricci – 2004 – Rotifera: Bdelloidea, in Fresh water Invertebrates of the Malaysian Region.Pg 121-126

    Edmondson, W.T. 1959 – Rotifera. Pages 420-494 in W.T. Edmondson (ed.), Ward and Whipple’s Fresh-water Biology, Second edition. John Wiley and Sons, Inc. New York, NY.

    Fontaneto D., Boschetti C. and Ricci C. – 2008 – Cryptic diversification in ancient asexuals: evidence from the bdelloid rotifer Philodina flaviceps. J . Evol. Biol. 2: 580–587

    Haigh S. B. 1963 - Notes on the study of bdelloid rotifers. Quekett J. Microscopy, 29: 133-138.  

    Hendrik Segers – 2007 - Annotated checklist of the rotifers (Phylum Rotifera), with notes on nomenclature, taxonomy and distribution. Zootaxa 1564- Magnolia Press

    Hyman, Libby H.1951 – The invertebrates – Acantocephala, Aschelmintha and Entoprocta, McGraw-Hill book Co.

    Ricci C. and Melone G. – 2000 – Key to the identification of the genera of bdelloid rotifers. Hydrobiologia 418: 73–80.

    Ricci C., Melone G. and Walsh E.J. – 2001 – A carnivorous bdelloid rotifer, Abrochtha carnivora n.sp. Invertebrate Biology 120: 136–141.

    Claudia Ricci, Manuela Caprioli and Diego Fontaneto 2007 – Stress and fitness in parthenogens: is dormancy a key feature for bdelloid rotifers? BMC Evolutionary Biology, 1-7(Suppl 2)

    Ricci Claudia y Giulio Melone1998 – The Philodinavidae (Rotifera Bdelloide): A special family. Hydrobiología 385:77-85

    Ricci Claudia, Russell Shiel, Diego Fontaneto & Giulio Melone– 2002Bdelloid rotifers recorded from australia with description of Philodinavus aussiensis n.sp.

    Bdelloidea specialists: In view of the possibility of annoyances to the specialists by mechanically sent spam, finding their e-mails addresses is left to the initiative of the advanced amateurs.  Communication methods are mentioned in their bibliography, and most titles are published in Internet in PDF format.

    Caprioli, Manuela  (Italy)    

    De Smeth, Willem  (Belgium)    

    Fontaneto, Diego (Italy, UK)      

    Melone, Giulio (Italy)       

    Ricci, Claudia (Italy)         

    Schmid-Araya, J.M. (UK)  

    Segers, Hendrick (Belgium)       

    Song, M. O. (S. Korea)

    Sarma, S.S.S.  (Mexico)



1 – eyes at the end of the rostrum in Rotaria, bright field – Oliver Barth

2 – head of Philodina – electronic flash – Charles Krebs

3 - another head of Philodina – bright field, electronic flash, by Charles Krebs, showing the red eyes

4 – Philodina megalotrocha – DIC – a picture by Abel Lear

5 – Another DIC image of the head of a probable Philodina, by Abel Lear

6 – Adineta cf. tuberculata, bright field, Michel Verolet

7 – a view of the head of A. cf. tuberculata, by Michel Verolet

8 – dorsal view of the head of the same, bright field, Michel Verolet

9 – bright field at its best, detailed anatomy of the head of Adineta, showing the complex rostrum Michel Verolet

10 – The new megapixel digital cameras allow detailed pictures of active animals, and even movies at a good resolution. Adineta, by M. Verolet

11 – Habrotrocha attached to a bryophyte, picture by M. Verolet

12 – one of the attached examples - a stretched Habrotrocha, M. Verolet

13 – Rotifer’s eggs glued to an algae filament, M. Verolet 

14 – One of the eggs, the ramate trophi confirm it is from a bdelloid 


Comments to the author, Walter Dioni , are welcomed.

Editor's Note: This three part article by the author was first published in French on the Microscopies Magazine website.


Studies on genetoxicity, carcinogenicity and reproductive toxicity haven’t shown positive results, neither in experimental toxicological nor epidemiological studies made in hospital workers.
With thanks to
Aydin Örstan.

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