formalin, no mercury fixatives.
The title image is a very young mayfly (Ephemeroptera) larva, fixed in 70% alcohol and mounted in PVA-Glycerol. Imaged with the 4x planachromatic objective on a National Optical microscope with a COL (circular oblique illumination) filter. The field image width was 3.4 mm. This and all the other images in the article were captured with the digital camera integrated onto the microscope.
I want to
discuss here three very well known fixing methods of quite
general use which can be complementary to the formulas
that I presented in the
first part. They
are fixatives for the use of biologists and microscopists,
rather than histologists. Although it is true that certain
fixatives used by the histologists can be very useful for
'microzoologists' and 'microbotanists', I think it is
better to present the histological fixatives without toxic
products in another article, if I have success in the
tests which I am currently making. (I speak of course of
non-commercial histological fixatives, because there are
several commercial products which are sold at very high
prices for the professional laboratories, and which are
Chydoridae, fixed with 70% alcohol, wet mounted in
1:3 glycerin-water. Three image mosaic. Captured
with the x10 objective. Half reduced. Background
Young Culex mosquito larva. Fixed with 70% alcohol
many months ago. Mounted in PVA-Glycerol.
Captured with the 4x objective. Width of the field
of view 3.4 mm. COL filter.
Alcohol 70% is a standard fixative for many zoological groups, the arthropods in particular. It behaves well with insects, arachnids, crustaceans and larger animals. It is safe (if you do not drink more than two glasses of wine per day!) and the fixed samples are very durable. But it has a great defect. It dehydrates specimens and contracts the tissues a lot, and, employed alone, it is unusable with the smaller microfauna which most of the time are unrecognizable right after fixing. You can judge by the included pictures that it is not the best fixative for the smallest fauna.
If you want to have your sample fixed in 70% alcohol your technique will vary according to the specimen. But the general rule is to estimate the volume of the material (plankton, sediments, algae, etc.) to be fixed, including in the estimate the sample water volume. Then you add sufficient alcohol at 90% or 96% to obtain the desired percentage. The table below is a guide to make dilutions at 70%. Do not be surprised by the figures. The table takes account of the quantity of water in the added alcohol. The Total volume is included as an aid for you to place the sample to be fixed in a suitable bottle.
alcohol at 70% could be appropriate only for samples that
you can drain almost completely. Insects, small fish,
tadpoles, or even entomostraca collected with your net and
well drained. As the water of the sample will dilute your
alcohol, do not forget to do one or two changes with fresh
alcohol after 12 to 18 hours.
Alcohol is a "macroscopic" fixative. It does not fix the nucleus well. To improve fixing, acetic acid is added. The acid fixes very well the nucleus and moreover it expands a little the tissues, counteracting the action of alcohol.
Wolman has proposed a formulation to fix the sections made with the freezing microtome:
Acetic acid..............................5 ml
This formula is certainly of a more general use.
But, if you neither intend to undertake histology, nor make taxonomic descriptions with very exact dimensions, you can use only alcohol.
A chlorophycean algae, with a gelatinous envelope
of a species unknown to me. x40 objective.
Somewhat scaled up. Very cropped image
Two specimens of Euglena deses, objective
x40. 70% alcohol. Mounted in the same. Captured
inside the empty valves of a cladoceran. Cropped
Methyl alcohol (methanol) which once was cheap, was formerly used mainly as a denaturing agent of "good" ethanol, to prevent ingestion. Do not drink methyl alcohol, it is fatal. But in many recent works it is recognized that methyl is really as good as or better as a fixative than the ethylic, and it is now proposed as a total or partial substitution in traditional formulas.
One of them is Carnoy’s, which is today named Methacarn, with its modified formula:
Becuse of its fast penetration, Carnoy, and now
Methacarn, are used for chitinized organisms, and tissues
used in genetic investigations (vegetable roots, anthers,
buttons of flowers, testicles and ovaries, eggs of
insects, etc.). But also to fix smears of cytological
materials, or free living and parasitic protozoa. The
alcohols used in these formulas, both the ethyl as well as
the methyl are recommended to be absolute. Apart from
being very expensive, the absolute alcohols are really
almost impossible to be maintained in the amateur’s
laboratories. They absorb water from the air almost from
the moment at which the bottle is opened. 96% alcohol is
almost as good.
Actually some researchers reject the ethylic and use exclusively the methylic (a little cheaper) or the isopropyl alcohol (a little more expensive).
As they are interchangeable you can see that the commercial alcohols denatured with another alcohol are perfectly adequate for amateur microscopy. It is not the same when the additive is a vegetable essence or another oily substance which mixed with water becomes opalescent.
that hot water (heat really) was recommended for the last
30 years of the former century (yes, the twentieth!), but
was only really accepted within the last 20 years of the
century. Most helminthologists usually employ it. It is
said that the treatment by hot water fixes the animals in
their normal aspect, in extension, without deformation,
and keep their true dimensions, which is very important
for good taxonomy.
Diatom, the day after being fixed with hot water.
Euglena deses. x100 0I. Hot water.
Cropped pictures from wet mounts sealed with
You can put your creatures on the slide in a drop of water
and heat from beneath with a match or a lighter. The
objective is to obtain a temperature in the neighborhood
of 50-60ºC. The tardigrades, the trematodes, the small
cestodes, nematodes, leeches, and others
micro-invertebrates respond very well to this operation.
With the larger helminthes you can even heat the fixative
and pour it on the worms in a dish.
For the smallest protists you put the sample, preferably concentrated by filtration, in a container which accepts four times the same volume. You wait until the animals (rotifers, gastrotriches, nematodes, protozoa, etc.) resume their normal activity, and you suddenly add a double volume of almost boiling water. As Edmondson notes (1959), organisms in the region where you poured the water are certainly overheated, and the edge of the container did not receive enough heat, but a ring located between the two zones will present animals in a state of perfect extension. Yes, you do have the task to seek them out drop by drop, but Edmondson notes that it is the only method which makes it possible to fix some specimen of Notommata (a very difficult to fix rotifer) with its "auricles" spread out well.
Epiphanes, captured x40, mosaic of two
Mytilina, captured x40, amalgamated with
Cyclopid copepod, captured x40. Mosaic of 10
images. Original is 1028 x2000 pixels.
I have lost the original pictures of these two
beautiful rotifers.So I present only the two icons
Click over the image
to see a 55% reduction of the original
Generally after fixing by hot water, one makes a post fixing with a conventional fixative (alcohol 70% for entomostraca, lactocupric for the smallest invertebrates, etc. according to the species present in the sample). I always do this, and I advise you to do the same.
The professionals and some amateurs as well, fix some materials, (primarily for histology), with the microwave oven. This certainly requires a "laboratory oven" or the careful calibration of your domestic one. It is possible to do it. I calibrated mine.
An amoeba at 100x killed by the boiling
But the microwave times for the very small quantities of the materials used are so short, and the adjustment so difficult (because of the irregular distribution of radiation in the space of most of the domestic ovens), that I have given up after having cooked many nematodes, rotifers and protozoa. The oven is better used to heat the water or the fixative which you would use, or to dry the border sealing your coverslips (for this last application be very careful, start with a time of not more than 5 seconds of exposure at 100% power). If you want to heat a larger sample ensure that the rise in temperature (even if it is reached in 2 minutes) is gradual and not instantaneous as when you add almost boiling water, the surprise factor is lost and your subjects can contract.
crystals………………….. 5 g
Keep in brown or opaque bottles. Add 0.5 ml to 100 ml of
All those who work with plankton samples know the so-called "Lugol's solution" which is employed to fix such samples. It is really Rhode’s fixative (Lugol's solution almost 10 times concentrated + acid acetic, or potassium acetate) which is employed. It is very suitable to preserve phytoplankton, protozoa and rotifers in the plankton, to which iodine (a heavy element) adds weight, and cause their quick precipitation, allowing its concentration.
Euglena prob. oxyuris, x100 OI
Collage of microflagellates x100 OI, taken from
several pictures. Click the
image to see a labeled copy
Phacus pyrum, x100 OI
The planktologists take their samples, put sub-samples in
special cylinders and let all the organisms precipitate to
the bottom. The cylinder bottoms have the thickness of a
coverslip and allow the observation and counting of the
organisms with an inverted microscope.
Don't you have an inverted microscope? Well, here is one of the techniques which you can apply to study the very interesting ultra-microscopic phyto and zooplankton that pass through the usual plankton nets. Use a bottle or other tall container of 1 or 2 lt. preferably a glass one, and with a flat bottom or conical bottom. I have even used inverted bottles of soda beverages, without its bottom. Add 5 ml of Rhode's to the sample, mix it well and leave in darkness for 2-3 days. With a flexible tube improvise a siphon and discard supernatant water without agitating the sediment, leaving at the bottom a tenth of the initial volume. Now stir up, and pour the liquid in another suitable tall and cylindrical flat or conical bottomed bottle. After two days draw off most of the liquid, and take drops of the sediment at the bottom to examine them under your microscope.
You will be astonished by the great quantity of micro phytoplankton species and even by the very small micro zooplankton which you can now detect.
Chaetonotoidea gastrotrich x100 OI.
Click the image to
see a labeled and bigger copy. This is not a
normal image of a fixed gastrotrich. Normally they
roll-up and become almost illegible. This one was
very compliant. This is a central optical section
that shows most of the anatomy details of these
The samples fixed with Rhode’s can be preserved for a few weeks, protected from the light. But the organisms take a brown color which can obstruct the observation. To restore to them their transparency add some small drops of a 5-8% potassium hyposulfite solution.
For those who will carry out only some tests I give here a formula which has given me good results, but which depends on the quality of the medicinal tincture of iodine that you can find in your pharmacy.
I start from a commercial solution of formula:
crystals…………….……. 1.2 g
I add 50 ml of white commercial vinegar (5% solution of acetic acid)
My final formula is:
Which is more or less a Rhode’s diluted 4 times, by
taking account of the really active ingredients. For 100
ml of sample I add 2 to 4 ml of this alternative formula.
If you want to test this way study the formula of "your"
medicinal tincture of iodine and proceed in the same
A miscellanea of decanted bacteria and
NOTE: when you work with the other fixatives, the cilia of ciliates and other invertebrates are almost invisible. You can often highlight them by adding a simple trace of Rhode's to your fixed wet mount. Be careful: only one trace. This can highlight the cilia, the nucleus and flagella also.
other safe formulas and methods, that generally have more
specialized uses, but we have reviewed various
alternatives without any dangerous ingredients, which when
used with wisdom will allow you to work with an extensive
number of microalgae and microinvertebrates
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