To produce prepared microscopical slides of good quality often the refractive index of the mounting medium is often required. In case of stained sections the refractive index should coincide with that of the stained material, that means, the unstained material should be invisible in the mounting medium to suppress the blurring "refraction image". On the other hand, in case of diatoms the refractive index should be significantly higher than 1.45, which is the refraction index of the frustules. The difference should be 0.3 units at least. When investigating cruder forms like radiolarians or foraminifers, a smaller difference is desired to increase transparency. Unfortunately, Abbé refractometers are outrageously expensive, but the much simpler methods will also do..
Fix a vertical stick of 30 cm on a plate of wood of about 20x20 cm and cover the plate afterwards with a sheet of graph paper. Then fix a cavity slide horizontally tothe stick. Cover one quarter of the cavity with a coverslip and fix it in this position with a hot mounting medium (Fig.1). Now the cavity together with the coverslide forms a small hollow prism. Later, one drop of the liquid to be investigated will be put into this prism.
Measuring procedure
Fill the cavity with the liquid under discussion and move your eye in such a way that the zero-mark on the graph paper is seen at the very rim of the cavity, but just outside (blue line). Then move along the graduated line with the tip of a pencil and stop when the tip just appears inside the rim (red line). Note the position the tip of the pencil has touched. Using a calibration curve the value so obtained defines the refractive index. If the index of the medium is smaller than that of glass the two lines displayed in the graph are swapped. In this case the value obtained must be counted negative. If both indices are identical, both lines coincide. Of course, first a calibration curve must be established. This is done by using liquids of a known refractive index. Usually this calibration curve is a straight line.
| Medium | nD |
| Water | 1,333 |
| Ethanol | 1,361 |
| Isopropanol | 1,378 |
| Chloroform | 1,449 |
| Toluene | 1,496 |
| Methylbenzoate | 1,517 |
| Clove oil | 1,544 |
| a-Bromonaphthalene | 1,656 |
| Methyleniodide | 1,744 |
This method requires a microscope with a mechanical stage and a condenser with a high aperture (three-lens system) which can be adjusted vertically with a gear drive, an eyepiece with a reticule (graduated scale), an object slide prepared in the way explained above and a special filter: Fix one halve of a razor blade to a filter glass in such a way that the cutting edge corresponds exactly with the diameter and put this filter into the filter support of the collector with the fixed blade in an upside position. Use a clear green filterglass or a yellow filterglass in combination with a sodium vapour lamp, if available, to suppress colour fringing.
Measuring procedure
The measurement is done with a 20x objective, if possible with a 40x objective or a 60x objective. In theory high magnification is preferable, but often blurring through colour fringing decreases accuracy. So the optimal magnification must be empirically tested.
Adjust the objectslide in such a way that the optical axis of the system passes the cavity from outside, focus on the upper surface of the slide, open the condenser diaphragm and move the condenser vertically, so that the sharp edge of the blade is distinctly in focus. Remember the position of the edge on the reticule of the eyepiece. Move the object slide until the optical axis of the system just passes the rim of the cavity from inside. The position of the edge will then switch to the left (negative difference) or to the right (positive difference, fig.2). Remember the new position. The difference of the two positions thus obtained presents the refractive index. Of course first a calibration curve must be established.
You need a cell as shown in fig.3; the distance of the inner surfaces should not exceed 0.5 mm. Treat the surface of the objectslide gently as well as the surface of the coverslide with an abrasive, then rub some black ink into the scratches thus obtained and clean the surfaces afterwards. These prepared surfaces must face one another. The coverslide must be fixed with two clamps, otherwise it will float, if liquids of high density are inspected. Fixing the slide with a mounting medium is not a good idea because in this case it is difficult then to clean the cell.
Measuring procedure
You need a microscope with a graduated micrometer drive. Make sure that this drive is linear and independent of the starting position of the stage! When measuring the rotation of the micrometer drive, keep in mind that the zero-position may be passed!
The method itself is simple: Determine
the virtual distance between the two surfaces by focussing on the black scratches.
Using a calibration curve you will easily find out the refrative index.
Measurement of the refractive indices of mineral crystals
The principle is very simple: Mix ethanol and bromonaphthalene (e.g.) to produce a series of media with a known refractive index, then find out the medium in which the mineral crystal just disappears visually. Unfortunately, this method is very inaccurate.
Using the "Line of BECKE" is much nore accurate. If you look at a grain of sand mounted with water you can see a bright line surrounding this grain. It is called "the Line of BECKE". If you lower the eyetube this line moves into the the surrounding water, the medium with the lower refractive index (LLL-rule: lowering leads to lower index). In case of Bromonaphthalene this line will move inwards instead. Varying the mounting medium, you can fix the refractive index quite accurately, because, if there is no difference, there is still a weak line which is displaying many colours. This is the consequence of dispersion: In case of short-wave light (blue and green) the refractive index is always higher than in case of long-wave light (yellow and red), the difference depending on the media. Let us assume that the indices are identical in case of green light, and that the diffraction of the sand is higher than that of the mounting medium. Then the blue line will be inside the grain whereas the red line will be outside. The method explained here is very useful, but it is useless in case of very high refractive indices, as no proper mounting media are available then.
The
refractive index, measurement and application
If you are interested in this topic we refer to:
The
refractive index, measurement and application in microscopy and in polarized
light microscopy
pdf-file 0.5 MB, 25 pages, 24 figures,
German language.
High-refractive mounting media (PLEURAX / NAPHRAX / ZRAX)
Information
published on the net has been checked by synthesising the above mentioned mounting
media several times with slightly different methods. In case of NAPHRAX an improved
synthesis is given. Here are the results:
Dr.G.Rosenfeldt
INTRODUCTION
The refractive index of a substance depends solely on the number of easily movable pairs of electrons per volume unit. The higher this number is, the higher is the refractive index. Therefore the refractive index increases in this order: polar aliphates < non-polar aliphates < non-polar aromates. In addition large atoms increase the refractive index markedly: oxygen < sulfur, nitrogen < phosphorus < arsenic, chlorine < bromine <iodine. For non-polymers this means: the higher the refractive index of a monomer is, the more toxic, the more volatile and the more sensitive it is to oxidation. The toxic and volatile methylene iodide is a good example of this rule, and also the ingredients of extremely high refractive mounting media: triphenylarsin or arsenic sulfide. So it is improbable to sythesise polymers whose refractive index exceed significantly that of PLEURAX when using "civilian" precursors.
PLEURAX
WARNING! WHEN SYNTHESISING PLEURAX SOME TEN LITERS OF H2S ARE SET FREE. THIS GAS IS AS TOXIC AS PRUSSIAN ACID! SO THE METHOD "GARDEN PLUS WIND FROM THE REAR" IS A LETHAL RISK! THE SYTHESIS MUST BE DONE IN AN APROPRIATELY EQUIPPED LABORATORY WITH AN EFFECTIVE FUME HOOD.
Melt 110 grams of crystalline phenol in a 250 ml beaker on a magnetic stirrer. Then add 40 grams of powdered sulphur and slowly heat the mixture up to 150 oC (internal thermometer!). Add a spatula tip (approximately 100 mg) of anhydrous (!) sodium carbonate (catalyst) and raise the temperature to 170 oC, stirring continuously. At 160 oC the reaction starts with gentle foaming and H2S is set free. Cover the beaker with a piece of cardboard to avoid the evaporation of the phenol. Check the production of H2S with lead acetate paper from time to time.
Stir for 4 hours. From time to time take a small sample with a glass rod and dissolve the product in a test tube containing about 5 ml of isopropanol. The sample must be solved completely, otherwise sulphur will still be present. After about two hours the reaction is nearly complete resulting in a dark brown resin, which is yellow in thin layers. Continue heating for four hours and check the production on H2S with wet lead acetate paper. If even then some sulphur should be left add some more phenol and sodium carbonate and keep heating. The reaction temperature is not critical, but should not exceed 170 oC, otherwise too much phenol will evaporate and sulphur will spoil the product. When the reaction is finished remove the cardboard and heat for another hour to remove phenol. The volume will decrease significantly. The resulting product still has the smell of H2S, but this odour disappears with time. Some phenol in excess is no problem but will decrease the refractive index.
Allow the resin to cool down to 100 oC, continue stirring, then add 50 ml of isopropanol. After a homogeneous solution has formed, pour it into 50 ml vials, filling them to two-thirds. If the viscosity of the cold mixture should be too high, add some more isopropanol and heat in a laboratory oven to obtain a homogeneous solution.
Yield of pure resin: about 60 grams.
After MELLER, MIKROKOSMOS, slightly modified.
SOME COMMENTS ON PLEURAX
1. Phenol is usually coloured slightly red by oxidation products. These impurities do not hamper the synthesis, they even act as a catalyst. Freshly distilled in vacuum phenol is colorless, but in this particular case not reactive enough.
2. Allow exceeding sulphur to crystallise, then pour the now sulphur-free PLEURAX into another vial.
3. PLEURAX is soluble in isopropanol and acetone, but not in toluene or xylene.
4. When preparing diatoms, put a small sample on an object slide, let it dry completely, cover it with one drop of isopropanol to remove the air from the frustules, then cover with PLEURAX, add a coverslide and remove the solvent by heating. Only dry PLEURAX has the high refractive index wanted. The yellow colour is no drawback.
5. The long term stability of PLEURAX is unknown. PLEURAX consists of benzene rings with OH-groups connected via sulphur bridges. Such a polymer may be susceptible to oxidation: sulphur bridges may be oxidised to SO2-bridges (clouding of the slide), phenolic OH-groups may be oxidised to quinones (darkening of the polymer). Whether such changes actually occur in the course of time is unknown. Protecting the slides with a ring of varnish is recommended.
6. PLEURAX is said to have a refractive index of about 1.73. According to my measurements the refractive index is 1.68. This diversion may be a consequence of the grade of polymerisation dependent on the quantity of sodium carbonate used (smaller quantities should enhance the grade of polymerisation). PLEURAX has a high dispersion and shows fluorescence when blue light is used.
7. PLEURAX is unsuitable as a mounting medium for UV-microscopy because wavelengths below 400 nm are blocked.
8. The synthesis is safe and requires neither much time nor much equipment. The product is ready for immediate use without further treatment.
9. Due to the high toxicity of H2S an effective fume hood is required!
10. The synthesis of PLEURAX is preferable in comparison to NAPHRAX.
NAPHRAX
Improved synthesis
Substances
Glacial acetic acid
200 ml
Naphthalene 50 g
Paraformaldehyd 12,5 g
p-Toluenesulfonic acid (monohydrate) 5 g
Apparatus
500 ml flask, water bath, centrifuge, magnetic stirrer, distilling apparatus, heating mantle
Sythesis
Mix naphthalene, toluenesulfonic acid and glacial acetic acid and heat in a water bath until a homogenous mixture is obtained, then add paraformaldehyde, distribute the powder by gentle shaking and heat for 72 hours in a water bath at 95 oC. Cover the water with 1 cm of oil to prevent evaporation. Immerse the flask only partially. Insert a plug and put a piece of cardboard between the plug and the neck to allow vapours to escape, Shake the flask gently from time to time..
The praformaldehyd is completely dissolved after a few hours and after about 18 hours a colourless liquid resin starts to precipitate, which becomes solid even at 95 oC. Towards the end of the reaction the resin is honey yellow and opaque. Allowe the mixture to cool, pour off the liquid phase and wash quickly with toluene. Loss of material must not be feared because the resin dissolves very slowly in cold toluene.
Purification / Method 1
Dissolve the resin in toluene at 95 oC and heat until a clear honey-yellow solution is obtained. Add a teaspoon of marble grains for deacidification and stirr at room temperature (magnetic stirrer) at least for three days. The white powder precipitated during this time is centrifuged (filtration is not recommended). Then the toluene is removed by distilling (heating mantle!) along with a small amount of water and some acetic acid. The result is a honey-yellow, clear, hard resin. This is covered again with about 1 cm of toluene and again heated up to 90 oC . The solution so obtained is ready for use even if some traces of acetic acid should be still present.
Addendum: Wait for another two weeks to allow precipitation (the white precipitate will adhere on the inner surface of the flask), then pour the now crystal-clear solution into small vials.
Purification / Method 2
Pour the crude product, dissolved in toluene, into a separating funnel and add about 100 ml of water. Shake gently. After a few hours three layers will separate: the top phase contains the resin dissolved in toluene, the bottom phase is a mixture of water and acids. In between you get a stable white paste-like emulsion. Sincs this paste cannot pass the tap of the separating funnel the toluene layer must be siphoned off. Stir the separated resin solution for several hours with granular calcium chloride to absorb water, then the toluene will be largely removed by distilling. Then proceed as described in the section "Method 1". The solution is free of acids, it secretes no precipitate even after a long time, but this method is associated with a loss of material because of emulsion formation.
SOME COMMENTS ON NAPHRAX
1. The method published recommends 20 ml HCl conc. and 5 ml of H3PO4 conc. as a catalyst. The resin so obtained is nearly black and the solved resin exhales HCl for a long time. The method does not avoid the formation of the white precipitate. After deacidification and the removal of the precipitate NAPHRAX thus produced is quite usable though - in thin layers the dark colour is of no disadvantage - but the resin may still contain reactive CH2Cl-groups which may lead to aging. The use of p-toluenesulfonic acid represents a significant improvement!
2. The entire synthesis can be done in an apartment. But the synthesis is very time-consuming, the raw product must be purified in any case and a variety of laboratory equipment is needed.
3. The composition of the white precipitate could not be determined yet. The IR-spectrum indicates a mixture of several substances, paraformaldehyde is definitely absent.
4. When preparing diatoms, put a small sample on an object slide, let it dry completely, cover it with one drop of toluene to remove the air from the frustules, then cover with NAPHRAX, add a coverslip and remove the solvent by heating (100 oC). Only dry NAPHRAX has the high refractive index wanted. Thin layers of this mounting medium are nearly colorless.
5. NAPHRAX has a refractive index of 1.66, shows less dispersion than PLEURAX but is stimulated to fluorescence by blue light .
6. NAPHRAX is suitable for UV-microscopy, as wavelengths down to 340 nm are transmitted.
7. NAPHRAX consists of naphthalene rings linked together by CH2-groups - so the structure is similar to polystyrene. Such hydrocarbons are chemically extremely inert, the resin should have a high long-term stability. It is however important to remove the white precipitate completely (see above)!
ZRAX = NAPHRAX !
Original link: http://www.sas.upenn.edu/~dailey/zrax.pdf . This link is now inactive, and a new one could not be found.
ZRAX was distributed by Professor Dailey, but no information concerning the synthesis is available. The above source leads to the assumption that ZRAX is nothing else than NAPHRAX, synthesized under special conditions and carefully purified . Furthermore, the source provides the information that NAPHRAX may not have a long-term stability.
Meanwhile Mr.Matthias Burba gave me a small original sample of ZRAX, so an infrared spectrum could be made. This yellow resin is obviously identical with NAPHRAX produced with p-toluenesulfonic acid as catalyst. Whether the production method is completely the same must remain open to dispute. (The slightly different shapes of the IR-peaks can be explained by the different thicknesses of the samples - what is essential are the positions of the absorption bands).
The refractive index of ZRAX is 1.68 and thus is slightly higher than that of NAPHRAX. This difference suggests a higher degree of polymerization, combined with a slightly higher density.
Dr.G.Rosenfeldt
Optical staining - an impressive gimmick
You need a clear coloured glass filter fitting to the filter-support of the condenser, a condenser that can be adjusted vertically by means of a gear-drive, some circular cardboard-pads (4 mm - 12 mm they can easily be made with a core drill), coloured transparent varnish as used for glass paintings, black varnish and a rotatable stage used to protect microscopic slides with a ring of varnish. For testing purposes use starch dispersed in water.
First try to obtain a good dark-field illumination: Fix a cardboard-pad with a drop of glycerol in the middle of the glass filter and open the diaphragm fully. For weak objectives (up to 20x) it is easy to get a good dark field illumination this way. Once the appropriate diameter of the pad is established remove the pad, clean the filter and draw a ring of black varnish of exactly the same diameter around the centre of the filter using a slide ringing table (slide ringer). When the varnish has dried completely cover the inner opening with transparent varnish of different colour (see left fig.). The result is a dark field image with the background now not black but coloured. In the example given objects are displayed yellow against a green background. If the background is too bright reduce the internal opening by another black ring of varnish - this re-adjustment of the background brightness is critical!
Much more impressive results are obtained using a coloured sector disc (see right fig.). Such filters were available many decades ago, but then they were no longer produced, since this illumination is inferior to proper dark field illumination because of its lower contrast. Nevertheless, this kind of illumination is often preferable because the contrast of dark field illumination sometimes is too extreme. The colour combination is of course a matter of taste, adjacent rainbow colours are recommended.
The illustrated pdf_file
(6.8 MB, German language 
) is a short guide with various tips, which are not only important for
beginners. This small publication means to encourage beginners and amateurs
to make paraffin sections, to process them and to stain them.
The preparation method described here is suitable only for small worms, insects, arachnids and other small animals with a more or less stable cuticle. So first try to get adequate specimen! To do so, use the device outlined here ("BERLESE-funnel"). Fill it with a sufficient quantity of decayed leaves and humus from the layer which is located between the upper layer of dried leaves and the topsoil. The heat of the bulb and the increasing dryness of the upper layer of the funnel forces small animals to move downwards where they finally end in alcohol of 80% - what a nice way to pass away! Methylated spirit of 80% is sufficient. After about 24 hours the procedure is completed.
A BERLESE funnel can be easily made from cardboard, but sheet metal is preferable since this material transports heat much better to the lower part of the device. The lower opening should be closed with a wire mesh. Textile nets are inadequate as the animals will not pass this material.
Safety note: Never leave the funnel unattended as accumulation of heat in the then dry upper layer may cause fire!
To produce prepared micro slides you need the BERLESE mounting medium. This is a highly concentrated solution of chloral hydrate and gum arabic in water, which penetrates in a short time into the objects and dissolves almost all soft tissues, making the objects transparent. This process takes several days, sometimes some weeks, and it may happen that the objects become completely opaque for a few days first. The BERLESE mounting medium can be bought in shops for school supplies. We do not recommend to prepare the solution yourself.
The preparation method is very simple: As BERLESE solution does not tolerate alcohol, put the objects into water for one day to remove the alcohol completely, then position the wet objects on an object slide, cover them with plenty of BERLESE mountant and cover with a coverslip. In case of thicker objects put some splinters of coverslips between coverslip and slide. The bleaching process usually is completed after a few days, but then it takes at least another month until the mounting medium is solid. So keep the slides in a horizontal position. The objects cannot be stained because BERLESE dissolves or distroys almost any dyestuff, only soft tissue staining with "nuclear fast red aluminum sulfate" and similar mordant dyes (BECHER-dyes) are possible, but not very useful, as BERLESE solution leads to a severe damage of soft tissues. The coverslides need no ring of varnish. The prepared slides are stable, but temperatures below zero may cause irreversible damage, because the chloralhydrate will form crystals. In that case remove the coverslide by simply covering the whole slide with water for some days, then make a new prepared slide.
Glycerol gelatine ("GG") melts at about 60 oC. Since multiple melting cycles decreas the quality melt GG only once with hot water and then distribute the GG in small jars, which must be closed carefully. That way you can use small quantities each time. To avoid air bubbles during preparation remove the molten GG with a glass rod and not with a dropper pipette. The frequently recommended method to melt small blocks of GG directly on the object slide produces numerous air bubbles. Never heat GG with a flame or allow it to boil!
Adequate objects: Desmids, filamentous algae, moss, pollen, spores, protonemata, reproductive organs of moss, small crustaceans, rotifers with stable cuticle, small insects, small spiders, mites, ticks.
Preprocessing: The objects are covered with a solution of 4% formalin (10 ml of water plus 1 ml of formalin conc.) for one day. After having removed the chemical fixative the objects are transferred into a flat dish and are covered with at least 30 ml of a solution of 5 grams of glycerol in 100 ml water and then they are put in a dust-free place. After about one week most of the water will have evaporated. Occasionally the objects must be moved into the centre of the dish to avoid desiccation! This preprocessing is necessary to avoid shrinkage. Finally pure glycerol (80% for medical use) is added. In pure glycerol (anhydrous glycerol is unnecessarily expensive and less useful as well) the objects are stable and can be stored.
Sometimes it is preferable to transfer the objects after they have been washed directly from the fixative to an objectslide and to cover them with glycerol-water. When the water has evaporated they must be covered with molten GG; that way the GG will not be diluted by glycerol.
Staining: Only alizarinviridin and nuclear-fast-red when applied together with special mordants render stable stainings: In case of alizarinviridin chromium aluminium sulfate must be used, in case of nuclear-fast-red aluminium sulfate. Such solutions are for sale. The objects are covered with the diluted solution (about 1:10) for one day. Afterwards they are washed several times and then covered with glycerol-water. The objects will never be over-stained as these dyes mainly stain the surface and are not concentrated in the tissue.
Mounting: You need small coverslips (12 mm) and large coverslips (20 mm). Round coverslips lead to more beautiful prepared slides but are much more expensive. In addition a mounting medium solved in toluene or xylene is needed. Very sharp "watchmaker´s tweezers" are very helpful.
The objects are positioned on the small coverslide and then covered with molten GG. If the GG becomes solid, heat it very carefully with a small spirit lamp. Now the coverslip is turned upside down and placed on the larger coverslip to form a sandwich-structure. If GG oozes out this is no problem. Let the coverslips dry for some weeks and check them every few days. In case of shrinkage add some more GG.
After drying remove all GG from the rim of the larger coverslip with a razor blade and cleanse the surface with some alcohol. After that, put a big drop of mounting medium, solved in toluene or xylene, on an object slide and place the sandwich - the smaller coverslip facing downwards - on this drop. Store the prepared slide horizontally and allow the mounting medium to dry.
Notes: If the GG still should contain too much water, it will evaporate after some time, for no sealing in this world is totally impermeable! This drying process will inevitably spoil the prepared slide by sucking mounting medium into the GG. So allowe the GG to dry completely! The mounting medium cannot seal the inner part of the slide hermetically, it is there to prevent the intrusion of bacteria and fungi.
Mounting with glycerol: Professionals often use glycerol instead when applying the "sandwich-method" explained here, because then the time-consuming drying process can be omitted and the prepared slides can be finished immediately. But if glycerol is used the glycerol layer between the coverslides must be very thin, otherwise drops of the mounting medium will penetrate. In addition, the mounting medium must show a sufficient viscosity and the solvent must evaporate quickly in order to prevent the penetration of the resin into the glycerol. So resins solved in toluene are preferable. Such slides must be kept strictly horizontal, of course.
Simple staining methods for botanical sections
Chemicals and dyes: Distilled water, isopropyl alcohol, xylene, mounting medium solved in xylene, DELAFIELD´s hematoxylin, chrysoidine, aniline blue, safranine, methylated spirit. You also need staining blocks and dropper pipettes.
Suitable objects: Lignified stems or twigs, about 5 mm in diameter.
Preparation: Make sections with a razor blade (brand blades!) and collect them in a staining block filled with distilled water. It should be stressed that it is impossible to get complete cross sections this way - unfortunately not only beginners try that! Try to produce small wedge-shaped sections instead. Such sections do not look very nice, but they are sufficiently thin at the "edge" of the wedge and show all details wanted.
Monochromatic staining with DELAFIELD´s hematoxylin
Put the sections into a small staining block, wash with distilled water (no tap water!) and cover them with diluted DELAFIELD´s hematoxylin for about 5 minutes. Stir gently. Other preparations of hematoxylin are inadequate here. Remove the dye stuff with a dropper pipette, then wash several times with tap (!) water. The colour of the sections will change from light brown to dark violet or dark blue with hematoxylin being fixed on the tissue at the same time. This reaction is due to the alkalinity of tap water. Therefore distilled water, rainwater and tap water with nearly no alkaline are insufficient. In that case add some sodium hydrogencarbonate. Then wash the sections three times with isopropanol (2 minutes each time), wash with xylene two times, mount with a resin solved in xylene (e.g. Canada balsam) and cover with a coverslide. If a stove is available dry for 48 hours at 60 oC.
Dichromatic staining with DELAFIELD´s hematoxilin and chrysoidine
Stain the sections with DELAFIELD´s hematoxylin as described above. Then remove the water and cover with a solution of chrysoidine (1 gram chrysoidin dissolved in a mixture of 50 ml methylated spirit and 50 ml distilled water). Wait until she sections are stained dark orange, then wash quickly three times with isopropanol. Since the sections are stained too intensively now, add some drops of water and stir gently sucking the liquid with a dropper pipette to and fro. The isopropanol, as it contains a small amount of water slowly extracts the chrysoidine. When blue and orange areas are distinctly different stop the process by removing the diluted isopropanol and adding pure isopropanol. Wash twice, then wash twice with xylene and cover with Canada balsam or a similar mountant. Non-lignified cell-walls are stained blue, lignified cell-walls are stained orange. This method is safe as the blue staining of hematoxylin is stable during the process.
Dichromatic staining with safranine und aniline bue
Dissolve 1 gram of safranine or aniline blue respectively in a mixture of 50 ml methylated spirit and 50 ml destilled water..
First stain the sections with safranine for 5 minutes until they are dark red, wash them with isopropanol three times, then add some water then to the osopropanol and extract the safranin slowly until the sections are a dark pink. Add aniline blue for about one minute, wash again with isopropanol, add some water to the isopropanol and extract aniline blue until the sections show distinct areas stained pink or blue respectively. Stop the process with pure isopropanol, wash twive, then wash twice with xylene and cover with a mountant solved in xylene. The method needs some intuition, as the extraction of aniline blue effects safranine as well, so do not remove too much of the safranine during the first step.
Since all methods given here are carried out in staining blocks and the solvents are added with a dropper pipette, the consumption of dyes and solvents is very low.
Production of NIGLYTIN
As NIGLYTIN is not available any more, the original recipe for this mounting medium is unknown. The recipe given here is that of "new NIGLYTIN", but this medium seems to possess similar qualities.
Mix 7 grams of gelatine with 60 ml of distilled water, wait some hours, then heat the liquid in a water bath until a clear solution has formed. This solution is mixed with 50 ml of glycerol (80%), 2 grams of "nigrosine water-soluble" and 0.6 gram of phenol. Stir for some time at 70 oC (water bath), then filter at the same temperature in an oven. The product has the same qualities as glycerol gelatine.
NOTE: Use only gelatine free of insoluble impurities. This must be checked in a preliminary test by microscopic inspection.
Suitable objects
Blue-green algae (cyanobacteria), filamentous green algae, all kinds of planktonic blue-green algae.
Fixation and pretreatment
The objects are treated with formol (1 ml of formaldehyde 40% per 10 ml sample), ar prefeably with PFEIFFER´s mixture (remove the water and cover with PFEIFFER). After 2 days remove the chemical fixative, then cover with 50 ml of diluted glycerol (5 grams per 100 ml distilled water), transfer the material into an open jar with a flat bottom and let the water evaporate slowly in a dust-free place. Finally add concentrated glycerol and store the objects in small specimen glasses.
PFEIFFER´s mixture
| Formaldehyde 40% | 100 ml | |
| Vinegar made from wood | 100 ml | |
| Methanol (not ethanol !) | 100 ml |
Since "vinegar made from wood" is no longer available, use 100 ml of vinegar and add 3 ml of creosote to the mixture (300 ml).
How to use NIGLYTIN
Transfer a very small amount of the objects on an object slide, remove most of the glycerol and mix with a very small drop of liquid NIGLYTIN (melt it at about 60 oC on a water bath). After mixing the two media very thoroughly with needles cover the mixture with a round coverslip. Press this coverslip gently. The NIGLYTIN should become somewhat translucent (bluish black, not "pitch black"). It is of utmost importance that the objects touch the inner surfaces of the coverslip and the object slide as well. Sometimes little pieces of metal (e.g. nuts) put on the coverslide are useful. Allow the NIGLYTIN to dry completely, remove NIGLYTIN which has oozed out, cleanse with alcohol and protect the prepared slide with a ring of varnish.
Sometimes it is better to transfer the objects directly after the fixation on an object slide and to cover them with diluted glycerol. When the water has evaporated they are mounted with NGILYTIN then. This method helps to avoids flaws.
Note: The objects must touch the coverslip as well as the object slide, otherwise they will be masked by black NIGLYTIN. Therefore filamentous algae should form only one layer (see above)!
Some examples
If the method is applied properly, the algae stand out brightly against a dark background.
The real significance of NIGLYTIN is to make slime fully visible. Even experienced microscopists are often dumbfounded when they suddenly realize that certain algae (especially colonies of cyanobacteria) show a large brightly lit halo of mucus that was earlier completely invisible before! In this respect the method is even superior to the phase contrast method.
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Spirogyra |
Spirogyra Brightfiel, GG |
Spirogyra Darkfield, GG |
Spirogyra Phase contrast, GG |
The NIGLYTIN preparation clearly shows a halo of mucus which remains invisible in brightfield and darkfield. Phase contrast gives an extremely bright edge but ultimately displays only the cell wall.
Using NIGLYTIN requires some experience: If the slide shows bright flaws (which cannot be completely avoided), the glycerol was not properly mixed with the NIGLYTIN. If we see "black algae on a black background", the NIGLYTIN layer is too thick - between the slide and the coverslip only one layer of objects is allowed, the objects touching the slide and the coverslip as well.
Sampling
Beginners
should concentrate on recent species first. Foraminifers are exclusively marine
organisms, usually benthic. We find the tests concentrated in the sands of the
so called "high water mark", that means in that area of the beach
which is abundantly covered with debris and dried seaweed. Collect approximately
100 grams of the upper layer of sand (1 cm) with a spoon, or rather with a small
trowel, and put the sample into a plastic bag. If the sample is dry, it can
be immediately processed. If it is wet wash it very thoroughly with tap water
and afterwards with distilled water. Damp samples must be kept wet because salt
crystals will destroy the tests.
Fossil species can be found in tertiary clays and in chalk layers of the cretacous era, but not each sediment of the tertiary era contains foraminifers - ask specialists!
Tertiary clays are broken into small pieces of about 1 cm length. These pieces are covered with hydrogen peroxide of 12% (use a big jar because of foaming!) The penetrating peroxide reacts mainly inside the small lumps producing gas bubbles of oxygen which gently press the sample apart. When the sample has become pulp, elutriate the fine clay particles, wash several times with tap water, then with distilled water and dry the sample on blotting paper. Professionals rinse the sample through a set of sieves (mesh size 2 mm to 0.05 mm), but such sets are expensive.
Chalk is also broken into small pieces, then covered with water-free (!) acetic acid ( "glacial acetic acid "), which is mixed with 10 grams of anhydrous (!) copper sulfate per 100 ml to absorb the water formed by chemical reactions. Water free acetic acid dissolves the matrix of the chalk without destroying the foraminifers, although the tests are sensitive against acids! When the process has finished pour off the acid and wash quickly (!) with tap water until there is no longer any acidic reaction (pH paper!). Elutriate fine particles, wash the sediment with distilled water and dry on blotting paper. A set of sieves is recommended.
Selecting foraminifers
Studying foraminifers is not expensive! You need a simple binocular microscope, some PETRIE dishes and a fine pointed sable paintbrush, also about 100 "PLUMMER cells" for storing the specimens selected.
Cover the bottom of a PETRIE dish loosely with a small portion of the sample, put it on a dark surface and pick out the interesting objects under the stereo microscope by touching them with the damp paintbrush. Transfer them into a PLUMMER cell. It is more convenient to use a special picking tray with ists bottom being marked like a checkerboard to make a quantitative selection easier. Even better is a special picking tray with a perforated bottom: Make a small frame for a PLUMMER cell, place the dish with its first hole directly above the cell, then push each foraminifer into the hole with a needle. Move the dish to the next marked square and do the same.
As a sample does not only contain foraminifers but many other organisms (small shells, small mussels, parts of echinoderms, elements of sponges and corals, skin teeth of sharks, conodonts, ostracodes and much more), the beginner should first separate everything. Then try to distribute the elements found to different PLUMMER cells. All cells are labelled with the date and the locality and the bulk of the sample should be kept in a glass jar for subsequent investigations. Later it is recommended to fill one PLUMMER cell with all foraminifers of one location to get a quantitative section and to prepare cells containing only tests of identical species.
How foraminifers are identified
Foraminifers always show internal compartments that can be easily recognized in transparent forms. Opaque forms must be soaked with castor oil, which makes them transparent. Many tests confusingly resemble small shells (Fig.1, Fig.2), but the difference can be seen when inspecting the aperture of the test: shells always have a large aperture while foraminifers never have one. The aperture is either small (Fig.3 , Fig.4) or instead of an aperture there are only a few pores or slots to allow the plasma to leave the test. Foraminifers which build their tests by incorporating sand for example, are very difficult to identify as foraminifers - this can only be done only through experience.
How to improve experience
A beginner should not immediately try to determine the species found! It is much better to make a collection of PLUMMER cells first and to enjoy the abundance of different forms of objects. Later join a group of specialists. You can find these groups on the internet.
Sooner or later you have to buy a more powerful stereo microscope. The optical quality of the device is important, but also a good "ergonomic design" so that working without stress is possible. An (expensive) trinocular tube is recommended to always have a digital camera at hand.
One more tip may be added: Magnificent pictures of foraminifers are obtained by using a dark-field reflected light microscope. Such devices are very expensive - perhaps it is possible to get access to such a device by contacting a geological institute.
Finally, here is a remark concerning the SEM images of foraminifers of this site: To study foraminifers a scanning electron microscope is not required, it is even a disadvantage, since you can see the objects only from outside and you cannot move them into any direction (which is very important for their identification!). A visual examination under a stereo microscope is not only cheaper but even more informative! Only in case of photographic representation the SEM is more powerful because of its extraordinary depth of field and the possibility of digital post-processing.
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