Erdtman's (1960) acetolysis technique is the most popular method, which is followed almost throughout the world. The different steps involved in this technique are given below:
1) Collection and preservation of pollen material: It involves collection of anthers from mature flower buds just before opening and anthesis of flowers by using clean forceps and preserving the anthers or sometimes anthers with a portion of filament of stamens in clean glass vials with 70% glacial acetic acid. Some palynologists recommend 70% ethyl alcohol for preserving the anthers in place of glacial acetic acid. In this condition, the anthers packed with pollen can be stored indefinitely.
2) Anthers are transferred to a centrifuge tube and washed thoroughly with distilled water. The anthers are crushed with a glass rod and the mixture is filtered through cheesecloth or a sieve (c. 180 ^m mesh). The lower fraction is recollected in the centrifuge tube.
3) Freshly prepared acetolysis mixture consisting of acetic anhydride and concentrated H2SO4 (sulphuric acid) in the ratio of 9:1 was poured in the centrifuge tube containing washed pollen. Care has to be taken to add conc. H2SO4 to acetic anhydride drop by drop to avoid excess heating. The acetolysis solution reacts violently with water. Glacial acetic acid washes are necessary to replace water with acetic acid, which does not react violently with the acetolysis solution. Although acetolysis procedure varies from palynologist to palynologist, the following procedure seems to be most suitable. From the first to last acetic acid wash, caution has to be taken so that no water comes in contact with the solution. Only distilled water has to be used for washes to avoid any contamination.
4) The centrifuge tube containing pollen in suspension of an acetolysis mixture is placed in a water bath and heated at 70°C for 5-10 minutes, stirring the contents with a glass rod intermittently. The time required for acetylation varies from material to material. Pollen after completion of acetolysis process turns light to golden brown in colour.
5) Acetolyzed pollen suspension is centrifuged and supernatant liquid is decanted.
6) Pollen sediment in a centrifuge tube is immersed in glacial acetic acid for a few minutes and the superfluous liquid is decanted after 5 minutes centrifugation (c. x 2000 rpm). This is washed with distilled water 2-3 times.
7) The pollen material is ready for mounting on the slides for microscopic examination. However, it can be stored in this condition in vials containing 50% glycerine.
8) Acetolyzed pollen material is mounted in either glycerine jelly, polyvinyl alcohol, lactophenol, Canada balsam, corn syrup, or synthetic resin: piccolyte. The choice of the mounting medium depends on the individual palynologist. It is proved beyond doubt that molten glycerine jelly prepared by using the following method is the most ideal one. Pollen mounted in glycerine jelly remains unaffected even after 4-5 decades. Water solubility and viscosity of glycerine jelly makes it an ideal mounting material for pollen.
9) After mounting the cover glass on acetolyzed pollen in glycerine jelly, edges of the cover glass are sealed with paraffin wax or nail polish to avoid desiccation.
As mentioned earlier, the assets of acetolysis of pollen grains include bringing transparency to and expansion of pollen to facilitate examination of pollen morphological characters. However, there are some disadvantages of this technique. The pollen grains sometimes get over expanded and become unfit for original size measurements. It has been reported that fossil pollen and modern pollen when subjected to acetolysis expand to different degrees. Hence, it is advisable to use unacetolyzed pollen or spores when size determinations are critical.
Sometimes discolouration of pollen is required, which is achieved by chlorination of sample as follows:
Add 5 cc of glacial acetic acid to a centrifuge tube containing pollen material and stir the contents. Then add 1-3 drops of saturated solution of sodium chlorate with a 0.5 cc pipette. Follow this with 1-2 drops of conc. HCl. The solution changes colour in 1-2 minutes as a result of liberation of chlorine gas. Centrifuge sample, decant chlorination mixture, add water, transfer to the acetolysed fraction, stir and follow the other regular actolysis procedure.
Kisser's method of preparation of glycerine jelly is usually followed by palynologists. The following ingredients are required for glycerine jelly preparation:
Gelatin 50 gms
Distilled water 175 ml
Glycerine 150 ml
Gelatin is first soaked in 175 ml cold water for 2-3 hours. Superfluous water is discarded and gelatine is heated till it melts. 150 ml of glycerine is added to molten gelatin and while still hot it is filtered through spun glass pressed into the lower part of the heated funnel. Glycerine jelly has its natural light golden yellow to brown colour. It is stored in Petri dishes or a suitable glass container in a refrigerator.
The acetoloysis procedure described above is good for fresh pollen or spores. However, when fresh pollen material is not available, palynologists have to rely on herbarium material. Several methods have been proposed for pretreating pollen and spore material extracted from herbarium specimens and then subjecting it to Erdtman's acetolysis method.
Dahl (1952) suggested softening the anthers extracted from herbarium material in 10% NaOH or KOH and later subjecting them to lactic acid. Canright (1953) preferred to mount dry pollen from herbarium specimens in lactic acid, which expands the pollen.
Normally the acetolysis procedure for preparing pollen slides takes about 30 minutes. Several palynologists have suggested quicker methods for pollen preparation. Ikuse (1956) preferred to dissect the pollen from the anther in a drop of 95% alcohol; followed by heating to evaporate the alcohol, and then staining with either gentian violet or fuchsin in 95% alcohol. The material is further washed and dehydrated in absolute alcohol followed with a drop of xylene before mounting with Canada balsam.
Avetissian (1950) recommended a simplified method of acetolysis for preparing modern pollen on the slide. It is a quicker method, which takes 3-4 minutes compared to 25 to 30 minutes for Erdtman's acetolysis procedure. It is especially valuable for use with small quantities of pollen as there is very little loss in the procedure. The whole process can be watched under the microscope. The following are the various steps involved in this time-saving procedure.
1. Place the anther on a slide and cover with a drop of the acetolysis mixture (acetic anhydride and conc. H2SO4 in the proportion of 9:1).
2. Dissect pollen from anther with a needle. Scatter the liberated pollen in a thin layer over the slide in a spot about the size of the cover glass.
3. Heat the slide over a spirit lamp and observe the progressive colour change under the microscope. Stop heating when the protoplasmic contents are removed and exine is clear to dark brown.
4. Cool and wash the preparation with a drop or two of alcohol.
5. The pollen can be stained with fuchsin, gentian violet or methyl green.
6. Mount in glycerine jelly.
TEM AND SEM OF POLLEN FOR MORPHOLOGICAL STUDIES Transmission Electron Microscopy (TEM)
The exine stratification is best studied by making ultra-thin sections and examining them in transmission electron microscope. The sections are post-stained in, for example, lead citrate and uranylacetate to achieve good contrast.
Scanning Electron Microscopy (SEM)
In order to see exine surface details better, scanning electron microscope is a useful instrument. The exine architecture and the inside of exine and apertures may be studied with this microscope. Due to the depth of focus in SEM, three-dimensional views of the objects are observed.
DETAILED PROCEDURE OF PREPARATION OF POLLEN GRAINS FOR SEM STUDIES
The method proposed by Falk (1980) is found to be most suitable for preparation of pollen grains for SEM studies.
The pollen grains are first subjected to acetolysis and they are transferred to 20% alcohol. Gradual step-by-step dehydration of pollen grains is carried out by transferring acetolyzed pollen through series of alcohol concentrations like 30%, 50%, 70%, 95% and finally in 100% acetone for a period of 30 minutes. The dehydrated pollen grains are then dusted on the smooth surface of an adhesive tape pasted onto the stub. These stubs with pollen grains mounted on them are further dehydrated by placing them in an oven at 60° C for a period of 30 minutes.
The stubs are then coated with gold in a sputter coat (JEOL JSM 1100). The coated pollen grains are observed in the SEM (JEOL JSM 5600 LV) and photographed.
B: TECHNIQUES USED FOR RECOVERING MICROFOSSILS
INCLUDING FOSSIL POLLEN AND SPORES FROM SEDIMENTS AND PREPARING THEM FOR MICROSCOPIC EXAMINATION
Fossil pollen was observed as early as 1836 by Goeppert and in 1839 by Ehrenberg. Since then many techniques have been invented and used by palynologists. As mentioned early it is difficult to have a standard uniform method as many times the method depends on the choice of the palynologist, type of material and objectives. However, common goals for their techniques are outlined below. It should be remembered that the main purpose of these techniques is to extract or recover pollen and spores from the variety of sediments such as peat, lignite, coat, shale, clays, soil in which they have been entombed in the past.
The techniques used for recovery of microfossils differ with various samples depending on the geological age, lithology, chemical composition of the cementing material, the presence and amount of organic matter and the degree of their carbonization. However, successful results in this regard can be obtained if the following guidelines and objectives are taken into consideration.
1) To get rid of the matrix material as far as possible without loss of organic matter.
2) Concentration of the organic material as much as it may be representative for different types of samples and the forms recovered are not damaged.
3) Processing of samples should take the least time to achieve best results.
Irrespective of type of sample, certain essential steps are involved in the recovery of microfossils, which have been shown in the following flow chart.
Concentration/Preparation for microscopy
Precaution has to be taken to see that the rock material to be analyzed for pollen contents is crushed carefully, neither is it ground nor are the pieces left so large, as chemicals may not to be able to penetrate and react. Crushing of the sample should not be so violent so as to destroy microfossils preserved in it. Carbonates are broken by treating the samples with dilute HCI, whereas silica and other minerals are dissolved in HF (Hydrofluoric acid). The humus and other organic matter is dissolved by weak alkali treatment.
The following procedures are to be used for recovering microfossils from specific types of sediments. These techniques essentially involve maceration of samples.
It is a type of loosely bonded sedimentary rock, which is loaded with organic matter including pollen and spores with only a minimum of sand or clay. The most common method for collecting peat samples is to take cores from subsurface area of peat deposits by using Hiller type of core borer as illustrated in Fig. 9.1. Several palynologists employ different techniques but the one suggested by G. Erdtman and followed by several palynologists including Lagerheim, Wodehouse and Faegri and Iversen, appears to be simple and the most efficient.
The technique involves boiling a piece of peat sample in 10% NaOH or KOH on a slide until the water evaporates. Add a small amount of glycerine, mix thoroughly and transfer the material to a clean slide, mount a cover glass over it. The procedure can be performed in a small beaker. Erdtman's acetolysis technique including bleaching or chlorination can be used for
recovering well preserved pollen and spores from peat samples. If necessary, demineralization of peat sample can be done prior to alkali treatment or acetolysis procedure.
It is geologically older than peat and occurs in early Tertiary sediments. It is also referred to as brown coal. The richest lignite deposits in India occur in Neyvile lignite mines in Cuddalore sandstone series of the Miocene age.
Most lignite samples can be macerated by soaking in dilute KOH (10%) for few hours to several days depending on the hardness of the material. The lignite sample can be boiled for 5-10 minutes to dissolve the humic materials. According to Clair A. Brown, 10% KOH is corrosive to some pollen and spores and therefore use of 5% KOH is recommended.
The following step-by-step procedure can be followed for maceration of lignite.
1. Boil 1 gm of powdered lignite in a beaker or small Erhlenmeyer flask in 20 cc of 5% KOH for 5-10 minutes.
2. Pour into centrifuge tubes, centrifuge and decant. Add remainder and repeat.
3. Resuspend in distilled water, centrifuge and decant. Repeat once or twice or until liquid is nearly clear, or until alkali is washed out.
4. Residue can be mounted with or without staining.
5. Alternately, as per Iversen's suggestion the residue can be dehydrated in glaciad acetic acid and subjected to the acetolysis process.
Traverse (1955) modified the above procedure by prior demineralizing lignite sample in HF and further bleaching in 5-7% sodium chlorite in dilute HCL, followed by acetolysis and mounting macerated sediment in glycerine jelly.
The following procedure for maceration of lignite for the recovery of microfossils adopted by Rao and Vimal (1952) was used for miocene lignites from the Warkali formation in Kerala, South India. This procedure appears to be most appropriate for Indian lignite samples.
1. Soak lignite in conc. HNO3 for at least 12 hours.
2. Wash with distilled water several times.
3. Treat with 10% KOH for 2 hours.
4. Remove silica by heating residue in HF for 1 hour.
5. Rewash and mount residue in pure glycerine, seal with Canada balsam.
Coal is a very compact stratified organic sedimentary rock also classified as a perfect example of compression fossil. In addition to carbonized plant matter it contains several minerals, oils, etc. Coal samples when macerated show that they are loaded with microfossils such as gymnospermous pollen grains, pteridophytic spores, leaf cuticles, vascular elements, etc.
There are three major maceration techniques for liberation of pollen and spores from coal, namely the Schultze method, nitric acid combinations and bromine-nitric acid process. Each of these methods has undergone minor modifications by various palynologists. Schultze was primarily interested in the chemistry of wood and coal, treated some coal with a mixture of nitric acid and potassium chlorate, and followed this maceration with a bath of ammonia. This mixture of nitric acid and potassium chlorate used for maceration and oxidation of coal samples came to be known as Schultze's solution.
The macerated sediment revealed a good quantity of vascular elements, pollen and spores. This fortuitous discovery by Schultze (1855) has been a material aid to coal palynology. The use of potassium chlorate (used in explosives) which accelerates the oxidation process is not always necessary.
Kosanke (1950) had successfully used Schultze's maceration technique for coal samples. This method can be also used for other sedimentary rocks such as shales or sandstones. However, microfossil recovery from carbonaceous shales and sandstones requires treatment with HF.
The following steps are involved in Kosanke's maceration procedure:
1. Place coal fragments or powdered coal in Schultze's solution. (One part of saturated, aqueous solution of potassium chlorate and two or three parts of cold, concentrated nitric acid. This solution oxidizes the coal and different coals require different lengths of time.
2. After oxidation is complete, wash out the acid and add 10% KOH. The potassium hydroxide dissolves humic materials and liberates pollen and spores. The time varies from 15 minutes to 12 hours.
3. Wash the residue through a 65-mesh sieve.
4. Stain spores with Safranin Y for 10-12 hours.
5. Wash the sediment, decant, centrifuge, and run into solvents for mounting in diaphane.
Tschudy (1958) modified the maceration procedure as mentioned below. This procedure seems to be very successful in recovery of higher percentage of fungal spores, pollen and spores from the coal samples.
The following steps are involved in this procedure:
1. Take 1 gm of pulverized sample.
2. Soak over night in 52% HF to remove non-organic materials.
4. Treat residue with Schultze's solution (75% nitric acid and crystalline sodium chlorate.) Oxidation is rapid. Stop at end of 15 minutes. Centrifuge and decant.
5. Wash with water, centrifuge, and decant.
6. Treat with a mixture of 80% acetone and water for 15 minutes. Centrifuge and decant.
7. Wash through several changes of water until supernatant liquid is clear.
8. Transfer to sample vials with equal volume of water.
9. Mount for microscopic study.
Use of bromine for maceration of coal was recommended by Bharadwaj (1957). The steps involved are:
1. Crush 15 to 20 gm coal into pieces 2 to 5 mm sizes.
2. Place them in wide-mouthed, glass-stoppered one-litre bottles. (Set up 6 numbered bottles at one time.)
3. Place in acid-fast sink under hood. Equip sink with frame arranged so that horizontal bar with setscrews centre on top of each glass stopper.
4. Add 5 to 8 cc of bromine to each jar, shake thoroughly, place in sink and allow to act over night.
5. Next morning fill sink with running water to a depth of 6 cm. If temperature of water is above 5 to 10°C, add ice.
6. Open bottles and add 2 to 4 cc of fuming nitric acid to each jar. Adjust setscrews so that stoppers are not too tight. (The worker should wear rubber gloves, a rubber apron, and gas mask.) Stoppers should be tight if treating rich coals as higher pressure is necessary.
7. After every 20 to 30 minutes, add 15 to 25 cc of nitric acid until the jar is 1/3 full. Shake jars frequently to allow uniform maceration. Maceration takes place in 2 to 8 hours, depending upon the kind of coal.
8. Add small quantities of water at 5 to 10 minute intervals, when maceration is complete, until jar is full.
9. Pour contents of jar on Müller gauze (silk screen) and wash simultaneously with a thick spray of water. Add water to jar and pour until all contents are removed. Wash until all acid foam is gone.
10. Invert sieve over a large porcelain dish and work residue into the dish. Decant excess water.
11. Divide residue into two parts A and B. A. Boil 2 to 4 gm in 10% KOH. B. Soak remainder in cold 10% KOH for 5 to 10 minutes.
12. Centrifuge, wash, decant boiled fraction until clear. It contains microspores.
13. Resieve and wash residue B until water is clear. Dry on sieve at 50°C. This fraction contains megaspores and cuticles.
14. Mount these microfossils on a microscope slide with suitable mounting medium.
Surange et al. (1953) recommended the use of Harris' bulk maceration process to recover megaspores from coals of India as per the following procedural steps.
1. Wash coal in running water then place in commercial HNO3.
2. Wash off acid after the coal has macerated.
3. Heat residue in 10% KOH and allow to stand for at least an hour.
4. Remove alkali by rapid decantation.
5. Transfer residue to screen and place under running water. Care is needed as the megaspores swell when heated in alkali and become very delicate.
6. Pick dry megaspores from screen or paper under screen; or pick wet megaspores directly from the water.
7. Pick dry megaspores under binocular microscope.
8. Treat individual megaspores in HNO3 to reduce black colour.
9. Remove silica fragments by placing megaspores in 20% HF over night. Wash off acid.
10. Mount spores in glycerine jelly.
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