Lake Sediment Analysis

The following methods and protocols are all related to the chemical and physical properties of soil.

Unless stated, the methods are currently being used in the Laboratory and have Risk Assessments associated with them. If you are planning on carrying out any work in the lab you must read and sign the Risk Assessment first. Please contact the Laboratory Supervisor for details.

Core Extrusion

The method of core extrusion depends upon the type of corer used. Kajak, Glew and similar cores up to 50 cm long can be extruded on the screw-threaded extruding rig allowing sampling intervals of 1 mm if required. Mini-Mackereth and Fat Livingstone cores are extruded using the pole and pin and can therefore only be sliced at 0.5 cm intervals or coarser at the moment.

The project leader is responsible for specifying the sampling intervals required for each particular core. Core slices are usually placed directly into whirlpak bags following extrusion. This has the advantage of avoiding the need to transfer dried sediment from Petri dishes to bags at a later stage for storage and also prevents the sample from losing water before wet density analysis can be done. The conventional Livingstone cores are extruded whole in the field and carried back in 1m lengths of split piping wrapped in cling film.

These cores should be wrapped in tubes of polythene and heat-sealed if they are not to be analysed immediately. Glew, Mackereth and "Fat" Livingstone extrusion techniques will normally be demonstrated by one of the technical staff if asked.

Sediment Description

Sediment colour is described using the Munsell soil colour charts as the core is extruded. Colour and the point of any colour change is noted in the core log book which also holds the routine sediment analysis data. Troels-Smith description of the sediment may also be needed, in which case the project leader will indicate the degree of detail required.

Use of Freeze Drier

For some analyses, e.g. lipid analysis, it is essential that sediment is freeze-dried rather than oven-dried. Freeze drying is preferable to oven drying because it removes the water content without greatly altering the physical structure of the sediment. As a result, sediments containing clay are much easier to handle after freeze drying (they remain friable) and diatom breakage resulting from contraction of the sediment during drying, is also reduced.

Operating the MODULO 4k Freeze drier

Preparing samples. All samples must be frozen in the lab freezers before they can be freeze-dried. The samples should be placed in containers which allow as much of the sample to be exposed to the vacuum as possible. Petri dishes are ideal for sediment.

Preparing the freeze-drier. Switch on the freezer unit (main power socket and on a switch located on the front of the freezer unit) and leave for around 10 min. allowing the temperature to stabilise between -55 and -60oC. Also, at the same time, switch on the vacuum pump (main power socket and switch on the left-hand side of the pump unit) ensuring that the drain valve is closed and the flask unit is sealed. This action heats the oil to a working temperature and makes freeze-drying more efficient. After the temperature has stabilised and the oil heated, switch off the pump (leave the freezer on) and release the vacuum by opening the drain valve. Before use, check the level and quality of the oil through the window on the front of the pump. The oil level should be between the upper and lower marks and should be a clear brown colour.

Loading. After the vacuum has been released, remove the top of the perspex flask. Take pre-frozen samples from the freezer and transfer them to the flask as quickly as possible to prevent melting and subsequent water boiling. In the case of Petri dishes, samples are placed on plastic trays with the lids removed or slightly open. As each tray becomes filled, levels are built up using small glass phials as props between trays. The trays can be lifted into place using needle-nosed pliers available in the lab. Using Petri dishes with the lids slightly open, 7 samples per tray can be dried, making a total of 56 samples on 8 trays.

Switching on. After loading, the unit should be sealed by replacing the perspex flask lid and ensuring that the drain valve is closed. The drain valve requires only slight tightening - over-tightening will damage the valve. Switch on the pump as above.

Freeze-drying. The pressure in the flask should decrease with time indicating that freeze-drying is taking place and that water is being removed from the samples. When the pressure drops to 0.08 - 0.1 mbar, as indicated on the pressure gauge, the sample should be dried (visual inspection can also be used to determine the stage of drying). Drying time varies with the type of material and generally increases with 1) increasing water content, 2) increasing thickness of the sample and 3) decreasing surface area in proportion to sediment size. As a rough guide, for clay-rich sediments in Petri dishes, samples can take up to 48 hours to dry and for sandy sediments, up to 36 hours.

Switching off. Switch off both the vacuum pump and the freezer unit. Then release the pressure in the flask before removing the samples. To release pressure, the drain valve should be opened to allow air to enter the chamber. It is advisable to place your finger over the drain inlet/outlet whilst opening the valve to assess how much air is entering. It is important to allow air to enter very slowly to prevent a rush of air from dispersing dried sediment around the chamber causing contamination. When the flask has reached atmospheric pressure, the lid and the samples can be removed. Make sure that there is nothing attached to the drain inlet/outlet before releasing the vacuum.

Clearing out the ice. After the samples have been removed, open the drain valve fully and attach a rubber pipe to the valve inlet/outlet putting the other end into a container. Ice formed during freeze-drying in the condensation chamber will melt and the water will pass out through the drain valve. Although it is not strictly necessary to do this after every run, it is advised as it improves the efficiency of future runs. The ice can be left to melt overnight or to speed the process up, warm water can be poured over the ice in the chamber. This should be done with care - always ensuring that the angled pipe in the condensation chamber is in place.

Cleaning the freeze drier. After each run, clean the trays, flask and chamber thoroughly using a damp paper towel and dry. Never use solvents to clean components of the freeze drier.

Safety notes

Care should be taken whilst loading/unloading the freeze-drier as this involves working from a stool
Users should handle the perspex flask carefully and check regularly for any abrasions or fractures which might impair the strength of the chamber. Remember, during freeze drying the flask is under a high vacuum any weaknesses could cause the chamber to implode.
Do not touch the inside of the refrigeration chamber whilst the fridge is on or immediately afterwards as inside temperatures can be as low as -60^oC

Diatom Preparation

It is necessary to remove any organic matter from diatom samples in order to make microscopic identification easier. The cleaning process also allows unwanted mineral material to be removed and the concentration of diatoms to be adjusted [a full discussion of diatom analysis is available in Battarbee (1986)].

Safety

Hydrogen peroxide is a very powerful oxidising agent. Contact with the skin should be avoided and rubber gloves and eye protection should be worn when handling it. If hydrogen peroxide comes into contact with eyes, skin or clothing, wash the spillage under running water. Spills on bench tops, floors etc. should also be diluted with water before mopping up with paper towel. Mopping up concentrated hydrogen peroxide with a paper towel can cause fires.

Method for the preparation of samples using a waterbath

This method is particularly suitable for large numbers of sediment samples (Renberg, 1990). It can also be safely left without any risk of explosion and there is less risk of samples boiling dry. It may not be suitable for samples which tend to react vigorously with hydrogen peroxide, such as large epilithon and epiphyton samples.

In this procedure centrifuging the washed sample may be replaced by settling out in a cool place overnight. This is convenient when handling a large number of samples in glass test tubes and reduces the breakage of fine, filamentous diatoms. However, if the samples are prepared in plastic centrifuge tubes they may be centrifuged if preferred.

Equipment

Plastic centrifuge tubes in a rack, water bath filled with distilled water, hydrogen peroxide (H₂O₂) 30% (100 volume), distilled water, hotplate, 50% hydrochloric acid (HCl).

Digestion procedure

Place approximately 0.01 grams of dried sediment (or 0.1 gram of wet sediment) in each tube, weighing to four decimal places if diatom concentrations are to be calculated. Moistening dried sediment with a few drops of H₂O₂ will help its dispersal when the rest of the peroxide is added.
Add 5 mls 30% H₂O₂ to each tube and place in a rack in the water bath (in a fume cupboard) at room temperature. If the sediment does not react violently with the sediment, the temperature of the water bath can be increased to 80^oC. 'Blank' tubes containing only H₂O₂ can be placed at intervals in the rack and analysed to check there is no cross-contamination between the tubes during digestion. Evaporation from the water bath is reduced using floating plastic spheres.
Heat samples, checking the level of H₂O₂ from time to time, until all organic material has been removed. This process may take several days, depending on the organic content of the samples. Do not allow the samples to dry. Also, keep the water level in the bath topped up with distilled water. The heater cuts out and a red alarm light comes on if the water level drops too low.
Remove the tubes from the bath and add just 1-2 drops of 50% HCl to each tube, which will eliminate any remaining H₂O₂ and any carbonates. The fizzing which occurs also helps to unstick any diatoms which may have become attached to the side of the test tube.
Top up test tubes with distilled water and centrifuge for 4 minutes at 1200rpm. Alternatively leave to settle out overnight at 4^oC. The resulting supernatant liquid is then decanted and the diatoms are resuspended in more distilled water.
Repeat this washing process four more times, either allowing the diatoms to settle out overnight at 4^oC between each wash or by centrifugation. 1-2 drops of weak ammonia (NH₃) solution added to each sample with the final wash will help keep any clays in suspension and will also prevent the diatoms from clumping together when making up slides.

Method for the preparation of samples on a hotplate

This method is particularly suitable for preparing bulky samples such as epiphyton. It tends to be quicker than the water bath method (see below) but needs closer supervision. If beakers are allowed to boil dry, diatoms can be difficult to dislodge from the glass. There is also a risk of explosion when very concentrated, hot hydrogen peroxide rapidly oxidising samples with high organic content. For this reason, it is essential that the fume cupboard window is fully lowered if the heating samples are left unattended, that the window is lowered to a safe working height and that eye protection is worn whilst working at the fume cupboard.

Equipment

100ml or 250ml beaker for each sample, hydrogen peroxide (H₂O₂) 30% (100 volume), distilled water, hotplate, 50% hydrochloric acid (HCl), centrifuge tubes, centrifuge.

Digestion procedure

Place about 0.1g (dry weight, 1 g wet weight) sediment into a beaker. If diatom concentration is to be assessed, the sediment should be weighed to three decimal places.
Add about 20 mls H₂O₂.
Heat on a hotplate set at 90^oC in a fume cupboard until all organic material has been oxidised (1-3 hours). Coarse plant material in macrophyte samples may be removed after half an hour.
Remove the beakers from the heat. Add a few drops of HCl (50%) to remove the remaining H₂O₂ plus any carbonates and wash down the sides of the beaker with distilled water.
Allow to cool in the fume cupboard (chlorine is generated from the HCl) and pour into centrifuge tubes, leaving any coarse sand in the beaker. Top up with distilled water.
Centrifuge at 1200 rpm for 4 minutes.
Decant off the supernatant and resuspend the pellet by tapping the base of the tube. Top up with distilled water and centrifuge as before.
Repeat the washing process at least three times. Clay may be removed during the last wash by adding a few drops of very weak ammonia solution (1%) to the sample. The clay is then decanted off with the supernatant. The sample is now ready to make into slides.

Diatom Slide Preparation

Diatom slides are usually made up by allowing the diatom suspension to settle out on a cover slip overnight, as described below. This produces an even spread of diatoms over the coverslip but it can take up to two days. It is possible to speed up the procedure (resulting in lower-quality slides) by gently heating the coverslips after the diatoms have been allowed to settle for 30 minutes. This may result in some clumping of the diatoms but the slides can usually still be counted.

Equipment

Hotplate, round glass coverslips 19mm diameter, glass slides, 1 ml pipettes for each sample, Naphrax diatom mountant, rigid metal tray, distilled water.

Procedure

Dilute the cleaned diatom suspension to a suitable concentration. It takes practice to get the concentration right. The suspension should look neither totally clear nor milky. Fine particles in suspension should be just visible when the suspension is held up to the light.
Place metal settling out trays with coverslips in a position where they will not be disturbed, away from dust sources and air currents.
Using the 1 ml pipette, place up to 0.5ml of well-mixed diatom suspension on each coverslip, cover the tray to keep off dust and leave to dry. This may take up to two days.
Heat a hotplate in a fume cupboard to 130^oC.
Place 1 drop of Naphrax on a glass slide and invert the coverslip with the dried diatoms over the drop.
Heat the slide on the hotplate for 15 minutes to drive off the toluene in the Naphrax.
Allow the slide to cool and then check that the cover slip does not move when pushed with a fingernail. If it does move then the slide will need to be heated a little longer.

Use of Microspheres in the Determination of Diatom Concentration

It is useful to add a known number of microscopic markers (in the same size range as diatoms) to a known amount of sediment to determine the concentration of diatoms. This also enables the concentrations of individual taxa to be determined, thus avoiding the distortion caused when individual species are expressed as proportions or percentages of the total count.

In the Department of Geography, we use DVB (divinylbenzene) spheres with a mean diameter of 6.4m. Unfortunately, they may not survive the digestion process, or if they do the difference in density between the spheres and diatoms results in the possibility of some spheres being lost in the washing process following digestion. For these reasons they have to be added to the samples just before the slides are prepared. It is important to use DVB spheres since other plastics dissolve in the toluene mountant. Although glass spheres have the advantage of being able to survive digestion, they are less easy to count because they come in a much larger size range than plastic spheres and are also more easily confused with chrysophyte cysts under the microscope.

The DVB spheres are bought as a concentrated suspension. This is sonicated to disperse the spheres before dilution with distilled water to make a concentrated stock suspension which is calibrated using a coulter counter. This is then diluted further to make 1-litre batches of working concentration at around 5x106 spheres per ml. The addition of a small amount of ammonia solution helps to keep the spheres from clumping. To prevent the growth of micro-organisms a very small amount of mercuric chloride is added (<3 mg/L) and the prepared suspension is kept at 4oC.

Procedure

If diatom concentrations are required, the microsphere markers are added after the last wash just before the slides are made. Use the suspension (x10⁶ concentration) stored in the 1-litre conical flask in the cold room. Do not use the stock solution kept in the white polythene bottle in the lab fridge. Because of the toxicity of mercuric chloride, it is important to wear gloves when handling the microsphere suspension and to carefully clean up any spills.

Initially add 1.5-2 mls of 5 x 10⁶ suspension for every 0.1 grams of dry sediment digested, being careful to shake and sonicate the microsphere suspension before each use to disperse the spheres evenly. A suspension of 5x10⁵ spheres per ml is easier to use for sample sizes of 0.01 grams dry weight when using the waterbath method. In this case, it may be advisable to include some replicate samples since the smaller sample size makes this method less suitable for quantitative work.

Make a test slide to check the diatom-to-microsphere ratio (ideally 1:1) and calculate the required amount of microsphere suspension before adding spheres to a whole batch of samples. When making up slides containing DVB microspheres, care must be taken not to heat them above 130^oC.

Pollen Analysis of Terrestrial Sediment

Notes on Sample Preparation

Samples should be stored wet at <4^oC.
If sampling from an unsliced core, all the samples should be taken at one time to prevent having to re-open the bags and drying out the core material.
Beware of contamination - clean all spatulas etc. thoroughly between samples.
If the core is long but covers a short time period, a large sample volume (1cm³) may be necessary. However, if the core is short, covering a long time period uses 0.5cm³.
If using absolute pollen techniques (quantitative), add weighed tablets of the exotic taxon to a known volume of sediment in a labelled 50 ml boiling tube.

Removal of Calcium

Slowly add about 10ml of 10% HCl to the sample. When effervescence stops, place the tube in a boiling water bath and stir well until effervescence again stops. If the tube threatens to froth over, reduce the foam with a squirt of acetone. Centrifuge and decant.
All centrifugation should be at 3000 rpm. for 4 minutes and about 2ml of methanol should be added to reduce the specific gravity and reduce the losses in decanting.
If sediment is highly calcareous, it will be necessary to centrifuge, decant and add a further 10ml of HCl. It is essential to remove all calcium carbonate at this stage, as the later addition of HF will result in the formation of insoluble calcium fluoride.
After the supernatant has been decanted, thoroughly mix sediment at each stage.
Wash with distilled water, centrifuge and decant.
If the samples are extremely calcareous, this process should be repeated 2 or 3 times.

Removal of Humic Acid

Add 10ml of 10% KOH. Place in a boiling water bath for no longer than 5 minutes. Stir occasionally.
Record the darkness of the supernatant as a measure of the degree of humification of the sample.

Removal of organics

Strain and wash the sample through a fine mesh screen (170-180um) into a 50ml polypropylene tube. Wash the residue on the screen thoroughly with a jet of distilled water
Centrifuge and decant.
Place the coarse residue trapped on the screen into a labelled petri dish and examine it under a low-power microscope for seeds, fruits, moss remains, large pieces of charcoal, etc.
Wash and centrifuge at least 5 times with distilled water until no trace of brown colour remains in the supernatant, remembering to mix thoroughly each time after decanting. This removes many small organic and inorganic particles.

Removal of silicates

Add 10% HCl, stir, centrifuge and decant.
If sediment contains mineral matter, treat it with 40% HF. In a suitable fume cupboard add about 10ml HF and place the tube in a boiling water bath for 20 minutes.
Stir occasionally with a polythene rod.
Remove the tube from the water bath, add methanol to reduce the specific gravity and centrifuge.
Decant carefully into a sodium carbonate neutralising solution.
Half-fill the tube with 10% HCl and place in a water bath for 20 mins. Stirring occasionally, centrifuge and decant.
If the supernatant is yellow (or green) repeat this procedure - it is better to add fresh HF rather than leaving the tubes in the water bath for longer!
Add about 10ml water, stir, centrifuge and decant.

Removal of water

It is essential to remove all water before the acetolysis step as the acetolysis mixture reacts very violently with water.
Add 10ml Glacial Acetic Acid, stir, centrifuge and decant.
Repeat this step to ensure all water is gone.

Removal of cellulose (acetolysis)

Add 9ml acetic anhydride and 1ml concentrated sulphuric acid (Erdtman's acetolysis solution) and place in the water bath for a maximum of 3 minutes, stirring after 1.5 min.
Remove the tube from the bath and fill with it Acetic acid, stir, centrifuge and decant.
Wash with a further 10ml Acetic Acid. (Stir, Centrifuge, Decant!)
This removes the soluble cellulose products of the acetolysis.

Staining and mounting

Wash the sample into 15ml glass centrifuge tubes using water.
Centrifuge and decant.
Add 9ml water and 1ml KOH to obtain the correct pH for staining (about pH7). Stir, centrifuge and decant.
Add 1 or 2 drops of 1% aqueous safranin and 10 ml water. Stir, centrifuge and decant. Do Not Overstain!
Add about 10ml of Tertiary butyl alcohol. Stir, centrifuge and decant Stir, centrifuge and decant
Wash sediment into a labelled vial with the minimum amount of TBA, centrifuge and decant
Add silicon oil (200cs viscosity equal in amount to the remaining residue, stir well and leave unstoppered in a fume cupboard (lightly covered) to allow residual TBA to evaporate.
Slides are prepared by pipetting a drop of the suspension onto a slide and covering it with a square coverslip. The coverslip is then held in place using clear nail polish.

Fly Ash Particles

There are two types of fly-ash particles - carbonaceous particles and inorganic ash spheres. Carbonaceous particles are made, surprisingly enough, principally of elemental carbon and so strong acids can be used to remove the unwanted sediment fractions (see Safety below). Inorganic ash spheres on the other hand are composed of the same sorts of compounds as many sediment minerals i.e. oxides of silicon, aluminium and iron. Consequently, the technique for extracting these particles is less ruthless but also leaves more material at the end of the preparation.

It is important to remember that these are supposed to be a quantitative-techniques and so care should be taken to ensure that particles have settled out properly before pipetting off supernatant liquid and when transferring sample residues from one container to another. Also, it should be remembered that fly-ash particles are in the air all the time, especially in large cities like London and so all containers must be kept covered unless they are in the fume cupboard with the extractor on. This is especially important when doing preparations from lakes in 'clean' areas.

Safety

The preparation procedure for carbonaceous particles described below uses some pretty nasty acids and so the utmost care must be taken when handling these chemicals.

In particular hydrofluoric acid (HF) can give very bad burns and the information on how to treat these (in the lab user manual) should be read before HF is used. A protective PVC apron is available for use, and safety glasses, gloves, and lab coats should also be worn at all times. The other acids (hydrochloric and nitric) are perhaps not as nasty but should also be treated with the same care.

If a small amount of HCl or HNO₃ comes into contact with the skin this should be washed under running water for a few minutes. For larger quantities - follow the same procedure, but seek medical advice as soon as possible and report this to the Laboratory Supervisor and the Departmental Safety Officer (Mark Maslin). For contact with the eyes - use the eye-wash bottles to irrigate the eye and then follow the same procedure as above.

Making up dilute acids

When making up dilute acids from concentrated ones remember to always add acid to water NOT the other way around. For example, when making up 1 litre of 6M HCl from concentrate HCl, put about 450ml of distilled water into a 1-litre volumetric flask, then carefully add 500ml of concentrate. HCl and then finally top up the flask with distilled water from a wash bottle until the bottom of the meniscus is level with the line on the neck of the flask.

Carbonaceous Particles from lake sediments

Carbonaceous Particles - Procedure

Afternoon before booked time

Accurately weigh 0.1-0.2g dried sediment from selected samples into labelled 12ml polypropylene tubes on a 4-figure balance. (All weighings should be done on such a balance and recorded). (This is the mass of the sediment 'M')
To each tube, add 1.5ml of concentrated nitric acid. Cover and leave in a safe place overnight. This allows for any reactive organic material to be removed prior to heating and saves the loss of any sample by the excessive reaction when the sample is heated later.

Day 1

To each tube, add another 1.5ml of concentrated nitric acid. Place in the waterbath. Heat at 80°C for 2 hours (not including heating up time).
Remove tubes from water bath and top-up with distilled water. Centrifuge at 1500 rpm for 5 minutes.
Pipette off the supernatant nitric acid taking care not to disturb the sample pellet at the bottom of the tube. Removed acid must be neutralised before it can be discarded. Please follow all laboratory protocols for the neutralisation and removal of chemical waste.
Add 3ml hydrofluoric acid to each tube and return to the water bath. Heat at 80°C for 2 hours (not including heating up time).
Remove from the waterbath, top-up with distilled water and leave, covered, overnight in the fume cupboard.

Day 2

Centrifuge at 1500 rpm for 5 minutes. Pipette off the supernatant liquid taking care not to disturb the sample pellet at the bottom of the tube. This discarded supernate must be neutralised as above.
Add 3ml 6M HCl to each tube and heat in a water-bath at 80°C for 2 hours (not including heating up time).
Remove from the waterbath. Top up the tube with distilled water and centrifuge at 1500 rpm for 5 minutes.
Pipette off the HCl taking care not to disturb the sample pellet at the bottom of the tube (again neutralising the acid before discarding)
Wash the sample with distilled water. Centrifuge at 1500 rpm for 5 minutes. Pipette off the liquid taking care not to disturb the sample pellet at the bottom of the tube.
Repeat this washing and centrifuging once more.
Label and weigh a small sample vial (c. 10ml is sufficient) and an air-tight lid for each sample. (This is the mass of the empty vial 'VE')
Transfer the sample residue to these bottles and re-weigh. (This is the mass of the vial + sample 'VS')
Place a cover slip for each sample onto a metal settling tray. Pipette a drop or two of each sample onto the respective coverslip. Heat gently (lowest hotplate setting) to evaporate the water.
Re-weigh the sample bottles. (This is the weight of the vial after subsampling 'V_SUB')
When the water has evaporated, mount the cover-slips onto standard microscope slides using an optical mountant such as 'Naphrax' or similar.

Counting

SCPs should be counted using a light microscope at x400 magnification. All the SCPs on the cover-slip should be counted (number 'N') and care should be taken to ensure the full area of the dried sample droplet(s) is covered.

The concentration of SCPs in the sediment sample is thus:

SCP concentration = 100N/E * M - where E is the percentage of the final suspension evaporated onto the coverslip:

E = 100*(VS - V_SUB)/(VS - VE)

The SCP concentration is in units of 'numbers of particles per gram dry mass of sediment' (or gDM^-1).

Inorganic Fly Ash

Inorganic Ash Spheres from lake sediments:

0.1 - 0.2g of dried lake sediment is accurately weighed into a 12ml glass test tube and 2ml of 30% H2O2 is added to each sample. This is left covered overnight at room temperature.
An additional 5ml of 30% H2O2 is added then added to each sample. This is heated in a water-bath at 80 - 90°C for 3 hours.
Cool, top-up each tube with distilled water and centrifuge for 5 minutes at 1500 r.p.m. Pipette off the supernate.
To each tube, add 5ml 0.3M NaOH and heat in a water-bath at 80 - 90°C for 3 hours. This stage removes biogenic silica (e.g. diatoms, chrysophyte cysts etc.).
Cool, top-up each tube with distilled water and centrifuge for 5 minutes at 1500 r.p.m. Pipette off the supernate.
To each tube add 5ml 3M HCl and heat in a water-bath at 80 - 90°C for 1 hour. This stage removes the less robust carbonates and bicarbonates in the sediment.
Cool, top-up each tube with distilled water and centrifuge for 5 minutes at 1500 r.p.m. Pipette off the supernate.
Repeat the washing and centrifuging procedure.
Label and weigh a small sample vial with an air-tight lid for each sample. (This is the mass of the empty vial 'VE')
Transfer the sample residue to these bottles and re-weigh. (This is the mass of the vial + sample 'VS')
Place a cover slip for each sample onto a metal settling tray. Pipette a drop or two of each sample onto the respective coverslip. Heat gently (lowest hotplate setting) to evaporate the water.
Re-weigh the sample bottles. (This is the weight of the vial after subsampling 'VSUB')
When the water has evaporated, mount the cover-slips onto standard microscope slides using an optical mountant such as 'Naphrax' or similar.

Counting

IASs should be counted using a light microscope at x400 magnification. All the IASs on the cover-slip should be counted (number 'N') and care should be taken to ensure the full area of the dried sample droplet(s) is covered.

The concentration of IASs in the sediment sample is thus:

IAS concentration = 100N/E * M

where M is the mass of sediment and E is the percentage of the final suspension evaporated onto the coverslip:

E = 100*(Vs - V_SUB)/ (VS - VE)

The IAS concentration is in units of 'number of particles per gram dry mass of sediment' (or gDM^-1)

Note

Therefore, all containers must be kept covered unless they are in the fume cupboard with the extractor on. Special care regarding the possibilities of cross-contamination between samples must be taken when undertaking preparations from lakes in 'clean' areas.

Cation Analysis

The following method has not yet been fully developed. It is recommended that several reference samples are analysed at the same time. Also include "reagent blanks" (adding no sample to the beakers but all the reagents).

Equipment

PTFE beakers
Hot Plate
50ml Volumetric Flasks

Reagents

Hydrofluoric Acid
Nitric Acid
Hydrochloric Acid
5M Hydrochloric Acid
De-ionised Water

Procedure

All beakers and flasks should be acid-washed prior to use.

Accurately weigh approximately 50mg of sample into the PTFE beaker.
Add 2mls of Nitric acid and 8ml of hydrofluoric acid to each beaker.
Place on the hotplate at 60°C for one hour.
Increase the heat and evaporate to dryness.
Allow to cool slightly and add 5ml of HCl to each beaker.
Gradually increase the heat and evaporate to dryness.
Add 10mls of 5M HCl and warm gently to dissolve. Cool.
Make up to 50mls with de-ionised water in a volumetric flask.
Decant into suitable containers and store in a fridge until analysed.

These samples are now ready for analysis using either ICP or AAS.

Wet Density

The density of the sediment is determined by its composition. In a sediment core variations in density down a core indicate fluctuations in sediment composition suggesting more than one sediment source. Sediment density is also required for calculations of sediment accumulation rate if the core is to be dated.

Equipment

The density of the wet sediment is measured using a 2cm³ capacity brass phial and a balance weighing to 4 decimal places. You will also need a fine metal spatula.

Procedure

The clean phial is weighed empty to 4 decimal places and then carefully filled with wet sediment. Any air bubbles are removed by tapping the base of the phial on a firm surface and the surface of the sediment is then smoothed to be level with the top edge of the phial.

The phial is then re-weighed and the weight of the sediment is divided by two to determine the density as grams per cm³. Replicate measurements on one sample can be carried out to assess the consistency of the analyst.

The sediment may then be used for dry weight or loss on ignition analyses, or returned to the original sample, in which case care should be taken to avoid contamination with other samples. The phial is carefully washed and dried before measuring the next sample.

Loss on Drying (Percentage Dry Weight)

The percentage of weight loss when sediment is dried is determined by the sediment composition. Generally, sediments with a high organic content weigh less when dried than an equivalent wet-weight of sediment with a high mineral content.

Equipment

Clean, dry porcelain crucibles, spatula, balance weighing to 4 decimal places, oven set to 105°C, desiccator.

Procedure

Weigh an empty crucible to 4 decimal places. Add between 1 and 2 grams of wet sediment being careful to keep sediment away from the rim of the crucible where it may become dislodged and fall out during handling. Re-weigh the crucible and place it in the oven for at least 12 hours or overnight.

Using tongs, remove the crucibles from the oven and allow them to cool in a desiccator (to prevent the re-absorption of moisture) before re-weighing. The percentage of weight remaining after drying can then be calculated. The same sample can also be used for loss on ignition analysis.

Loss on Ignition

The percentage weight lost on ignition gives a crude measure of the organic content of the sediment. Generally, percentage loss on ignition values show an inverse relationship with percentage dry weight values.

Equipment

Clean, dry porcelain crucibles, spatula, balance weighing to 4 decimal places, furnace set to 550°C, desiccator.

Procedure

The dried (overnight at 105°C) sediment samples in crucibles are placed in the furnace and kept at 550°C for 2 hours. It is easier to pack the furnace (it will take up to 80 samples) when it is cold in which case a further ¾ hour should be allowed for heating up. After 2 hours the furnace door should be opened to allow some of the heat to disperse before removing the crucibles with long-handled tongs and placing them on an asbestos mat. When they have cooled slightly they should be placed in a desiccator and allowed to cool fully before re-weighing. The percentage of the dry weight lost on ignition can then be calculated. The remaining ash sample may then be used for carbonate analysis.

Percentage Carbonate Content

The carbonate content of the sediment can be roughly determined by heating the sediment to 950°C. The amount of carbon dioxide lost in the process, as carbonates are converted to oxides, can be used to determine the original carbonate content of the sediment.

Equipment

Clean, dry porcelain crucibles, spatula, balance weighing to 4 decimal places, furnace set to 950°C, desiccator.

Procedure

The weighed ash samples (see method for loss on ignition) are placed in the furnace and heated to 950°C for 4 hours. The furnace door is then opened to allow the heat to disperse (or left closed to cool overnight) and the crucibles are removed and placed in a desiccator as described above. When cool the samples are re-weighed and the difference between the ash weight and the weight lost at 950°C is multiplied by 1.36 (the difference between the molecular weights of CO2 and CO3) to derive the carbonate content which can then be expressed as a percentage of the dry weight.

Calculation of the above can be carried out using COREDAT.

Carbonaceous Particles from Soils (terrestrial)

Taken from Yang, H., Rose, N.L. & Battarbee, R.W. (2001)

Accurately weigh c. 0.2 g soil into a 50 ml PTFE beaker and add 6 ml concentrated nitric acid.
Heat on a hotplate at 100 °C until the liquid is reduced to c. 1 ml. It is important to avoid the digestion from becoming dry to prevent SCP fragmentation. During heating, 'rock' the beakers every 5 - 10 minutes in order to prevent the material from sticking to the wall of the beaker.
Remove the beakers from the hotplate, allow them to cool and add 10 ml of distilled water to each.
Transfer this suspension to a 12 ml polypropylene tube and centrifuge for 5 minutes at 1500 rpm.
Pipette off the supernatant liquid, being careful to avoid disturbing the pellet of sediment residue at the bottom of the tube. Removed acid must be neutralised before it can be discarded. Please follow your laboratory protocols for the neutralisation and removal of chemical waste.
Rinse the walls of the beaker and transfer the washings to the tube. Repeat the washing and centrifuging procedure until the beaker is clean and all sediment material is in the polypropylene tubes.
Add 3ml hydrofluoric acid to each tube. Heat in a water-bath at 80°C for 2 hours (not including heating up time).
Remove from the waterbath, top up with distilled water and leave, covered, overnight in the fume cupboard.
Centrifuge at 1500 rpm for 5 minutes. Pipette off the supernatant liquid taking care not to disturb the sample pellet at the bottom of the tube. This must be neutralised as above.
Add 3ml 6M HCl to each tube and heat in a water-bath at 80°C for 2 hours (not including heating up time).
Remove from the waterbath. Top up the tube with distilled water and centrifuge at 1500 rpm for 5 minutes.
Pipette off the HCl taking care not to disturb the sample pellet at the bottom of the tube (again neutralising this acid before discarding)
Wash the sample with distilled water. Centrifuge at 1500 rpm for 5 minutes. Pipette off the liquid taking care not to disturb the sample pellet at the bottom of the tube.
Repeat this washing and centrifuging once more.
Label and weigh a small sample vial with an air-tight lid for each sample. (This is the mass of the empty vial 'VE')
Transfer the sample residue to these bottles and re-weigh. (This is the mass of the vial + sample 'VS')
Place a cover slip for each sample onto a metal settling tray. Pipette a drop or two of each sample onto the respective coverslip. Heat gently (lowest hotplate setting) to evaporate the water.
Re-weigh the sample bottles. (This is the weight of the vial after subsampling 'V_SUB')
When the water has evaporated, mount the cover-slips onto standard microscope slides using an optical mountant such as 'Naphrax' or similar.

Counting

The concentration of SCPs in the sediment sample is thus:

SCP concentration = 100N/E * M

where E is the percentage of the final suspension evaporated onto the coverslip:

E = 100*(VS - VSUB)/(VS - VE)

The SCP concentration is in units of 'numbers of particles per gram dry mass of sediment' (or gDM-1).

Note

It is important to remember that these are quantitative techniques and so care should be taken to ensure that particles have settled out properly before pipetting off supernatant liquid and when transferring sample residues from one sample vessel to another. Also, it should be remembered that fly-ash particles are deposited from the air and are present in high concentrations in many urban areas. Therefore, all containers must be kept covered unless they are in the fume cupboard with the extractor on them. Special care regarding the possibilities of cross-contamination between samples must be taken when undertaking preparations from 'clean' areas.

Pollen Preparation Protocol for Marine Fossil Pollen

Samples are typically between 6 and 9 grams (if volume is needed, use water displacement in a 5 cm3 measuring cylinder). For the Marine Isotope Stage (MIS) 5e project use 7 grams split into 3 equal parts (~2.33 grams each) and place into 50 ml plastic centrifuge tubes. In the prep sheet provided, note the exact weight of each sample, the tube number and an estimation of how much sample is left (if any). Note: Marine samples are usually dry. In order to facilitate their disintegration, break down the sediment using softly a pestle without removing the plastic bag.

Add one tablet of lycopodium per sample (i.e. only in one of the 3 tubes) and write down the exotic concentration and batch number (this is used to quantify the concentration of pollen within the samples).

Removal of calcium

Add 50 ml of 10% HCl. Marine samples are extremely calcium carbonate-rich so add HCl very slowly. Try to get the entire sample in contact with the HCl, by using the whirl mixer and sometimes the caps (be careful when opening the caps, since the reaction produces gases). When the effervescence stops, fill the tube up to the 50 ml mark and place the tubes in the water bath (~70°C) for 1 hour. If the tube threatens to froth over, reduce the foam with a squirt of acetone or methylated spirits, which break the surface tension. This acidifies the samples to pH1. Centrifuge at 3000rpm for 5 mins and decant.
Wash with H20: decant off the "yellow" or "brown" clear liquid supernatant in one fluid movement from tubes in the fume cupboard, and thoroughly mix the solid residue left in the bottom of the tubes. Add slowly distilled water up to the 50 ml mark. Note: Make sure that all the sediment is mobilised and broken down (use the caps and if needed a glass rod). Centrifuge; whilst this is spinning prepare sieves i.e. place a 180μm sieve into a 1-litre beaker. Decant off the cleaned sample.

Removal of humic acid

Add 35 ml of 10% KOH and place in the water bath for 4 mins. This takes the pH up to alkaline (pH14) taking humic acids into solution. Do not whirl mix or decant.

Removal of organics

Pour each sample through the sieves, and wash with distilled water (using a spraying jet and a nozzle restrictor on the washing bottle). Sieving should be carried out wearing safety glasses as even a small amount of KOH can seriously damage your eyes. Transfer all the produced liquid into the tubes, centrifuge and decant (if there is any remaining liquid in the beaker, this step will have to be repeated; try to split the remaining amount into equal parts so that all your samples have less than 2.5 grams of sediment since larger samples might not get completely digested by the chemicals). If there is anything interesting on the sieve keep it in a small vial.
Wash with H20 until the liquid that is being decanted off is clear. This removes humic acids (brown) and clays (usually grey) and might take up to 15 washes. Only in case the samples are very “hazy” exceed 13 washes. When all the KOH has been removed (either after the 1st or the 2nd wash) add 2 drops of 10% sodium hexametaphosphate (Calgon); this helps break the electrostatic bonds.

Removal of silicates

Wash with 50 ml of 10% HCl. This is to acidify the samples before HF, assist the action of HF on samples and make sure there is no residual carbonate. Prepare a large beaker containing a saturated solution of sodium carbonate (which neutralises HF) in the fume cupboard.
Add 30 ml of 40% HF and place in the water bath for 1 hour. Stir twice every 20 mins using a plastic pipette. After this time put caps on tubes, centrifuge and decant. HF removes silicates from the samples.
Wash with 35 ml of 10% HCl and place in the water bath for at least 40 mins. This helps to remove silicate residues and fluorosilicates (heating increases the solubility). Centrifuge and decant.
Add another 30 ml of 40% HF and place in the water bath for 1 hour. Stir twice every 20 mins using a plastic pipette. After this time put caps on tubes, centrifuge and decant.
Wash with 40 ml of 10% HCl and place in the water bath for at least 1 hour. Centrifuge and decant.
After decanting the supernatant liquid and using the pipette get a small amount of sample and prepare a slide by adding distilled water. Check the slides at a light microscope under x40 magnification for the number of remaining silicates. If large pieces and more than 3 can be seen in each field of view repeat the HF and HCl steps but with smaller amounts. If samples are silicate free, put the sample back in the tubes using distilled water.
Transfer samples into 1 tube, clean black dry debris from the walls of the tubes and add a small amount of 10% HNO3 to remove sulphides and pyrites. I aimed for 3ml of concentrated HNO3 and 30 ml of H2O, but larger amounts can be used as long as you keep the proportion 1/10. Place in the water bath for 4 minutes. Centrifuge and decant. Note: some of the sample stays on the tube walls, so before centrifuging use methylated spirits to incorporate it with the rest of the sample.
Wash with H2O, centrifuge and decant. Meanwhile, put the wash bottle of TBA into the water bath to melt (remember to open the lid since TBA expands with heat).

Staining and mounting

Add 22 ml of 1% KOH (2 ml of 10% KOH and 20 ml of H₂O). Centrifuge and decant. The purpose of this stage it to take the pH from slightly acid to slightly alkaline. (Safranin turns blue in acidic solutions!!).
Wash with H₂O. If the supernatant liquid is brown repeat this step until it becomes clear (maximum 3 washes).
Add 3 drops of Safranin and 50 ml of H₂O. Centrifuge and decant.
Add 10 ml of TBA, centrifuge and decant (TBA removes water from samples so silicone fluid/oil can be added).
Transfer into vials: label vials (site once and depth three times since TBA erases marker labels). Put a small amount of TBA into TUBE 1 (not vial 1) and Whirl mix as normal. Then pour into VIAL 1, and repeat until all the residue is in vial 1. Once completed for all the samples place the vials into the centrifuge ‘tube holders’ (add tissue paper at the base of the holders to make them “softer”) using tweezers and spin at 3000 rpm for 3 mins. If you cannot transfer the entire sample at once, centrifuge, decant and then add the remaining sample.
Decant TBA from the vials and add silicon oil (roughly equal to the quantity of the residue) with a disposable pipette. Mix well with a cocktail stick leaving one in each vial, place safely in a beaker lined with tissue and cover to prevent contamination. The TBA will evaporate off over a couple of days, the samples need to be checked and stirred to make sure that they have not dried out and stirred during this period.
When all the TBA is evaporated put labelled caps on

Lake Sediment Analysis

Operating the MODULO 4k Freeze drier

Safety notes

Safety

Method for the preparation of samples using a waterbath

Equipment

Digestion procedure

Method for the preparation of samples on a hotplate

Equipment

Digestion procedure

Equipment

Procedure

Procedure

Notes on Sample Preparation

Removal of Calcium

Removal of Humic Acid

Removal of organics

Removal of silicates

Removal of water

Removal of cellulose (acetolysis)

Staining and mounting

Safety

Making up dilute acids

Carbonaceous Particles - Procedure

Afternoon before booked time

Day 1

Day 2

Counting

Counting

Note

Equipment

Reagents

Procedure

Equipment

Procedure

Equipment

Procedure

Equipment

Procedure

Equipment

Procedure

Taken from Yang, H., Rose, N.L. & Battarbee, R.W. (2001)

Counting

Note

Removal of calcium

Removal of humic acid

Removal of organics

Removal of silicates

Staining and mounting

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