On the 28th July 2021 I collected Tony Pattinson’s Swift Microtec-100 Inverted microscope from Beconsfield services. A cautious meeting of bodies in the light of our long self isolation. Tony has moved on to better things and I was very willing to help him make space in his lab.
I’ve said this a million times but microscopes are like handbags and shoes – you never have the exactly right one for what you want to do, so you have to buy another one.
My interest at this point was to enjoy some pond dips, and to have a good working distance in which to manipulate stuff. The inverted provides me with the opportunities I wanted. A plus point is that Tony is a skilled ‘modifier/ innovator’ so this came with a great LED conversion, a mechanical stage that can take either a slide or a 60cm Petrie dish, and an adaptor to an M42 thread in the Camera port. All I had to do was get an M42 to EOS M3 adapter – but the VERY thin one, the camera port already had optics that work.
I have never used an Inverted before so busily set about working out my protocols, a very enjoyable process.
Then came the fateful message from Robert Ratford asking if I would like to have a table at Quekex on 3rd October.
I considered the fun I had had and that it is not everyday that you get a chance to use a different sort of microscope, so I agreed to an interactive exhibit where people could actually have a go.
Was this a good idea? Well maybe I had not thought through what they might look at. Actually I really had not thought that through.
After a little thinking I decided I would look around locally and see what I could find. I settled on sampling local water areas. These are the locations I chose together with information about them that I think is relevant to what I might find.
I live in Hertfordshire, Stevenage to be exact. The groundwater here sits on top of, and in, a very deep chalk stratum. There’s a bit of glacial mud on the top but when you dig down you hit chalk. The water areas I chose are all at the spring line, No big rivers or sea inlets for me, just little trickles and ponds. One of the little trickles in the River Bean which is a designated a Chalk Stream – you can see more at https://www.riverbeane.org.uk/.
For your information Chalk Streams are not at all that common. 85% of the world’s Chalk Streams are found in the England from Yorkshire down to Devon.
Obviously my dipping will reflect that.
So to my dipping stations:
In the Town Centre Gardens is a lake. It is on the site of Bedwell Mire and is fed by a spring that delivers 1.5 litres a minute, and another stream that rises at Coreys Mill a mile or so away, right on the watershed between the Thames catchment and the Ouse catchment. This lake is in a landscaped park and the outlet for it has been engineered through a runnel under the dual carriageway and eventually discharges in a tributary of Stevenage Brook, which is itself a tributary of the River Beane. The Beane drains into the River Lea and thence out through the Olympic Park site into the Thames at London 30 miles away.
In Whomerly Wood less than a mile to the east, is what was a Duck Decoy pond before the New Town was built. Now it is a rectangular pond, home mainly to sedge, duckweed , footballs and shopping trolleys. I have never seen this pond dry out. In long hot summers the level drops but there is always water in it, I conclude, therefore that it too is spring fed. There is a small trickle exiting from the south-west corner. Inevitably this too will end up at Stevenage Brook which skirts the southern edge of the town. The wood that surrounds this gem is mainly Hornbeam Elm and it can be very shady, much of it was planted to supply London Bakers with firewood, before they changed to using coal. There are many woods like this skirting London and now they are well established ‘ancient’ woods. A year or so ago I was part of a working party that cleared the pond and the shade growth on the Southern edge. Mallards sometime nest here, as do moorhens.
Moving further east we come to a feature which is definitely not spring fed. In 1971 series of lakes were built – right up here on the watershed, Fairlands Valley Park. They are fed by road run off and rainfall, with a consolidating settlement pond, and an oil weir at the north end, and a pretty substantial dam at the south. This has a self-priming siphon that drains into a small stream that previously served the meadows when it was farmland. Of course this stream eventually makes it into Stevenage Brook. There are several areas with a toal of 16 acres of water.:
- A ‘river’ with sedges and Phragmites reeds
- An Environmental lake which is gated to prevent untoward public activity, there are two islands,
- the Millenium lake which has one large island and is a recreational lake with events such as Fireworks (on the island), Model boat days, etc.
- and finally, abutting the dam, 11 acres of sailing and fishing lake.
All of the lakes abound with wildfowl, although they are mostly very common varieties. Recently Kingfishers have been see on the cascades between the lakes, and once an escaped Pelican took up residence on the fishing lake for several weeks. These are a very popular public resource. (in snowy winters the slopes in the Valley and on the dam are the Stevenage Ski Slopes.)
I chose to sample at places where duckfeeding and dog access is encouraged in the Millenium Lake backwater, and the Environment lake Education Access
Further east we come to Aston End Brook which now follows along the edge of Gresley Way. It has many characteristics of a chalk stream and drains farmland to the east of Stevenage. This crosses under a road and the bridge area collects quite a lot of debris, at that point I sampled it,
To the south the land rises and the terrain is a bog with Marestails and nettles. This is a Nature Reserve where a small but very unusual dome of peat is associated mire vegetation. This small area of wetland has developed where water appears in the form of a spring. The permanently wet conditions have allowed the development of peat, slowly growing over many years into a domed structure. This unusual habitat is technically called ‘rheotrophic hangmire’ and is the only one in Hertfordshire. The top of the peat dome supports unusual spring-line fen and mire vegetation with prominent tussock sedge and marsh marigold.
The mire also drains into Aston End Brook.
Just under two miles to the east Is the River Beane and both Aston End Brook and Stevenage Brook feed into the river.
What I got.
I have sampled these areas a couple of times and was rather disappointed with the results. The Duck Decoy pond gave the most active specimens. The mud even yielded sulphur bacteria (identified for me by Tony who regularly gets them from his pond.) There were Amphopods – shimps, too.
The Town Centre Pond produced a sparse population of copepods and various algae and diatoms.
I have previously sampled Fairlands Lakes and harvested a blue Stentor, but no luck this time, neither in the mud nor near the surface.
Aston End Brook by the Bridge yielded a Pea Mussel and more amphipods.
The mire was disappointing. Back in February 2020 when I was part of a working party actually on the mire I found Pea Mussels by the spring source. This time I fished over the fence – it is not safe to venture into the mire on your own, in places it is over 6 feet deep, and you can’t tell just by looking, only when your pole starts to disappear.
My sampling of the River Beane was minimal. Between Walkern and White Hall it is suffering from leakage and most of the flow is underground . The river is easy to access, but not very productive.
This left me wondering what people at Quekex could observe.
Lisa and Nigel Ashby kindly dipped Keston Ponds, near Bromley, and The Tarn, at Mottingham – both south of London, and their samples were much more interesting.
Keston Ponds was a very popular dipping location for Quekett members a hundred years or so ago.
On the day people did have a go at the Inverted Microscope – mainly with the Keston Ponds dip. and all-in-all I think I achieved what I set out to do. What I do know is that having started out with a scanty knowledge of my local sources, I now know a whole lot more, I have also got to grips with using the Inverted, it was a good buy.
The Microscope
It is over a year since my last entry. The lack of entries has been more due to lack of time than nothing to say. Everything being equal I now plan two different ones. Both provoked by acquisition of new (to me) microscopes and exhibiting at Quekex 21 last Sunday.
I am lazy and never get round to things I thought would be interesting to do. But today I am tackling one of those.
I have a mount of Fluorite (CaF2) with inclusions, Cornwall, by J. A. Bottomley and dated 1996. The crystal is mounted bare and the adhesive might be glass bond, others in the collection are mounted with that.
It is prismatic and about an inch by a half, and perhaps 1.5mm thick – sorry about mixed units. Glass-like and clear and with distinct inclusions.
These are shown by two diagonal parallel bands. One coherent and dense, the other rather like a trail of bubbles or particles. There is also a fan of very fine needles. And in two places distinct prismatic crystals – also clear
Some facts about Fluorite
Fluorite is the only common mineral with four directions of perfect cleavage. This perfect cleavage combined with the mineral’s isometric crystal structure frequently cause it to cleave into perfect octahedrons
Fluorite is very easy to identify if you consider cleavage, hardness, and specific gravity. It is the only common mineral that has four directions of perfect cleavage, often breaking into pieces with the shape of an octahedron. It is also the mineral used for a hardness of four in the Mohs Hardness Scale. Finally, it has a specific gravity of 3.2, which is detectably higher than most other minerals.
Although color is not a reliable property for mineral identification, the characteristic purple, green, and yellow translucent-to-transparent appearance of fluorite is an immediate visual clue for the mineral
Fluorescence
In 1852, George Gabriel Stokes discovered the ability of specimens of fluorite to produce a blue glow when illuminated with light, which in his words was “beyond the violet end of the spectrum.” He called this phenomenon “fluorescence” after the mineral fluorite. The name gained wide acceptance in mineralogy, gemology, biology, optics, commercial lighting, and many other fields.
Fluorite typically glows a blue-violet color under short-wave ultraviolet and long-wave ultraviolet light. Some specimens are known to glow a cream or white color. Many specimens do not fluoresce. Fluorescence in fluorite is thought to be caused when trace amounts of yttrium, europium, samarium, or other elements substitute for calcium in the fluorite mineral structure.
[I need to check it with a blacklight torch for fluorescence – have ordered one from Silverline (£8.35) l = 350 nm]
Most fluorite occurs as vein fillings in rocks that have been subjected to hydrothermal activity. These veins often contain metallic ores which can include sulfides of tin, silver, lead, zinc, copper, and other metals.
Fluorite is also found in the fractures and vugs of some limestones and dolomites. Fluorite can be massive, granular, or euhedral as octahedral or cubic crystals. Fluorite is a common mineral in hydrothermal and carbonate rocks worldwide.
I thought this could be a suitable subject for some experiments with lighting.
Set up
This was using a camera stand, no microscope and a Canon EOS M3 camera with remote shutter.
I created a temporary stage with a sheet of ground glass to give me space for transmitted illumination.
I also called into action a programmable array of 64 Coloured LEDs, previously developed for Rheinberg illumination.
There are issues with the LED array in that the light is not uniform even when filtered. However I can change the colour and colour distribution by a simple button click on the programming Arduino.
I imaged the crystal using combinations of white, yellow, blue and no light, some with a patch stop over the core of the array. None of the other combinations gave usable images. Using the white illumination I then placed a polaroid filter under the slide, then added an analyser at extinction.This did not significantly help.
I swapped out the 64-LED array for a Flat White LED Celing Panel. Then tried this with polarised filters. There was no significant transmission at extinction but looking at the slide there was an enhanced 3D view of some of the structures.
LED all white
Yellow with blue patchstop
Polarised light
Polarised light – at extinction
LED Flat panel
Flat-Polar
Flat-Polar from the side
(taken with Pixel 3a)
The 64 array LED setup is not really very helpful, the fact that the LEDs give spots of light detracts from the ability to change the pattern. The flat white ceiling panel gives the best image with few artefacts. The Fluorite has insignificant birefringence.
Fluorescence
Still left to do is the test fluorescence, will do when the lamp arrives.
The lamp, a small black light torch arrived, but the beam was way too bright to differentiate what was in the specimen. So I ordered a 365nm UV pass filter to block the blue wavelengths.
I laid the torch down on a black surface and attached the filter in the lightpath (Black tack is good stuff). Using two small foam blocks and placed the slide so that the beam passed through obliquely. I was pleased to see that yes parts of the slide do fluoresce.
7th August 2020
I shared a link to this on QueketMicro on Facebook and Jonathan Crowther made several interesting comments. One of which suggested putting the UV filter on the camera rather than the torch. The resulting effect (using the camera stand and Eos M3) was very different to putting the filter on the torch.
The image has been post processed in Corel Paint Shop Pro XVIII. Mainly to accentuate the detail visible and clear the back ground.The crystalline inclusions are not as clear as before
I now have another query to find and answer for -‘Why does this system give such a pink colour, not at all visible in the previous system?
28 August 2020
As previously reported I have some Oamaru diatomite. Some of this was passed by various means to Michel Haak who cleaned it up and let me have back some strew slides. So now I have slides Nos. 413-417 in my cabinet. They’re nice clean slides and I decided I would catalogue, as best I could, everything on them, just because I felt like it.
Technical Information
I started with slide no. 413 and tracked across it using my Zeiss standard, with Carl Zeiss x40 Fluotar PH2 objective and Leitz Wetzlar x10 eyepiece. This gave a small field of view, but as much detail as I can get without oiling up. Using my Canon EOS M3, set at infinity and f4.5, I photographed every object that took my fancy, including the anonymous fragments, and a reference scale bar (100 div=1mm). Having set up I kept all settings uniform for the whole exercise, except where noted. This was done to make all images comparable. [However at first I had the illumination set to Phase contrast but Changed to Brightfield at object 73. Stacking was done either with Zerene Stacker, or Helicon Focus. Stitching where necessary with ICE]
999 images and 214 objects later I stacked and stitched, as necessary ,and set about giving things names. The general frquency of objects was:
Silicoflagellates
I always see a lot of rather nice siliceous meshed structures. I thought they were sponge spicules, but a bit of on-line searching showed me that they are the skeletons of Silicoflagellates. This is a group I’ve never previously heard of, although I have seen these things in many marine strews. So it’s nice to now know that they are marine planctonists that are both photosynthetic and heterotrophic. They have been around since the early Cretaceous and are still present today. My material comes from the late Eocene to early Oligocene and is about 32-35 million years old. In the gallery I have included pictures of what I think are Naviculopsis constrictor and remind me of a sort eye of a double needle. Distephanus crux with its spikey needles, Bachmannocena apiculata rather similar to the Distephanus crux, and Corbisema hastatar, a rather empty triangle, you seem to get a lot of these.
Radiolarians
Another frequent non-diatom presence are the Radiolarians. These are holey bundles with variable longish spikes and generally a central capsule. In life they would have housed protozoa in the plancton. This time I shan’t be concentration on them.
Fragments
Then there are numerous fragments of plated structures, generally perforated and sometimes with one distinct edge. There are also meshed fragments which may well be diatomaceous in origin, or may not.
Spicules
Spicules are frequent. These spikey, glass structures often come from sponges of many sorts, where they are embedded in the sponge to give it form and shape, or protection. Some of them look like vicious gothic weapons or maybe Reindeer galloping through the sky. I have a plan to collage images of these for Christmas Cards one year, but not yet. I did find out, though, that one very dark blob that bears a strong resemblance to a bath sponge is also a spicule – a sterraster one. They’re not infrequent and I always thought they were some sort of radiolarian.
Diatoms
Finally the whole point of the exercise is to identify the diatoms in the strew. Oamaru diatomite is famous for the variety of diatoms it contains. Some are really common, and, to be honest, it can be tedious imaging your tenth Coscinodiscus. Others hold a rarity value and these are what I’m really interested in finding.On this slide I found a Rutillaria radiata, I’ve never seen one of those before. Another intriguing one, to be honest it made me think of a pair of knee-joints, was Briggera capita in girdle view.
Now the mystery object
There were 11 other new-to me examples. One of them is the reason for this blog entry. I called it ‘Looks like a shark’s tooth’ because it does. This chevron shaped object has one complete tip that is so very like those found on Triceratia. unfortunately the other two points seem to be fractured. It is very dark and has a pitted surface, but clear distinct edges. On the internet the best match I could get was with Triceratium bicornigerum, but it wasn’t that good a match. A search through the remaining slides from Michel Haak, and others also from the same specimen failed to yield any other, similar objects.
I’ve acquired a pdf version of Schmidt’s Atlas, from the diatom-forum.io group, thanks.
So, easy peasy, just find a match, or not.
The nearest thing I have come up with, so far, is Triceratium alternans, Even that is not great.
But if I match the Schmidt image with one given by Stuart Stidolph in his paper A Record of some Coastal Marine Diatoms from Porirua harbour, North Island, New Zealand. And then match my one, the level of variation could possibly be acceptable. But the Stidolph images show something much larger than mine, Schmidt did not give scale bars. I’m not convinced I’ve pinned it down.
Now if anyone, especially an experienced diatomist ever reads this and can help me identify the ‘shark’s tooth-like’ diatom, I’d be delighted.
Gallery:
Silicoflagellates.
Radiolarians
Sterraster spicule.
Rutillaria radiata
Briggera capita
Comparison off my specimen with Schmidt and Stidolf specimens
References
Schmidt. A (1874-1959), Atlas Der Diatomaceen-Kunde, Tafel 78 no.14. pdf of 1972 version
Stidolph Stuart R. (1980) A record of some coastal marine diatoms from
Porirua Harbour, North Island, New Zealand, New Zealand Journal of Botany, 18:3, 379-403,
At Microscopium I bought a Beck Stereo which is much better suited to diatom picking than my other scopes.
At Quekex 2019 I was talking to people about the best way to clean up my acid washed diatomite extract, and I have been reading various methods.
My first essay into this was to give a small sample an extended soak (several days) in Sulphuric acid, followed by repeated dH2O, shake, settle, decant and do it again, to remove surplus acid.
I have also prepared some clean slides. (massage with liquid soap, rinse thoroughly in hot water, distilled water, then Ultrasound in water with EtOH at 30 degC for 600 seconds. Dry on clean, lint free cloth (Clean old handkie) store in dust free box).
Next I made up a Gelatine wash for the receiving slides. 5% leaf gelatine in dH2O with 1 crystal of thymol per 100ml. When required this was liquefied in a constant temp bath at 35degC. 1 drop applied to a cleaned slide and rubbed with clean thumb until nearly dry, whisked sideways with a clean lint free cloth, to displace any inadvertent lumps. dried and stored in a dust free box.
A drop of the rinsed Oamaru diatom extract was placed on a slide and, using a glass needle made last year, some specimens were hand-picked to a receiving slide. Apparently freshly made glass needles have too much residual static.
Sounds straightforward doesn’t it.
Most of it was, but it is really tricky to avoid dust. The Gelatine prepared slides are very sticky – everything sticks firmly to them. Geting the hand-picked diatoms off the needle was tricky but I found that blowing down the capillary did the job. I wasn’t able to move them about afterwards and broke a rather nice Aulacodiscus
and smashed a lovely Trigonium in the attempt. I did manage to get another one though.
I had forgotten the exact detail for fixing in Pleurax, and could not locate my notes. In the end I used a glass sheet on the hotplate to help to even out the temperature. I dried the receiving slide and a freshly cleaned coverslip on the hotplate at about 40degC then I set it at about 100 degC to add the Pleurax, but that was too hot. I got bubbles – too many
and they also destroyed another very nice girdle view of a Trigonium
The two diatoms I have got mounted are not clean enough, and one is broken.
More practice is needed.
One consolation is that along the way I did photograph a very nice glassy spicule through my new microscope.
I think this is a variation on a Tetraxon spicule (based on Fig 3-3 p84 of Invertebrate Fossils, by Moore, Lalicker and Fischer that I also bought at Microscopium).
One thing I did do was have a lot of fun doing all this.
In the Amateur Microscopy Facebook group Wojtek Plonka posted about an LED gizmo he had constructed that used programmable LEDs. You can program it to work from a Arduino as a ‘stand-alone’ device that you can switch between modes (of your own devising). He was using a NeoPixel 7 Jewel with an Arduino to mimic Rheinberg illumination.
I always struggle to get Rheinberg filters to work for me – just don’t get the central spot in the right place. This gizmo seemed to be quite flexible so we (me and E) made one up.
I’m still having an issue with positioning under the stage so I looked at the videos of Kevin Webb’s talks in 2015 – both at Northampton Natural History Society’s Microscopy exhibition and at Quekex 2015. His trick is to use an inverted microscope so there’s plenty of space for organising the light source. However with similar LED light rings he produced some amazing images, ones with detail that surpasses those you can get with conventional illumination.
This is something with potential and is worth spending some time on.
Thus far I have removed the condensers from either the Zeiss Standard or the Novex B and blue-tacked the circuit board with the jewel on to the condenser carriage – lining it up by looking down a tube and centreing the middle pixel. But I can’t rack it close enough to the stage to get into the dark ground position because the PCB is too big and snags on the undercarriage. I don’t want to cut it down until I’m sure of the best size……
I have also played with colour configurations and intensities – I need low intensity to be able to see what I’m looking at, and high intensity for photography, especially if I introduce polarising filters or dark ground, which attenuate the light strength. By using a potentiometer in the circuit I can control the intensity.
So here’s what I have managed to do so far…
A Seabuckthorne scutiform scale illuminated by half red, half green and a null centre LED. This does not achieve Rhenberg, but the refraction of the coloured light does show up some of the details.
The same set-up all I did was to press a button on the Arduino controller to switch to a Yellow ring with a blue centre – classic Rheinberg configuration.
This, on the other hand is how it looked with all white LEDs and a diffuser to help suppress fringing.
With the standard Phase Contrast Condenser in place and the usual illuminant I got this:…
Which is, in my opinion, a much nicer image, much clearer.
I’m fiddling with this system to try to help with the refining of the Oamaru diatoms. They do stand out with the LED lighting, and every little helps.
At Quekex2018 Win van Egmond suggested a way to use the all white setting of the LED Jewel with a stack to produce Dark Ground Illumination. So, naturally I have had a go at his suggested device: two Petri dishes stacked. The top one with a black mask and central hole. The lower one with a black stop (actually this needs to be slightly larger than the hole in the mask to eliminate parallax issues.)
The sides are wrapped with reflecive silver paper (Kitchen foil) and this should give a dark ground surface on which to put the picking tray. The whole thing – wrapped in black duct tape sits under a dissection microscope and the LED is placed under the lower dish.
It works quite well and cost me nothing as I already had all the parts. I do like Heath-Robinson constructions – in making them you end up having to understand what you have done.
I’ve now got some diatom picking skills and tools. They’re not perfect but do work much of the time.
I have a problem which is that having found a nice specimen and picked it what next?
Initially I put them onto a clean slide – a tip from the microbe hunter forum is to mark the receiving slide, on the obverse, with rings. This is a good idea because it helps locate the pickings when done. But the slide is clean and dry, as are the diatoms. I have tried going straight to Plurax after heating the slide etc to get the diatoms to stick. But there are at least two reasons not to to this.
The less urgent one, but the one that bugs me most is that, generally, I can achieve only ONE good specimen per slide.
The more urgent one is that I need to improve my Plurax fixation and control the temperature much more than I have. It has been a bit hit and miss so I get VERY uneven distribution of the mountant and BIG holes in it. I can cure that one with practise and diligence.
I am waiting for the ingredients for an adhesive to arrive and that should help me get a better outcome too – more about that later.
So what is the ‘frustration’ – it’s finding nice specimens, photographing them and then not being able to make good permanent mounts, and probably losing them on the way to such.
Yesterday, in a dried strew of the acid washed residue, I came across this one.
Trinacria simulacrum. Grove, E and Sturt, G 1887 [ according to my identification from Oamaru diatoms]
It is sitting in its strew on a slide in a slide carrier waiting for me to get a better technique, etc. And there’s no guarantee that when I go to pick it off
1. it will still be there or
2. I won’t break it.
THAT is frustrating. But at least I have a picture of it.
I also console myself that my sampling is of a minute fraction of the material I have. If I can find one of anything in the sample there must be a great deal of them in the bulk material. i.e. thay can’t be THAT rare. So if I lose one I’ll probably find another as I work through it.
I’d better get on with improving my techniqus.
In a different strew – actually not a strew but a dried out aliquot in a 55mmm Petrie dish there is this:
From what I can see I think it is probably a Stictodiscus novaezealandiae Grunow, A in Schmidt, A 1888 – same source.
It looks pretty much entire and if I add some distilled water to the right pick I might be able to float it free of the brash. Note all the conditional words there.
But today, while waiting for the delivery of materials, I think I’ll start afresh with a good boil in HCl of a new dollop of the mud. Wish me luck.
Trigonium articum Cleve, P. T. 1868.
I have been experimenting with picking off individual diatoms.
- I tried an acupuncture needle attached to part of a mechanical stage – not very effective.
- Next a 1ml pasteur disposable pipette – this one had a very small hole and worked quite well The new 1 ml ones are coarser – not so good.
- My best shot so far is a glass micropipette stuffed into the coarser 1ml Pasteur.
I’ve always enjoyed working with glass, but you do need it to be soda glass in the domestic environment.
I got myself some melting point tubes, pulled them out over a tea light and gently snapped them, wrapped in tissue, between my thumb nails. They’re surprisingly effective, with reasonable shaped ends.
Then under the stereo microscope on the lowest magnification, but with the x2 Barlow lens in place. I used this to pick off individual diatoms onto prepared coverslips.
The diatom suspension (from the filtration experiments – both 250 mu and 75 mu residues separately) was in a 55mm Petridish illuminated by two ‘angel-eyes’ LED rings strapped together to provide a low cost lateral illumination that worked extremely well – but did get hot. This had the effect of contrasting the diatoms against the dark backround – making them easier to see.
.
The picked diatoms were examined under a magnification of 400 to check their suitability and the selected ones mounted in Pleurax.
This is what I have, so far:
And I have refined quite a few techniques.
So I had some sediment derived from the Oamaru Cormacks diatomite. Cleaned up as described below, but still a little grubby. It consisted of siliceous fragments and diatoms.
The challenge to get out intact, or nearly intact, diatom frustules from brash was to separate objects of the same size made of the same material without damaging them.
Vibration.
Place a suspension of the material in a liquid on a vibrating surface (in this case a Boston Acoustics Woofer loudspeaker laid horizontally). Subject it to vibration at between 15 and 45 Hz for a period of time.(without damaging the loudspeaker).
I tried this using an online tone generator , 50mm glass Petrie dish and distilled water.
I made a dH2O suspension of one drop of the sediment to give me a slightly cloudy mixture. Aliquots were then subect to a range of vibration and time. Samples taken from the centre, one third radius and two thirds radius where popped onto slides and examined, first wet at x10, then when air dry at x40.
Disappointment.
The ratio of desired object to brash was really low, more so than in the untreated material.
I am currently making some measurements across the range to confirm my first impressions but would infer that the vibration gave me more brash – so probably fragmented a lot of the material further.
On consideration vibration is used to disrupt particles, and although this method may work well with living diatoms that are proected by the cell contents, these fossil ones are actually rather brittle.
Another consideration is that the sound waves are propagated on an up-and-down axis. the ripples in the suspension will also be up-and-down, so why would this method segregate the particles? However, as I said, the method apparently works with living diatoms. Obviously there is more physics to be explored in this system. I will probably not follow it through, my life is limited and I really wouldn’t have the time for this game.
Below are pictures of the only decent objects from this experiment at 26Hz for 3 minutes.
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- Actinopyctus like
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- Coscinodiscus superbus
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- Diploneis
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- Xanthiopyxis
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- Hemiaulus
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- strange object
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- could be Isthmia
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- Raphoneas
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- Navicula oamaruensis
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- oval
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- Xanthiopyxis
They were mostly rubbish but one or two delights made it through the process.
At 15Hz for 10 minutes there were no usable objects.
Filtration.
A friend alerted me to a set of micron filters available from a Marine Aquarist at relatively reasonable cost. So I got some- ostensibly 250, 125, 75, 50, 25, 10 and 5 microns – and passed some of the diluted suspension through the series. The greatest residues. by a long way, were in the 125 and 10 micron filter. The final filtrate still showed some signs of milky white particles, microscopically they are not interesting in this context.
The experimentation is in its early days, but the fraction retained by the 125 micron filter has yielded a greater density of desired size objects, less brash than any of the vibration fractions.
Identifcation where attempted is provisional and made using Nigel Charles’ excellent site.
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- Triceratium – maybe Morelandii
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- amorphous circular
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- Azepeitia maybe
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- circular
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- Coscinodiscus
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- disc
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- mystery object
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- round diatoms
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- Triceratium – maybe undulans
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- Triceratium of some sort
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- Triceratium or Trigonium
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- Triceratum – rectangular
Wet preparation.
Finally I put a 50mm Petrie disc under the x16 objective and had some really good fun shaking it gently.
The two most gratifying results were to watch a Coscinodiscus swing from valve to girdle view:
It really does bear an uncanny resemblance to the O2 Dome.
And to get three aspects of a Biddulphia elongata:
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- Biddulphia elongata – valve view
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- Biddulphia elongata – oblique view
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- Biddulphis elongata – girdle view
These remind me of an mummified body.
What is most satisfying is the variety of objects I am seeing, and from minute drops of the suspension.
If you were one of the people to whom I gave a drop of suspension at the QMC President’s Address, these diatoms came from the same mixture. (unfiltered and unvibrated)
I haven’t included the weird and wonderful spicules that are also there, maybe that will be in a different post.
Jacky's microscopy 