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The Trials of Identification

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:

distribution on Slide 413


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.


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.


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 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.


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 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.


  1. Silicoflagellates-thumbSilicoflagellates.
  2. RadiolariansthumbRadiolarians
  3. Sterraster Sponge-spiculethumb
    Sterraster spicule.
  4. Rutillaria radiatathumbRutillaria radiata
  5. Briggera capita thumbBriggera capita
  6. Object from Oamaru DiatomitethumbTriceratium alternans E u varr variabile BrightwTriceratium alternans Bailey

Comparison off my specimen with Schmidt and Stidolf specimens


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,

New Microscope, New Energy

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.

spicule low powerAE

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.

LEDs and all that

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.

EOS M3_9999_13.JPG

So here’s what I have managed to do so far…

07-Hippophae x100 Reinberg R_G stack6HFthumbnail

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.07-Hippophae x100 Reinberg B_Y stack10 HFthumbnail

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.

07-Hippophae x100 LED-White-diffuse stack5 HFthumbnail

With the standard Phase Contrast Condenser in place and the usual illuminant I got this:…

2018-10-15 23-11-07 (C)thumbnail.jpg

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.)


EOS M3_9999_10   EOS M3_9999_12


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.

EOS M3_9999_6

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.


Tantalising frustration

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

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:

Stictodiscus novaezealandiae

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.


Today’s star prize.

Trigonium arcticum Cleve P T 1868

Trigonium articum Cleve, P. T. 1868.


Time to get permanent.

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.

Getting out the best bits

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.


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.


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.

They were mostly rubbish but one or two delights made it through the process.

At 15Hz for 10 minutes there were no usable objects.


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.

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:

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.

Cleaning up then – Next!

We had some good clear weather that gave me the chance to carry one aliquot of the Oamaru diatomite through the whole range of acid cleaning, and a second a good way through the process. I won’t bore you with the details but the idea is to:

  1. dissolve carbonates which would interfere with the later steps by forming insoluble salts;
  2. remove organic material by solution;
  3. dissolve everything else except the siliceous material;

which involves some nasty chemicals. Hence the need for clear weather and a ventilated cabinet.

At the Reading Convention I bought a hand centrifuge from the sale of member’s effects and it has been most helpful in speeding up the separation of residues from liquids. I must have spent several hours winding the handle on that day – all good exercise.

The aliquot that was most ready for taking through this process was, unfortunately, one I had boile dry-just- in a previous cleaning. I had checked and the diatoms were still there it was the mud that had burned. However I considered it worth carrying on as this is probably just a ‘first pass’ experiment to reveal the issues that might be involved.

It worked really well. The only downside is that as the sample started out charcoal colored rather than creamy white, the final outcome is rusty coloured – obviously it needs some extra treatment to remove the residue from the burn.

Every time I have separated residue from supernatant I have check a sample or two of the supernatant for the inadvertent presence of diatoms.

With the second aliquot somewhat behind the first in treatment but not tainted by burnt material, these liquids have tended to be opalescent, even after 20 minutes of centrifuging at one cycle per minute and a half – the fastest I can sustain for any period.

Consider the likelihood of an intact diatom being present in any of the samples I have looked at.

  1. the starting material was the supernatant formed when 1 kg of the rock was freeze-thawed in distilled water (approximately an equal volume of water), Each aliquot I have was taken after partial breaking up and was about 150ml.
  2. After taking the sample it was diluted with distilled water and boiled with soap, then passed through a cycle (something like four times) of settling, decanting the supernatant and redilution to remove the mud released.
  3. The resulting solid fraction was then heated with an equal volume of the acid and treated as above, This was done for three different acid mixtures.

The principle is that the diatoms are made of a glass like material and are heavier than mud and other contaminants. They are acid resistant – because they are glassy. They should settle into the solid fraction. However some of the diatoms are very small in which case other factors come into play. I am not sure of the physics but slight surface contamination might lead to flotation due to a charged surface, or something like that. This means that some – probably the smallest ones may well remain insuspension and so be found in the various supernatant fractions. This is where the centrifuge would help as it works mainly on  density (I think). Also every time material is processed, even with a lot of care, some will be lost. I have found that my early training in quantitative analytical biochemistry helped me to retain a lot of it, but there will always be losses.

All of these factors are ones that could result in me losing diatomaceous material from the solid fraction.

The samples I have examined microscopically are taken as one drop from the material, spread on a slide. I get about 25 drops to one ml. When I have used a heavy deposit I have diluted the material in the ratio of one drop to about 2ml and taken 1 drop of that onto the slide – about a 50 times dilution. If I find any diatoms in a sample with that degree of attenuation there must be loads in the original material – see how I use exact scientific terminolgy 🙂  . When I look at the supernatant specimens and they have a very low count of diatoms I feel justified, in this instance, of considering those an acceptable loss. So I will not pursue those.

The sediment seems to be composed of a lot of broken material, however it has been around for 35,000,000 years or so so that is not unexpected. In fact it’s a bit of a miracle if anything survives intact.

2018-04-28-01.03.35 ZS PMax

This is typical with needlelike things – spicules not diatoms – but among the detritus there is one circular diatom with an ornate border, one apparently empty ring and a rectangular oval with a dark area in the centre. All of these are diatoms from Oamaru but pinning down their exact name is a bit more difficult.

Here’s another version of the ornate one:

2018-04-28-00.54.26 ZS PMaxcropped

There are quite a few of these Biddulphia too:

2018-04-28-00.55.32 ZS PMax2018-04-28-01.01.19 ZS PMax

If you look carefully there is even one hiding here in the centre of the image:

2018-04-28-01.05.07 ZS PMax

So they must be realtively common in the sample. – remember this is one drop from a massive dilution of a bit of mud washed of the surface of a rock 35m years old.

Occasionally there is a different type:

2018-04-28-01.02.27 ZS PMax

this triangular fragment might be a Triceratium – for example T. castellatum West var. castellatum.

2018-04-28-01.06.12 ZS PMax

And these are ubiquitous – I’ve found them in every sample I’ve looked at.

Although I’m ignoring the spicules they are not without interest.

Here are a couple, they’re very three dimensional and this was taken from a wet mount so it was rather difficult to get a good clear image for stacking.

The first one has a clear spike at right angles to the cross, the second is a VERY spiky object and the third is probably not a spicule, but it is an interesting shape and I have no idea what it might be.


I need now to separate the whole elements from the detritus. For this I plan to try a method I saw demonstrated at the Reading Convention in 2016 – Tony Pattinson had a tone generator and a loudspeaker. I have my lovely husband currently considering where we shall get the appropriate parts from, confounded a little by the liquidation of Maplin.

Still working on the Pleurax mounts of partially cleaned Oamaru diatomite.

This is very addictive, a single slide just keeps giving, and I stil have to get rid of additonal rubbish. But today’s crop is herewith, again the file names give a clue to what I think they are, courtesy again of Oamaru Diatoms..

I contacted oamaru diatoms  about one of the images and was saddened to hear that Nigel Charles passed away in January and the site is being maintained but not extended by a friend. It is such a useful site and he must have put hours of work into it, Thank you Nigel and thank you friend for keeping it going.

At x40 I found:


and x16 gave me these items:

I am also getting better at cleaning up the images.

Pleurax makes a difference

Even though the Oamaru material is only partially processed I didn’t want to dispose of the slides I’ve been looking at, anyway I need to continue practising to make permanent mounts. So I dried the two acid wash slides on my travel iron and mounted them in Pleurax.  This was an excellent idea because the refractive indices of diatoms and mountant complement one another and everything is so much clearer. For example Aulacodiscus-maybe x40

I haven’t previously achieved this level of clarity (stack of 14 images in Zerene stacker PMax) with this material.

Other objects I have tentatively identified are:

Biddulphia novaezealandiae a fascinating shape, and another view of what may also be a Biddulphia.

There are several forms which seem to be something like Hemiaulus, and one that I am sure is H. polycystinum

And another group which seem to have features of Melosira expectata

A Tubularia

Actinoptychus, plus a boat-shaped one I haven’t yet identified.

And then there are objects like the spiny spicules

and spool-shaped items


I then realised that the thistle shaped items freely scattered through the detritus are probably Pterotheca, it’s so easy to miss something unobtrusive.


This whole exercise is very interesting.