A few weeks ago I managed to get a microscope slide of bandage from Mummy ‘1770’. Mummy cloth slides do appear occasionally and as I didn’t have one in my collection I thought it would be interesting to image. Mummy 1770 is in Manchester University and in the mid 1970s was unwrapped for study (more on that here). I presume this slide was made as part of this work. The was prepared by R. Yewdall and the slide is shown below.

Looking at the slide I got to wondering whether this would be good sample for imaging with different wavelengths of light. I was assuming the bandage had been dyed, and that using longer wavelengths might make any dyes more transparent. I did a quick and dirty set of experiments imaging the bandage with 3 different wavelengths on my microscope;

450nm – white LED light with a 450nm, 40nm FWHM bandpass filter.

560nm – white LED light with a 650nm, 10nm FWHM bandpass filter.

>830nm – halogen light with a 830 Heliopan longpass filter.

Imaging was done on my Olympus BHB microscope using a 2x Olympus Fl Splan NA 0.08 objective and photographed using a monochrome converted Nikon d850 camera. The full frame is shown and the field of view is about 3.5mm across.

With the 450nm blue light, the fibers are very optically dense, showing as black in the image. It is difficult to make out much of the structure of the bandage here. With 650nm light the fibers are now becoming more transparent and a bit more of the weave can be seen. With IR light of >830nm is used any dyes present are much more transparent and more can be seen of the weave structure itself. There are also some very dark specs visible in the bandage, which could not not be made out when imaged with the shorter wavelengths. These are presumably dirt that woven into the bandage, or have come from the breakdown of the mummy over time.

Choice of illumination has a huge impact on the final image, and simply by changing the wavelength of light being used can emphasize different aspects of the subject. Thanks for reading, and if you’d like to know more about my work I can be reached here.

Last year I was asked if I wanted to write an article for a special edition of the International Journal of Cosmetic Science which was to be celebrating the career of the well known skin research Professor Tony Rawlings. I’ve had the pleasure of working with Tony on a couple of occasions, and I’d like to think I learned a lot from him (although perhaps not as much as I should have). In my early days at P&G working on skin I got involved with what was at the time a relatively new technique – In-vivo Confocal Raman Spectroscopy of skin. This is a device which allows optical profiles of different chemicals within the Stratum Corneum, painlessly and quickly and without the need to cut biopsies out of people. After all my time and experience in the dermatology field I strongly believe that this is the most powerful tool for understanding skin to have arisen in a long time.

However, instead of just one article though I decided that two would be better. One article was to be about the development of In-vivo Confocal Raman Spectroscopy into a clinical tool which was something I was involved in back in 2004, in the early days of my research into skin. The other was to be a report on a study assessing a wide range of skin assessment methods, biophysical, spectroscopic and grading, to see how they compare for looking at dry skin and how Raman Spectroscopy provided new insight into it. This was something I ran back in 2006-7 but was never published at the time, however is a paper I’ve wanted to publish for a long time. Tony was involved with and provided many tough questions during the development of the Confocal Raman system for looking at skin, so this edition of the IJCS seemed like the right place to have this published. Despite it being a (good) few years since this was run, the techniques for looking at the skin remain fundamentally the same, so I thought it still warranted publishing.

I worked on these articles with Paul Matts, a friend and colleague of mine from my days in the Skin Care group of P&G and also intimately involved with the development of Confocal Raman Spectroscopy for the measurement of skin, and am excited to say they have just been published in the IJCS. Links below;

In vivo confocal Raman spectroscopy: The window into the skin

What defines dry skin? Correlating a range of skin hydration parameters with In Vivo Confocal Raman Spectroscopy

I understand they are going to be open for anyone to download, but if you are having issues accessing them and want to have a copy please contact me through the tab at the top of the site.

Congratulations Tony, what an amazing career and legacy. You’re a skin legend……

As always, it is great to see a new article being published. This one is in the Quekett Journal of Microscopy and is something I’ve written on initial prototyping of a new dark ground condenser for my UV research, based on the use of a fused silica Axicon lens.

Very early stages, but this shows promise and is one of many projects on the burner at the moment. As always, thanks for reading and if you’d like to know more about my work drop me an email.

As regular readers will know I have been doing a lot of microscopy research recently. I’ve ended up producing a lot of very high resolution images of different diatoms and I’ve been asked what I was planning on doing with them – was I going to produce a book or something else. I’ve settled on making a new website which is now live. It is called Diatomimaging.com (click on the link to access it).

The site is a virtual museum, with a blend of science and hopefully some art as well. On there are my images of diatoms, along with details on how they were taken. I’ve also included photos of the slide that they were imaged from. I’ve typically found that exisiting sites would share images of diatoms, but with little information on how they were taken, and often the details of the slide and slide maker were omitted. These slides and the subjects they hold are culturally and scientifically significant, with some diatom collection areas no longer even producing new material for people to work with (Oamaru in New Zealand for example).

My goal with the site is to share images of these amazing structures with the wider scientific community, but also make them available to anyone who finds beauty in nature. I would like to hope that it will prompt others with collections of these wonderful subjects to share what they have as well. Who knows, you may have a new species waiting to be discovered on a slide….

In my early days of UV microscopy, I got to wondering what the resolution was with my setup. I ended up buying a Newport USAF resolution test slide designed for really high end microscopes, and doing some tests with that to see what the resolution was when using 313nm, 365nm and 546nm light from a mercury xenon lamp (you can see that test here). However since then I have done more work on the microscope, and I have also started doing my diatom imaging. For looking at ready made diatom slides, where the coverslip and slides are glass and standard thickness, this effectively eliminates the ability to use 313nm light, and I am limited to 365nm and above. My preferred condenser (Olympus Aplanat Achromat) stops me from using 365nm light as it blocks it, and until recently I have been using filtered 450nm light as my preferred approach. Recently though I got myself a 395nm LED torch, the thought being to combine that with a 390nm Thorlabs bandpass filter. I can then use this with my preferred condenser for imaging diatom slides.

As an example of the types of images I can use this setup to create, the images below was taken on my Olympus BHB microscope using 390nm light with a 63x Leitz Pl Apo NA 1.40 objective with oil immersion, the Olympus Aplanat Achromat condenser, oil immersion, bright field (I’ve reduced the resolution sharing, but it gives an example of what can be done). The slide was by CN Walter.

This looked nice and sharp to me, but I wanted to get some more numbers to put behind this. I got my Newport test slide out again, and took three images where I changed the lighting – 390nm, 450nm, and unfiltered white LED light (400-700nm) – while keeping the rest of the setup the same. The objective was my 63x Leitz Pl Apo NA 1.40 with oil immersion, the condenser the Olympus Aplanat Achromat, oil immersion, bright field set to about NA 1.1. ISO100 for all images, although exposure time was varied so that overall image exposure was about the same for each one. Image was refocused for each one (although an Apo objective some minor correction of focus will pretty much always be needed with a NA 1.40 objective which has a tiny depth of field). Same small degree of sharpening I use for images from this objective, and finally an auto contrast. Shown below are crops from the centre of each image concentrating on the smallest sized features on the test slide.

The part of the images above to concentrate on is the ’11’ with 6 sets of bars underneath it. These dark bars are chromium deposited on the quartz substrate of the slide. Group 11 has 6 Elements below it, which become smaller and smaller. Elements 1-3 are numbered, and then 4-6 just have dots next to them (you can see 1-6 for Group 10 to help with visualizing what is going on). The bar/space widths are as follows;

Group 11, Element 1 – 244nm

Group 11, Element 2 – 218nm

Group 11, Element 3 – 194nm

Group 11, Element 4 – 173nm

Group 11, Element 5 – 154nm

Group 11, Element 6 – 137nm

There’s more info on the Newport Test slide here – mine is a Highres 2 with dark lines on a clear background.

There is a difference between the different light sources, with a higher resolution being seen with the 390nm light vs 450nm vs white light – the shorter the wavelength the better the resolution. It’s perhaps easiest to see by looking at Group 11, Element 6 (the smallest of the sets of bars below the ’11’ in the images). With unfiltered white LED light the bars of Element 6 all blur together, while they are just about visible with the 450nm light and 390nm light. The difference between 390nm and 450nm light is more subtle, but just about visible on these images.

One thing to note – the 63x Leitz objective is designed to be used with a 0.17mm thick coverslip and immersion fluid. However no coverslip was used here. The reason for this is a practical one – with a coverslip on the slide, a small amount of oil is needed between the coverslip and slide, and with the extremely small working distance of the 63x Leitz objective, there is a risk of grinding the coverslip into an extremely expensive test slide, and this is something I do not want to do. This will hamper the maximum achievable resolution a bit but in theory not too much as the refractive index of the immersion fluid should be close to that of glass, however it should allow for comparative testing between the different wavelengths.

On the plus point, resolution looks to be down below 200nm for this setup especially for the 390nm light.

Looking back at my original resolution testing, these longer wavelengths stack up very well against the deeper UV images in terms of achievable resolution. This seems odd, as shorter wavelengths improve resolutions (thank you Abbe). However the setup is different here to the previous images – here the objective and condenser are higher NA vs for the deeper UV images. The higher NA’s here improve resolution for a given wavelength. As always, the whole system needs to be considered when looking at resolution. Yes shorter wavelengths can improve resolution, but the overall effect depends on everything in the optical path. Know your kit….

Where does this leave us? The use of 390nm light is a long enough wavelength to allow for the use of standard optical elements in the microscope and standard prepared slides for imaging diatoms, and enables improved resolution compared with white light and even 450nm light. Downsides – well you’ll need a UV sensitive camera with the internal UV/IR blocking filter removed. The LED source I have is plenty powerful enough and was cheap – about 25GBP – however with cheap often comes ‘nasty’ and one quarter of the LED has already burned out after a few uses. Brightness is also not very stable, making capturing images for stacking a bit more annoying. Caveat emptor….. A more expensive UV LED setup like the types that Thorlabs make should improve the issue with brightness variation, but at a much higher cost. I could also make one myself, and may do that if I have the time and/or the inclination. I will certainly persevere with 390nm light though, and a replacement torch is on its way. If this new one burns out I’ll get some better LEDs and modify them myself.

As always, thanks for reading and if you’d like to know more about my work, I can be reached here.

Yesterday I found myself in need of an article published in the journal Diatom Research. As I do not have academic access to this journal, it was going to cost me a significant amount to download and read it (45GBP for 48 hour download access). I mentioned this on the Diatom Images Facebook page, and someone mentioned I should check out the International Society for Diatom Research, and think about becoming a member. Membership allows free access to the journal Diatom Research. And get this, annual membership is 40GBP. So by becoming a member for the year it costs me less than the prices of accessing one article from the journal, and I get as much use as I need from it, and it puts me in contact with folks with a similar interest. Win win (or should that be win, win, win).

After 2 years of doing my diatom photography, this was the first time I had heard about this group. If in doubt, ask about…..

Today’s post poses a question. When does the editing of images become too much? When doing microscopy, images have to be edited. Dust on the sensor needs to be removed, the background needs smoothing, noise needs reducing and images need sharpening. The question here though, is what is acceptable editing and what isn’t? A somewhat philosophical issue, but something I found myself thinking about with an image recently. Here’s the image, a Brightwellia coronata diatom from a strew slide of Totara, Oamaru, New Zealand, made by Ray Parkinson (reduced in resolution from the original 3500×3500 image for sharing here).

The image above was taken using my Olympus BHB microscope, using 450nm LED light, and a 63x Leitz Pl Apo NA 1.40 oil immersion objective. Brightwellia coronata is a beautiful diatom, and one which is very difficult to find undamaged. I have various strew slides from Oamaru which have broken ones on them, and some which are almost but not quite intact. A few days ago I was looking through this slide from Totara, as I’ve been having some good luck with strew slides recently. After looking through most of it, and seeing fragments I saw what looked to be an intact one, buried in the middle of a load of others. This was what I saw with a 10x Nikon Plan Apo NA 0.45 objective – the B. coronata is in the middle of the image.

As can be seen from the image above, its a busy slide and with quite a bit of debris across it. In addition to what looks to be a small B. coronata in the middle of the image, there are a couple of larger broken ones to the left and lower left of the one in the middle. In this low magnification image the main one looked to be intact, but there was some dirt at the edge of it at the 8 o’clock position. When I did a high magnification image of it using my 63x Leitz Pl Apo NA 1.40 objective, the piece of dirt was just covering the very edge of the diatom. Removing most of the dirt which wasn’t over the diatom was simple enough using the stacking software (Zerene) and blank image. However I started wondering, can I remove that piece of dirt in Photoshop by cloning a short section of the edge of the diatom that I can see and pasting it over the bit where the dirt was? Below are three crops of the area in question from the stacked image to show what I mean.

First is the stacked image from Zerene, the bulk of the dirt which is not on the edge of the diatom has been removed. It obscures the very edge of the diatom’s rim over roughly 3% of the overall circumference.

The diatom underneath did not look to be broken, it just looks like the piece of dirt is laid on top of it. The next image is with a section from the rest of the visible edge of the diatom cloned and pasted over this bit with the dirt and smoothed out (and I’ve removed some dust on the sensor – the circle at the lower right of the image above).

I then went through the final editing (removal of dirt from the sensor, denoising, sharpening, smoothing, contrast etc), and ended up with the image shown at the beginning of this article. The crop of the area of interest after all this is shown below.

And this is where I come back to my original question? When does editing of the image become too much? As a matter of work flow, I play around with brightness, contrast, sharpness etc all the time. I clone out areas where there is dust on the sensor, making a best judgement as to what to replace it with by looking at the surroundings. I remove dirt and other diatom fragments from the background of the image, and smooth it out so it doesn’t detract from the main subject. However what I don’t do with a broken diatom is replace parts of it to make it look whole again. At the end of the day I am trying to image what is there. This case is a bit of a middle ground. The diatom did not look broken from what I could see, and the dirt was covering a small part of the edge of it. I made a judgement call as to what the bit that was covered would look like and replaced it. It certainly makes the image look nicer, but is it still relevant as an image? This is one of those images where I could argue either way. If I were to put this image in an article I would feel the need to disclose that it has been edited (and how it has been edited), but this is just how I work.

Before I wrap up, here’s the slide.

Image processing is a necessary part of dealing with microscope photos. What we deem as acceptable comes down to a number of factors including our own personal preferences and where the final image will end up being used. I hope you found this interesting, and if you’d like to know more about my work I can be reached here.

Bit of an update today on a new piece of equipment for my UV microscopy work. A few days ago I was at a Quekett Microscopical Club meeting, and one of my microscopy friends approached me and offered me something they had recently got as part of a consignment of microscope parts – a condenser made by Beck and described as ‘Quartz condenser, NA 1.25, W.I.’. I nearly fell off my seat, as quartz condensers are rather rare. Anyway I jumped at the chance to have it, and this post shares some initial images of it, and shows the transmission through it in the UV.

Here’s the condenser.

It came in a mount which is slightly too big for my Olympus BHB but the condenser itself unscrews from the mount and is RMS threaded as shown below.

Last year I had a condenser holder made which is RMS threaded and fits my microscope, so this will fit just fine. It also has a small diameter so will fit through the hole in my stage.

Why the excitement? Normal glass blocks UV, especially the shorter wavelength end of the UV spectrum below 365nm. As such it is no good for imaging at 313nm or below. Quartz however lets UV through and is good with light down to at least 250nm. Unfortunately quartz condensers were both rare and expensive when originally made, and as far as I am aware no-one currently makes them, which makes finding them a challenge. While I keep an eye out for them in the usual places, finding them is uncommon, and usually needs an element of luck (as was the case this time). This one has ‘W.I.’ written on the side, which means it is designed for use with a drop of water as the immersion fluid, although for low NA applications (when the objective is below NA 1) it would probably be fine to use it dry.

As I had not seen one like this before, the first thing I did was measure the transmission spectrum through it using my Ocean Insight FX spectrometer, and got the following.

As expected, the transmission through it in the UV was good, not dropping at wavelengths below 365nm as would be expected for a normal glass one.

I have not seen one like this before, but I suspect it was originally designed for fluorescence work rather than UV microscopy (as mentioned for a couple of quartz condensers in this Beck catalogue) and is probably a simple Abbe construction. Simpler is normally better when it comes to UV.

Joining a club such as the Quekett is a great way to meet very knowledgeable and passionate folks, but also to find historically interesting items which you may otherwise never get to see, and I can well recommend it. I now look forward to trying this out for my UV microscopy work. As always, thanks for reading and if you’d like to know more about my work I can be reached here.

This is something which goes back to my early days of exploring microscopy in 2020. I bought a slightly mysterious lens, a 50x Leitz Pv NA 1.0 glycerine immersion objective, which has designed to be used with a 0.18mm thick quartz coverslip. I originally wrote about this here. At the time when I tested it I was slightly surprised to find that despite being designed to be used with glycerine as the immersion fluid and for use with a thin quartz coverslip, the UV transmission was not like a Zeiss Ultrafluar. Although offering some UV transmission especially at 365nm, it dropped at the shorter wavelengths and was blocking the light by about 320nm. At the time I didn’t expect this, as I had thought it would transmit further into the UV, however the objective had some unusual markings on it, and I wondered if this was a prototype, mock up or master copy for the factory perhaps just fitted with glass elements. Since then I have always kept a look out for another copy of the lens to test for transmission and compare with my original copy. However this is a rare objective, and it was nearly 4 years before I saw another one for sale. Today’s post shares the results of my transmission testing of this new copy, and discusses the implications of my findings.

Here are the two copies of the objective. My original one with ‘Muster-fasserei’ written on the side on the left, and the new copy on the right.

And the back side of the objectives.

There are some differences in style and labelling of the two objectives, but overall they should be the same – 50x magnification, phase contrast (Pv), NA 1.0, glycerine immersion and for use with a 0.18mm thick quartz coverslip on a 170mm tube length microscope. Except, according to the original Leitz literature, they were actually meant for a microscope, but a ‘microspectrograph’. However they do work as normal microscope objectives with a phase contrast ring in them.

So, the six million dollar question, how does the UV transmission of the two version compare? To check this I measured both of them using my Ocean Insight FX spectrometer and got the following graphs.

What does the graph above show? There are the two copies of the objective. The transmission through the original one with ‘Muster-Fasserei’ in blue, and the new copy in red. The transmission spectra for the 2 objectives are almost identical and certainly close enough for me to be happy that there are no major differences between them in terms of construction. Both have good transmission at 365nm, and then dropping in transmission below that, with them being essentially opaque by 320nm. Below 320nm we are down in the noise and although it says about 3% transmission this is just a quirk of the measurement process.

Where does this leave us? My original ‘Muster-Fasserei’ copy has the same transmission as the other copy, so optically I am happy that this is the same as the production copies. They have good UV transmission above about 350nm, but poor transmission below that, so they are not true UV lenses like Zeiss Ultrafluars or the Leitz UV objectives, but then they do not claim to be. Glycerine immersion normally would indicate UV work, and the 0.18mm thick quartz coverslip would also back that up. Now though I come back to the entry in the Leitz catalogue which described the objective for being for a ‘microspectrograph’. I suspect the use of glycerine as an immersion fluid was to reduce fluorescence as it is low fluorescence fluid. Also the quartz coverslip will be lower fluorescence than a glass one (I’ve looked at fluorescence of glass and quartz/fused silica here). Minimizing fluorescence of components in the optical train will be very important for a spectrograph which will be measuring light intensity. These were therefore probably designed to be used in a setup where low fluorescence was important, so perhaps for use with light with wavelengths from 350nm and up.

Answering questions about older microscope equipment can be a real challenge. Documents do not always exist online, and personal knowledge of folks who might have used or worked on older equipment is unfortunately disappearing rather too rapidly. This has taken a few years to get an answer but I feel I know more about them now than I did before, and as a scientist that is very important to me. As always, thanks for reading and if you’d like to know more about my work, I can be reached here.

Firstly, apologies for not posting a lot recently. My wife and I had a holiday in Tasmania in January, and I am still going through the photos from that (it’ll be done by the end of the month, fingers crossed). I have done a bit microscopy in my down time though, and it got me wondering, what would be my wish list for diatoms to image? I’m sharing some thoughts on this today, some of my wish list I have and others that I have yet to image. These images were done using my modified Olympus BHB microscope, and have been reduced in resolution for sharing here (the original are much higher resolution).

EDIT, 18th March 2024. I have managed to find an example of one of the diatoms am looking for – Truania archangelskiana – on a strew slide I had, so have included an image of that now.

I’m going to start with ones on the wish list that I have managed to find. Firstly one of the structurally prettiest diatoms I have imaged so far, Cerataulus subangulatus from the Oamaru deposit.

This was a slide made by Emiliano Bellotti and has a single example on slide. A beautiful structure as well as being a challenge to image.

Next we have Brightwellia Coronata, again from Oamaru.

This was on a strew slide from Allans Farm, Oamaru by Bernard Hartley, and is the most complete one I have found in the slides I have. It seems to be very prone to breakage, and finding a complete one is a ‘challenge’, although many Oamaru strew slides will contain fragments of them.

Next is Hydrosilicon mitra.

The slide maker here is JA Long, and the location is given as New Hebrides. It’s quite a rare diatom and extremely fragile. This one is the best I have with only a few broken ribs.

Next is Monopsia mammosa, another one from Oamaru.

Another slide by JA Long, and I do have second example of one which is more broken up. This has a couple of cracks, but that is about it.

This one I only imaged a few days ago – Actinoptychus affinis from Java.

This was a slide by Samuel H Meakin, and is almost complete apart from a bit of the edge missing. I love the patterning on this one.

Now for Triceratium nitescens on a slide by the maker RI Firth.

A beautifully shaped diatom from Barbados and one I was really happy to be able to image (the slide says ‘very rare’ and I believe that is true).

Now we get into ones that I am still actively looking for examples of, because I either only have a fragment of it, the one I have is not ideal for some reason, or ones which are so unusual I have only seen pictures of. To start with Charcotia decrescens from Antarctica.

This one is on a strew slide by Klaus Kemp, and is actually pretty good. I’m being picky here, it was at quite an angle in the strew, and made imaging difficult, so is one I keep an eye out for. I think it is also known as Chacotellia decrescens, and Actinocyclus actinochilus.

The next is one I only have a fragment of – Biddulphia pedalis from Oamaru.

Another slide by Samuel H Meakin, I have seen fragments of it, but so far have not tracked down a whole one and it seems to be quite a rare diatom. Also known as Grovea pedalis. Lovely details on it though. The image above was done using 365nm light and the short wavelength makes all the small features really pop.

The last one I do not have a individual sample of, but I have managed to find it on a strew slide by Bernard Hartley. It is Truania archangelskiana, and is shown below.

Reported locations for this one include Singiliewsky and Inza, Russia and Eidsbotn Fjord, Devon Island, Nunavut, Canada (seen here). An image of one can also be seen on the Photomacrography forum, here. A striking looking diatom and well worth looking out for for imaging.

I find imaging diatoms fascinating and rewarding, and never cease to be amazed by the varying forms. There are loads I could have listed here, but this gives a nice cross section of some of the more unusual ones. I hope you enjoy these images, and if you have any slides of these you would be open to finding a new home for, I can be reached here.