When I get the chance I make the time to give talks about the research that I do, and this last weekend I spoke at the Royal Photographic Societies Imaging Science Group meeting – Good Picture 2022.

The meeting brought together those of us with a passion for imaging, from widely different backgrounds, for a day of talks, and was held at the University of Westminster in London.

I gave a talk on the building of my UV microscope, and its use for looking at sunscreen emulsion structure and the imaging of the fine structure present in diatoms.

Involvement with organizations such as this helps drive the science forward and is something I recommend to anyone at any stage in their career. Face to face meetings have been hard, if not impossible, for the last few years, but things are starting to ease a little now, and it has been great to get back and see people again.

As part of running my company, I think it is important to give something back to the wider world. Wildlife is a bit of a passion of mine, and I often spend time photographing the birds in our garden. For a long time now I’ve been a volunteer with various wildlife related projects and organizations such as Surrey Wildlife Trust and the Woking Biodiversity Partnership, either helping out in person or through donations from my company. I also donate to charities which look after injured wildlife as humanity has a huge, and often detrimental, impact on the environment. I wanted to share a recent story as it highlights one of the reasons I do what I do.

A couple of months ago I went out into my front garden one sunday morning to go the bins. It was raining heavily, and as I came back towards the house I saw a bird under my car. Initially I thought it was a Sparrowhawk, but it turned out to be a Kestrel, a beautiful little bird of prey. I went towards it and it hopped off, unable to fly. My guess is that it was hit by a car as we live on a busy road. I went back inside and got a box and a blanket and went back out, spending about 5 or 10 mins gradually getting closer to the bird, and was eventually able to pick it up and put it in the box. I wouldn’t necessarily advise doing this – sharp talons and beak – but I had thick gardening gloves on.

I phoned the Wildlife Aid Foundation who are based about 15 miles away from me and they said ‘yes, bring it down to us’. I quick drive later and it was with them. It turns out that it had had a head injury, but they were hopeful of its chances. A few weeks later I was called up to say it had healed and was ready to be released and if I wanted to come and collect it so I could release it where I had found it.

Back I went and brought the Kestrel home and duly released it in the back garden (away from the traffic) where it was glad of its freedom – it was off as soon as the box lid was opened. Although I managed to grab a couple of quick photos of the release, it was all over very quickly and off it went.

The Wildlife Aid Foundation is one of the organizations I donate to when I can, however this very personal experience really brought it home to me as to why these places need to keep going and carry on doing the work they do. We all have huge financial pressures at the moment with everything that is going on in the world. If you can though, remember the wildlife when it comes to giving something to charity. Research has shown the beneficial effects spending time in nature has on our well-being, and with the rate at which we are destroying it, anything to help is a good investment in our future.

Coating of diatoms in a thin layer of metal or metal oxide has been reported for many years as a way of improving their visibility for microscopy (as examples, work I have discussed and published on slides from Horace Dall and John Dale. However it was never a common technique and is not widely used today. Recently I was sent a few slides from a fellow microscopist which contained diatoms which had had a thin (a few nm) coating of gold applied to them before mounting.

Her is an example image from one of the slides. It shows part of a diatom (likely Triceratium grande, or Triceratium favus). The resolution has been reduced for sharing here.

And the slide itself.

Imaging was done on my modified Olympus BHB microscope with a 60x Olympus Splan Apo NA 1.4 objective, and oblique illumination from below using white LED light. The image is a stack of 10 images (stacked using Zerene stacker) and photographed with a Canon R7 camera. The image shown has not been cropped – this was the full field of view.

Gold coating of of the diatoms certainly improves the visibility of the features, and I am looking forward to looking at the other samples. Thanks for reading, and if you’d like to know more about my microscopy or other aspects of my work, I can be reached here.

On saturday 15th October 2022, I attended the Quekex meeting of the Quekett Microscopical Club (of which I am a member). I had been asked to give a talk on my UV microscopy work on sunscreen and diatom imaging, and as it turned out I had also been awarded a Certificate of Technical Merit for one of my UV microscope images of a diatom (taken using 313nm light). A very successful and fun day and here’s a few pictures from it.

If you’d like to know more about my microscopy or other work, I can be reached here.

Given my ongoing research interest in UV microscopy, I am always looking out for information on the subject. A few days ago I found a brochure published by Bausch and Lomb which talked about objectives and other items they had made which were designed for use at 365nm. As I hadn’t heard about these before, and I haven’t found any more about these, I thought I would share it here.

The document was called “Ultra Violet Photomicrography at 3650Å Optics and Other Accessories”. Here’s the cover page.

3650Å is 3650 Angstroms and is the same at 365nm. These are glass objectives and are corrected so the focus point at 365nm is the same as that at 546nm, so that they can be focused in visible light and then used for imaging at 365nm. This was done to improve resolution, as a ‘half way house’ imaging in the UV with glass optics but not going to the extremes of using quartz optics and imaging down below 300nm.

They give an example of the comparison between a 546nm image and a 356nm one of a Amphipleura pellucida diatom, which shows the improvement nicely.

Here’s the information about the objectives.

Thing is, I had not heard of these before, nor have I ever seen any examples of them. If anyone has any, I’d love to see what they look like, and please feel to contact me here.

Always nice to see an article getting published. This time a piece in the Balsam Post, which is the newsletter of the Postal Microscopical Society. The article is on a set of aluminium coated diatom slides that the society has, and I looked at them with visible light and with 365nm UV using my UV microscope.

While I spend a lot of time working, it is nice to occasionally take a break and look at other samples using the equipment I’ve built. The quest for improved resolution in microscopy led me to looking and metal and metal oxide coatings for samples, and this resulted in me looking at the work of Horace Dall (who I have also written about before – here) and John Dale. The Postal Microscopical Society had a set of diatom slides by John Dale which had been coated in aluminium, so I requested those and offered to write a short article for their newsletter on what I found.

Organizations such as the Postal Microscopical Society and the Quekett Microscopical Club should not be overlooked when it comes to research. While not all the members will be actively involved in scientific research the collective knowledge of members is enormous, especially when it comes to historical work. I can do nothing but recommend anyone interested in microscopy look to join these groups. Giving something back by writing the occasional article is my way of saying thank you, and provides work which will hopefully be of help to others in the future.

As always, thanks for reading, and if you’d like to know more about my work, please feel free to contact me.

Short update today, as this is a work in progress. With UV microscopy, conventional glass lenses absorb the short wavelength UV, and become opaque. This means using quartz, fused silica, calcium fluoride, etc for lenses especially when going below 300nm. I have some commercially made UV objectives, but nothing with a low magnification – the lowest magnification I have is a 10x one (which equates to a 16mm focal length on my setup). Having a wider field of view is nicer for looking at big samples, but there is of course a trade off in resolution. This got me wondering if I could make my own objective using a simple fused silica single lens, which is the topic of this post.

To make it I went back to my old friends – Thorlabs – for the components. For those of you who don’t know, Thorlabs is an excellent optics supplier, a sort of Lego for optics geeks, and offer a huge range of components. I’ve written about them before here. For the initial attempt I decided to use a 40mm focal length, 25mm diameter, plano convex fused silica lens, as I had one already. This was mounted in a Thorlabs SM1 tube, and then a RMS to SM1 adapter so it could be screwed into a normal microscope objective port on the microscope. I also fitted another SM1 tube to act as a light baffle to help remove stray light. Here it is on the microscope, pointing at the sample.

The sample is my fused silica diatom slide for UV work. The diatom arrangement is about 2mm across, and is far too big even for a 10x objective.

Here’s how it looks using visible light (546nm).

And now with 313nm light.

The good thing is it does image the whole of the arrangement, but they are very soft images. Not a huge surprise, this was a single 40mm focal length plano convex lens after all. This was more a proof of concept than anything else. The dirt in the mount it more obvious at 313nm. Again not a massive shock as the shorter wavelength light shows more dust and dirt. Some (but not all) of the diatoms look darker at 313nm than at 546nm, and this is what I have seen with my imaging. There is also a slight change in size of the field of view when changing wavelengths – 546nm and 313nm focus at different places with this lens, so the sample needs moving as the wavelength changes. This is easier to see if I flip between the two images (click on the image for this to work).

Can I make my own objectives using stock components? Yes. Would I want to use a simple plano convex lens for my microscopy? No. Too soft and too low a contrast. I have got a couple of aspheric lenses on order from Thorlabs (one is fused silica the other in glass) to see how they work better – being aspheric I am hoping for a slightly sharper image from them.

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

Quick update today – the latest edition of the Handbook of Cosmetic Science and Technology has been released. I have a chapter in it on the use of UV imaging for skin and cosmetics, and it is great to have work in such an influential skin research textbook.

Congrats to the editors, as this one has been in the work a while (it started pre Covid).

Combining multiple images together which are taken with different focus points is a valuable tool for microscopy, as it allows thicker subjects to be imaged and for it all to be in focus. This focus issue can be a big problem with microscopy as especially with high magnification and high Numerical Aperture (NA) objectives, the depth of field can be tiny, often well under a micron. The process of of combining images like this is called Stacking. There are various piece of software to do this including Zerene Stacker, and Helicon Focus, and I personally use Zerene Stacker. Today’s post isn’t a ‘how to’ with stacking, as I am no expert in it, but I wanted to share a diatom image which I recently did for which stacking we very helpful, and to show you the effects of using it.

Firstly the final image, and then I say how it was done. This (I have been told by experts) is an Actinoptychus heliopelta diatom. It has been reduced in resolution for sharing here – the original was 5202×4823 pixels, and this copy is 1600×1483.

Also a crop from the original, and even this needed to be reduced in size to make it suitable for sharing.

This was photographed using my modified Olympus BHB microscope which I built for UV microscopy of sunscreens. The camera was a monochrome converted Nikon d800. The objective was a 60x Olympus SPlan Apo NA 1.4 oil immersion. Condenser was an Olympus Aplanat Achromat set to oblique illumination. The light source was a white LED one I made. The image was made from a stack of 18 individual photos at different focus points, combined in Zerene Stacker using what they call the Dmap process. I’ve also cleaned up the image, sharpened it, and tweaked the curves. As you can see though the majority of the diatom is in focus. The diatom was on a slide from the 2005 Meakin Collection (Petersburg, Virginia) which I got as part of a recent purchase.

So, why bother stacking? It comes back to the issue of sample thickness, and the depth of field that is possible with the objective. By taking multiple images, moving the stage vertically so that different parts of the image are in focus, during the stacking process the software then takes the ‘in-focus’ parts of each image and combines them together to produce the final image. To help visualize that, here are three images from the original stack, taken at different stage positions. These have not been cropped, but have been reduced in size for sharing. They have also not been cleaned up, as I did this at the end.

As you can see, the three images each have different parts of the diatom in focus, and one single image would not have captured the whole of the diatom AND got it in focus.

The software itself is straightforward to use, but it takes a little bit of experimenting the get the best settings that it uses for processing. This will likely depend on your setup, and to some extent personal taste when it comes to how you like your images. Give it go, and open up a whole new way of looking at your microscope images. To be clear the technique is not suitable for all subjects – if there is movement between each image then that can be a problem. This is why I do not use this for sunscreen emulsion imaging, as there is often some small degree of movement of the droplets.

As always, thanks for reading, and if you’d like to know more, please contact me here.

A long time ago I did a PhD in surface chemistry at the University of Durham. My professor (Prof Jas Pal Badyal), is still there and still doing great research. Some of my equipment is still in the lab, and I was lucky enough to visit there earlier this year and see some of the work the team are doing. During my PhD, some experiments were done which produced results which I never fully answered. However with my new interest in microscopy it has helped me look at my PhD again with different eyes, and cast light on some of my observations from over 25 years ago. Today I’d like to share these with you.

The first was purely a few nice images from a microscope slide I bought. During my PhD I worked with precious metal compounds. I’d dissolve them up in different solvents, spin coat them on to different substrates and then hit them with a cold gas plasma to reduce them down to metal layers. The goal of this was to find new ways to make catalysts on temperature sensitive substrates. At the time, I’d often get nice complex crystal structures and a few of these even made it into my PhD as rather poor quality photos. The microscope slide I got recently was of potassium platinum cyanide (a platinum compound) on glass. Under crossed polarized light, which is often used for the imaging of crystals, this is what it looks like.

The image above was taken using an 1x Olympus SPlan FL1 objective, on my modified Olympus BHB microscope. The whole circle is about 6mm across. I removed the condenser and just used the field lens to cast light onto the sample given its size. Very often I’d get complex dendritic structures like this with silver and palladium compounds, and I struggled to photograph them with what I had to hand at the time.

Here’s some more images from this slide as it is so beautiful and complex. These were taken with a higher magnification objective (a Nikon 10x NA 0.5 Fluor). If only I’d had this microscope back in my PhD….

The second thing is a bit more of a technical one, and the purchase of a book gave me some answers I could have done with when I was back in the lab. This came about because of a chance observation one day. I used to deal with gold chloride (AuCl₃) solutions. These were spin coated on to Nylon, and the result was the Nylon would slightly dissolve, and the gold salt and Nylon would mix together to form a thin layer. Normally I would then treat this with a hydrogen plasma to make a gold metal layer. To analyse the samples I used a range of different techniques, one of which was X-ray Photoelectron Spectroscopy (XPS) which involves using X-rays to irradiate a sample, kicking out electrons which are then collected and analysed. The energy of the electrons tells you about which elements are present. I noticed that with some of my gold chloride samples, if I analysed them before treating them with the hydrogen plasma, they would come out of the device looking different where the X-rays had hit the sample. They would be a different colour, often red or brown. Sometimes this sample would then change over the next day or so and become more ‘gold’ looking. Essentially, the X-rays were doing something to the sample. Here’s some of my original notes from my lab books from 1996 (and yes I still have my PhD lab books, and no my hand writing has not improved over the years). The little ‘letter box’ shaped things in the text are actually the coated Nylon samples from the experiment – I stuck them in the lab book where possible.

In the second image above, one of the samples looks to be transparent red, and this gives you an idea of what they looked like (this one never changed over time). When I originally did the work I suspected this effect was due to the X-rays breaking down the gold chloride molecule and forming nano gold clusters or colloids suspended in the Nylon polymer matrix. Depending on the precise nature of the X-ray treatment with some of these samples the gold clusters then migrated over time to form larger structures and eventually more coherent gold films. However this was a distraction from the actual work, as my main focus was plasma treatment, and I was never able to prove what was going on.

A few months back, I came across a book called “Colloids and the Ultramicroscope” by Zsigmondy, and translated by Alexander in 1909. This was for sale in the US, and being a bit of a nerd I bought it as ultramicroscopy was a technique I was interested in reading about as it was an approach to produce very high resolution images. Here’s the book.

It turned out that he had used this technique to look at gold colloids, and amazingly the book had some tables and colour plates of gold colloid solutions with different sized particles.

It does indeed look like small gold colloids have that distinct red colour which I observed – the smaller the particle the more intense the red.

Very often as scientists our experiments throw up questions which we cannot answer at the time. Unfortunately, in today’s deadline driven world, these are often seen as problems – things which slow us down and distract from the desired goal. But the key driver for a scientist is exploring and hopefully explaining the unknown. Sometimes this happens quickly, but as in this case it can take years, and inspiration often comes from unexpected sources.

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