I have been using a few techniques to prepare specimens to take microphotographs of, and have just added the word Microtome to my microphotography glossary, so I figured I’d share some of this information with you today via the podcast. It’s still young and doesn’t have that many entries, but if you are interested in microphotography, you may find my glossary useful, but nothing is useful unless you know it exists. Almost a year ago now, when I started to make photographs with a microscope, in the spirit of sharing what I am learning as I learn it, I started working on a glossary. It’s as much if not more for my own purposes than anything else, but I linked it to the Microphotography menu that I added under the Posts menu at the top of the website.
With just four entries now that I’ve added Microtome, the M category is in first place with the S category at three entries in second place, but the S category has a higher word count, so it’s not worried about falling behind. As you can see from this screenshot, I am adding various terms, but also included a number of stains that I’m also using, so we’ll touch on much of this today in my continued Microphotography series.
What is a Microtome?
So, if you are like me, and had little to no experience with microscopes, you may be wondering what I Microtome is. I first heard this word about eight months ago as I started to learn how to prepare my own plant specimens for the microscope. A microtome is basically a tool to cut thin sections of your specimen so that they can be studied, and in my case, more importantly, photographed, under the microscope. Anything opaque that is much thicker than a few thousandths of a millimeter basically prevents any light from passing through it, which can make it difficult to see and photograph the specimen with a light field microscope. You can use dark field as I’ve mentioned before, where you block out the light from directly below the specimen, and allow light to spill onto it from the sides, but in light field microscopy you need a certain amount of light to pass through your specimen.
For quite a while, I was using a makeshift microtome, which was essentially just two halves of an old-fashioned razor blade. If you place your specimen, say for example, the stem of a plant, on a piece of foam in water, and cut it with two halves of a razor blade pressed together, you will often find yourself with a very thin section of the plant between the two blades. Without the water to lubricate the cut and support the thin slice, it will often just break up as you remove it from the razor blade, so the water is important.
I didn’t document the broken razor process in photographs, so I have nothing to share, but I was relatively successful at creating sections of plants using this method. Here is a photograph of a section of an Azalea stem that I cut from the bush at the foot of my apartment steps. As you can see, the light is passing through parts of the section quite nicely, because it’s thin enough to cut many of the cells and fibers across their width allowing the light to pass through, but also, although this could be thinner, the material of the plant that is left is also allowing some light through in most areas. I also like how many of the microscopic hairs can be seen along the edge of the plant here.
The red color in this specimen comes almost exclusively from the Safranin stain that I applied. I probably should dilute it more, but I really like the strong colors that I get when I simply place a drop of the stain on top of the section when it sits in probably an equal amount of water on my microscope slide, so it’s about 1:1 water to safranin. I’m no chemist or biologist, but I read that safranine binds to acidic proteoglycans in tissue, so although I’m guessing here to a degree, I imagine this is why it strongly stains the starch sheath or endodermis, which is the bright red line that you see running up the middle of the photograph.
The problem with using two razorblades is that even though you hold the dull end of the blades, you still end up with lots of tiny cuts on your fingers, and the thickness is often uneven, with the section getting gradually thicker as the stem is cut. In a bid of overcome this, I bought a hand microtome, which is the device you can see in this iPhone shot from last week, when I spent a few hours cutting and photographing the stem of one of our lunchtime strawberries.
You can see the strawberry stem inside the foam dowel that I cut in half for a couple of centimeters, then cut out a bit of a groove that I placed the stem into, and then clamp the dowel into the microtome with the screw on its side. Then I wet the razor blade that you see in the foreground, and run it along the glass surface of the microtome, cutting off a thin section of the stem. You then rotate the base of the microtome to move the specimen up slightly and cut another section. The more you rotate the base of the microtome the thicker your sections become. The goal is to cut very thin sections, but it takes practice, and I’m still getting the hang of it myself.
Here you can see a Petri dish with some water in it, and some of the sections that I tried cutting from the strawberry stem. The one in the foreground was a little too thick, but the others were quite interesting to take a look at. Here also is a photo from a few months ago, showing a number of slides of sections of a cucumber stem, which are the round sections, and there are also a number of sections of the skin of the cucumber a little further into the cucumber from the stem.
I read that the best way to transport these sections from the water to the microscope slide is to use a fine-haired brush, but I haven’t got one of those yet, so I have been improvising by using a tiny chemist spoon to scoop my sections out of the water, or sometimes I just use a drop of water from a pipette to wash the section off of the razor blade dropping it directly onto the slide, where I then apply a drop of stain.
Staining and Preparation
The amount of time that you need to stain a specimen varies from stain to stain. I’ve also stained a blood sample using Leishman’s stain, so that I would see my own blood cells, and that required quite accurate timing. Most of the time with these plant segments, I don’t worry too much about the timing, and generally leave the stain on for anything from a few minutes to ten or fifteen minutes or so, as I continue to work getting more sections. I find that it’s more important to keep the sections moist, so I don’t allow them to dry up on the slide. I either leave them in the water in the Petri dish until I’m ready to stain them, or I drop the section straight onto a slide and apply a drop of stain straight away.
After leaving the stain on the specimen for a while, I generally just drop some purified water onto it a number of times, until the water starts to look clear. In the above photo there was still quite a lot of stain around some of the sections, but I’m now trying to wash more of it away. After placing a few drops of water on the slide, I use some lens cleaning paper to draw the water away. I then place a drop of Gum Media onto the specimen if I want to create a permanent slide or Glycerine for a semi-permanent slide, although I have found that with plant sections Gum Media works a little better as it doesn’t dry up the specimen as it hardens. I also have some Canada Balsam to try but haven’t gotten around to that yet.
Horses for Courses
You can see that I used various colored stains which I’d also like to talk about a little. As I experiment with microphotography and the various sciences that it enables me to dabble in, I’m finding it fascinating to try various stains to see how they interact with the specimens I’m photographing. The Safranin that I used on the Azalea stem is beautiful, but we generally don’t associate plant stems with the color red, so I also sometimes use a light green stain, as I did for some of the cucumber stem shots, such as this one, showing three of the vascular bundles at 100X life-size magnification.
Here, for example, I used a Janus Green stain, which is actually more blue than green, but you can see how it reacted differently to the outer skin or epidermis of the cucumber, staining the cells there blue, while leaving the cells in the periderm mostly clear, and making the chloroplasts clearly visible. To me, this is completely fascinating to be able to experiment like this, and see things that I would never have dreamt I’d be able to see with a $300 microscope, although granted, it does have upgraded objectives.
I also selected a more appropriate color stain for this next image. This shows the Protoxylem Vessels near the stem of a carrot, stained with Eosin Y, which is an orange-pink-colored stain, which I thought would look more natural for a carrot. Again, it’s fascinating that we can see what are essentially the veins of the carrot stems as they enter the carrot. In some ways, it’s pure luck that I got a section right at this point, but as we know, we make our own luck. Just doing this stuff and making many different slides to look at helps me to find things that I didn’t even know I was going to stumble across.
I also used the orange-pink-colored Eosin Y stain for this section of a Gingko leaf stem, that I picked up during a walk in the city a few weeks ago. This was stained and placed between two polarizing filters to bring out that almost jewel-like color that I really like. Although most of the time I find myself attracted to higher magnification portions of these sections, for this shot, I actually did four slightly overlapping 20 frame stacks and then stitched them together, using the new stitching feature in Capture One Pro version 22, which I’m really enjoying. That gave me a 100-megapixel photograph of something barely 2 millimeters in diameter. It doesn’t get much geekier than this.
To bring the staining conversation full circle, and show one last stain, here we are back to the Azalea stem that I started off with, but this time, instead of Safranin I used Methylene Blue stain. Again, I probably should have diluted it a little more, but I really do like the strong blue that this stain provides, and love that I can take the same specimen and make so many variations.
Also, one other thing that I sometimes do, is to simply invert the color of the photograph, as that can be quite effective too. Here is a before/after screenshot of a photo of the Azalea stem with light staining using Aceto Carmine, but because it was relatively uneventful, I inverted the color completely in Capture One Pro, by taking the black point and white point on the levels tool and moving them into the opposite positions. This is a quick and easy way to invert the color using the RGB Levels. Then essentially with a color negative, I flipped it into black and white. I’ve embedded a screenshot of the levels into the screenshot of the before/after so that you can see what I did.
OK, so we’ll wrap it up there for this week. I know this sort of photography isn’t everyone’s cup of tea, but it’s keeping me sane as the pandemic keeps me indoors, and I’ve actually developed a passion for this work that I didn’t expect when I started, so I hope you’ll bear with me as I occasionally jump down this microphotography rabbit-hole. And before we finish, I’d like to say a huge thank you to our new Patreon supporters Joseph and Trevor. Thank you to all of the patrons that are helping to keep the wheels on the MBP Wagon.
Over the last week, I’ve been on safari. Not in Africa, or even a cheesy safari park, but in three drops of water on a microscope slide. I spent multiple hours looking at a number of slides with drops of river water on them from the Tama River, which runs just five minutes from our Tokyo apartment. The experience was, to a degree, relatively life-changing. I was amazed by not only the number and variety of organisms found in just a few drops but possibly more intriguing was their resilience.
Not only did they seem to be perfectly happy to live in a glass jar for around four days before I put them back in the river, but they also seemed to be mostly fine with being tapped off their root or scraped off of a piece of reed, and sandwiched between two pieces of glass for hours at a time. They just kept going about their business, rummaging around, eating, and crapping, regardless of where they were.
I actually emptied the four jars of water and various organisms back into the river last Sunday, and I’ve spent most of the last week creating a video, which I’ve embedded into this blog post below. I’d have completed this a day sooner, but DaVinci Resolve decided it would be a good idea to delete four days worth of editing while I had breakfast on Thursday, and the rest of Thursday was taken rebuilding what I’d lost, so this has taken longer than necessary, but I’m happy with the results. The video is the main content for this week, but I did want to share a few photos with you as well, and talk about some of the creatures that I met on my microscope slide.
I’d also like to warn you before you look at the video, that it may actually disturb some people to know what is in plain old river water. I found the majority of the organisms attached some plant life that was put into my sample jars, and the majority of what I saw was, in my opinion, as cute as can be. But while looking at some cute plankton, we were occasionally visited by what looked like, in comparison, I giant almost transparent anaconda snake, with lots of little four-toed legs.
I’ve put two warnings in the video, shortly before the anaconda enters the screen, and I mention how long it will be there as well if you want to overt your eyes. I personally found it fascinating, like the rest of the organisms, but I imagine some people will not be overall happy with the visit. In fact, even the plankton going about its business might disturb some people, so only watch if this sort of thing interests you. Anyway, here is the video, starting with an explanation of the project and a bit of footage of fetching the samples, followed by around 30 minutes of footage through the microscope. As you watch, keep in mind that there are two main types of plankton. Phytoplankton, which are plants, and zooplankton, which are animals. Pretty much everything in this video falls into these two groups of plankton.
I hope you enjoyed the video. Let’s also take a look at a few of my favorite photos from the project, and I’ll explain some of the challenges faced as well. First up, here is a light field photograph of a Nematode otherwise known as a roundworm. These are very common, and can exist in both water and in soil, and are also able to live in acidic substances such as vinegar. They can be parasitic, and some species will bore into the soles of human feet if you walk barefoot in a river or on soil with Nematodes in it. I’m including this shot mostly to start explaining the difficulties of photographing plankton.
Because this was light field, with light being passed directly through the microscope slide, there was quite a lot of light to work with, especially as I’ve customized my microscope to add more light via an additional LED ring light around the original single LED light, but still, to try and freeze this Nematode, which tend to wriggle around all the time, I increased my shutter speed to a 1/500 of a second. With this shutter speed, because of the higher light levels, I was able to use an ISO of 6400, which is good compared to most of the following examples I’ll share.
I then switched to dark field microscopy, which, as I explain at the start of the video, is a technique that can be used by placing an opaque disk between the light source ant the subject, and adjusting the size of the disk, the distance of the light from the slide, and the diaphragm that controls the diameter of the light, so that the light spills onto the subject from the sides rather than passing directly through it, as in the previous image.
What you are looking at here is a Lapadella, a type of Rotifer, which in my Japanese Plankton book translates as a Rabbit Rotifer, probably because the two-toed tail that you can just about see in this image looks like rabbit ears. You’ll see that better in the following images, but I wanted to include this to show how dark field microscopy illuminates the surrounding phytoplankton, and in my opinion makes for much more pleasing images. There is also less to clean up. With the previous light field image I had to clean up the background quite a lot, but this image is pretty much straight out of the camera, with just a simple tone curve applied to increase the contrast a little. Also, to try and freeze this Rotifer as it swam around, I tried working with 1/1000 of a second shutter speeds, which required ISO 20000 to get this exposure, so I was starting to push the limits a little here.
For this next image though, I changed to the 40X objective lens, to get a closer look at the Rabbit Rotifer, but with that, because of the lower light that the lens collects at this magnification, the ISO jumped up to 51200 at 1/1000 so I had to run this through On1’s NoNoise AI software to clean it up. That still produces a usable image, but definition drops slightly using these settings.
I’m also fighting with the poor image quality that I get with my camera adapter, as well as the very shallow depth of field. I have been overcoming these restrictions by doing a lot of focus stacking, but for moving subjects, focus stacking isn’t possible. I got lucky with this next photo, as this creature, which I believe is a type of Copepod, stayed very still for a while as I shot a 39 frame stack. Shortly after I finished my stack he swam away at speed, so I was very lucky, but the stack also recorded trails in the debris that flowed past the organism as I made more frames, which I thought was quite effective as a photograph as well.
Because it was stationary though, I was also able to reduce my shutter speed to 0.3 seconds and use ISO 400, so the image quality is greatly increased with this settings and the focus stack. This was one of just a few images that I was able to shoot like this though, with the majority of the rest being single shots, struggling with shutter speed and ISO settings trying to make the most of what I was being presented with.
Having learned from the first few days that I really needed to get my ISO down lower than it was going, I reduced my shutter speed to 1/250 of a second, and accepted a little more subjected movement over grain for this next image, in which I was able to get three Rabbit Rotifers in the frame together.
They are providing a top, bottom and side view, so this is one of my favorite shots that really illustrates this beautiful life from.
In the next image we can see the outlined lorica which almost look like little wings, but it’s actually just the outer shell of the Rotifer being caught by the light and actually continues down to the base of the foot.
Still at 1/250 of a second shutter speed, my ISO was a 20000 for this shot, Once again here I ran this through On1 NoNoise AI to bring the grain caused by the large dark portion of this image under control.
This final shot is another of my favorites, shot with the 40X objective, the Rabbit Rotifer was stopping still for a few moments at a time, so I reduced my shutter speed to 1/125 of a second, and got a few shots where it was still, and relatively clear. There are also a couple of air bubbles which I was intrigued to find that the plankton has a very hard time with. The surface tension of the air is impenetrable despite seeing the Rotifer interact with the air a few times. It’s like a solid wall to them.
I don’t really have anything that I’m all that happy with of the Mouse Rotifer, which appears frequently in the video. They are slightly smaller than the Rabbit Rotifer, but their behavior is just like little mice, scurrying around in their environment, eating the algae on the phytoplankton. Anyway, we’ll leave it there for now. Please do watch the video when you have time. If you are listening to this you can jump directly to the video on Vimeo with the short link https://mbp.ac/universe.
Today I’m very happy and honored to be joined by one of the premiere microscope photographers in the field, Håkan Kvarnström. Having communicated with Håkan for the last four or five months, it was a pleasure to actually catch up with him for the Podcast. I needed a few months to learn enough to be able to have even a half-intelligent conversation with a micro-photographer at Håkan’s level. It was worth the wait though, so I hope you enjoy our conversation. I had it transcribed, for those that prefer to read, and you can use the above audio player too if you want to listen to us chat directly. Either way, I hope you enjoy my conversation with Håkan Kvarnström.
Martin: Håkan, it is absolutely an honor to have you on the show today. Welcome.
Håkan: Thank you, Martin. Delighted to be here.
Martin: I am very new to microscopes or micrography. It’s been three months or four months now since I got my first microscope. I have had no experience with them until that point. So, I am looking forward to learning some stuff from you today. But tell us a little bit about yourself, how you got into photography initially before micrography, microscope photography?
Håkan: Well, I’ve had different cameras since I was a kid. I grew up in a small village in Sweden and started photography quite early having cameras, mostly analog at the time. But the photography interest took off when I got kids and I wanted to start documenting celebrations, weddings, birthdays, and family events, and also a dedicated motorcycle biker, and I wanted to document the road trips. This way, I got into photography more and more over the years.
For the last few years, besides microphotography, I’m also starting to get an interest in portraiture and other types of normal photography. I’m also moving into medium format cameras, digital, of course. I’m not only focusing on microphotography nowadays.
Martin: I see. Well, you are obviously one of the first names that come to mind. As I’ve looked at your website and seen more of your images, I’ve realized that I had seen some of your images before I bought my microscope. You’re probably one of the forerunning people in the field. I know that, as I was looking through, and I’m thinking, “Hang on, that’s one of those photos that I showed my wife to say, ‘This is why I need a microscope.’” It really is amazing to be sitting here talking with you.
We’ll touch on some of the things that you would talked about a moment ago about your other photography. It’s great that you do other types of photography, and it also plays a part in how beautiful your microphotography is as well. We’re going to talk about all of that as we get through this. When did the microscopes come onto the scene?
Håkan: I’m a scientist by heart and education. It started maybe six, seven years ago when I was reading a local website with classified ads, and I saw a very nice microscope for sale. I did have a microscope as a kid. I sold it in my late teens. But all of a sudden, I got this itch again and realized I definitely need a new microscope to explore the micro world. I’m somewhat interested in both the microworld, but also the macroworld. I’m a dedicated Star Trek fan and like stars, the universe, galaxies. It was more of a lucky strike that I found a microscope before I bought the telescope.
The good thing about a microscope is that you can do it all year round. You don’t have to have good weather, Sweden can be quite cloudy, but the microscope, I can use all year round and not dependent on light pollution and stuff like that. So, it’s a very good hobby.
Martin: I have to admit, a big part of me jumping into this year has been the pandemic. We can’t really get out. I can’t go on tours, I can’t do a lot of the things that I usually would do. That was part of it. Then for me, Don Komarechka, a good friend of mine wrote a book that I reviewed. He talked about shooting some macro work with microscope lenses, the objectives. I was looking at that and I was thinking, “Oh, this is really cool, but I don’t think I want to get into botching things together. So, I’m just going to go buy a microscope.” It was then when I saw some of your work as well.
Håkan: Actually, I got his book last week as well. It’s an amazing book.
Martin: It is. Well, I was lucky that he sent me the PDF because he wanted me to review and write a few words about it. I was lucky to get an early sneak peek at it. You got into the microscope, you told me that you recently have started working with a 60x water immersion lens, and that to me, I was thinking okay, so I knew about– My microscope came with a 100x oil immersion lens, but I didn’t even realize that there was such a thing as a water immersion lens. Tell us a little bit about that, is it really difficult to work with liquids and lenses?
Håkan: The water immersion is quite good. The basic problem with microscopy is that the higher magnification you want, the more important it is to have a good level of refraction indexes. The whole light chain going from the condenser to the specimen, through the objective and so on, the higher refractive index you get, the more resolution you can squeeze out of the system in a way. The problem with the air gap, normally, if you have lower magnification objectives, you have an air gap between the specimen and the objective, but you can’t have that for 60x, 100x, and even higher magnifications because the image quality will degrade. Then, you use immersion. The most common way is to use oil. The problem with oil of course is that it’s very messy and you get oil everywhere. And you need to clean the objectives using special cleaning fluids and stuff, so it’s something I try to avoid, even though it gives you probably the best resolution in most cases.
Water immersion is a middle ground. You have lower refractive index, but it’s good enough to dramatically increase the resolution compared to using an air gap, and you don’t have to deal with a mess. You can just wipe it off with a lens tissue afterward. It’s very simple to use, very quick, no smell, no mess, and you get really good image qualities.
The other advantage of using water immersion is that, if you compared to oil immersion where you might have a working distance of maybe 0.15 millimeters, you get double that with water immersion. So, you can image much thicker specimens. If you have a small animal, for example, or if you have really thick algae, you can get a better image quality because of the increased working distance. I can definitely recommend using water immersion lenses.
Martin: This is me trying to learn from a master here, not so much providing something for the audience. I’m totally selfishly trying to learn something here. When you’re using either water oil, you’ve got your specimen under a coverslip and you’ve got the oil on top of the coverslip?
Håkan: Yeah. That’s the way. First of all, you have a glass slide where you put your specimen and then you put a drop of water on the glass slide and then you cover it with a coverslip. So, you have glass, water, glass, and then you put water on top of the coverslip again to go into the objective. You have an uninterrupted chain of high refractive indices going from the bottom of the cover glass all the way through the objective. That is what gives you the increase in resolution. There are also objectives that you can dip directly into the specimen, but those are more for in vitro procedures when you do live-cell imaging. Not very good resolution perhaps, but for specific purposes only.
Martin: I recently replaced the– well, filled the four slots on the turret of my microscope with the plan achromatic lenses. The highest magnification I got was a 60, but it’s an air 60, it’s not made for going in water. I understand what you’re saying. The jump from 40 to 60 even, and you can see that the image, the resolution, drops relatively quickly when you switch them out. But it is still really nice to be able to get in that close and still be able to see. I’ve still got the 100x oil objective, so I think I’m going to put that on and give that a try at some point.
Håkan: Yeah, definitely. In my view, the 40x air is difficult enough to use. I rarely go with an air gap higher than 40x magnification because it becomes almost impossible to get good image quality. I have a long working distance 50x objective, which is designed to be used without the coverslip. They are used for more macro-like work. You don’t really have an insect eye, for example, or something you want. But using a coverslip and then have a 60x air will not give you much quality, I would say.
Martin: I haven’t been using it with coverslips. I haven’t really used coverslips very much at all yet, but I’ve got a few questions about staining and some of the other things later. So, I’ll save that.
Håkan: It’s very important. It’s only the really low magnification objectives like for 4x and to some degree, 10x can be used without coverslips. Otherwise, they need to be specially designed at higher magnifications to be used without the coverslip. There are special objectives you need to use if you don’t have a coverslip on. Otherwise, you will not get any image quality at all.
Martin: Well, the 40x objective works pretty well, but the 60, I have found it’s pretty difficult to get anything decent with. It’s nice to have the plan achromatic, that makes a big difference over the original ones that came on my relatively cheap microscope. So, that’s nice just there.
Håkan: Agree. Microscope lenses are exactly like cameras, or normal photography, it’s not the camera body that is important. It’s the lenses. You have to put all your money and spending on the right lenses that will give you the quality you need, not the camera or the microscope stand as such.
Martin: Yeah. You talked in your email about the importance of camera techniques, and good composition and lighting. Tell us a little bit more about how that plays into your microphotography.
Håkan: That was one of the reasons as well as I started to learn more about more normal photography, about composition, about how to use lighting and flashes and how to work with the foreground, the background and the subject, and try to use well-known photography techniques also to improve the quality of the images from using the microscope. The basic principles on what makes a good photograph is still valid even though it’s a microphotograph. If you look at my images, many of them are quite have clean backgrounds and so on. But I’m starting now to change my artistic language, if you like to have more details in the background, trying to use different foregrounds, and also working with not overdo the focus stacking part, so everything gets sharp, but also try to make out of focus areas to use the bokeh, also in microphotography. By learning normal photography better and understanding the masters of photography over the years, I think I can also start making better microphotographs, which appeals to a wider audience perhaps, not only the micro nerds probably.
Martin: I am fully with you. I found that as I’ve done certain work with certain specimens that I’ve left a fair amount out of focus. Because we are all working with such shallow depth of field, pretty much everything has to be focused stack to a degree. But I agree, you don’t have to have sharp from front to back. It’s much better sometimes to just let it fade off into the bokeh as they say.
Håkan: Yeah. You are way ahead of me there in terms of maturity as a photographer, because I have an engineering background, and for an engineer, if you can make it sharp, you make it sharp. As an artist, you have to think differently. I guess I’m maturing later. I have recently learned what makes a good photograph. I’m learning all the time. Well, I haven’t really implemented that fully into my micrographs yet, but I’m working on it.
Martin: Well, there’s one of the photographs. You’ve sent me eight photographs to talk about, but one of them has really nice layers of focus with the specimens in the background, and a few in the foreground that is very much like you just said. You’re doing it, it’s there, and it’s beautiful.
Håkan: I’m trying.
Martin: You’re achieving.
Håkan: Trying to use the techniques I’ve learned in other fields of photography.
Martin: Yeah, we might as well right now start to work through your images. You’ve sent me eight shots. The first one, can I guess what this is before you tell me, just in case?
Martin: Is it fishing wire?
Martin: So, it’s a natural organism?
Håkan: It is.
Martin: Oh, okay. Then, I have no idea. I thought it was fishing wire. [laughs]
Håkan: Do you want to know what it is?
Martin: Yes, please.
Håkan: It is the hair from my 10-year-old daughter, which I tied in a knot and photographed.
Martin: It looks as though you’ve polarized some there as well, you’ve got some color coming through.
Håkan: Yes. I used polarized light. That’s one of the tricks you should learn, or everyone should learn using microscopy because it’s very cheap and very easy to do. You have to have a set of polarizers, and you can twist the polarizers to different angles and get different color effects.
Martin: Most of what I’ve done so far has been exactly that. It’s such a lot of fun.
Håkan: It’s a lot of fun. Then, I tied the hair in a knot and I used super glue to glue the knot, the hair strand to a piece of rubber, into each end to a piece of rubber. And I used super glue to get it to have exactly the right tension. The problem I had was sometimes I tied the knot too hard, and sometimes I tied the knot too loose. So, it took a while before I found the perfect knot. It’s very difficult to see because it’s so small, so you have to guess a bit and try it out. In the end, I got the results I wanted. I used a 20x objective for that one.
Martin: Is your daughter fair-haired then?
Håkan: She has white hair.
Martin: Okay, that’s why it looks so translucent. It’s such a beautiful shot. The coloring, not so much the color of the hair, but the background and everything, what did you use for the background?
Håkan: Well, I used the retarder. I have a retarder in the microscope as well. More or less, if you tweak the retarder, you can get any color you like. I usually try the different dials and try different angles to get the colors I like and want, and that’s where I end up. It’s trial and error more or less.
Martin: I’m getting loads of ideas here. This is such fun. I realize now, of course, as you rotate one of the two polarizers, you go from white to black, and then there are loads of shades between. I see now.
Håkan: Yes. It’s amazing to work with those filters. You can get any color you like, more or less. It looks more complicated than it is, I think when you are playing around with it.
Martin: Oh, yeah. I know now that I could have bought some for like $100 that fit onto my microscope, but I couldn’t find anything that I could use. So, I started with old polarizing filters, just sort of wedging them in here and there. And then, I bought some circular drill bits, and drilled circles out of polarizer filters and cleaned the edges up, and I’ve actually inserted those. One in the filter holder above my condenser. The other one, I take the head off of my microscope and put it in on top of the tube where the light comes up.
Håkan: Yeah, that works.
Martin: Yeah, it’s working pretty good.
Håkan: This one, I used to 20x objective, which was a long working distance, designed to be used without a coverslip. So, that’s why I got the resolution. You can actually see the scales on the surface of the hair.
Martin: That’s the next thing. I’ve got the plan achromatic 20x in my Amazon cart. It’s waiting for the next time, and they’re not that expensive. The ones that I’ve been getting, it’s like, I think it’s $60 for the next one. I’ll get that.
Håkan: It’s a good investment. If there’s anything you should spend money on in microscopy, it’s the objectives, so that’s the secret to everything. It’s different depending on what you’re aiming for, but if you want to do video, for example, it’s to have a longer working distance, so you can get more depth of field because video gets really boring if you only see one half of micrometer every time.
Håkan: And then you have to move back and forth to get focus. Then, I use longer working distance objectives to get more in focus. You lose resolution, but you gain depth of field.
Martin: Yeah. I’ve done a few videos because it’s been so amazing to see some of the stuff that I’m looking at. When you’re having to tweak the focus all the time, it’s much better when you use the 4x or 10x.
Let’s take a look at the next one. This is amazing. Tell us about the second image you sent.
Håkan: Yes. That’s a green alga. It’s one of the biggest green algae, it’s called a Microasterias, and it can be up to maybe 0.3 millimeters, 300 micrometers in diameter. That one is actually stained. So, I stained it with calcofluor white, which is a really useful stain. It stained the outer shell in a way in blue, so it radiates in blue light. The red one, you can see is the red parts, is the chlorophyll that autofluoresces. All chlorophyll autofluoresce when you shine them with ultraviolet light.
Martin: Oh, I see.
Håkan: So, that’s the natural light. Blue parts are stained.
Martin: Ultraviolet light, amazing.
Håkan: Well, it turned out to look like a man with two eyes and a nose.
Martin: How are you getting the light in there? Are you shining in from the side or have you got something in your–?
Håkan: All I have a fluorescence attachment to the microscope. So, it’s like a top light you use. Instead of illuminating the specimen from below, you illuminate it from above. There’s this small mirror cube with fluorescent filters in them. So, you radiate them with a very wide bandwidth light source ranging from 300 nanometers up to maybe 1000 nanometers. And then, you have filters to only let through specific wavelengths. Let’s say in this case, I used I think a 340-nanometer filter. So, specific stains go with a specific wavelength, then you can filter out the rest. Then, you get the black background because everything is filtered out except for the radiation or the photons with a specific wavelength designed for that stain. It’s a very useful technique to get higher resolution and to get specific features of a specimen to be visible, and you can take away the rest.
Martin: Wow. This is probably one of these shots that I would have said to my wife, “Look, I need a microscope because of this.” And now, I’ve found that sometimes they’re still out of reach because I need a particular type of microscope. But it’s all great, it’s such amazing work.
Tell us about the next one.
Håkan: The next one is also a green alga called Botryococcus braunii, is the Latin name for this. That is an interesting alga because it contains so much oil. I think 40% of the biomass is oil. When you put it between the cover glass and the slide, and the water starts to evaporate, it squeezes on the alga. When it squeezes on the algae or the weight of the cover glass, pushes out the oil in small droplets around the surface. So, you get this fantastic effect where you can see all the small oil droplets around the edges of the specimen. In fact, I think this type of algae can be used to produce biofuels, so you can grow them, even though they grow slowly, but still, there have been trials to do biofuel from these types of algae.
Martin: I was thinking that. That’s amazing. I would have guessed that they were air bubbles, but I thought, “He’s too careful to get air bubbles in there.”
Håkan: No, it’s oil. If you look closely at the image, you can see the channels inside where the oil flows.
Martin: It’s amazing, absolutely beautiful. The next one, this beautifully symmetrical shot. Tell us about this.
Håkan: Yeah, that’s Cosmarium, it’s also a green alga, and that is also polarized light. There I think I used some retarders and filters as well to get the sort of shining effect in the background. It looks like a world within. It looks like there’s some kind of a shell where you have a secret world inside the– you can actually see how transparent the shell is, and you can see all the chloroplasts inside. I think it’s very beautiful in a way.
Martin: Oh, absolutely.
Håkan: It’s so interesting that all these algae you have in the sea actually produce an equal amount of oxygen as the rain forests do. Even though you can see them if you take the diatoms and the algae you have in the sea, and also the cyanobacteria, are one of the major contributors to oxygen production and also the carbon capture in the world. They are amazingly important, even though most people have never seen them.
Martin: And that is why it’s so important that global warming makes the seas too hot and then this stuff starts to die off, or you end up with too much of one. We have this thing in Japan. I haven’t a clue what you would call it in English, but it’s like the red tide if I– do you get what I’m saying.?
Håkan: Yeah, I know what it is.
Martin: It’s a type of red algae, but it grows so thick, actually the algae itself changes the temperature of the water below because the light can’t get through it. It’s crazy stuff.
Håkan: It’s important with the balance, it can’t be too warm, it can’t be too cold, you have to be careful. Also, if it becomes too many of them, they fall to the floor of the sea and they start consuming oxygen when they break down and creating these dead bottoms, where the fish dies and so on. Finding the right balance is important, and I think these are too important to be ignored. You have to work on–
Martin: Yeah. Nature did a really good job of finding that balance until we started messing it up.
Håkan: Yes, unfortunately.
Martin: Oh, dear. Okay, the next one, this is simply amazing. It looks like wheat, the head of a wheat plant.
Håkan: Yeah, many people say that it looks like wheat, but it’s called Dinobryon, it’s also a Latin name. It’s actually a colony of– it’s called golden algae. They are quite interesting because they have these two hairs at the front, flagella. They can actually swim, so they can move around in the water. And you can see there’s a red spot in each one of them, and they can sense light, so they can swim towards the light.
Martin: Oh, wow. Are these classified fully as an animal then or–?
Håkan: No, it’s still an alga.
Martin: Still an alga, wow.
Håkan: And they are also very useful or they are very efficient in cleaning lakes and ponds because they eat bacteria. If there’s a bacteria-rich lake, if you have a lot of these, they will eat the bacteria and convert them to biomass instead. Very useful in many ways, but they nice smell bad, unfortunately.
Martin: Really? Oh, dear. Well, they don’t look smelly. They look absolutely beautiful.
Håkan: Yeah, looks can be deceiving.
Martin: I’m looking at the same photo that I see in your background. Is this a feather?
Håkan: Yeah, it’s a peacock feather.
Martin: Oh, wow.
Håkan: I think that’s 10x magnification. I also illuminate from the top, so I use LED lights, I think I had four different LED lights. And then, I use ping pong balls like a lightbox or what you call a softbox. I put one ping pong ball over the specimen, and then I put ping pong balls on each light to get the diffused light, really diffused light as possible. And then, I turned on also the normal illuminator on the microscope to get the very faint background light. Because I had such a low voltage on, it turns almost red or orange. That’s why you got the orange color.
Martin: Yeah, that’s amazing. Were the ping pong balls white?
Håkan: Yeah. I use white.
Martin: They’re great diffusers.
Håkan: I use two sizes. I have the normal ones, which I normally put for smaller specimens, and I cut a hole for the objective. So, the objective goes inside half of the ball.
Martin: Oh, wow.
Håkan: And then I have a bigger jumbo ball size for the bigger specimens. It’s a very good trick actually, you should try it.
Martin: Yeah. I can imagine it because it’s thin, and yet it’s a good shape. Obviously, probably 50% translucent or cell size. Amazing. I’m learning so much. This next one is the one that I was talking about earlier, where you’ve got the different layers of bokeh with the background containing stuff. Tell us about this.
Håkan: Yes, that specimen is called Xanthidium. It’s also some kind of Desmid. It belongs to the group of Desmids. It’s a green alga as well. In this case, I used focus stacking, but I use focus stack them in, I think three or four sections. I’ve realized that it’s very difficult to do focus stacking and get everything perfect at once. You need to break it down into several stacks. So here I use the stack with the surrounding algae, the smaller ones with one stack, and I used this one set of the stack for the foreground, the specimen in this case. And then, I actually took a single shot where almost everything was out of focus for the real background. And then, you need to combine these images into one image when you do the focus stacking, because you can do it in the stacker if you include the images, and then you do retouching in Zerene Stacker or Helicon Focus, for example. But it takes some work. It’s done at the same time but requires more manual work to get their final result.
Martin: I see. I’ve really not used the retouching module in Helicon Focus yet. I’ve had a few plays with it, but it always seems a bit clunky, I’d probably end up jumping into Photoshop to do that. But I see what you’re saying. I definitely can understand the necessity and the reasons why you would do that. One stack for a certain area, another stack for other areas.
Håkan: I think that’s a very good trick, because if you have two deep stacks, for example, if you do more macro-like work, when you have thicker specimens, you might have hundreds of photographs that you need to stack. The problem is that some blurry areas might become sharp again because you’re reaching deeper into the specimen. And then, the stacker gets confused. Is it sharp or should it be out of focus? And then, you get artifacts in the image, so then you really need to break it down. Okay, focus on the surface first, and then you get the surface clean, then you can focus on the other parts later, and so on, and I try to divide and conquer your way forward to get to a good final image. It can take hours. You can spend hours on this. But the results, I think, it’s getting better when you are doing it like that, especially for the deeper stacks.
Martin: That’s great advice, I’ll give it a try. I’m not a very patient postprocessor. The good thing with Helicon that I found is that once you’ve processed a stack, you can reprocess it with different settings pretty quickly. I’ve been playing with different settings and going through, but I get a little bit frustrated with the retouching because you click something, and then it takes a while even with an eight-core CPU iMac. So, that does get a little bit on the edge of my limits of patience with photo retouching.
Håkan: Normally, I use two stackers. I use Zerene Stacker, which is one commonly used software in photography. I use Helicon Focus, which is the other one. I think they’re both good. They both have strengths and weaknesses. The retouching module is better in Zerene. It’s easier to use. I use that quite a lot actually because let’s say you have a stationary specimen, but you have maybe bacteria swimming around or you have other small creatures swimming around, moving during your shoot, they will leave streaks when you stack, so you get black streaks on the image, a lot of black streaks. And then, I also have a trick, I always shoot a way out of focus image, which is very blurry, so I get the perfect background, and then I include that image into the stack. Even though I won’t use it for any of the focused parts, I can remove all the streaks by selecting that image as the source and then just painting away all the streaks with it out of focus background. So, that is a good trick to use if you want to clean up the background, and maybe you don’t have an image in the stack that is completely clean. That’s why I really go far beyond focus to just get that image into the stack as well.
Martin: That’s such great advice. I wouldn’t have thought of that. So, yeah, I’ll write that one down. Another great piece of advice. This final one looks like Christmas crackers. What’s this?
Håkan: That’s also a green alga called spirogyra. It’s very interesting because the chloroplasts have a spiral shape, so they are like a tube with spirals inside. If you see the middle parts, you can see them with pure autofluorescence, meaning you can only see the red shape, the red spiral. The other ones that are green as well have been stained. It was a lucky strike here as well. I took three different stains, and I mixed them in various portions in different bottles. And then, I added the spirogyra into those, and I mixed it. And then, I pulled strains up from the jars and rinsed them, put them on the slide. Some of them were stained, some were not, so I got this pattern. So, I tried to stretch them to straighten them out. And, of course, they are no thicker than a strain of hair. So, they’re really, really thin.
Martin: Wow. That’s amazing that you not only did that with the different stains, but you were able to get them all nicely spread out and everything.
Håkan: Well, if you have enough of them, and you pull them out on a piece of glass, you have to look, of course, to find that small portion. 99% of the slide looks like a mess. You need to look for the composition you want where something looks nice. And then, sometimes you’re lucky, and sometimes you’re not lucky. Sometimes, you need to crop a bit to get what you’re– But that’s the problem with microscopy as well, you don’t have a zoom. Based on the magnification you have selected, you have to live with the composition you get because you can zoom in, you can’t move closer, you can’t move away, and you have no zoom lens, so you have to live with what you get and just crop instead.
Martin: They call the 4x a scanning lens, don’t they?
Martin: It’s so amazing, just scanning across– I scan with the 10x as well, depending on what I’m doing. But just looking across a slide and seeing everything down there, and then zooming in– Well, not zooming, change your objective and getting a closer look, just amazing.
Håkan: If you’re moving up to maybe 40 or 60 or 100, it’s almost impossible to scan a slide because it will take forever. And by the time you’re done, the water has evaporated and you have to start over again. So, it’s very good to go back quickly to lower magnification to see what you want to focus on.
Martin: Yeah, that’s a good point. The thing with me so far is that most of what I’ve been doing has just been crystals, and they hardly change for days. So, it’s a lot easier to have fun with that. I can imagine the–
Håkan: Yeah. Now, that’s also a really interesting specimen to work with. Different chemicals and crystals and salts and whatever you have and to see– amazing shapes and patterns coming from those crystals, especially if you’re using polarized light.
Martin: Yeah. We watched a good movie on a Saturday afternoon. I have to admit that a tear dropped down from my eye, and I thought, “Hang on.” While it was still on my eye, I ran upstairs and got a slide, dropped it onto the slide, and then let it dry and polarized. It was amazing. All of the patterns that form just in the salts in a teardrop, it was pretty amazing.
Håkan: Yeah. I guess, it’s the salt as you said, that does the trick. It crystallizes.
Martin: Yeah. Almost like snowflakes in some parts. We’ve just seen some amazing specimens. W\here do you find them?
Håkan: Well, that’s the good part with microscopy as well as your hobby, that you don’t have to, or if you compare to nature photography, in general, you have to travel. People go to Iceland, Greenland, Antarctica, wherever they need to go to find good– There’s inflation in photography that you have to have all those exotic places to go to. I think with microscopy, you don’t have to go far. You can go outside and you can find the closest pond or you can find an insect or you can find a lake, you can have a net, and drag the net in the water for one, two, three minutes. And you will have endless of material to look at.
Even though I’ve been doing this for like five, six, seven years, every time I look, I see something I have never seen before. They’re millions of shapes and species in those samples. I am living close to the King’s castle here in Sweden, Drottningholm, it’s called. They have this English park where they have a huge number of ponds and small islands, and you can walk around, it’s open to the public so you can get specimens there. That’s the main source I use for all my findings.
Martin: So, that’s mostly freshwater, then?
Håkan: It’s only freshwater, I would say. I haven’t done much saltwater photography at all. I’m living close to a lake, which I’m using, and I live close to this park, where I take my samples. On occasion, I have also looked at more marine saltwater specimens, but not that much actually. Mostly freshwater.
Martin: That’s good to hear. I’ve got a river near me, although rivers are running water, there is some sort of like stagnant pools at the sides in the summertime, and there’s always algae in there. So, I’ve been trying to make a day when I can just go down with a couple of tubs and scoop some out and see what I can find.
Håkan: Yeah. But the trick, if you have running water, is to buy one of those sampling nets. So, you can put the net into the running water for an hour or so and come back and get the net. And then, you have maybe collected quite a few things.
Martin: I see. That’s a good idea.
Håkan: Getting more condensed, so you don’t have to look– You can only look at one drop at a time. It’s very good to have a condensed sample of material. Otherwise, you have to look forever.
Martin: I’ll be on Amazon after talking and see if I can find one. Excellent. What do you do to keep them alive? I know that you were saying earlier that you have live specimens on your slides. What do you do to keep them alive?
Håkan: Well, I use simple glass jars. I collect them, I try to figure out what I have, using the microscope to see what type of algae or what type of animals I have in there, and I try to find the right conditions for them. I read up. I tried to read books and see what they like, what type of nutrition they need, how much light they would like to have. Some species want a lot of light, like green alga, some want less. Like cyanobacteria, they don’t want much light. I have a table with wheels that I can push back and forth. And then, I have one growing lamp that I use so I can adjust the light by moving the table back and forth depending on what I’m growing at the time.
And then you use nutrition, like BG-11, which is a common one. I also buy nutrition from Carolina Biological in the US. They have this Alga-Gro, which I use. But in many cases, you can buy any plant nutrition bottles from the grocery store to kick off some rapid growth.
Martin: We talked earlier a little bit about the staining. What sort of things do you do to prepare? So, you’re going to sit down for a session and try and capture some of these algae? What sort of things do you do to prepare that slide for viewing and photographing?
Håkan: Well, the majority of the specimens I photograph, I don’t do any real preparation. I just take a drop and I put them on the slide. I put the cover glass on and I start photographing. The advantage with my microscope is I’m using also a contrast technique called DIC, Differential Interference Contrast, which is like optical staining. It uses differences in refraction index to highlight edges and highlight differences. So, you can get increased contrast just by using optical means. Therefore, you don’t need any stains.
But for fluorescence microscopy, you need stain because then something needs to be shining when you light them with ultraviolet light. So, therefore, I have used various, like calcofluor white, another one is called eosin, there’s also one called acridine orange, I think is called, which I use, which is like an orange color. And then, you put them into the stain for a few minutes, and then you rinse them in water, and then you put them on the slide and put a drop of water on and the cover glass on, and then you blast them with various frequencies of light only, and to see what happens.
But the most interesting part, I think, is to find specimens that fluoresce by themselves. You don’t have to stain them, you can just try it out with– you can buy a flashlight, which sends out around 300 nanometers or 350 ultraviolet light. And you can go around in the garden, try and see what gives red light, what gives some other color, blue light. And if you find something of interest, you bring it to the microscope, and you can use the same technique there to really see the specimen, shining and different colors by themselves.
Martin: Wow. I’ll have to try that.
Håkan: That’s a really good trick.
Martin: Yeah. I have some small ultraviolet, like keyholders with one LED on it.
Håkan: That is good enough, you can try that. If you take that one, and then you shine that onto some green alga or green leaf, and you should see the red immediately.
Martin: Well, I’ll take it out and have a play. You already mentioned a number of these to eliminate your specimens. I imagine that you can relatively easily do darkfield work with your– do you do darkfield at all?
Håkan: I do. The simplest one, of course, being bright field, which is just having a plain light source, and you don’t do anything with filters and stuff, and that gives you poor contrast. It’s difficult, and also, since many of the specimens you are trying to look at are more or less transparent. They don’t have any color. They are completely transparent, especially if they’re swimming in water. So, it’s very difficult to get good contrast. But there, you can use the various techniques to improve that, DIC being one of them, which unfortunately is quite expensive compared to many of the others. The most effective, I will say, and the most beautiful, perhaps in a way is darkfield, as you say. Having a black disk, a darkfield stop that you put on top of your condenser or inside your condenser to avoid direct illumination onto the specimen. It gets lights from the sides, from the top and it gets a completely black background and very colorful images. Not really many details but, like art, you don’t need all the details all the time. It’s more like the colors and the shapes. A darkfield is very simple to build yourself as well. You didn’t have to buy expensive equipment, you can just use what you–
Martin: The listeners will be able to hear it rattling around, but I bought a set of rings that go into my filter on eBay. Someone’s making them with a 3D printer. I’ve had a lot of fun with it. I tried making them myself, to begin with, with a marker pen and some clear files and everything, and then it got everywhere. I’m like, “Oh, no, I need to just buy some.” But they were like $30 for a full set.
Håkan: Yeah, but definitely worth every penny. It’s an amazing technique and it’s very simple to use as well. The problem normally with darkfield or in many cases is, it works up to the 20x objective. Above the 20x objective, it’s very difficult. You need special equipment, and then you need to use oil. You have to oil the condenser to the underside of the glass slide which creates a mess because if you then move the slide back and forth on the stage, the oil starts to drag off the stage. But if you have the patience and if you have the time, it gives you extremely good resolution, even at high magnification and the darkfield, so it’s a very beautiful image.
Martin: I’m going to have to talk to my wife and get permission to buy that 20x, that’s in my car because it’s definitely necessary. The gap between 10 and 40 is huge in size. It will be really nice to have something between–
Håkan: I would say the 20x is, I think, my most used objective.
Martin: I can imagine.
Håkan: 20x is really good. I think that’s extremely important to have. Definitely convince your wife.
Martin: It’s not a huge amount of money. I wouldn’t get onto her about that if she’s in a good mood later. And it’s crazy because it’s a business expense, but this is how it is, especially with the corona days.
Håkan: Yeah, I understand.
Martin: I’ve got a number of questions left, and I’m aware that you’ve got– we don’t want to keep you too long. So, we’ll try and jump through these a little bit. One of the things that we are going to talk about was depth of field. We’ve mentioned this a number of times. We are working with such shallow depth of field, is there anything that we haven’t already mentioned that you do to work with that or overcome it?
Håkan: Well, that’s the tricky part. The more resolution you want or need, the higher numerical aperture you need to have on your objective, and the less depth of field you’d get. So, it’s a combination that is not very good in a way, making it more and more difficult to get decent images. And also, then it becomes very sensitive to the distance between the specimen and the cover glass. The higher up you go, the more sensitive and more difficult it will be to get good images even though you’re using focus stacking. My general recommendation is to if you want to take pictures, don’t select objectives with too small depth of field. Use objectives with slightly lower numerical aperture, that gives you more freedom to move up and down and give more depth of field. Also, for video, it’s crucial, of course.
It’s very difficult because compared to normal photography, when you have meters or at least decimeters or inches of depth of field, here you have micrometers or half of a micrometer. So, it’s a constant challenge, of course, to work on those, especially at high magnifications. For the 4x and 10x, I think it’s fairly okay.
Martin: You could get away with single photos and lots of bokeh with the 4 or 10. I have a note as I hear about ISO. I’ve been finding that I’ve been using ISOs between– sometimes rather than letting my shutter speed get too low, increasing the ISO to like 800 or 1200, 1600. What sort of ranges are you working with, with your ISO?
Håkan: Well, it depends. Normally, the default mode I do– in most cases we have fairly stationary specimens, they don’t move very fast or it can be an alga, they are sitting still, so you don’t have to have a fast shutter speed. You can be quite slow. So, I set the ISO at 100, and then I just adjust the shutter speed to get decent lighting, to get the exposure right. The aperture is fixed, you can’t change that for your microphone. It is what it is. So, the only two things you can play with are ISO and shutter speed. So, I adjust the shutter speed, but if I have moving specimens like I shot some vorticella that was moving really fast, then I need to increase the ISO to get the shutter speed I need, and maybe go down to 1000, 2000, or something like that, then I need to go up fairly high. But I do have a 100-watt lamp on my microscope, so I have quite a good light. I do have a strong light– [crosstalk]
Martin: That’s the thing. Mine’s only got a cheap light. I need to see if I can actually change that out.
Håkan: Yeah, because that’s really important to have good lighting. You have really strong light, you can tweak it up dramatically compared to the 20 watts that you might have.
Martin: Yeah, I’ll have to take a look. I don’t know if I can even change it. I imagine I can with a bit of wrangling.
Håkan: When I do fluorescence photography, it’s different because those autofluorescence capabilities of these specimens, they’ve very weak lights, so I need to go up to maybe 3000, 4000 in ISO, and then I would need to work with noise reduction instead postprocessing, and also deconvolution to some degree because I don’t want to expose for like 30 seconds. I’m trying to avoid that. I try to do maybe one or two seconds but then increase ISO. It’s a mix, but you need to have a good camera. I use normal mirrorless cameras, I don’t use specific microscope cameras. It’s better to use regular cameras, which–
Martin: I decided that from the start. Even the expensive microscope cameras are like 20 megapixels. I wanted a little bit more wiggle room than that.
Håkan: I totally agree.
Martin: That’s the main thing with me at the moment. Without breaking the bank, and even this, it’s going to take a lot of wiggling around financially, but I’ve got to get a better camera adapter. That’s the weak link in my system at the moment.
Håkan: Yeah. But that’s the trick, to always try to find the weakest link and then to improve that to see if it can raise the level of quality. Unfortunately, they’re becoming more and more expensive. The weakest links disappear, they will be more expensive weak links to– [crosstalk]
Martin: Well, my microscope, my compound microscope or biological microscope cost me under 400 bucks, which is probably on the high-end amateur, not professional level. My stereo microscope was actually about 600. That’s probably a better microscope in its class, but a good adapter is probably going to cost me around $1,000, which is about the sum of both of my microscopes. So, I’m a little bit cautious about that.
Håkan: We talked about that before, but you might be thinking about designing your own as well, buying lenses. That’s a possibility as far as you get the cost on it.
Martin: You make it sound easy in your email. I can’t really get my head around that just yet. Maybe we’ll have to take that offline and see.
Håkan: Yeah, definitely.
Martin: Okay. Your work is incredibly artistic. You’re probably making the most artistic microphotography or microphotographs that I’ve seen. What are your thoughts on this form of photography as an art? You’re not just documenting stuff, you’re making beautiful art with it. I’m sure you’ve developed some thoughts on that.
Håkan: Yes, but I do find these specimens really beautiful. I think it’s so fascinating to see that, even though you can’t see them and they’re so small, they have such fantastic shapes and forms and colors. You can see shapes where on a micro level, like in spirogyra, for example, it goes all the way through the universe, all through the galaxies and the same shapes reappear from micro to macro level. I think the beauty of nature is amazing. I never get tired of looking at these specimens and to see them and to see them under the microscope in real-time and see them moving, see how they are bubbling with life if you look inside them, and you can actually see the photosynthesis ongoing in there-,
Martin: It’s amazing.
Håkan: -crystals moving around. I think they are really beautiful. In general, nature photography, if you consider this to be nature photography, some people don’t do that, but I do.
Martin: I do, too.
Håkan: Even nature photographers are struggling with getting their work seen as art. Landscapes are doing a better job, nature photography, animals are having more of a struggle. Some people like these images, some people think they are creepy. They don’t want to see the eye of a fly.
Martin: My wife won’t look at my bug shots.
Martin: She doesn’t want to see them at all.
Håkan: I think they’re beautiful, bug shots.
Martin: I do too.
Håkan: They’re amazing design, these bugs. The eyes and the different parts, the eight eyes of the spider, and so on and so forth. It’s amazing.
Martin: I’m waiting now to find– we have the cicada, they’re like little matchbox cars with wings. There was one dead on my balcony a few days ago, but it was in the middle of some pigeon crap. So, I decided to give that one a miss. But I’m looking forward to getting– there’s usually one or two drops and die on the balcony a year. So, I’m going to get some of the eyes and see if I can get some good specimens.
I think we’re just about coming towards the end of my list of questions. Can people find you online? I know your website, and can you tell us where to go and I’ll put links into the show notes?
Martin: Excellent. I look forward to continuing to learn from you as I’ve watched these accounts. I’ve been a huge fan of your Instagram account. I will put all of the links to your stuff in the blog post and the blog post for this is going to be at https://mbp.ac/750. Is there anything else that you wanted to share with the audience before we wrap up, Håkan?
Håkan: I’m not sure. I think we’ve covered a lot of interesting topics. I’m looking forward to talking to you some time again.
Martin: Yeah, absolutely. I will be picking your brains on the camera adapter as well.
Håkan: Yeah, let’s work on that, let’s see if we can find a solution. I will try to build mine now for the medium format cameras, and then see we can adapt it to your setup as well.
Martin: I eventually found one that I think is pretty much what I want. It’s a very wide field, it goes into the seamount rather than the smaller tube. It gives you a really big wide image pretty much to the edges of the circle. I’m trying to get them to loan me one for a week to see if it will actually work.
Håkan: Oh, that’s good. The important part is that you have your objective. Normally, objectives have an image circle of maybe 20 to 25 millimeters. Cheaper ones maybe 18 or 20 and the more expensive objectives 26.5 even. And then, you need to magnify that a little bit to fill the sensor. Otherwise, you will get dark corners on your images. To find those lenses that will magnify just enough to fill the sensor, but not too much, because then you are losing so much working area, so you need to find the sweet spot depending on the camera you have and the size of the sensor you have.
Martin: Well, the one that I’m looking at, they say it gets you just inside the image circle. If it does, I’ll be happy.
Håkan: For a full format camera, then.
Martin: Yeah. But it’s $900.
Håkan: Yeah, that’s the issue. It cost more than your microscopes.
Martin: Exactly. Okay. Well, we’ll wrap it up then. Really, thank you so much, Håkan. It’s been an amazing hour. I’d love to catch up again at some point. And yeah, we’ll talk about the other things in email and see if I can pick your brains a little bit more. But thank you.
Håkan: Thanks for having me. Very nice to be here.
Martin: Not at all. It’s been my pleasure. Thank you very much.
Håkan: Thank you, bye.
OK, so that’s it for the interview. I hope you enjoyed that fascinating conversation. I’ll leave you with one last very cool photo that Håkan sent me. Amazing!
Following on from my previous post about the moral dilemma I’m struggling with regarding killing an insect for a photograph, we’re going to put that aside for this episode, and simply talk about how I went about the process. If you don’t agree with this and feel uncomfortable listening or reading, please turn this off or close your browser now. I won’t do a lot of this kind of post, but I do think that there will be at least a portion of the audience that will find this useful, so here goes.
I’ve been quietly preparing to photograph insects more over the last few weeks, based in part on the very informative videos from insect photographer guru Allan Walls, so a public shout-out and thank you to Allan for making that stuff available. You can actually see most of what I used in this photo, so let’s walk through this item by item, and I’ll fill in some gaps as we proceed. Firstly, you’ll see the jam jar that I caught the fly in, and then poured in 90% alcohol and 10% water which both kills and cleans the fly. Allan Walls also recommends using an ultrasonic jewelry cleaner to clean off stubborn dust that can form on insects, but so far I haven’t needed this. Maybe at some point, I’ll pick one up.
I left the fly in the alcohol overnight, both because it was late when I caught it, and to ensure that any parasites that might have been inside the fly were also killed by the alcohol. Then, before I photographed it, I placed it in a Petri dish with some distilled water in it, to rehydrate the fly. I then selected a #2 insect pin which is the second to thinnest pin in that selection of pins you see in the white envelope on the right. You can also see the pin next to the Petri dish on that sheet of lens cleaning paper. I used a pair of very fine tweezers that you also see here to open up the wings of the fly a little, and also pull its legs away from its body a little too, then as it dried naturally in the air, I placed a tiny blob of superglue onto the insect pin and stuck the pin to the underside of the fly’s abdomen.
The fly was probably only 3 to 4mm in length, so I used the magnifying glass that you see on the left with the built-in USB-powered LED lights to see what I was doing. The paper is lens cleaning paper that I use to clean my microscope slides, and, of course, lenses, but reuse them like this when dried to guard against spilling alcohol, etc. on my work surface. Once they actually get dirty I throw them away. After preparing the fly, I use a soldering stand to hold the insect pin and position the fly under my Stereo microscope to start photographing it, as you see here.
I didn’t photograph every step of the shoot, because keeping the fly out of the alcohol or water gradually allows it to dry out, and the eyes start to collapse in, which doesn’t look good. For some of the images, I used a piece of black velvet below the fly to give it a black background. The microscope came with a black plate that replaces the semi-transparent circular plate in the above photo, but it’s so reflective that it appears almost white when illuminated from above, so I find that the black velvet soaks up the light much better. I am also using an LED ring light that attaches to the bottom of the microscope lens, providing really nice adjustable light for the subject.
Here is one of the first shots of the Housefly using my Stereo microscope and the 2X Barlow lens. My Stereo microscope has a zoom mechanism that provides 7-45X magnification with the 10X eyepieces, and the Barlow lens doubles that to a 14-90X magnification range. I haven’t yet calibrated my Stereo microscope to find the magnification of my camera adapter with this scope, but it’s probably around 2X, so we’re actually looking at around 24-180X magnification in the photo with the 2X Barlow lens in place.
I learned an important lesson with this first photograph though, and that is that you have to be very careful with the amount you adjust the focus when shooting images of insect eyes to focus stack. You can probably notice the four slightly darker rings around the top half of the eye that is caused by too large a gap between my focus stack images. To prevent this I turned on the “Focused” checkbox in Helicon Remote, which shows the area of the specimen that is in focus in blue, as you can see in this photograph of my computer screen as I worked.
Once I’d identified the problem and the cause, I started to place my thumb on the back edge of the focused area on my screen and then adjusted focus for the next image with my finger marking the previous edge, and I only moved the focus halfway back from there, so it was half overlapping the previous image. When you shoot the eyes like this, you don’t get the nasty focus error rings that you see in the earlier image of the Housefly.
At times like this, it helps to have an extra hand to trip the shutter, so I used the built-in Mac OS Voice Control feature, found under System Preferences Accessibility options. There were also times when I was photographing the fly with my compound microscope when I was holding a flashlight to illuminate the subject, and I also have to adjust the focus of the microscope between each frame for the stack, so I set up a voice command to enter the keyboard shortcut to take a photo in Helicon Focus every time I said “new photo”. I also configured a command to create a new stack every time I said, you guessed it, the words “new stack”.
There were also times when although I was not holding a flashlight, or marking the focus zone with my thumb, but it just kept things more stable by not physically touching my keyboard. It helps reduce vibration on my work surface, so the voice control came in very handy.
There is one limiting factor working with a Stereo-microscope for photography, in that, because they are designed to provide a stereoscopic view of the specimen, there is a parallax shift as you adjust focus when redirecting the image from a single lens to the camera port. This means that as I adjust the focus the subject gradually drifts across the screen, and will eventually move so far that there is nothing left to stack, so I am still limited to a certain depth of field. In this photograph of a scarab beetle, which I luckily found dead outside of my apartment recently, I initially tried to stack enough images to get the entire beetle in focus, but the parallax shift caused the beetle to move so far over to the right that I ended up with only half a beetle after stacking. To avoid that, I stopped my stack around halfway down the beetle, and I actually like the out-of-focus abdomen anyway, so all was well. This still took 40 frames to create this image, with around half of the scarab beetle in focus.
Anyway, having figured out how to photograph the eyes on the housefly without the nasty focus rings, I also shot a few stacks that resulted in some images that I am relatively happy with, such as this one, off slightly to the left of the face. I darkened down the body in this and a few of the images, as there are areas that don’t look great, but also there were a few stacking issues that left a few too many remnants, but I actually am mostly interested in the face of the Housefly.
To get this look relatively naturally I used an adjustment brush in Capture One Pro and brushed out the background in a number of different layers, probably up to five or six per image, and allowed some of the layers to overlap with the back of the eyes a little so that it looked like the fly was poking into a spotlight rather than being more fully illuminated from the start. I reduced the Exposure and sometimes used a tone curve adjustment on the layers to gradually darken the background.
To the right, above, I’ve also included a shot where I didn’t totally darken down the body, and again, without the focus rings in the eyes, so I’m pretty happy with this. Both of these images were shot at around 60X magnification through the eyepieces of the microscope. It’s a little bit scary to first look into the eyepieces at this magnification, but seeing all of the detail in this creature is incredibly fascinating.
Here are two final images to finish with. The first was shot with my Compound microscope, illuminating the Housefly with an LED flashlight, at 100X magnification. The second and final image is from the Stereo microscope again, and I had started to run out of time as the eyes were gradually losing their luster and starting to concave very slightly in a few areas.
As I mentioned in my previous episode about the moral aspect of doing this, this fly was pretty much doomed the moment it came into our apartment, but I don’t feel 100% happy with killing it for these photographs. I’m struggling with the decision a little bit, although I do find it fascinating. For larger insects, I’m pretty much going to rely on finding them dead after they’ve lived their lives, and see how much I can make of that. As I mentioned earlier, I’m not going to do a lot of this kind of episode, so if you don’t like it, stay tuned anyway, as we’ll be back to my regular work more often than not as we move forward.
Today we’re going to take some time to think about the moral dilemma of killing insects for photographs. This can be a very polarizing subject, and I know that many people have strong feelings against killing insects for photographs. I personally hadn’t killed an insect for a photograph until two days ago, and this came after a conversation with a good friend of mine about this subject that kind of help me to understand my own position on this complicated subject. I’m still very much undecided about my stance but figured this was as good a time as any to talk about this, and I’m also interested to hear your thoughts as well.
As this can be controversial if you can feel a flame war coming on, I ask of you three things. The first is to hear me out. Listen or read this entire post before commenting. The second is if at the end of the post you feel like attacking me, walk away from the computer for a few hours and then come back and write your comment after you’ve cooled down a little. And even then, don’t attack me. I’m sharing my views in good faith, and you are in my house now. If you attack me in a disrespectful way, I reserve the right to delete your post. I’m fine with you sharing alternative views, but do it with respect, or you’ll be wasting your time. The third thing that I ask, is if you are a visitor, please give your real name when commenting. If you comment anonymously, the chances are I’ll just delete your comment.
So, here’s how I feel about killing insects for photos. I’ve been photographing insects for many years. Not often, but I’ve been doing it for probably around 20 years now. I have mostly photographed live insects in the wild, and occasionally tried to photograph live insects at high magnification, and generally failed. They move around too much, making focus stacking pretty much impossible, so the results are generally not what I want.
The Hypocricy Aspect
For many years I’ve felt that I would not kill an insect just for a photo. There was and still is a bit of a moral dilemma behind this decision, but in addition to that, I actually didn’t think I could physically kill an insect, but hypocrisy kicks in here when I say that in some ways it’s the size of the insect that concerns me. If I think about it, I will slap a mosquito on my arm, which generally results in its demise. We have these annoying little tiny flies that appear from nowhere in the summer and walk across my laptop screen. These also generally find themselves stuck to my thumb, then scraped onto a piece of used paper or tissue in our waste paper basket. So, I won’t kill an insect for photography, my love and passion, and profession, but I will kill one for trespassing in my apartment. That is completely hypocritical.
Other People Kill My Meat
Another huge chunk of hypocrisy that surfaced as I wrote my thoughts down for a friend is that in the regular state of the world, as in, we are not being attacked by aliens or zombies and I’m not in survival mode, I generally rely on other people to kill my meat. I don’t eat meat or fish every day, but when I do, the weight of killing and preparing that meat falls on someone else’s shoulders. I can and have plucked chickens and skinned rabbits, and I am fine with gutting a fish. Come to think of it, I’ve killed a few fish too when I was a boy, caught with a fishing rod from a holiday town pier, and eaten that morning for breakfast. The rabbits and chickens though were killed by someone else.
Now, going back to the aliens and zombies scenario, if we were in a life or death situation, and my survival depended on killing an animal, I probably could, but I would not find it easy. My friend and I agreed that killing for sport is not something that we agree with. But if the animal is going to be eaten, without waste, then it’s more acceptable, in my opinion. I realize that there may be vegetarians or vegans listening with other views, and I apologize if any of this offends you, but I need to get this out as we work our way back to the topic of killing insects for photographs, which I’ll try to do now.
So, I realized that I’m completely hypocritical when it comes to my stance of not killing insects for photos, yet I will squish a mozzie or fly for no greater crime than sucking my blood or walking across my computer screen. I generally don’t kill spiders in the house, though larger spiders that make me or my wife uncomfortable generally get ushered outside. It’s actually rare for us to get common houseflies in our apartment because we have mosquito screens on all of our windows and we don’t leave the doors open unattended. But a few nights ago, there was a fly sitting on our bedroom floor when I went in to turn the air-conditioning on. It was probably on the floor to keep cool, but it flew up and over onto our curtain as I walked closer.
Housefly in a Jar
Having already thought a lot about this recently, I grabbed a jar that I just happened to have put aside and caught the fly. Before we talk about the fate of this particular fly, I should tell you that no housefly has ever gotten out of my apartment in the past unless they fly out before being noticed. As a general rule, if a fly comes into our apartment, it will either be swatted, or sprayed, but its exit mechanism is generally via the trash. So here again, I found myself in a hypocritical dilemma, but, I figured if this animal was going to die, is it really wrong for me to benefit from its demise? Am I any more guilty for killing it and taking its photo than I would be for killing it and sticking it in the trash can?
As I mentioned earlier I’m not really OK with hunting for sport or fun, but if the animal will be eaten without waste, then I have no problems with that. By photographing the fly, surely I’m making more use of it than I do by simply disposing of it. This was the thought process over a relatively long time recently, that resulted in the fly from my bedroom sitting in a jam jar in my studio. Initially, I thought I’d just leave it in the jar for the night, but then I started thinking that it would stress the fly. I know this might also sound hypocritical as I’d decided to kill it, but I did not want to cause it any unnecessary stress.
This, by the way, is why I don’t really agree with putting insects in the fridge to slow them down enough to photograph them, or using chemicals to put them to sleep, for example. If we’re going to cause them physical discomfort, we’ve already crossed the line. I did cross the line with the fly. I said a few words of apology with my hands together for what its worth, then I opened the jar and poured in around one centimeter of pure alcohol mixed with 10% water. The fly died instantly, arguably with less trauma than having its backside smashed through its brain in a deftly swatting action.
So, I’d done it. I’d killed a fly and I was going to photograph it the next day. I didn’t feel 100% OK with this. I woke up early the next morning with a pang of guilt, but I went on to photograph the fly over a few days, and I can live with my decision, but this doesn’t necessarily mean that I am now going to become a mass murderer. At this point, my thought process is unchanged in that either I or my wife would have killed that fly anyway. I still, at this point, do not think I could kill anything much larger. I know that size should not be important, but for me, it is. My friend, who I will not name just now because I didn’t get permission to do so before he left for a short holiday with friends, talked about bacteria dying as part of his job. I’m sure that most people, although I’m not going to assume all people, are happy to let that one slip. I mean, you can buy yogurt that comes with bacteria in it, and even when we wash our hands were killing bacteria. We’re actually also killing insects as we wash away little fleas that people have on our skin, but we won’t go there right now.
Sometimes It’s Unavoidable
So, for me, at this point, it seems that size plays a part in my ability to kill an organism. I’m not too worried about killing tiny insects, but I feel a lot of resistance towards killing anything much larger. The point with the previous paragraph is that if we are going to say that all life should be respected, then really, how far can you take that? Just washing your face or leaning on the arm of a chair carries a good chance that you’re killing something. It’s just so small that you don’t know you are killing it, but does that make it OK? I’d say that it has to because it’s unavoidable.
One other thing that I’d like to mention is that I recently photographed a scarab beetle that I found dead at the bottom of my apartment stairs. I am fine with that too. I’m relatively OK working with insects. I am just not comfortable killing anything much larger than a fly, so making use of an insect that has already lived its life is in many ways the ideal solution for me. I don’t pick up anything that’s decaying, but if something is upside down in the middle of the pavement the chances are it’s just died, so I’ll pick that up and photograph it, generally after giving it an alcohol bath to kill anything that might be stuck to the insect or embedded inside it. I’m not sure I could handle a parasite bursting out of an insect while I look at it through a microscope at 50 to 100X lifesize.
Could I Kill an Ant?
I mentioned to my friend recently that I think I could probably just about bring myself to kill an ant for a photograph too, and this is what got us talking about the size thing. The reason that I have not yet done that though, is because I would have to go looking for one. Although I’ve killed ants that got into our kitchen back in England, we don’t get ants in our Tokyo apartment, so I would have to enter their realm in order to get me an ant, and although I think I will do it at some point, I’m still struggling with that to a degree.
I feel as though the hypocrite is still there. Why does size matter when it comes to my ability to take the life of an insect? Come to think of it, when I see the number of insects stuck to the front of my car after a summer drive, even killing larger insects is not necessarily something I’ve never done, but there is the problem of intention. I don’t drive my car with the intention of killing insects, and for numerous reasons, I wish I could avoid it.
Maybe I’m actually just a big wuss, and I’m too scared to kill anything with any more presence than a housefly, and I’m just wrapping that up as a moral dilemma? I’d actually say that this is around 20% true, but the other 80% of me really just would feel too guilty. For the time being, I’ll be patient and continue to wait until I find dead specimens of larger insects. There was actually a cicada dead on my balcony a few days ago too, but it had been mingling with pigeon crap, which is nasty stuff, so I decided to give that a miss.
That’s Where I Stand
OK, so that’s where I stand on this. It’s not necessarily a bold stand, and I’m not pretending to be either a saint or necessarily consider myself a sinner. I should also mention that I don’t necessarily condemn anyone that takes a different stance. If you have no problems killing insects for photographs, I’m actually probably a little bit envious if anything. There’s that 20% wuss coming out again. And, if you strongly believe that we should not kill insects for photographs, I do get that too, but having just euthanized a housefly, I’d be hypocritical to pretend that I’m completely on your side of the fence.
Why Do You Stand?
I would like to hear your views, hopefully from both sides of the fence, or even if you are like me, sitting mostly on the fence. Once again, please don’t blindly attack me. Even if you have strong views on this, relay them calmly in the comments below, and give your name. I don’t like talking to anonymous people. You know my name, please use yours. And if you want to say something that can’t be associated with your name, or if leaving your name annoys you, then just close the browser and walk away, or you’ll be wasting your time, as I will delete your comment.
In the next episode, I’ll talk about some of the methods I used to prepare the housefly to photograph it and share some of the photographs. We’re talking housefly at between 50 and 100X magnification, so if you think that might disturb you, you might want to skip next week. Personally, I think the shots are beautiful, but then I might be a bit weird in that respect. I’m sure you’ll decide for yourself.
It’s been a crazy week as I tried to pull in a little bit of microphotography while working on a second update to my new MBP Fine Art Border Tools plugin for Photoshop. I’m almost ready to submit a new version that includes text-based watermarks in addition to the graphical watermarks already available. I have a few more creases to iron out, but it’s shaping up OK and I hope to get it into the system during this week.
There has also though been a nice little addition to my Microphotography world too, that has led me to a number of conclusions that I wanted to discuss, and also get onto the main topic which I’m really still learning myself, which is measuring minutia with the aid of a microscope Calibration Slide or Stage Micrometer, and as you’ll see, I’ve created some overlays that I can apply in Capture One Pro that enable me to show the size of specimens or the field of view in my photos when necessary. This is more for scientific presentation than artistic, but it’s nice to be able to do this, so I figured I’d share what I’m up to.
So, the addition I just mentioned is a set of Plan Achromatic lenses for my compound microscope. As I mentioned when I bought this microscope, it was positioned as a somewhat high-end but still very much amateur microscope. At just under $400 compared to the thousands or tens of thousands of dollars for real high-end gear, you can probably appreciate that my microscope is good for hobbyists and serious home users, but not great compared to what the big boys and girls would buy. Because of this, the more I worked with it, the more I am learning about what I need to do to really start getting better image quality.
Something that had been frustrating me a little was that I was having to do really long image stacks to get everything in focus, and as I continued to study this field, I found out that my achromatic objective lenses that came with my scope have a tendency to get soft or blurry as you move away from the center. I could overcome this to a degree by stacking more, but the edge really never got totally sharp, in some images, so I looked around to see if there was anything available to cure this.
I found that there is a type of lens called a Plan Achromatic lens, which creates a flat image, so that when viewing through the eyepieces and supposedly for my camera too, the image would be flat to the edge. After hankering for a few weeks, I decided to take the plunge and buy a set of lenses when the company I bought them from, Amscope, in the US, had a Fourth of July sale. I was able to get a 4X, 10X, 40X, and a 60X objective lens all for less than $200, excluding postage to Japan. I decided to go for a maximum of 60X objective and replace my 100X Achromatic objective lens with that because the thought of using oil to see with the 100X lens scares me, plus, most of what I’m looking at still doesn’t require a cover-slip, and applying oil directly to the crystals I’m photographing would make a bit of a mess.
After a mysterious four-day stop-over in Calgary, my new Plan Achromatic lenses arrived a week after I ordered them, and I was pretty amazed by the difference. Through the eyepieces, the difference is huge. They seem brighter, but the bigger benefit is that they are sharp as tacks from edge to edge, so I was very happy to have upgraded. However, when photographing with the new lenses, although the flatter image made it possible to create sharper photographs with smaller image stacks, I noticed that the right side of the frame was still a little bit soft, but only on my camera. As a test, I rotated the camera in the camera port and the soft area moved with it, meaning that the flaw that remains is in my camera adapter, but then I always knew that this was the weak link. At some point, I’ll pick up a better adapter, but for now, I can overcome that deficiency with a few extra images in my stacks.
I needed to rephotograph my Calibration Slide with these new lenses, so I’m going to continue now and talk about how I’m using that. First of all, this is a photo of the Calibration Slide, shot with a regular macro lens for illustration purposes. On the left side, there is a 1cm wide micrometer numbered in 0.1mm divisions, and to the right of that, inside that bold circle, there is a cross-shaped micrometer 1mm wide and 1mm high, with larger divisions which are 0.1mm apart, and between them are 0.01 mm markers, which we can convert to 10 microns per division. On the right, there is a dot with a diameter of 0.15mm and finally a smaller 0.07mm diameter dot.
So, to put this into perspective, let’s take a look at how I’m using the two micrometers on the left. I haven’t really used the dots much, so I won’t talk about them. I should also note that to properly calibrate my microscope for scientific purposes, I should use an eyepiece that contains what’s known as an ocular micrometer, but I don’t have one of them, and I’ll explain my process, and why it’s not a big deal for me. First of all, here are four images, shot with my new Plan Achromatic lenses. There are a few things to note before we move on. Firstly, I’ve cleaned up each image, and inverted the color, so that they are white on black. This makes it easier to overlay them on an image to measure the subject, as I’ll show shortly. Also, I’ve marked these images with the magnification of the Objective lens, which doesn’t take into account the magnification of the ocular lens or camera adapter. I generally use my 10X magnification eyepieces, although I also have some 25X magnification eyepieces. Also note that the micrometer that you see in the first image shot with the 4X objective lens shows 4.3 mm of the 10 unit 1cm wide micrometer, but the other three images show the center of the 1mm wide micrometer, which we can see in full with the 10X objective, but only the center of it with the 40X and 60X objective lenses.
In a scientific situation, I would compare the units on an ocular micrometer with the units on the stage micrometer to calculate the size of a specimen that I’m viewing, so that I could make notes about the specimen for future research. But for me, I want to know the size of the specimens in my photographs, as it’s the photograph that is the main purpose of my use of the microscope. Measuring anything requires that we have a reference that won’t change, and that is why I prepared these images. Consider that the images were shot with a camera that is mounted via an adapter to the triocular port directly above the slide. I have two spacers on my adapter that move the camera far enough away that I am photographing the inside of the image circle, generally allowing me to see small black triangles in the four corners. This means that I am actually photographing at a higher magnification than my objective lenses.
As long as I don’t crop my image though, if I overlay the photograph for the objective lens in Capture One or any image editing software that enables me to overlay an image, and the two images are the same size, anything that I can see in the photo can be directly measured with the respective micrometer. Here, for example, is an image of some polarized Sodium sulfite crystals shot with the 10X objective lens with its respective micrometer overlay at 33% opacity. We know that this entire scale from side to side represents just one 1mm. The tall increments are 0.1mm apart, with the smallest divisions being 0.01mm or 10 μm (microns). Based on this, we can measure the width of the crystals, with the larger ones being around 50 microns and the smaller around 30 microns wide.
Here now is another image shot with the 40X objective lens, using the center of the same stage micrometer. The first thing we can learn is that we are looking at 440 μm or 0.44 mm of the world. Also, we know that the camera sensor is 36 mm wide, which is 36000 μm, so if we divide 36000 by 440, we get 81.8 which tells us that the magnification due to my camera adapter’s relay lens and spacers is actually a hair under 82X, not the 40X of the objective lens. This means that my camera adapter is magnifying the image 2.0455 times. To check, we can do the reverse calculation to anticipate the size I’m seeing with my 60X objective lens, by multiplying 60X by 2.0455 to get 122.73. This should be close to the magnification we can calculate using the same formula that we used for the earlier images.
Here is a photograph with my new 60X objective lens, which, with the stage micrometer we can see measures 290 μm wide. If we divide 36000 (the width of the sensor in microns) by 290 we get 124, so my 60X objective is providing 124X magnification with the adapter spacers, which is slightly larger than my check calculation, but I’m estimating the find 10 microns as being 5 microns for each of those two gaps, and that may be slightly off. There may also be some variance in the marked magnification of the objective lens, so I am now confident that my camera adapter is providing approximately 2.05X magnification for the image size on the sensor.
The last calculation that I had to sleep on before understanding is how come I’m seeing an image that is approximately 400X with my 40X magnification objective, multiplied by the 10X of my ocular lens or eyepiece. I’m seeing an image that is roughly double area that I’m seeing in my camera image, but one is 82x magnification and the other is 400x magnification. I tried to figure this out yesterday and did a bit of searching online, but I couldn’t find a definitive answer, so I went to bed feeling relatively stupid.
My web searches told me that the calculations I’ve explained so far are correct, which is great, but I also learned that you can use the same calculation to calculate the magnification of a printed image. It still didn’t click until I got up this morning, and I reverse calculated a 400X magnification by multiplying 400 by 440, the width of my 40x objective field of view in microns, and that gives us 176000. This means if I was to print an image shot with the 40x objective and my 2.05 magnification camera adapter at 17.6 cm we’d be looking at a 400x magnification image. The magnification on the sensor is accurate, but we are not going to view the images at 36mm wide. There is always going to be some magnification as we project the digital image onto a larger viewing media, be it the computer screen, or a print. I’ve tried to layout most of what we just covered in the below graphic.
Let’s do one last calculation to extend this, just for fun. So let’s say I print one of these images at 18 x 24 inches, one of my favorite print sizes. We know that 24 inches is 609.6 mm. To avoid using really big numbers, we can convert 440 microns to millimeters, which is 0.44 mm, so if we divide these two numbers we get 1385, which we now know to be the magnification of my digital image when printed at 24 inches wide. Of course, we then have the magnification due to viewing distance, but let’s stop there. I’m sure you get the picture.
One last thing that I wanted to share with you is that although I left the 50-micron squares in the center of the stage micrometer photos that I shared earlier if I need to measure something with the smaller 10-micron divisions, I can actually move the overlay around in Capture One Pro for a more accurate measurement. As you can see in the below image, the hexagonal crystal in my 40X objective photo is approximately 93-microns wide.
OK, so we’ll start to wrap it up there. If you are looking for an upgrade for a microscope with Achromatic lenses, I can definitely recommend picking up some Plan Achromatic lenses. The total cost of my microscope including this upgrade is now running at just under $600 and there are some models available that come with Plan Achromatic lenses at around this price, but many are much more expensive, so I’m relatively happy with the path I’ve taken still. And, I can probably sell my original four lenses for around $100 if I’m patient, so that will help me to recoup some of my outlay. I think I’d prefer to keep the 100X oil objective though, just in case I need that later, as I don’t think I’ll use it often enough to buy a Plan Achromatic objective to replace it. Before that, I’ll probably get a 20X Plan Achromatic objective as the jump from 10X to 40X is quite large and I often find myself wanting something in between.
My next job is to figure out how to accurately measure objects on my stereo microscope, which will be infinitely more complicated as it has a zoom mechanism, so that should be fun. Note that I’ve added a Microphotography link to the Posts menu above that automatically pulls together everything I’m posting related to microphotography, so check that out if this subject is of any interest to you.