I have a handheld digital microscope that I use to take pictures and videos of invertebrates. For slow moving or relaxed subjects its quite easy, for miniscule, fasting moving, panicking creatures it can take ages to get them into view and focus. I most often put them in an empty petri dish and chase them around with the scope. Is there any better way of doing this? I tried using a micro zip top bag today to put a beetle in and it made things more doable haha
A couple suggestions for using petri dishes:
- use mini-size dishes (35 mm diameter)
- pad the dishes with medical gauze or paper towels stacked so that your specimen is pressed lightly against the lid to limit mobility (disadvantage: you have to turn the specimen over to photo underside)
Chilling (e.g. with a picnic ice pack or in your refrigerator) can be a way to make subjects more cooperative. I have never tried this with really tiny specimens (<2mm) but I have found it helpful for many small hymenopterans. Different species respond differently to cold, so it may not work in all cases.
I find that while putting subjects in a container is helpful for preventing them from flying away, the container often makes it difficult to get good photos that show tiny details, even if said container is made of thin, highly transparent, non-reflective material.
Just to clarify, these mini handheld ‘microscopes’ are not working in what most people think is the microscopic magnification range. Despite the claim (no matter the brand) that they achieve magnification of 1000X, it’s really more lIke what you get with say a 40X microscope objective.
That said, they can be a very useful and very affordable iNat tool with practice and an understanding of their limitations. And a fair bit of patience!
Another important thing to consider is that many of these cheap tools will claim 4K image output, but really, they all use 2MP sensors and what you get as a 4K image is simply that output digitally resampled up to 4K range. More pixels, but not more detail.
But even the lower true HD output can supply postable results. Also, most offer video or stills.
The difference in image quality between a still and a frame from a video is zero, so I almost always shoot video (set to the sensor truth of HD resolution) and go back in later to find decent frames. Is far easier and more productive than hunting for in focus stills. Video output is very choppy because the frame rate is very slow. Typically 12 fps.
And I use these scopes mostly for slow moving or deceased (found) subjects, and almost always without the stand. Also, get a small led white video panel to prove more illumination (from the side more) to improve noise reduction.
As for water stuff, a lower cost alternative to petris, and a smaller work stage? Clear plastic coin collector cases.
As for calming down aquatics, I’ve heard others use the fridge trick, but I’ve also heard it can be tricky and even fatal for some creatures. I haven’t done anything like this, but I’m definitely interested in trying too.
Thanks for bringing this topic up! Would love to see what you come up with.
Some previous suggestions for microcrustaceans:
https://forum.inaturalist.org/t/is-there-a-way-to-slow-down-microcrustacea/22693
Since the responses seem to be making different assumptions about what kind of small invertebrates you are trying to view, perhaps it is worth asking: Are you primarily interested in aquatic or non-aquatic specimens?
The options available to you will also depend on whether you need a set-up that is suitable for the field or whether you will be viewing the specimens at home.
My suggestion of refrigeration was based on experience with small flighty insects; I doubt it is quite as effective for aquatic invertebrates who are likely adapted to living in fairly cold water. (Chilling is also not a practical solution if you will be out in the field on a hot day, as an ice pack is unlikely to remain suitably cool for hours on end no matter what kind of insulation you have available.)
I have found that for aquatic organisms the first thing to do is remove most of the water (e.g., with an eyedropper) so that the space in which it can zoom around is as small as possible.
I do lot’s of super-macro ant photography, and so far the most practical solution in the field has been baiting. Different species like different baits, and for some it won’t be practical, but for many ants I just put a drop of sugar water in front of them and that usually gets them to hold still long enough to properly focus.
I find obs of my beetle group of interest and taxonomic expertise, in which folks have the specimen in a vial or a rush of liquid, in a ziplock bag, and so on. I do appreciate that folks don’t want to kill the insect, but let’s be realistic for a moment? The structural details used by taxonomists to identify insects are very miniscule and any distortion makes identification all but impossible. Maybe to genus but that is hardly very informative scientifically. To be realistic, insects generally need to be collected and photographed under a microscope. Personally, I do the whole insect curation thing, and look for specialists who can do the IDs. Odonata and Lepidoptera I do not collect, though micro moths should be collected and spread for ID.
The link that Chris shared contains a fairly thorough treatment. The article includes:
- Mechanical Methods of Slowing and Restraining Microscopic Animals
- Thermal Methods of Slowing Microscopic Animals
- Narcotizing Microscopic Animals
This doesn’t apply universally since there are plenty of insects that are very easy to identify. And “id to species” is also not the only reason people make observations. Than there is also the issue that killing/presverving can itslef cause plenty of distortion. Ants curl up when dead and it’s actually quite hard to uncurl them without damaging important features.
Yeah, there are a lot of insects that can be identified without microscopy.
I agree with the point that insects photographed through the walls of a container are often difficult to identify, because in most cases the containers are not translucent enough for clear photos or the photographer has not successfully gotten their camera/phone to focus on the insect instead of the container. So I don’t recommend this as a method for iNat observations unless the observer has practice taking photos under such conditions.
But for insects that are identifiable from photos but too active to photograph easily, sometimes putting one in a container is the easiest and most effective way to get it to stay still long enough to take photos that aren’t blurry, or to get photos of macroscopic features that are easily recognizable but not generally visible on a moving subject.
There are plenty of equally cheap digital microscope availables with 5MP sensors, so it obviously pays to shop around.
It should be possible to achieve at least 15 fps with most cheap devices. This may be partly determined by the software, rather than just the hardware. I use mpv, as it has an enormous number of options available for getting the most out of the input.
I find this quite surprising. For me, a good quality stand is absolutely crucial. I usually let live specimens run around on a short sprig of plant material whilst holding my hand tightly against the microscope/stand. Eventually, the specimen will pause for a few seconds, when I then try to get the desired shots. This invariably requires that everything is held still at the crucial moment - the smallest movement will usually result in blurring, given the short depth of view.
Almost all my specimens are terrestrial arthropods. For immobilising, I most often use zip-top bags. Occasionally, I also use a lump of sticky-tack to keep the subject fixed in place for high-magnification shots - but obviously this is only really suitable for fairly robust specimens with hard exoskeletons, such as beetles. It can often be very effective, though, as it’s possible to move the specimen right up to the microscope lens (like a miniature steadicam rig)
In my experience, it’s rarely necessary. Patience will win out in most cases. The only times I would consider resorting to killing is when very fine details like surface micro-sculpure are needed. But really, I’d much rather keep every specimen alive and release it as soon as possible, even if that means I lose the chance to record a species I haven’t seen before. Discovering that I can’t identify something with the resources currently at my disposal is still knowledge gained.
I would find it very hard to justify all the effort of collecting, recording and returning specimens, if I didn’t learn how to identify everything myself. I want to read the whole book, not just be told the ending…
If you want to view them from the side instead of the top, try a small acrylic box for fish. You couldn’t use this under a microscope, but for larger creatures, you could get photos from the side.
Also for water viewing tanks? Spectrometer sample cells. Online prices are fairly cheap and the glass is flat and optically excellent.