The most basic form of microscope is the hand lens. I use a ×10 lens; they are also commonly available in ×5 and ×20. I have found if I am careful I can actually get somewhat usable photographs by holding my hand lens between my phone camera and the specimen, and playing around with angles and distances. This requires holding my phone in one hand, and both the lens and the specimen in the other — a bit difficult, but it allows a lot of control over angle and lighting. Alternatively, if I can still get a decent angle, I set the specimen on my knee and just hold the lens. These are the techniques I use in the field and it can be enough to photograph the conidiophores of Peronospora.
For more detail and magnification you will want to use a proper microscope setup. However, you may not always want to mount a specimen on a slide. You can often get decent images by simply placing plant tissue under the microscope and having a look. This would be easiest with a stereo microscope, but I have found it works okay with my optical transmission microscope (i.e. a microscope designed for slides). This technique is particularly useful when you cannot see what part of the leaf to mount on a slide, when the pathogen structures are at a low density across a wide area. Obviously it works best when these structures are relatively large.
This is when you take several images focused at different depths and combine them using software to create a new image that has a larger depth of field. This can be particularly useful for microscopy without slides as the depth of field can be very poor if you are using a microscope designed for slides, where everything is flattened by the coverslip. It can also be useful with slides. Focus-stacked images are not primary data and you should always clearly indicate that you have used this method, especially in scientific publications. There is a useful blogpost on focus stacking here.
There are three possible ways to look at pathogens with spores on the outside of the leaf:
Each of these methods has pros and cons, and the best technique generally depends on what taxonomic group you are looking at. The first does not damage the specimen in any way, so I often use this first, especially if I am not sure what I am looking at or if I don't know where the spores are densest on the leaf.
This can be useful for finding good patches of conidiophores, particularly for Ramularia which can be difficult to see under a hand lens. I then know where to target with the sticky tape method described below. I put a clean glass slide on the microscope stage and place the leaf on top. I use a gooseneck lamp (the kind normally used with a dissecting microscope) to shine light directly onto the surface of the leaf. I can generally get good images at ×40 and ×100 with this technique.
For a few species, this is almost the only way to view the spores. Ramularia that infect grasses, such as R. holci-lanati, produce very few conidiophores. Using other techniques will often lead to these being lost and not visible on the slide, but this tecnique does not damage the specimen and allows the spores to be seen and the identification confirmed.
This is the way I normally make slides for rusts and downy mildews. I hold the infected part of the leaf over a glass slide and gently scrape the spores onto the slide with a razor blade. It can be difficult to avoid cutting into the leaf itself so this takes some practice. I then drop a small amount of water onto the spores on the slide and add the cover slip on top. With rusts I often add the water first and wet the blade slightly, which allows the spores to stick to it.
Often multiple different stages of rusts will be growing on the same leaf, visible as slightly different colours. It can be useful to isolate these from each other and look at each in detail. Chris Preston suggests using very fine tweezers to pick spores from a particular part of the leaf, and drop them onto a drop of water on a glass slide. I have also used tweezers to pick individual downy mildew conidiophores from a leaf, which can be useful when they are very few, although it can damage them quite severely.
Downy mildew conidia can become very tangled with this method. Björn Sothmann uses a pair of fine forceps instead to remove just a few conidiophores and places them in a drop of water on the slide. He then arranges them neatly using a needle while looking through the eyepiece lenses under the lowest magnification. Once they are arranged he puts a coverslip on top. I have tried this technique but it is rather difficult and takes a lot of practice. Björn has used it to produce much nicer micrographs of this group than I can, for example here.
Groups like white moulds, powdery mildews, and Cercospora have rather delicate structures, the detail of which is lost if they are scraped off with a blade. For example, whether spores are produced singly or in chains is an important character in Ramularia. Instead, I press a piece of sticky tape onto the infected area of leaf. I then place the tape sticky-side-down onto a drop of water on a slide. More water is needed for this than for other techniques. I then place this under the microscope and look normally. The glue on the tape produces a slightly granular texture across the image, and tends to go cloudy over time once it is wet, though the speed of this varies between makes. The kind I use is “Sellotape Super Clear”.
The leaf spot fungi are particularly difficult to view under the microscope. Most of them produce spores tightly packed into small pycnidia embedded in the leaf tissue (left-hand figure). Pycnidia should be visible under the hand lens as small black or brown dots, but are sometimes near-invisible.
It is best to cut leaf spots with a razor blade or scalpel while they are still dry: this prevents the cut bits sticking together and allows you to make extremely thin sections. Hopefully some of these will slice through pycnidia. You can then add water and place a coverslip on top. The spores of Septoria and most other leaf spots are small, very thin, and colourless (right-hand figure), so you should be methodical and patient when looking for them.
Sometimes despite your best efforts you will find all of the pycnidia on the slide are intact, or the conidia are trapped inside the ones that have been sectioned. To fix this, place the slide on a flat surface and using the blunt end of a pencil or something similar, deliver a sharp but gentle tap to the coverslip. Be careful not to overdo this and break the coverslip! This method can break open pycnidia and force the spores out.
Often spores will fall out of a pycnidium to the bottom of the slide, so focusing the microscope on the bottom can make it easier to find them. Sometimes they will be more concentrated around an open pycnidium and following the trail can be a way to find pycnidia.
It can be difficult to see the septa in the conidia of some species of fungi. This is an important feature for identification so this can be an issue. Outer Hebrides Fungi has an example of conidia of Septoria arundinacea in Lugol's solution here, showing how it can be used to make the septa very clear and easy to count.
Entyloma produces its teliospores throughout the affected leaf tissue. I have found the best way to look at these is to again chop a small piece of tissue up, but then to place the cover slip on top and “smear” it by pressing firmly and evenly across the coverslip and moving it gently up and down with respect to the slide.
SEM is different to the other types of microscopy on this page in that it does not use light. Instead, a beam of electrons is fired at the sample and the resulting signals are used to understand the shape of the sample and some aspects of its composition. In order for this to work the sample is first coated with a very thin layer of a metal, often gold. Obviously, this is relatively expensive: at my university, the cost to the department for a single run of four samples is at least £20. The scope I have used in my department is a JSM-IT200, which I can say is a joy to use and very beginner-friendly. It is very hard to find prices for these things online but I think an equivalent machine new would cost somewhere in the mid-tens of thousands of pounds and up, rather outside of the price range of any individual who hasn't won the lottery. They are really machines meant for large research organisations like universities. Furthermore, without a library of SEM images to compare to, it can be difficult to interpret them when all you can compare them with is light micrographs and written descriptions based on light micrographs.
On the other hand, SEM gives you a level of detail you will not even get close to with light microscopy. This could be very useful for rust ID if there was a reference library for all of the species known on a given host. Currently, this does not exist. In summary, I think it would be useful if people studying rusts made an effort to build this reference library, and I think anyone describing new rust species should make an effort to include SEM images of spores in their description (not unreasonable given it only takes a few hours and maybe 20 quid). That said, SEM is not something anyone just trying to ID something should bother with.