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Valuable teaching websites for microscope use:
Now that you have your microscope, if you haven’t already looked at How to Use the Microscope to learn the parts of a microscope and how to operate it, do so now. We'll be reviewing some of this below as we apply the information to our task, but it would be good for you to get the overview first. That page is from The Biology Corner, a website for science teachers.
In the Tape Testing for Mold topic, we’ll be talking about how and where to tape sample, but meanwhile, let’s get the background work done for looking at a slide under the microscope.
Setting up a microscope slide
Microscope slides are 3”x 1” pieces of clear glass. They are available for around $5/box on Amazon. It doesn’t matter if you get them with or without a frosted end. You’re not going to be writing on the frosted end and permanently storing the slide.
If you were looking at a tape sample under the microscope, you’d first put the tape on a microscope slide.
In order to better view any mold on the slide, you’d first put a drop of stain on the slide before positioning the tape. The stain gets absorbed by the mold, thereby coloring the mold. I’ll come back to the stain below.
Stick one end of the tape to the slide, and then smooth out the rest of the tape on the slide.
Blot the slide on a piece of paper towel to remove extra stain. You don’t want to accidentally get the microscope lens wet, or you will have to blot the lens with a paper towel and then sit for 10 minutes while the lens dries off.
Position the slide on the microscope stage.
Focusing the microscope
Knobs are attached to the underside of the stage so that you can move the stage backward and forward and from side to side, viewing different areas on the slide.
Two sets of knobs are located on each side of the microscope towards the lower back. The larger knob is stiffer to turn and is for coarse focus. You turn that to get in the ball field. Once the coarse focus is set, you rarely have to touch the big knob again, unless someone else accidentally turns it.
If you have trouble focusing with the coarse focus knob, maybe there is nothing to look at on the slide. Hold the slide up to the light to locate the debris. Make sure the debris is over the hole on the stage where the light shines through.
Once you have located and focused your sample, then you want to fine focus the image to better see the details. The smaller set of knobs is for fine focusing. Since there is a set of knobs on both sides of the microscope, you can use either your left or your right hand to focus.
Focus the eyepieces to your eyes
What if the view through the left eyepiece is clear but through the right eyepiece is fuzzy? You have to focus both eyepieces to your eyes. Here’s how to do that: Look through the right eyepiece and use the fine focus to make the image of what you are viewing crisp and clear. Then close your right eye and view the image with your left eye. The left should be just as clear as the right. If the left is blurry, turn the base of the left eyepiece until the image is clear. Then you will be able to see clearly with both eyes. (Take glasses off to focus.)
If you only see a partial picture when looking through your left eye, slide the eyepieces closer together or farther apart to match the distance between your eyes.
Other knobs
On the right hand side, if you are right-handed (and on the left, if the microscope is adjusted for a left-handed person), there should be two knobs (one on top of the other) which are used to move the stage back and forth and from side-to-side. Use these to move around the microscope slide.
Spending a little more for a microscope with these knobs is worth the money. With cheap microscopes, you have to move the slide back and forth with your fingers.
There are other little knobs around the stage that have their own jobs. Touching these little knobs is rarely, if ever, needed after the dealer has them in adjustment. You can try them and see what they do but remember the reference point that they came set to, so you can return the setting to that point.
I trained myself to look at and identify mold. You can do the same. I worked with the book, Identifying Filamentous Fungi – A Clinical Laboratory Handbook, by Guy St-Germain and Richard Summerbell, Star Publishing Company, CA, 650-591-3505.
Let me know how I can help. Perhaps you would like to send me photos of what you are seeing under the microscope to confirm the identification. Check the master list of photos in the next section.
Gather your supplies:
You’ll find ¾” clear, shiny transparent tape in any office supply store. Don’t use Magic Tape, because the light won’t pass through it, and you’ll see just a cloud-like blur. Avoid invisible tape.
Buy 3”x 1” microscope slides on-line. They come in boxes of 100. Frosted ends don’t matter, unless you want to make permanent slides and label them. I haven’t found out how to prevent permanent slides from going cloudy, even after adding alcohol to the stain.
I re-use slides. After the tape is removed, the slide is wiped off with a damp paper towel. There should be no writing on the slide. An identifying number is written on the tape.
I used to soak the used slides in hydrogen peroxide overnight to kill any mold on them. Then, through experience, I learned that soaking wasn’t needed. Just cleaning off a dirty glass surface with a damp paper towel was sufficient.
Stain your sample.
Mold spores are easier to pick out from background debris if you put a drop of colored stain on the slide under the tape. I use a fuchsin (pink) stain, but many microbiologists prefer cotton phenol blue, a blue stain which may be better for photography. The fuchsin costs less and doesn’t give me a minor headache, like the blue stain does.
You might have trouble purchasing stain, because some supply houses only sell to businesses. If you can’t locate stain or it’s too costly, send me a stamped, self-addressed envelope, and I’ll send you a little fuchsin powder. A little goes a long way, and the supply I bought from Clarkson Laboratory & Supply, 1-619-425-1932, for $27, should last for years to come, even sharing it.
I was originally told to add distilled water, vodka (my guess is to keep down the growth of bacteria), and glycerin to a tiny bit of the fuchsin powder, like the amount on the head of a pin. Since then, I’ve at times been close to running out of the solution on a job, so I added tap water to the little bit of remaining stain solution, and this worked fine, just being a little lighter in tone than usual but workable. Now even when first mixing the stain, I just add tap water.
I don’t carry fuchsin powder around, because it stains very easily. Imagine a client’s beige carpet with bright pink stain on it. Instead, I keep a second little medicine bottle (with a dropper), containing water and fuchsin concentrate (fuchsin and water) in my office.
I add a few drops of this solution to a second medicine bottle filled with tap or distilled water. It’s a homemade stain, but it’s cheap and it works. I used to pay about $45 for a small medicine bottle of prepared fuschin stain, until I learned I could buy the whole bottle of fuschin powder for $27.
I hold the medicine bottle up to the light to gauge the color of the water, so that it’s not too light or too dark. You’ll soon get a feel for what shade you prefer for viewing mold. I add 3-4 drops of concentrate to a second medicine bottle.
Don’t spill the bottle of stain!
The bottle of stain can be set in a container so that if the bottle tips, at least the stain is held in the container.
Screw on the top when the bottle is not in use. It’s easy to get lazy on this subject, but after you tip the bottle over accidentally a few times and are relieved that the top was on, you discipline yourself because you know that accidents happen. A client’s cat once knocked an expensive piece of equipment off a table, fortunately not damaging it.
Clear medicine bottles with glass droppers are available on Amazon.
Protect the table with a cover. Twice I had a bottle of stain tip over, and I was very glad to have a plastic leaf bag underneath and not the finished surface of the client’s dining room table.
Taking tape samples
Take about 3 inches of clear, glossy tape and stick one end to an index finger and the other end to your thumb, sticky side of tape facing outward.
You could either press the tape straight down onto a surface with your two fingers or slide your index finger back toward the middle of the tape so that you press on the surface with your index finger. Just the middle of the tape should have visible debris on it. The two ends should still be sticky.
If you pick up a wood splinter or other larger material on the tape, knock it off. A 3-dimensional piece of grit will just make a bump under the tape.
Some surfaces have a lot of dirt on them. Brush crumbly matter off the tape.
It’s hard to see mold spores amidst a jumble of dust and other particulates, even if the light could get through solid pieces. If you just touch the tape lightly to the surface so that not much dirt adheres, you should see spores more easily. Tip: Instead of sampling the dusty top of a surface, sample the underside.
If there’s too much debris on a slide, maneuver the view to areas that aren’t so dense with debris. If those areas are free of spores, then there is a better chance that the dirtier areas are fairly clear, too.
Often I’ll touch a tape with a lot of debris (such as from an AC filter) to a culture plate to see how much mold grows. I might not get a good look for spores under the microscope, but spores will grow out from debris.
In one sample where I hadn’t seen spores under the microscope, plenty of Aspergillus niger colonies grew, which was a lesson that when there is a lot of debris, you just might not be able to see the mold spores.
Examining a tape touched to the inside of ductwork, either through an access hole or inside a vent, may also provide a clue about the amount of mold in the system.
What you should have now is a 3” piece of tape that has nothing but fingerprints on the ends and some debris in the center of the tape. Even a half inch or less of debris is fine, less than the size of a dime. When the debris is magnified 400 or 600 times, the area could look as big as a football field.
Now that you have your tape sample, you’re ready to stain the sample.
Using the eyedropper, place a drop of stain in the center of the microscope slide.
Stick one end of the tape near the white or opaque end of the slide. Position the end so that when the tape is flat on the slide, the debris on the tape will be approximately in the center of the slide.
Hold the free end of the tape with one hand. With a finger of the other hand, smooth the tape down lightly over the stain. You might have to lift the tape and smooth it again to spread the stain out under the debris.
When enough of the debris has stain on it, smooth the entire tape down to eliminate most of the air bubbles.
If any stain seeped out from under the tape, blot it with a tissue, so the microscope stage doesn’t get wet from the stain.
If any tape is hanging over the end of the slide, either fold it back or slice it off against the slide. Otherwise, the tape may stick to the microscope stage and interfere with moving the slide around.
Now that we’ve gone through preparing a single sample, let’s back up a minute and talk about how to take multiple samples and not mix them up. Here’s my method:
I use a gallon-size zip-lock bag, writing the client’s name and date on the white label. Then, as I take a sample, I use a fine-point Sharpie to write the number on one end of the tape. This permanently marks the tape. In a notebook, I use a ballpoint pen to identify the location for each number.
For example, 1. inside kitchen sink cabinet; 2. half bath sink cabinet; 3. main bath sink cabinet on right; 4. main bath sink cabinet on left; 5. base molding under hall window, 6. discolored area on bedroom ceiling, and so on.
I don’t carry the microscope slides around to reduce the risk of glass breakage.
What I often do is to make my list of where I want to sample first. Then I’ll follow my list in taking samples. Line the numbered tapes up on the outside of the bag, making tabs at one end of the tapes to make removal easier. Don’t place tapes over the white label on the bags, because the white may come off.
At the microscope
Set up a comfortable viewing area on a table or desk near an electrical outlet. One of my office chairs is one with the backless seat where you sit on one upholstered pad and your knees go on the lower pad. Over extended periods of microscope work, I appreciate different positions from my computer chair.
Place the stained microscope slide on the stage, using the clip to secure it.
Look at the numbers on the three or four objectives on your microscope. Those numbers tell you the magnification of the lenses. On my microscope, there is a 4x, a 10x, and a 40x. Remember to multiple the number on each objective by the number on the eyepiece lenses. So, with my 15x eyepiece, the 4x objective really shows an item 60 times as large as life. The 10x shows 150 times, the 40x shows 600 times, and the 100x shows 1500 times (with the 100x, you have to use immersion oil to get clarity).
Here’s some help with focusing the microscope:
What if someone turned the coarse focus knob by mistake so that the focus is way off and you can’t get anything to focus? Use the 4x objective first. Focus using the coarse adjustment knob, which is easier to do at lower magnification, and then move to the fine focus knob. Once you have a focus at 60x, you can turn to higher magnifications and should be able to regain your focus at each one with just the fine adjustment knob.
60x is 60 times larger than life but still too small to adequately view mold. With practice, you’ll be able to pick out some mold spores and certainly large areas of mold. Stachybotrys spores are very tiny at 60x, but you can still identify them as Stachybotrys, once you know what you are looking for.
Here’s a common mistake. Do you see large round or irregular areas on the slide? They are probably air bubbles. I mistook them for microorganisms the first time I saw them under a microscope, and lots of my clients have, too. Mold spores are much smaller than these large air bubbles.
Some cheap tape, such as from Dollar General, results in more tiny air bubbles than better quality tape.
Now move on up to the next objective. On my microscope, that’s the 10x objective, which would give me 150 times magnification (10 x 15). Now the mold spores are a little larger and easier to see.
Remember that you shouldn’t have to touch the coarse focus knob again, because the sign of a microscope in good adjustment is that you can go from magnification to magnification with just minor fine focusing.
On you go now to the objective you’ll use the most, the 40x one which gives you magnification 600 times as large as life with 15x eyepieces.
Sometimes no matter how hard I try to focus, the slide remains blurry. If all else fails, see if you placed the slide upside-down on the stage, with the tape on the underside of the slide. I’ve done this more than once.
If you accidentally get stain on the objective’s lens and have to wipe off the stain, it might take a few minutes for moisture to dry. Images will be fuzzy until the lens dries. Wait a bit and then the clarity of the images should return.
At 600x, you’ll be able to see mold. If you know what to look for, you’ll find it sooner or later. Let me share my early experience with you.
May as a beginner with the microscope
I had had some background biology and microbiology science classes in college and grad school, and more when I started environmental home inspection studies, beginning in 1994. Those were the days before mold became a hot topic, and not much class time was devoted to the subject of mold inside houses.
Around 1996, I found a reference book mentioned above which would become my classroom. Identifying Filamentous Fungi: A Clinical Laboratory Handbook, by Guy St-Germain and Richard Summerbell, can be ordered from Star Publishing Company, 1-650-591-3505 (though they have an newer book out since then). There are pictures and descriptions of what mold looks like under the microscope, so you can match what you’re seeing with the pictures. The book gives several ways of identifying mold, and I spent hours and hours with that book and use it to this day as a reference. Your progress will be much faster, because I am going to tell you some shortcuts. You’d enjoy the book, but you’ll find enough here to get you started.
The first mold I identified microscopically was green bread mold, Penicillium. I was able to focus in and see the chains of spherical spores easily. I knew that I needed to find a fruiting body that produces the spores to be sure I was seeing Penicillium. There were so many spores on the slide I was viewing that I couldn’t see fruiting bodies for quite a while. Finally, I figured I’d look around the edges of the mold, where the growth wasn’t so dense. Suddenly, I saw a branch-like fruiting body with chains of spores coming off the tips. How exciting! How delicate, how beautiful! My first positive identification! I was on the way.
Since those days, I have identified a lot of the molds in the St-Germain-Summerbell book. When I started, I’d see a new mold and then leaf through the book to see if I could identify the mold. I’d look for the fruiting bodies, for the shapes of the spores, for the colors of the colonies (if I was growing them), and for other identifying characteristics that the book listed. How exciting to see Trichoderma, Mucor, Rhizopus, Syncephalastrum, right under my microscope! And to find other molds, one after the other. Some of these molds have to be grown in Petri dishes from air samples; you typically won’t find them on tape samples.
The good news is that there are only about a half dozen common indoor molds associated with dampness conditions. Thus, while I had a lot of gratification from learning to identify other molds, most of what I saw at houses was Alternaria, Aspergillus, Aureobasidium, Chaetomium, Cladosporium, Penicillium, Stachybotrys, and sometimes Trichoderma.
These common indoor molds can be found through tape sampling. Unless you decide to do culture plate air sampling business, you probably won’t find too many fungi beyond these.
Some molds are not in my reference book. The St-Germain-Summerbell book deals just with molds that are known to affect human health, particularly through infection. Allergic or asthmatic symptoms or neurological concerns are not addressed in the book. So the book is limited, but it’s a good place to start for the pictures and descriptions. An Internet search on various molds brings up an endless list of sites relating to mold and to specific kinds of mold. Include “microscope picture” and the name of the mold in your search terms, and you’ll be in business.
Many molds don’t produce spores, and all you see under the microscope is a bunch of branches. These may be “sterile fungi” or “non-sporulating fungi,” which may or may not be allergenic. Little is known about them, and there are many, many species, often with minor variations. Labs generally don’t further differentiate them, if they are listed at all. As with any mold, clean off surface growth and try to prevent re-growth. One microbiology professor called them “the white fuzzies.”
The field of mycology (a branch of botany dealing with fungi) is vast. One day I read about a book identifying molds growing on telephone poles. Checking out telephone pole fungi sounded like a fun thing to do, so I ordered a copy of the book. Forget it. Trying to get conversant with the tiny distinguishing aspects of 1000 similar-looking molds was definitely not fun. I’ll leave telephone poles to the microbiologists. Nor do I try to distinguish among even the common species of common molds, such as Aspergillus. I leave that up to the microbiologists, should a client need to know. Sometimes the report comes back listing some species that aren’t even in my reference books. There are whole books written just on Aspergillus species. There’s also a website, www.aspergillus.org.uk.
Using immersion oil to get higher magnification
Skip this section if you’re not planning to work with 1500x magnification (the 100x objective x 15x eyepiece). I seldom use 1500x, but since many microscopes will come with a 100x objective, I’m including basic information on how to use it.
If you want to use the 1500x magnification capability, then purchase a bottle of Type A immersion oil from the place where you bought your microscope or on-line.
Focus the slide at 600x. Then, rotate the objectives slightly to the left, so that you have access to the stained sample on the slide.
Before the 100x objective snaps in place, place a drop of the immersion oil on the top of the tape covering the stained sample on the slide.
Now slowly move the bottom of the 100x objective right into the oil.
Work the fine focus knob for your 1500x view. Tool around and look your fill. When you have finished looking, move the 100x objective out of the oil and remove the slide.
Don’t expose any of the other objectives to the oil. Once you are viewing a slide at 1500x, that’s it, you can’t go back to a lower magnification for that sample without first cleaning the oil off the tape. If you do go back to 600x without cleaning the slide, you’ll have two objectives to clean, not just one.
After you finish using the 100x objective, clean off the oil with lens cleaning paper or a tissue. One technician told me a trick of the trade, i.e., that saliva, applied to a lens cleaning paper, helps clean off the last of the oil residue. I’m not real talented in getting immersion oil off the 100x objective, so if you’re having trouble, too, welcome to the club.
Because cleaning off the immersion oil was a nuisance, my compromise for when I wanted to see images larger than 400x, was to go to the 15x eyepieces and get 600x (15x times 40x) magnification without the use of oil. Now I mostly just use the 600x, which is adequate for our purposes.
You may think you have adequately cleaned a lens, only to find the image is very cloudy. Wait a few minutes. It should clear once it dries thoroughly.
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Next up: my cheat sheet of Mold under a Microscope or back to the Mold Topic Index.