Sick Bees – Part 13: Simple Microscopy of Nosema for beekeepers
It is greatly surprising to me that with the great interest by beekeepers in Nosema ceranae, how few actually make the effort to monitor the levels of this parasite in their colonies! Even more surprising is that, despite the considerable expense, many blindly treat their colonies without having any idea as to whether their bees are actually infected!
In my last articles, I addressed the importance of monitoring infestation levels of the honey bee parasite Varroa. Now I’m going to move on to the next common parasite—nosema—of which similar monitoring allows one to make informed management decisions. It is far better to learn to monitor nosema levels yourself than it is to depend on sending the occasional sample off for testing!
Checking for nosema infection level does not require laboratory expertise, and the cost of a good microscope can be quickly recouped by not wasting your money on unnecessary treatments, or from avoiding colony loss. Unfortunately, many beekeepers are intimidated by the thought of learning how to use a microscope, get frustrated due to unfamiliarity with the necessary techniques, or have trouble identifying the spores. I hope in this article to guide you step by step through the entire process of monitoring for nosema.
The photo above shows all the equipment you will need. I highly recommend the Omano OM36L microscope (shown), which has binocular eyepieces, battery power for field use, and a mechanical stage for moving the slide around (Microscope.com offers a “Beekeepers Special” for $349). For some reason, the optics of this particular scope really make nosema spores stand out! My advice is to pay the money for a decent scope, as this is likely the only one that you will ever purchase, and you don’t want to be stuck with one that does not live up to your expectations.
You can save money by getting a monocular (single eyepiece) scope (the OM136C costs $179 for the basic model). Whatever you buy, I do recommend that you get a scope with an adjustable condenser.
Updates (in red) July 2014: I no longer take samples from the entrance for a number of reasons:
- In cool weather or with weak colonies it is often difficult to collect enough foragers.
- Entrance samples contain a disproportionate amount of highly-infected older bees.
- It is difficult to compare results with others who have taken samples from under the lid.
- My current opinion is that under-lid, or upper frame samples are better indicators of the actual degree of infection of a colony.
- It is easiest to collect bees with a vacuum (search “Suckabee” at ScientificBeekeeping.com), but they can also be swept with a brush into an open jar of alcohol.
- If there are not enough bees, stand in front of the hive for a minute or two, then step aside and allow the rush of returning foragers to land.
- Or, block the entrance with screen (not a solid block) and return in a few minutes.
- Blow into the entrance to get guard bees to rush out (wear a veil!).
- If you can’t take bees from the entrance, then take them from under the lid, but realize that your spore counts will be substantially lower, by about tenfold. However, sampling of bees from inside may give you a better idea as to whether the colony is seriously infected!
- Samples can be kept in alcohol or frozen indefinitely until you process them.
- I now simply lift the lid and brush bees into a ziplock bag or jar of alcohol. If necessary, I pull an outside frame to obtain enough bees.
It is far better to view many samples quickly than to spend a lot of time with fewer samples, due to the inherent variation in samples from hive to hive, and week to week. I’ve switched to the really quick and clean “ziplock method,” which I learned from labs in Canada and Australia.
Dump the drained bees onto a white plate, then spread them out into a single layer, so that their legs aren’t all stuck together.
Samples smaller than 50 bees can be badly skewed by one highly-infected bee. A single bee may contain 500 million (500M) spores. That means that it alone will contribute an average spore count of 10M spores per bee to an entire sample of 50 bees, even if not a single other bee is infected! Therefore, the larger the sample size, the more accurate the results. Don’t place too much stock in the count from any single sample!
At this point, you can if you wish, filter the liquid through cheesecloth or a nylon stocking in order to remove most of the trash, but I generally find this to be unnecessary. If there are any bee parts under the cover slip, use a fresh drop, as the parts will hold the cover slip up and skew your spore count higher. If any water puddles around the cover slip, blot it off with a paper towel so that it doesn’t get on the microscope lens (this is really important—don’t ever shove a wet slide onto the platform, as it will crud up the lens).
Now here’s the beauty of the ziplock method—once you’ve gotten your spore count done, you can simply zip the bag shut and toss it into the trash—no mess or washing up! It only takes me 2-3 minutes per sample turnaround, and a minute of that is simply waiting for the spores to settle.
The following directions are specifically for the OM36, but will apply to most scopes.
- (Applies only to the first slide). Rotate the lens turret so that the 4x lens (the shortest one; with a red ring) clicks into place. The degree of magnification is the product of the 10x eyepiece (ocular) lens and the 4x objective lens in the nosepiece (turret)—in this case giving a magnification of 40x. At this magnification you can easily view bee body parts, but nosema spores would only be pinpricks.
- Place the prepared slide (with a cover slip over the liquid, and any wetness blotted off) onto the stage, clipping it into the spring-loaded holder. Click on the lamp (at back of the scope base), and turn the lamp brightness to about the “4” setting. Adjust the slide location so that the light shines up through the center of the “gunk” on the slide.
- Use the coarse focus knob to adjust the lens to about 7/8” above the slide.
- Now look through the eyepieces, and turn the coarse focus knob back and forth slowly until the bee debris comes into focus.
- Adjust the diaphragm (the size of the hole through which the light passes) lever toward the dark range, so that the debris looks “natural” and has clear texture.
- Adjust the distance between the eyepieces until you see only a single, round image.
- Looking through your right eye only, use the fine focus to adjust image until it’s sharp.
- Now, looking only through your left eye only, turn the knurled ring on the left eyepiece until the image is sharp. You have now customized the scope for your particular eyes and interpupillary distance.
- Now rotate the turret to snap the 10x lens (the next longer one; with a yellow ring) into place. Increase the light with the diaphragm lever if necessary. Slowly turn the fine focus knob back and forth a bit until the debris pieces come into focus. Now you are viewing at 100x magnification, at which nosema spores are barely visible. Feel free to explore the slide at any time by using the stage adjustment knobs (note the since a microscope inverts the image, that the image moves “backwards” relative to the movement of the actual slide).
- Now rotate the turret to snap the 40x lens (blue ring) into place, and adjust the fine focus slightly —the lens will barely clear the cover slip! Be careful not to focus down too far and crunch into the cover slip! At this magnification (400x) nosema spores are easily visible, but still small.
- Use the stage movement knobs to locate a pollen grain or bee hair. Now adjust the diaphragm lever again to the optimal light level so that those objects are clear to see.
- Now focus down (top of knob going toward the back of the scope) to the lowest level that objects are in focus and look for nosema spores. You must wait at least 60 seconds from when you first prepared the slide in order to allow the spores to settle—you can watch them as they fall to the bottom and suddenly come into focus!
- Once you find spores (you may not find any in your sample), move the fine focus until they “glow.” Then adjust the condenser (this focuses the light beam) to the point where the glowing spores are bright against a relatively dark background. You can now fiddle slightly with the adjustments to get the best possible image in which the nosema spores stand out.
- Once you’ve made all the above adjustments, you can leave them set. Subsequent slides can simply be placed on the stage, and the only necessary adjustment will be the fine focus. Whew!
Nosema spores have a few distinctive characteristics that will confirm your identification:
1. Nosema spores are still quite small even at 400x!
2. The spores are distinctive elongated ellipses—similar in shape to vitamin or fish oil capsules (but variable).
3. They will all be about the same size (N. ceranae spores are somewhat variable, especially bee to bee).
4. Most of the spores will settle to rest at the bottom of the liquid, and will thus all come into focus at the same level.
5. Note: in fresh bee preps (those not preserved in alcohol) the organisms in the gut are still alive, and the nosema spores will often jiggle and move about slightly.
There are two distinctive characteristics that will confirm the identification of nosema spores—these can be best observed by jiggling the fine focus knob back and forth slightly as you view the spores.
6. The spores will be clearly outlined with a smooth, dark elliptical line,
7. then the outline will fade, and the centers will glow brightly. A spore must have both of these characteristics, as other objects will also have oval outlines or glow, but won’t do both.
With practice, your brain develops a “search image” for the spores, and they begin to jump out at you from the background debris.
8. Nosema ceranae looks somewhat different than N. apis to the experienced eye—apis is a bit larger and broader, and the ends of the spores are “blunter.”
I’m going to assume here that you ‘re going to simply do “field of view” counts on a simple glass slide, as I don’t feel that tedious hemacytometer counts are generally justified unless you are compiling data for research.
Use the stage adjustment knobs to “take a trip” around the slide. Pick an area to view that has a representative spore density.
If you do decide to get a hemacytometer, I recommend a Reichert Bright Line—order through a lab supply, but make sure that they call the manufacturer directly and ask them to ship one with an extra dark background. Note that in the above photo, I’ve adjusted the scope such that the spore “outlines” are in focus, but the centers of the spores are not glowing much. Compare it to the following photo:
For Nosema apis, the treatment threshold was considered to be a mean spore count of 1M per bee. On the other hand, there is considerable debate as to what constitutes a worrisome spore count for Nosema ceranae. At the time of this writing (9/16/2011) a few million spores (up to about 25 in a field of view) would be considered by many to be “normal” for field bees for much of the season, perhaps spiking to several million (100 or more spores per field of view) during spring when there are heavy pollen flows, but then dropping to near zero during summer. I cannot make recommendations, but post the latest information at ScientificBeekeeping.com.
I have found no particular relationship between nosema and dysentery, other than that colonies exhibiting dysentery are more likely to transmit nosema spores.
I’ve shown many beekeepers how to quickly process bees for spore counts. I suggest that every bee club purchase a scope, and assign one member to gain proficiency at its use. At the break during your meetings, you can easily process a great number of bee samples in a few minutes (have everyone bring a counted sample in a ziplock bag). Such sampling will allow beekeepers to actually track nosema levels throughout the season, and determine whether it appears to be a problem in your apiaries.
You can also buy a microscope camera that will transmit the image on the slide to your laptop or to a digital projector for all members to view at the same time (as opposed to having a queue waiting at the scope). I’ve tried several digital cameras — be forewarned that the image will not be quite as clear as when viewed directly through the microscope lens, but definitely worthwhile as a training aid. Microscope.com offers the OptixCam series, of which I found the OCS-3.0 ($329) easy to use and adjust.
Now that you’ve forked over your hard-earned cash for a shiny new microscope, treat it well for a long life. A microscope is a precision tool full of delicate parts. Don’t ever bang or drop a scope — those tubes may contain 15-20 lenses that can be jarred loose! A wise practice is to always carry a scope with both hands. Keep the scope covered or in a case when not in use. Dust, moisture, skin oils, and bee guts are the scope’s enemies! Wipe off any liquids with a soft cloth moistened in isopropyl alcohol. Never touch the lens surfaces with your fingers or regular tissue paper. Use only microscope lens paper or a Q-Tip, moistened in lens clearer or alcohol.
Now that you know how to determine spore counts, I will be following this article with a deeper look into our current state of knowledge of Nosema ceranae.