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Protocol for rapid preparation of nosema spore samples

by Randy Oliver

Updates 4/13/09


A hemacytomer is unnecessary unless you are collecting data for statistical analysis. The “field of view” count with a plain slide is all that is necessary for management decisions.

If you wish to purchase a hemacytometer, I suggest a Hausser Bright Line. Order from www.fishersci.com, catalog no. 02-671-5, Hausser Scientific no. 1492. However, there is great variability in the darkness of the metallized counting surface. Make sure that you specify that the hemacytometer is hand selected by Hausser as “# 1492.darker” and drop shipped to you.

Taking Samples

1. Take samples of bees from colony entrances at midday. The point is to sample the oldest bees in the colony. If samples need to be taken from inside the hive, realize that the resulting spore counts may be 10-20 times lower than those from field bees, and cannot be compared to entrance counts.

2. Do not pick up “crawlers”, as they may be disproportionally infected. Sample size should be at least 25 bees—50 is better, 500 or a pintful would be even better. (Subsamples of 25 bees from a large sample have great variability in spore counts (Oliver, unpub data)). Samples may be stored in 70% isopropyl or ethyl alcohol, however, freshly preserved samples are easier to count.

3. The easiest ways to collect samples are with a modified vacuum (see The Suck-a-Bee) or by scooping bees off the inside of the net each time you move.

Spore Counting

(this looks like a lot of instructions, but with practice only takes about two minutes to prepare the slide)

1. Strain bees from the alcohol (a fork works well) and dump onto a white platter. Separate bees with a fork, then count into groups of 10. Record the total count of bees in the sample. For large samples, see [1].

2. Place the whole bees (or abdomens only) into a ½ pint Mason jar and add water at the rate of 1 ml per bee.

3. Screw the blender blade assembly from a household blender onto the Mason jar. For field sampling, 12 volt blenders that plug into the cigarette lighter are available.

4. Blend at “chop” for 5 sec, pause until blades come to rest, and repeat. Watch to make sure that no bees get stuck out of the liquid (rare). There should be no intact bee abdomens after the first burst.

5. Remove the jar, invert, and shake to wash down all bee parts into the liquid.

6. Remove the lid and dip out a sample of the liquid with a 10 microliter loop. The sample cannot be allowed to settle, since the spores quickly sink. If the sample sets, stir it vigorously to resuspend the spores.

7. Immediately rinse the blade assembly, before the slurry dries [4].

8. Some samples (but not others) have considerable foam, which is problematic. However, a bubble-free sample can be taken by tipping the jar to the side, and scooping the foam off the surface with a spoon until foam-free liquid is seen. You may need to allow the foam to rise first for a few minutes. If so, don’t forget to stir the liquid vigorously to resuspend the spores!

9. [Updated 10/02] At this point, you can simply put a drop of the slurry onto a plain slide, drop on a cover slip, and tap it down lightly with your finger tip. If there are any large bee parts holding up the slip, you must do it over, as they will hold the slip up, and the volume of fluid will give you a higher spore count. Then simply wait 60 seconds, and grade the spore count by triage: none, a few in the field of view, or “too many.” “Too many” is roughly more than 25 total spores in the field of view, which would be equivalent to a hemacytometer count of about 5 million spores per bee.

10. Or, apply the sample to one side of a hemacytometer [2]. Dip the loop a second time to obtain another sample, and apply to the other side of the hemacytometer.

11. Check the slide—if the liquid appears “white” under the cover slip, there is too much foam for counting. Clean and reapply.

12. This prep will contain significant debris (due to the grinding of whole bees), but with practice, bubble-free samples are the norm. If there is excessive debris that interferes with counting, swirl the slurry to resuspend the spores, and take more samples with the loop.

13. Wait a minimum of two minutes for spores to settle before counting.

14. Count 5 squares of 16 subsquares in the standard zigzag pattern. Count spores [3] touching the center of the triple lines to top and left, but not bottom or right. Count both sides of the hemacytometer.

15. Some squares will have too much debris. However, there are 25 squares available for counting. If a square appears to have too much debris, shift to the adjacent square left or right (as appropriate) and count the first square that looks “clean.” In practice, this is only necessary for about one square in 4 or 5, and does not appear to affect the count.

16. [Updated 10/02] Cantwell’s “standard method” boils down to this: dilute each bee or bee gut contents with 1 ml of water. Stir, and put a droplet into the hemacytometer. The “standardized spore count” is the mean number of spores in the smallest squares of the reticule x 4 x 10^6 (4 million). This will give you the mean spore count per bee.
If you see, on the average, one spore per smallest square, then you would have a spore count of 4 million per bee.
If you count the spores in 5 squares, then divide the total count by 5, and then multiply by 4 million. If you count 5 groups of 16 tiny squares, then that is 80 squares total, so you would then divide the total count by 80 x 4 million (this is the most common count).

General Notes

[1] For large samples, “count” the number of bees volumetrically. See The Nosema Twins 3—Sampling. Wet bees pack at the rate of 3 bees per ml. Thus you would add 3 times the amount of water that you have of packed bees.

[2] I use a Reichert Bright Line hemacytometer. I’ve noted that this model varies greatly in brightness of background. You may wish to try several.

[3] N. ceranae spores are variable in shape and size. Therefore, I only count those of appropriate size, that show an oval shape, with glowing center, and a dark rim when the fine focus is adjusted up and down. I do not count questionable spores. Avoid the temptation to recount if count was “unusually” high or low.

[4] Rinse the Mason jar in plain tap water, do not use soap, as soap residue may cause foaming. If enough time has passed for any slurry to dry, soak it, then wipe it clean under running water. I have tested the efficacy of a simple tap water rinse of the blade and jar assembly. When refilled with 30 ml of water, and reblended and shaken well, the spore count of the water was only 2 spores in 800 small squares. As a further note, there are surprisingly few spores released into the alcohol of a sample bottle.

Category: Microscopy

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