The images in the  previous mosquito instalment were made with the use of a Greenough microscope. Comparatively large specimens like mosquito larvae are well suited for use with a Greenough style binocular. This type of microscope is functionally, two monocular microscopes positioned close enough that each objective may observe the same specimen and the unique image of each be seen by the eye above each ocular. The wide field of view and exceptional depth of focus provided by the fully parallel optical paths is generally better than can can be had by the use of any single objective microscope.

For people without access to a stereo microscope larve may be observed with the lowest power objective available. In many cases (as in the inexpensive Bausch & Lomb ST used for this post) this will prove to be the 10x objective which will regretably not afford viewing the entire larvae. If the objective is divisible as in the commonly found B&L 10x then remove the front element, otherwise put the lowest power on hand in place. Use a low-power or wide-filed ocular where available.

It’s certainly arguable that for live viewing one really needs a low power stereo or disecting microscope, but a steady hand will do. Let’s work from the assumption that all that’s on hand is the B&L ST (Standard Teaching) microscope, plain glass slips and square 22mm coverslips. Be aware that the 10x objective and 10X ocular combination give a very limited field of view. A third instar larvae will span the visible field thrice over, and that’s just the first problem!

Mosquito larvae are thick, too thick to just take one up in an eyedropper, deposit it on the slip and cary on. The weight of the coverglass would crush and distort the larvae sevearly and it would tend to slope away from the head of the larvae presenting the real risk of leaving that portion high and dry while surface tension pulls the water away towards tail end. Something is needed to support the coverglass (apart from the larvae itself). Specially made well slips would work for smaller larvae, but they’re expensive and unnessicary; all that’s required is something to support the coverglass. Grease and wax are both good options, but wax is a touch more difficult. Don’t worry about using up that expensive high-density vacuum great used for groud glass joints, plain old white petroleum (Vaseline) will do nicely.

Grease:

1. Smear a small bit on the heal of one palm, make the spot roughly equal in size to that of the coverglass to be used.
2. Cafefully but firmly draw one edge of the coverglass over the greased spot on the heal of your hand. Use a bit of pressure (as if you were trying to wipe up the grease, which you are) but be mindfull of breaking the coverglass.
3. Repeat the manouver with the opposite edge, carefull to use the same face of the coverglass.

The grease will form two ridges, more than enough to support the coverglass.
Wax:

1. Soft dental wax is best but common parrafin wax will work as well.
2. Warm a few cubic millimeters by rolling it betwix index and thumb.
3. Press the warmed wax onto the far side (make believe there’s a specimen upon the slip and keep well away from it) of a clean slip as hard as you dare without breaking the slip.
4. Use a scalple to cut away two thirds of the wax and again roll and press the wax to the slip. Choose a point roughly 10mm distant to form the second point of a triangle with the first bit of wax.
5. Cut away one half the wax and again roll and press, completeing the triangle.

The wax supports are a good deal harder to get thin enough for most uses but can be a good option for anyone overly worried of breaking a few covers as they get a hang of the grease method.