University of Texas

A technique for spreading maize microsporocyte pachytene chromosomes for silver staining and EM viewing of synaptonemal complex core and lateral elements

--M. P. Maguire

The following describes a successful procedure for a number of maize stocks which combines parts of techniques set forth separately by Stack, Holm, and Jones and collaborators (mostly for other organisms) and contains a few original alterations. This technique produces relatively intact, complete-complement configurations which seem generally free of distortion.

New microscope slides are coated in advance with plastic by dipping them in a chloroform plastic solution (4g broken Falcon petri dishes: 400ml chloroform) and standing them on end in a test tube rack to dry. For this procedure plastic coated slides must then be treated with a glow-discharge unit to render the plastic surface hydrophilic, at a time no more than about three days before use. (This step may be omitted in procedures which call for use of a swelling medium containing a detergent.)

Fresh anthers at pachytene stage are macerated in a deep depression slide in 5µl of a freshly prepared ice cold medium: 2.5% sucrose, 1% polyvinylpyrrolidone and 2.5mM acid EDTA, adjusted to pH 4.6 - 4.7 with KOH; after maceration 60µl of an ice cold solution (6% paraformaldehyde, 1.5% sucrose, adjusted to pH 8.6) is quickly added. The depression slide is then covered and placed over an ice water bath for at least 30 minutes. The paraformaldehyde solution may be prepared in advance and kept in a refrigerator for 5 or 6 days. If the pH declines below 8.4, it should be discarded.)

After 30 minutes to 1 hour, the depression slide is removed from over the ice bath, and quickly warmed to room temperature on a laboratory bench. The contents are then micropipetted to plastic coated slides, sucking out the liquid from around the anther remnants. The plastic coated slides are then vibrated for 10 seconds by touching an electric vibrating engraver to the surface of the ground glass end, and these slides are then allowed to air dry, leaving them overnight at room temperature (protected from roach and ant demolition). Then the dried down preparations on the slides are rimmed with nail polish (to prevent loss of plastic during fixation and staining), and the drying process is completed by placing the slides for 3-4 hours on a slide warmer at 37 C. These slides can be stored indefinitely before fixation and staining (as described here, almost entirely the procedure of S. Stack and L. Anderson, J. Hered. 78:178-182,1987).

Immediately before staining, slides are treated with an ice cold, freshly prepared solution (4% paraformaldehyde, 1.5% sucrose, adjusted to pH 8.6) for 10 minutes (changed for fresh solution after the first 2 minutes). Then slides are briefly washed in 0.4% photoflo and air dried. Slides are then individually placed on props over 1mm of distilled water in petri dishes, and one drop of a freshly prepared 50% water solution of silver nitrate is added to each from a Pasteur pipette. A siliconized coverslip is applied to each slide, and petri dishes are covered and placed overnight in an oven at 60 C. Coverslips are then readily floated off, and slides are air dried and ready for scanning. This procedure may produce a rather dark stain so that reducing the staining time somewhat may be desirable.

Slides are scanned with phase contrast microscopy (with at least a 20X to 25X objective). Copper grids (Pelco IGC 50) are made slightly sticky by dipping them in dichloroethane (in which a short piece of scotch tape has been briefly swished), and drying them on parafilm. Such grids are carefully positioned on good synaptonemal complex configurations. (It is a good idea to tilt the slides slightly at this stage to determine whether the grids are securely positioned so as not to slip during additional manipulations.) The plastic film is then scored in a circle around and about 2mm from the grids. Next the grids on their plastic rafts are floated on a drop or two of distilled water. (The water is placed at the edge of the scoring and with luck will creep under the plastic). Then the slide is carefully immersed at a slant under the surface of distilled water in a bowl in such a way that the rafts carrying their grids are floated on the surface of the water, and the slide is then withdrawn. Plastic rafts with grids are then picked up on weighing paper (push down on the rafts from above, pushing them momentarily below the surface, and deftly invert and withdraw the weighing paper so that the plastic is on top of the grids on top of the weighing paper.) The weighing paper is propped on edge for drying. Later, after careful removal of the dried grids from the weighing paper, it helps to examine them on a microscope slide with phase microscopy and map the positions of the configurations to be observed. Store the grids in grid boxes, and you are ready for the EM.

Some cautionary notes: Coating slides with plastic film works best in relatively low humidity (below 60%). In high humidity I consistently get a cloudy plastic film.

Floating plastic rafts (with grids) off of slides works best at relatively high humidity (above 60%). In low humidity, apparently, static electricity frequently inspires grids just floated to make incredible flip dives below the water surface back to the slide from which they have just been removed, before it can be withdrawn. This is enough to wreck an otherwise placid disposition. I am told by others that sometimes grids resist being stored in grid boxes by jumping back out again as fast as you can put them in; I have not seen this action yet, and do not really need it.

Additional directions: Plastic petri dish pieces are quickly dissolved in chloroform under sonication.

Maceration medium pH adjustment is performed with KOH and HCl, avoiding addition of sodium which is considered toxic to plant cells.

A 25mM acid EDTA stock solution is prepared in advance (and stored indefinitely). Dissolution is accomplished by adding 4 or 5 pellets of KOH; pH is then adjusted to 4.1 with HCl and KOH.

Paraformaldehyde is dissolved by addition of 1N NaOH (2ml to every 100ml of solution) and heating while stirring to no hotter than 50 C. The solution is then cooled to about 4 C, and the pH is adjusted to 8.6 with formic acid, NaOH and sodium borate.

One additional piece of advice: Do not undertake this kind of silver staining unless you are fortified with grim determination, but if you are, you should eventually be duly rewarded.

Please Note: Notes submitted to the Maize Genetics Cooperation Newsletter may be cited only with consent of the authors

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