Fixation of well dried (at RT) PB smears can vary from 1-10minutes;
automated systems tend to use about 1-2 minutes and use the solution only
once. Manual systems- most labs would use about ten minutes, and for these
precautions against absorbtion of water via humidity must be taken, usually
being replaced twice daily, but more frequently at those times of the year
when humidity is high. The first signs of unacceptable water content in the
Fixing Methanol, will the appearance of clear refractive spaces on the
biconvave surface of erythrocytes perhaps only a few cells per High powered
field, but will increase further as the water content increases, to the
extent that the films are no longer diagnostic. Replacement when you see
more than say 1-2/HPF might not be a bad idea. This artifact may also be
seen in some automated systems where stain pack is not turned over very
quickly, and rather than replacing the stain pack, economy of reagent can be
maintained by manually fixing the slides before thay go on the machine, this
is particularly so for the older Hematek grey models. Caution:- longer
fixation times are required for Bone Marrow smears, viz 15-20mins and always
use fresh methanol for these.
Most persons using Giemsa prefer to stain the smear first with May Grunwald
or Jenner stain, either using it neat or diluting 1:2 with Buffer. This
pre-step gives better granule definition and clarity, and also changes the
traditional reddish purple of nuclei with plain Giemsa to a blue purple as
seen in Wright's.
The selection of Sorensen's buffer will vary form 6.4-7.2, with the lower pH
being most popular with wright's rather than Giemsa. The aim is to select a
pH that produces a colour balance that readily allows the user to
differentiate between normochromic and polychromic red cells and distinguish
toxic granulation when present, this is usually pH 6.8. If looking for
malarial parasites, then a pH of 7.2 best serves the exercise as it allows
better contrast to detect chromatin dots, trophozioites etc.
Dilution of the Giemsa solution is best immediately before use and will vary
from 1:8-1:12 depending upon your protocol. As a general rule of thumb the
higher dilutions require longer staining times of about 20mins and the lower
between 6-12 minutes depending upon tthe quality of the Giemsa. It was
frequently claimed that the longer times gave better definition, but I must
admit that I've seen short timed smears that are every bit as good. For many
years good quality Giemsa would be stable after dilution for about 6-8hrs,
for the last 2-3 yrs, the best you can hope for 3-4hrs. After dilution the
solution starts to deteriorate, with the appearance of flocculant and a
subsequent loss of staining ability or strength- as the time progresses you
may need to compensate by increasing your staining time, but after 3 hrs you
will need to replace it.
Recipes for Giemsa vary, whether it be that of Hayhoe or Dacie & Lewis, and
measurements may be by weight or volume. In any case those that have a 50%
by volume content of Glycerol (Analar or USP) are the most stable, and under
no circumstances ever heat you r glycerol to more than 45C. (Most texts list
56C), as when you get above these temps. there is a risk of oxidation even
in the stock solution, I use 45C as a cut-off point to give me a safety
margin. Dye content will also vary from 0.45-0.8%;-Lillie's comments should
considered here. After standing for up to 5 days, filtration to remove
undissolved material is essential.
Differentiation by giving your slides two rinses in buffer of two minutes is
fairly standard, but you can overdo it, and a single rinse of say three
quick dips  may in fact suffice. It will depend upon your giemsa solution
and tastes.
If overstaining is a problem then consider adding Methanol to your buffer
rinse starting at 5% and adjust to your tastes, followed by a water rinse to
remove solvent.
Regards Mike (downunder)