Protein Protocols Protein Protocols

Rapid Staining with Nile Red 247
11. Focus the camera (Polaroid or CCD) with the help of lateral illumination with a white
lamp, place the optical filters described in Subheading 2., item 9, in front of the camera
lens and, in the dark, turn on the transilluminator and photograph the gel (see Note 10).
Finally, turn off the transilluminator.
12. Develop the different Polaroid films for the time indicated by the manufacturer (see Note
11). Spread the Polaroid print coater on the surface of the 665 positive immediately after
development. (The positive prints of the 667 and 669 films do not require coating.) The
665 negatives can be stored temporary in water but, before definitive storage, immerse the
negatives in 18% sodium sulfite and agitate gently for about 1 min, wash in running water
for 5 min, dip in a solution of wetting agent (about 5 s), and finally dry in air (see Note 12).
13. Scan the photographic negatives to determine the intensity of the Nile red stained protein
bands if a quantitative analysis is required. The quantitative analysis can also be performed
directly from the image obtained with a CCD documentation system (see Note 13).
4. Notes
1. Isoelectric focusing gels do not contain SDS and should be treated with this detergent
after the electrophoretic run to generate the hydrophobic SDS–protein complexes specifically
stained by Nile red (6). In general, for systems without SDS we recommend an
extensive gel washing (20 min in the case of 0.75-mm thick isoelectric focusing gels) with
0.05% SDS, 0.025 M Tris, 0.192 M glycine, pH 8.3, after electrophoresis. The gel equilibrated
in this buffer can be stained with Nile red following steps 6–9 of Subheading 3.
2. To reduce the background fluorescence after the staining with Nile red it is necessary to
preclude the formation of pure SDS micelles in the gel (see Subheading 1.). Thus, use
0.05% SDS to prepare both the separating and stacking gels and the electrophoresis buffer.
This concentration is lower than the critical micelle concentration of this detergent
(~0.1% [3]), but is high enough to allow the formation of the normal SDS–protein
complexes that are specifically stained by Nile red (4).
3. Use 2% SDS in the sample buffer to be sure that all protein samples are completely saturated
with SDS. Lower concentrations of SDS in the sample buffer can produce only a
partial saturation of proteins (in particular in highly concentrated samples), and
consequently the electrophoretic bands could have anomalous electrophoretic mobilities.
The excess SDS (uncomplexed by proteins) present in the sample buffer migrates faster
than the proteins and form a broad band at the bottom of the gel (see Note 5 and ref. 4).
4. Storage of protein samples prepared as indicated in Subheading 3.2. at 4°C (or at
lower temperatures) causes the precipitation of the SDS present in the solution. These
samples should be incubated in a boiling-water bath to redissolve SDS before using
them for electrophoresis.
5. The bottom part of the gel (i.e., from about 0.5 cm above the bromophenol blue band to
the end) should be excised before the staining of the gel. Otherwise, after the addition of
Nile red, the lower part of the gel produces a broad band with intense fluorescence. This
band is presumably caused by the association of Nile red with the excess SDS used in the
sample buffer (see Note 3). In the case of long runs, bromophenol blue and the excess
SDS band diffuse into the buffer of the lower reservoir and it is not necessary to excise the
gel bottom.
6. Nile red is very stable when dissolved in DMSO (see Subheading 2.1.), but this dye
precipitates in aqueous solutions. Since the precipitation of Nile red in water is a rapid
process and this dye is only active for the staining of SDS–protein bands before it is
completely precipitated (4,5), to obtain satisfactory results, it is necessary: (a) to perform all
the agitations indicated in these steps (in order to favor as much as possible the disper-