DETECTION OF PROTEINS
SECTION III
The protocols in this section require that detectable proteins be previously separated using
either one-dimensional (UNIT 10.2) or two-dimensional (UNITS 10.3 & 10.4) electrophoresis. In
UNIT 10.6, detection by Coomassie blue and silver staining of protein-containing bands or
spots in a gel is described. Alternatively, proteins in a gel may be electrophoretically
transferred to a blot transfer membrane and detected by staining with India ink or colloidal
gold (UNIT 10.7) or by immunoblotting (UNIT 10.8). Another method is via biosynthetic
labeling of the protein of interest using [35S]methionine (UNIT 10.18), followed by autoradiography (APPENDIX 3).
Staining Proteins in Gels
UNIT 10.6
The location of a protein in a gel can be determined by either Coomassie blue staining
(see Basic Protocol 1) or silver staining (see Basic Protocol 2). The former is easier and
more rapid; however, silver staining methods are considerably more sensitive and thus
can be used to detect smaller amounts of protein. Rapid staining procedures are provided
for each method in Alternate Protocols 1, 2, and 3. Support Protocol 1 describes how to
photograph stained gels.
Fluorescent staining has become a popular alternative to traditional staining procedures,
mainly because it is more sensitive than Coomassie staining, and often as sensitive as
silver staining. The unit therefore includes a protocol describing SYPRO Orange or
SYPRO Red staining of proteins in SDS-polyacrylamide gels (see Basic Protocol 3);
variations on that procedure for proteins in nondenaturing gels are included as well (see
Alternate Protocol 4). SYPRO Ruby staining of two-dimensional gels is also described
(see Basic Protocol 4). Support Protocol 2 describes the photography of the fluorescently
stained proteins.
Additional fluorescent techniques have been developed to monitor phosphorylated and
glycosylated proteins. Alternate Protocol 5 describes the staining of phosphoproteins in
minigels and larger, two-dimensional gels with Pro-Q Diamond phosphoprotein gel stain,
while Support Protocol 3 tells how to image and document the findings from the
phosphoprotein staining. Support Protocol 4 details a method for selectively detecting
phosphotyrosine residues.
Alternate Protocol 6 details the differential fluorescent staining of glycosylated and
nonglycosylated proteins, and Support Protocol 5 describes how to photograph these gels.
COOMASSIE BLUE STAINING
Detection of protein bands in a gel by Coomassie blue staining depends on nonspecific
binding of a dye, Coomassie brilliant blue R, to proteins. The detection limit is 0.3 to 1
µg/protein band. In this procedure, proteins separated in a polyacrylamide gel are
precipitated using a fixing solution containing methanol/acetic acid. The location of the
precipitated proteins is then detected using Coomassie blue (which turns the entire gel
blue). After destaining, the blue protein bands appear against a clear background. The gel
can then be stored in acetic acid or water, photographed, or dried to maintain a permanent
record.
BASIC
PROTOCOL 1
Analysis of
Proteins
Contributed by Joachim Sasse and Sean R. Gallagher
Current Protocols in Molecular Biology (2003) 10.6.1-10.6.25
Copyright © 2003 by John Wiley & Sons, Inc.
10.6.1
Supplement 63
Materials
Polyacrylamide gel (UNIT 10.2A)
Fixing solution for Coomassie blue and silver staining (see recipe)
Coomassie blue staining solution (see recipe)
Methanol/acetic acid destaining solution (see recipe)
7% (v/v) aqueous acetic acid
Whatman 3MM filter paper (optional)
Gel dryer (optional)
1. Place the polyacrylamide gel in a plastic container and cover with 3 to 5 gel volumes
of fixing solution. Agitate slowly 2 hr at room temperature on an orbital shaker or
rocking platform.
If agitation is too rapid, the gel may break apart. Use fixing solution only once.
2. Pour out fixing solution. Cover the gel with Coomassie blue staining solution for 4
hr and agitate slowly.
Use staining solution only once.
3. Pour out staining solution. Rinse the gel briefly with ∼50 ml fixing solution.
4. Pour out fixing solution. Cover the gel with methanol/acetic acid destaining solution
for 2 hr and agitate slowly.
5. Pour out destaining solution. Add fresh methanol/acetic acid destaining solution and
continue destaining until blue bands and a clear background are obtained. Store the
gel in 7% aqueous acetic acid or water.
Do not store gels in fixing solution because protein bands will eventually disappear. Gels
can be stored up to 1 year at 4°C in a sealable plastic bag.
6. If desired, photograph the gel (see Support Protocol 1).
7. If desired, dry the gel to maintain a permanent gel record. Place the gel on two sheets
of Whatman 3MM filter paper and cover top with plastic wrap. Dry in a conventional
gel dryer 1 to 2 hr at ∼80°C.
Alternatively, place gel on filter paper and dry according to instructions supplied with gel
dryer.
ALTERNATE
PROTOCOL 1
RAPID COOMASSIE BLUE STAINING
Protein bands stained using this protocol can be detected within 5 to 10 min after adding
rapid Coomassie staining solution. Because the Coomassie blue concentration is lower
than that used in Basic Protocol 1, the gel background never stains very darkly and the
bands can be seen even while the gel remains in the staining solution. Another difference
is that isopropanol is substituted for methanol in the fixing solution. This method is
slightly less sensitive than the standard procedure (see Basic Protocol 1).
Additional Materials (also see Basic Protocol 1)
Isopropanol fixing solution (see recipe)
Rapid Coomassie blue staining solution (see recipe)
10% (v/v) acetic acid
Staining Proteins
in Gels
1. Place a polyacrylamide gel in a plastic or glass container. Cover the gel with 3 to 5
gel volumes isopropanol fixing solution and shake gently at room temperature. For
a 0.7-mm-thick gel, shake 10 to 15 min; for a 1.5-mm-thick gel, shake 30 to 60 min.
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2. Pour out fixing solution. Cover the gel with rapid Coomassie blue staining solution
and shake gently until desired intensity is reached, 2 hr to overnight at room
temperature.
Bands will become visible even in the staining solution within 5 to 30 min, depending on
gel thickness. The gel background will never stain very darkly.
3. Pour out staining solution. Cover the gel with 10% acetic acid to destain, shaking
gently ≥2 hr at room temperature until a clear background is obtained.
4. If necessary, pour out 10% acetic acid and add more. Continue destaining until clear
background is obtained. Store gel in 7% acetic acid or water, or in plastic wrap at
4°C.
It is usually unnecessary to add additional destaining solution.
5. If desired, photograph (see Support Protocol 1) or dry (see Basic Protocol 1, step 7)
the gel.
SILVER STAINING
Detection of protein bands in a gel by silver staining depends on binding of silver to
various chemical groups (e.g., sulfhydryl and carboxyl moieties) in proteins. The detection limit is 2 to 5 ng/protein band. In this procedure, proteins separated in a polyacrylamide gel are successively fixed in methanol/acetic acid and glutaraldehyde. After
exposure to silver nitrate, the gel is treated with developer to control the level of staining.
When the desired staining intensity is reached, the gel is fixed, photographed, and dried.
BASIC
PROTOCOL 2
Materials
Polyacrylamide gel (UNIT 10.2)
Fixing solution for Coomassie blue and silver staining (see recipe)
Methanol/acetic acid destaining solution (see recipe)
10% (v/v) glutaraldehyde (freshly prepared from 50% stock; Kodak)
Silver nitrate solution (see recipe)
Developing solution (see recipe)
Kodak Rapid Fix Solution A
Whatman 3MM filter paper (optional)
Gel dryer (optional)
NOTE: Wear gloves at all times to avoid fingerprint contamination.
1. Place a polyacrylamide gel in a plastic container and add 5 gel volumes of fixing
solution. Agitate slowly ≥30 min at room temperature on an orbital shaker.
2. Pour out fixing solution. Fix the gel with 5 gel volumes of methanol/acetic acid
destaining solution for ≥60 min, agitating slowly.
No actual destaining takes place in this step; fixation continues using the same solution
used for destaining in Basic Protocol 1.
3. Pour out destaining solution. Add 5 gel volumes of 10% glutaraldehyde and agitate
slowly 30 min in a fume hood.
CAUTION: Wear gloves and work only in a fume hood.
4. Pour out the glutaraldehyde. Wash the gel at least four times with water, ≥30 min for
each wash and preferably overnight for the last wash. Agitate slowly with each wash.
Analysis of
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Supplement 63
5. Pour out the last wash. Stain the gel with ∼5 gel volumes silver nitrate solution (to
cover the gel) for 15 min with vigorous shaking.
CAUTION: Dispose of ammoniacal silver solution immediately by flushing with copious
amounts of water, as it becomes explosive upon drying.
6. Transfer the gel to another plastic box and wash five times with deionized water,
exactly 1 min for each wash. Agitate slowly with each wash.
7. Dilute 25 ml developing solution with 500 ml water. Transfer gel to another plastic
box, add enough diluted developer to cover the gel during agitation, and shake
vigorously until the bands are as intense as desired. If the developer turns brown,
change to fresh developer.
Development should be stopped immediately when gel background starts to appear.
8. Transfer to Kodak Rapid Fix Solution A for 5 min.
If necessary, swab gel surface with soaked cotton to remove residual silver deposits.
9. Pour off Rapid Fix Solution and wash the gel exhaustively in water (four to five times).
10. Photograph the gel (see Support Protocol 1).
Gels should be photographed as soon as possible because there may be slight changes in
color intensity and increases in nonspecific background. The silver-stained proteins remain
clearly visible for at least 18 hr.
11. Dry the gel to maintain a permanent record (see Basic Protocol 1, step 7) or store in
sealable plastic bag (will last 6 to 12 months).
ALTERNATE
PROTOCOL 2
NONAMMONIACAL SILVER STAINING
This nonammoniacal silver staining procedure uses more stable solutions and detects
certain proteins not stained using the preceding protocol (Morrissey, 1981).
Additional Materials (also see Basic Protocol 2)
5 µg/ml dithiothreitol (DTT; APPENDIX 2)
0.1% (w/v) silver nitrate solution (store in brown bottle at room temperature up to
∼1 month)
Carbonate developing solution (see recipe)
2.3 M citric acid
0.03% (w/v) sodium carbonate (optional)
1. Place a polyacrylamide gel in a glass or polyethylene container and add 100 ml fixing
solution. Agitate slowly 30 min at room temperature on an orbital shaker.
Times are appropriate for all gel sizes.
2. Pour out fixing solution. Immerse gel in methanol/acetic acid destaining solution and
agitate slowly 30 min.
3. Pour out destaining solution. Cover gel with 50 ml 10% glutaraldehyde and agitate
slowly 10 min in a fume hood.
CAUTION: Wear gloves and work only in a fume hood.
Fixing in glutaraldehyde ensures that the gel will not bleach (and will probably last longer)
and is important for retention and detection of very small proteins.
Staining Proteins
in Gels
4. Pour out glutaraldehyde. Wash the gel thoroughly in several changes of water (or in
running water) for 2 hr to ensure low background levels.
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5. Pour out water. Soak the gel in 100 ml 5 µg/ml DTT for 30 min.
Treating proteins with DTT results in more reproducible silver staining.
6. Pour out DTT. Without rinsing, add 100 ml 0.1% silver nitrate solution and agitate
slowly 30 min.
7. Pour out silver nitrate. Wash the gel once quickly with a small amount of water, then
twice rapidly with a small amount of carbonate developing solution.
8. Soak the gel in 100 ml carbonate developing solution and agitate slowly until desired
level of staining is achieved.
9. Stop staining by adding 5 ml 2.3 M citric acid per 100 ml carbonate developing
solution for 10 min and agitate slowly.
Pay attention to the volumes of carbonate and citric acid solutions (steps 8 and 9). These
must be balanced carefully to bring the pH to neutrality. If the pH is high, the reaction will
not stop; if the pH is too low, the gel will bleach.
10. Pour off the solution. Wash the gel several times in water, agitating slowly 30 min.
11. Photograph the gel (see Support Protocol 1) and/or store by soaking 10 min in 0.03%
sodium carbonate and wrapping in plastic wrap or sealing in a heat-sealable bag.
RAPID SILVER STAINING
This protocol (Bloom et al., 1987) is based upon the preceding nonammoniacal silver
staining method. It is rapid and gives low background but may not be quite as sensitive
in detecting very small proteins because there is no glutaraldehyde fixation.
ALTERNATE
PROTOCOL 3
Additional Materials (also see Alternate Protocol 2)
Formaldehyde fixing solution (see recipe)
0.2 g/liter sodium thiosulfate (Na2S2O3)
Thiosulfate developing solution (see recipe)
Drying solution (see recipe)
Dialysis membrane soaked in 50% methanol
Glass plates
1. Place a polyacrylamide gel in a plastic container and add 50 ml formaldehyde fixing
solution. Agitate slowly 10 min at room temperature on an orbital shaker.
Times indicated are flexible and are appropriate for a 0.75-mm × 5.5-cm × 8-cm, 12.5%
acrylamide slab gel. Each gel is placed in an 8 × 14–cm plastic container.
2. Pour out fixing solution. Wash the gel twice with water, 5 min for each wash. Agitate
slowly for each wash.
3. Pour out water. Soak gel 1 min in 50 ml 0.2 g/liter Na2S2O3, agitating slowly.
4. Pour out Na2S2O3. Wash the gel twice with water, 20 sec for each wash.
5. Pour out water. Soak gel 10 min in 50 ml 0.1% silver nitrate solution, agitating slowly.
6. Pour out the silver nitrate. Wash the gel with water and then with a small volume of
thiosulfate developing solution.
7. Soak the gel in 50 ml fresh thiosulfate developing solution and agitate slowly until
band intensities are adequate (∼1 min).
Development continues a little after stopping (next step), so do not overdevelop here.
Analysis of
Proteins
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Supplement 63
8. Add 5 ml of 2.3 M citric acid per 100 ml thiosulfate developing solution and agitate
slowly 10 min.
Pay attention to the volumes of thiosulfate developing and citric acid solutions (see
Alternate Protocol 2, step 9).
9. Pour off the solution. Wash the gel in water, agitating slowly 10 min.
10. Pour off the water. Soak the gel 10 min in 50 ml drying solution.
11. Sandwich gel between two pieces of wet dialysis membrane on a glass plate. Clamp
edges of the plate with notebook clamps and dry overnight at room temperature.
SUPPORT
PROTOCOL 1
GEL PHOTOGRAPHY OF COOMASSIE- OR SILVER-STAINED GELS
Many research facilities have photographic services that are well versed in gel photography and can provide excellent documentation of experiments. The following guidelines
and equipment should be used to achieve optimal results. Any good single-lens reflex
(SLR) camera attached to a copy stand will work. If larger-format cameras are available,
sheet film will give spectacular resolution. The light box must produce relatively even
lighting.
Kodak T-Max 400 film is a fine-grained panchromatic half-tone film that provides
extremely high resolution and that works well for 35-mm gel photography. For contrast
enhancement of Coomassie blue–stained gels, photographing the gels through a deepyellow to yellow-orange filter (Wratten #8 or 9 or a yellow-orange Cokin filter) is
recommended. Silver-stained gels are photographed with a blue-green filter (Wratten #58
or a blue Cokin filter). Develop according to manufacturer’s instructions. Medium-contrast resin-coated paper is recommended for printing.
The instant films from Polaroid are ideal for fast, high-quality photographs of gels for
laboratory notebooks and publications. Typically, type 57 (4 × 5–in., MP4 camera) or type
667 (31⁄4 × 41⁄4–in., DS 34 camera) black and white film is used for documentation at a
shutter speed of 1⁄60 to 1⁄30 sec with an aperture of f16. Type 55 and 665 positive/negative
films provide not only a high-quality print, but also a fine-grain, medium-format negative
that can be used to produce multiple photographs of the gel in any size; for these films,
use a shutter speed of 1⁄4 to 1⁄2 sec and an aperture of f16. Aperture settings of f11 or higher
should be used for sharp photographs. If the picture is too light, increase the shutter speed
or go to a higher f number. If the picture is too dark, decrease shutter speed or go to a
lower f number (adapted from Polaroid Guide to Instant Imaging).
A simple test for the effectiveness of a filter is to place the gel on a light box and observe
the gel through the filter. Increased contrast of the bands should be obvious. Silver staining
can produce bands of various colors, which can present a problem in filter selection.
Kodak X-Omat Duplicating Film is a useful alternative because it yields black banding
in spite of the coloration caused by the silver staining (Anderson, 1988).
BASIC
PROTOCOL 3
Staining Proteins
in Gels
FLUORESCENT STAINING USING SYPRO ORANGE OR RED
Fluorescent dyes have a number of advantages over traditional protein stains. SYPRO
Orange and Red protein gel stains, outlined in the procedure below, can detect 1 to 2 ng
protein/band. This is more sensitive than Coomassie brilliant blue staining and as sensitive
as many silver staining techniques. Staining is straightforward, with less hands-on time
than typical silver staining protocols, and is complete in <1 hr. Stained proteins can be
visualized using a standard 300-nm UV transilluminator or a laser scanner. Although the
protocol below is limited to one-dimensional SDS-PAGE, alternative kits for native,
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isoelectric focusing (IEF), and two-dimensional SDS-PAGE applications are available
from the supplier (Molecular Probes). Coomassie Fluor Orange protein gel stain, a
premixed fluorescent stain, is also available from Molecular Probes.
The staining properties of the two SYPRO dyes are similar, and both are equally suitable
for use in most procedures. The SYPRO Orange gel stain is slightly brighter, whereas the
SYPRO Red gel stain has somewhat lower background fluorescence. For those using a
laser-excited gel scanner, the authors recommend the SYPRO Orange stain for argon-laser-based instruments and the SYPRO Red stain for instruments that employ green He-Ne
or Nd/YAG (neodymium:yttrium-aluminum-garnet) lasers. Both dyes are efficiently
excited by UV or broad-band illumination and, with the proper filters, work nicely with
CCD camera archiving systems.
Materials
SYPRO Orange or Red fluorescent staining solution (see recipe)
7.5% (v/v) acetic acid
0.1% (v/v) Tween-20
Additional reagents and equipment for one-dimensional SDS-PAGE (UNIT 10.2A)
1. Prepare and separate proteins using SDS-PAGE (UNIT 10.2A), but use 0.05% SDS in
the running buffer instead of the usual 0.1% SDS.
Gels run in 0.1% SDS show the same sensitivity for staining as those run with lower SDS
concentrations, but require either more time in staining solution or a 10-min rinse in water
before staining to reduce the background fluorescence that is produced by dye interaction
with SDS. Gels run in 0.05% SDS show no change in the migration of proteins and can be
photographed sooner because they require less time in the staining solution to clear the
SDS from the gel. Gels run in SDS concentrations <0.05% or in old running buffer exhibit
poor resolution of bands and other problems, so it is essential that the SDS stock solution
used to prepare the running buffer be fresh and be at the proper SDS concentration.
Do not fix the proteins in the gel with methanol-containing solutions. Methanol removes
the SDS coat from proteins, strongly reducing the signal from SYPRO Orange or Red stains.
2. Pour SYPRO Orange or Red fluorescent staining solution into a small plastic dish.
For one or two standard-size minigels, use ∼50 ml staining solution. For larger gels,
use between 500 and 750 ml staining solution.
Staining dishes should be cleaned and rinsed well before use, as detergent will interfere
with staining.
Alternatively, use Coomassie Fluor Orange protein gel stain which is available as a
premixed staining solution.
3. Place gel into the staining solution. Cover the container with aluminum foil to protect
the dye from bright light.
The staining solution may be reused up to four times. However, due to reduced sensitivity,
the use of fresh staining solution is recommended.
Several alternate methods of staining can be used. (1) Gels may be stained in sealable
plastic bags. However, it is still important to use the proper amount of staining solution.
(2) For low-percentage gels and for very small proteins, increasing the staining solution
from 7.5% to 10% acetic acid will result in better retention of the protein in the gel without
compromising sensitivity. (3) Protein gel stains can be dissolved 5000-fold into the cathode
(top) running buffer to stain proteins as the gel runs. The dye moves through the gel with
the SDS front, so that all sizes of protein are stained. Staining does not influence relative
migration of proteins through the gel. This method results in poorer protein staining than
the standard poststaining method, and requires the same amount of time because the gel
must be destained for 15 to 40 min in 7.5% acetic acid to reduce background fluorescence.
Analysis of
Proteins
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4. Gently agitate at room temperature for 10 to 60 min.
The staining time depends on the thickness and percentage of the gel. For 1-mm-thick 15%
polyacrylamide gels, the signal is typically optimal after 40 to 60 min staining. Once the
optimal signal is achieved, additional staining time (several hours to overnight) does not
enhance or degrade the signal. Gels can be left in stain for up to a week with only a small
loss in sensitivity; the detection limits under these conditions are ∼2 to 4 ng/band.
5. Rinse briefly (<1 min) with 7.5% acetic acid.
This brief rinse removes excess stain from the gel surface to reduce background fluorescence on the surface of the transilluminator or gel scanner (see Support Protocol 2).
6. Store gel in staining solution, protected from light.
The signal decreases somewhat after several days, but, depending on the amount of protein
in the bands, gels may retain a usable signal for many weeks. Gels may be left in staining
solution overnight without losing sensitivity. However, fixation in acetic acid is relatively
mild, so, for low-percentage gels or very small proteins, photographs should be taken as
soon as possible after staining, before the proteins begin to diffuse.
Gels may be dried between cellophane membrane backing sheets (Bio-Rad), although there
is sometimes a slight decrease in sensitivity. If the gels are dried onto paper, the light will
scatter and the sensitivity will decrease. Other types of plastic sheets are not typically
transparent to UV light. Store dried gels in the dark to prevent photobleaching.
7. Destain gel by incubating overnight in 0.1% Tween-20.
Alternatively, incubating in several changes of 7.5% acetic acid will eventually remove all
of the stain. Incubating in methanol will strip off dye and SDS, but will also precipitate
proteins.
8. Photograph gel (see Support Protocol 2).
ALTERNATE
PROTOCOL 4
FLUORESCENT STAINING ON NONDENATURING GELS USING SYPRO
ORANGE OR RED
Proteins can be stained after native gel electrophoresis (UNIT 10.2B) by dissolving SYPRO
dyes in water and then following Basic Protocol 3. Staining proteins in nondenaturing
gels is highly protein selective and will generally be less sensitive than staining proteins
in SDS gels; however, because there is essentially no background fluorescence, photographic exposures can be very long. If it is not necessary to maintain the protein in a native
state after electrophoresis, the best sensitivity can be achieved if the gel is soaked in 0.05%
SDS for ∼30 min and then stained with a solution of SYPRO dye diluted in 7.5% acetic
acid.
BASIC
PROTOCOL 4
Staining Proteins
in Gels
FLUORESCENT PROTEIN STAIN USING SYPRO RUBY FOR 2-D GEL
ANALYSIS
Fluorescent gel stains are gaining popularity due to the combination of simplicity and
sensitivity that they offer. SYPRO Ruby protein gel stain is an ultrasensitive, fluorescent
stain for the specific detection of proteins separated by polyacrylamide gel electrophoresis
(PAGE). This stain, designed especially for use in 2-D PAGE (Fig. 10.6.1; UNIT 10.4), has
proven to be the most sensitive protein gel stain for standard 1-D SDS-PAGE (UNIT 10.2A)
and an excellent stain for isoelectric focusing (IEF) gels (UNIT 10.3) as well. SYPRO Ruby
protein gel stain achieves comparable sensitivity to that of the silver-staining techniques
(see Basic Protocol 2 and Alternate Protocols 2 and 3) and offers substantial processing
advantages including the absence of overstaining, a linear quantitation range of over three
orders of magnitude, and less protein-to-protein variability. In addition, the SYPRO Ruby
10.6.8
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Figure 10.6.1 Typical two-dimensional gel stained with SYPRO Ruby gel stain.
staining protocol below stains glycoproteins, lipoproteins, calcium-binding proteins,
fibrillar proteins, and other difficult-to-stain proteins, and does not interfere with subsequent analysis of proteins by Edman-based sequencing or mass spectrometry. The stain
can be used with many types of gels, including 2-D gels (UNITS 10.3 & 10.4), Tris-glycine
SDS gels (UNIT 10.2A), and Tris-tricine precast SDS gels and nondenaturing gels (UNIT 10.2B).
SYPRO Ruby stain is also compatible with gels adhering to plastic backings.
Materials
1-D or 2-D polyacrylamide or IEF gels (UNITS 10.2-10.4)
Fixing solution for SYPRO Ruby staining of 2-D polyacrylamide gels (see recipe)
Fixing solution for IEF gels: 40% (v/v) methanol/10% (w/v) trichloroacetic acid in
H2O
SYPRO Ruby protein gel stain (Molecular Probes; see recipe)
10% (v/v) methanol (or ethanol)/7% (v/v) acetic acid
2% (v/v) glycerol
Analysis of
Proteins
10.6.9
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Plastic staining dishes of appropriate size for gels: polypropylene (e.g.,
Rubbermaid Servin’ Savers) or PVC photographic staining trays (e.g.,
Photoquip Cesco-Lite 8-in. × 10-in., for large-format 2-D gels)
Orbital shaker
Additional reagents and equipment for photography of fluorescently stained gels
(see Support Protocol 2)
Fix, stain, and wash the gel
1a. For 1-D polyacrylamide gels: Proceed directly to step 2 (no fixation required).
1b. For 2-D polyacrylamide gels: Fix for 30 min in an appropriate fixing solution for
SYPRO Ruby staining of polyacrylamide gels.
It is also possible to use serial combinations of these fixatives. Note that the combination
of ethanol and acetic acid can result in the formation of ethyl acetate, which is not only
toxic but may interfere with identification of proteins by mass spectrometry.
1c. For IEF gels: Fix for 3 hr in 40% methanol/10% trichloroacetic acid, and then perform
three washes, each for 10 min in distilled water, before proceeding to the staining
step.
2. Transfer the gel to a clean polypropylene or PVC dish of appropriate size. Incubate
the gel in the appropriate amount of undiluted SYPRO Ruby protein gel stain with
continuous gentle agitation, e.g., on an orbital shaker at 50 rpm. For maximum
sensitivity in 1-D or 2-D gels, incubate the gel for at least 3 hr. For IEF gels, incubate
the gel overnight.
The authors have found that polypropylene dishes, such as Rubbermaid Servin’ Savers, are
the optimal containers for staining because the high-density plastic adsorbs only a minimal
amount of the dye. For small gels, circular staining dishes provide the best fluid dynamics
on orbital shakers, resulting in less dye aggregation and better staining. For large-format
2-D gels, polyvinyl chloride (PVC) photographic staining trays, such as Photoquip
Cesco-Lite 8 inch × 10 inch photographic trays also work well. Glass dishes are not
recommended. The minimal staining volumes for typical gel sizes are: 50 ml, for 8 cm ×
10 cm × 0.75–mm gels; 330 ml, for 16 cm × 20 cm × 1–mm gels; 500 ml, for 20 cm × 20
cm × 1–mm gels, or ∼10× the volume of the gel for other gel sizes.
Using too little stain will lower the sensitivity. For convenience, gels may be left in the dye
solution overnight or longer without overstaining. Do not dilute the stain, as diluted stain
will result in decreased sensitivity. Do not reuse the staining solution, as this will result in
a significant loss of sensitivity.
3. To reduce background fluorescence and increase sensitivity, transfer the gel to a clean
staining dish and wash it in 10% methanol (or ethanol)/7% acetic acid for 30 min.
For polyacrylamide gels, perform this wash once; for IEF gels, perform the wash
three times.
This transfer step helps to minimize the deposition of stain speckles on the gel. To reduce
organic waste, stained gels may alternatively be washed in distilled water, although this
method does not reduce background fluorescence to the same extent. The gel may be
monitored periodically using UV epi-illumination to determine the level of background
fluorescence.
4. Optional (for permanent storage): Incubate the gel in a solution of 2% (v/v) glycerol
at room temperature for 30 min. Dry the stained gel using a gel dryer.
Note that proteins present at very low levels may no longer be detectable after gel drying.
Staining Proteins
in Gels
5. View and photograph the gel (see Support Protocol 2).
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PHOTOGRAPHY OF FLUORESCENTLY STAINED GELS
SYPRO Orange or Red Stains
Photographing and archiving the gel is essential to obtain high sensitivity. The camera’s
integrating effect can make bands visible that are not visible to the eye. Place the
fluorescently labeled gel directly on a standard 300-nm UV transilluminator or a bluelight transilluminator (e.g., Clare Chemical Dark Reader). Plastic wraps, such as Saran
Wrap, should not be used, as they fluoresce naturally and will fluoresce even more when
exposed to SYPRO Orange or Red stain. This results in a large background signal, making
it impossible to achieve good sensitivity. Pharmacia PhastGels have a polyester backing
material (Gelbond) that is not only highly autofluorescent, but also binds the SYPRO
Orange and Red protein gel stains, producing additional background fluorescence.
Consequently, the plastic backing should be removed before trying to visualize bands.
Pharmacia markets a gel backing remover for use with their Phast Transfer system. The
surface of the transilluminator should be cleaned with water and a soft cloth after use, to
minimize the buildup of fluorescent dyes.
SUPPORT
PROTOCOL 2
When using a Polaroid camera, use Polaroid 667 black-and-white print film and a SYPRO
protein gel stain photographic filter (Molecular Probes) to obtain the highest sensitivity.
Do not use standard ethidium bromide filters, as they will block much of the light and
lead to lower sensitivity. Supplemental UV blocking filters are not usually required.
Polaroid 667 film is a fast film with an ISO rating of ASA 3000. The use of different film
types may require longer exposure times or different filters. Exposure time will vary with
the intensity of the illumination source; with an f-stop of 4.5, 2 to 5 sec is typical for
SYPRO Orange, and 3 to 8 sec is typical for SYPRO Red. Noticeable photobleaching can
occur after several minutes of exposure to ultraviolet light. If a gel becomes photobleached, it can be restained by simply returning it to the staining solution.
Charge-coupled device (CCD) cameras provide good sensitivity. Contact the camera
manufacturer for the optimal filter sets to use. For those using a laser-excited gel scanner,
the SYPRO Orange stain is recommended for argon laser–based instruments, and the
SYPRO Red stain is recommended for instruments that employ green He-Ne or Nd/YAG
lasers.
SYPRO Ruby Stain
The SYPRO Ruby protein gel stain has two excitation maxima, one at ∼280 nm and one
at ∼450 nm, and has an emission maximum near 610 nm. Proteins stained with the dye
can be visualized using a 300-nm UV transilluminator, a blue-light transilluminator, or a
laser scanner. The stain has exceptional photostability, allowing long exposure times for
maximum sensitivity.
UV or blue-light transilluminator
Proteins stained with SYPRO Ruby protein gel stain are readily visualized using a UV
or blue-light source. The use of a photographic camera or CCD camera and the appropriate
filters is essential to obtain the greatest sensitivity. The instrument’s integrating capability
can make bands visible that cannot be detected by eye. It is important to clean the surface
of the transilluminator after each use with deionized water and a soft cloth (like cheesecloth); otherwise fluorescent dyes such as SYPRO stains, SYBR stains, and ethidium
bromide will accumulate on the glass surface and cause a high background fluorescence.
The authors use a 300-nm transilluminator with six 15-W bulbs. Excitation with different
light sources may not give the same sensitivity. Using a Polaroid camera and Polaroid
667 black-and-white print film, the highest sensitivity is achieved with a 490-nm longpass filter, such as the SYPRO protein gel stain photographic filter (S-6656), available
Analysis of
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from Molecular Probes. The authors typically photograph minigels using an f-stop of 4.5
for 1 sec. Using a CCD camera, images are best obtained by digitizing at ∼1024 × 1024
pixels resolution with 12- or 16-bit grayscale levels per pixel. The camera manufacturer
should be consulted for recommendations on filter sets to use. A CCD camera–based
image analysis system can gather quantitative information that will allow comparison of
fluorescence intensities between different bands or spots. Using such a system, the authors
have found that the SYPRO Ruby gel stain has a linear dynamic range over three orders
of magnitude. The polyester backing on some premade gels is highly fluorescent. For
maximum sensitivity using a UV transilluminator, the gel should be placed polyacrylamide-side-down and an emission filter, such as the SYPRO protein gel stain photographic filter, should be used to screen out the blue fluorescence of the plastic. The use
of a blue-light transilluminator (UVP Visi-Blue Transilluminators) or laser scanner will
reduce the amount of fluorescence from the plastic backing so that the gel may be placed
polyesterside down.
Laser-scanning instruments
Gels stained with the SYPRO Ruby protein gel stain can be visualized using imaging
systems equipped with lasers that emit at 450, 473, 488, or 532 nm. For the analysis of
stained proteins, note that SYPRO Ruby dye sometimes generates small speckles of
precipitated dye on the gel. The speckles have diameters ∼20% the size of the smallest
stained protein spot, making them very easy to distinguish. Analysis software for 2-D gels
will ignore small speckles if the minimum spot size of the program is set appropriately
(determined by trial and error).
For the identification of individual protein spots, it should be noted that SYPRO Ruby
protein gel stain does not bind covalently to proteins. Edman-based sequencing or mass
spectrometry data can be obtained after staining, with no interference from the stain.
Accurate mass spectrometry has been performed on a spot containing as little as 75 fmol
of stained protein.
ALTERNATE
PROTOCOL 5
FLUORESCENT PHOSPHOPROTEIN GEL STAINING FOR SELECTIVELY
STAINING PHOSPHOPROTEINS IN POLYACRYLAMIDE GELS
Fluorescent phosphoprotein gel staining provides a method for selectively staining
phosphoproteins in polyacrylamide gels. It is ideal for identification of kinase targets in
signal transduction pathways (see Chapter 18) and for phosphoproteomic studies. This
fluorescent stain allows direct, in-gel detection of phosphate groups attached to tyrosine,
serine, or threonine residues. The stain can be used with standard SDS-polyacrylamide
gels or with 2-D gels. Blotting is not required, and there is no need for phosphoproteinspecific antibodies or immunoblot detection reagents. The protocol delivers results in 4
to 5 hr. The stain is also compatible with mass spectrometry, allowing analysis of the
phosphorylation state of entire proteomes with detection of as little as 1 to 16 ng of
phosphoprotein per band. For individual phosphoproteins, the strength of the signal
correlates with the number of phosphate groups and is linear over three orders of
magnitude. The fluorescent stain with its ∼555/580 nm excitation/emission maxima can
be detected by use of a visible-light scanning instrument, a visible-light transilluminator,
or a 300-nm transilluminator.
Staining Proteins
in Gels
Materials
Protein-containing sample of interest
Methanol, spectroscopy grade
Chloroform, spectroscopy grade
Appropriate 1× sample buffer for electrophoresis (UNITS 10.2-10.4)
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Table 10.6.1 Examples of Commercially Available Phosphorylated and
Nonphosphorylated Proteins for Use as Controls
Mol. wt.
(Da)
Number of
phosphate
residues
Lower limit of
detection
Riboflavin-binding protein
α-Casein
29,200
23,600
8
8
1-3 ng
1-2 ng
β-Casein
Ovalbumin
24,500
45,000
5
2
1-2 ng
4-8 ng
Pepsin
Carbonic anhydrase
35,500
30,000
1
0
8-16 ng
Not applicable
Bovine serum albumin (BSA)
66,000
0
Not applicable
Protein
Phosphoprotein standards (PeppermintStick Phosphoprotein Molecular Weight
Standards, Molecular Probes; also see Table 10.6.1)
Fixing solution for phosphoprotein gels (see recipe)
Pro-Q Diamond phosphoprotein gel stain (Molecular Probes; see recipe)
Phosphoprotein gel destain solution (see recipe, or purchase from Molecular
probes)
Polystyrene staining dish (e.g., a weighing dish)
Orbital shaker
Additional reagents and equipment for polyacrylamide gel electrophoresis (UNITS
10.2-10.4), SYPRO Ruby protein gel staining (see Alternate Protocol 5),
Coomassie blue stain (see Basic Protocol 1), silver staining (see Basic Protocol
2), and imaging and documenting phosphoprotein-stained gels
Prepare samples by desalting and delipidating
1. Place a 150-µl sample containing ∼150 to 300 µg of protein in a 1.5-ml microcentrifuge tube. Add 600 µl of methanol and mix well by vortexing, then add 150 µl of
chloroform and mix well by vortexing. Finally, add 450 µl of distilled water and mix
well by vortexing.
A delipidated and desalted sample is essential for adequate separation of the proteins by
electrophoresis and subsequent staining by Pro-Q Diamond phosphoprotein gel stain.
2. Microcentrifuge 5 min at ~12,000 rpm, room temperature. Discard the upper phase,
keeping the white precipitation disc that forms between the upper and lower phases.
Add 450 µl of methanol and mix well by vortexing.
3. Microcentrifuge 5 min at ~12,000 rpm, room temperature. Discard the supernatant
and dry the pellet in a Speedvac evaporator for 10 min. Resuspend the pellet in
standard 1× sample buffer for electrophoresis.
Separate by electrophoresis
4. Separate the proteins using standard polyacrylamide electrophoresis techniques
(UNITS 10.2-10.4) along with phosphoprotein standards. To ensure detection of less
abundant phosphoproteins, use approximately the same mass of protein that would
be used for a typical Coomassie blue dye–stained gel (see Basic Protocol 1).
Use known phosphorylated and nonphosphorylated proteins as controls to help verify the
phosphorylation status of the unknown protein. Table 10.6.1 lists several commonly
available proteins that can serve as positive and negative controls
Analysis of
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Fix the gel
5a. For minigels: Transfer gel to a staining dish (plastic container or large plastic
weighing dish). Immerse the gel in ∼100 ml of fixing solution for phosphoprotein
minigels (containing acetic acid) and incubate at room temperature with gentle
agitation, e.g., on an orbital shaker at 50 rpm, for at least 30 min. If needed, repeat
the fixation step once more to ensure that all of the SDS is washed out of the gel. If
desired, leave gels in the fixing solution overnight.
If reusing a plastic container, clean thoroughly and rinse it with 70% ethanol.
Adhere strictly to the volumes and times specified in this protocol for fixation, washing,
staining, and destaining. Replicating the protocol is essential for consistent gel-to-gel and
day-to-day comparisons.
5b. For larger 2-D gels: Transfer gel to a staining dish (plastic container or large plastic
weighing dish). Immerse the gel in ∼500 ml fix solution for phosphoprotein 2-D gels
(containing trichloroacetic acid) and incubate at room temperature overnight with
gentle agitation, e.g., on an orbital shaker at 50 rpm. Perform a second fixation, for
1 hr, to ensure that all of the SDS is washed out of the gel.
In both of the above fixation steps, the second fixation is especially important if non-electrophoresis-grade SDS has been used.
Wash the gel
6a. For minigels: Incubate the gel in ∼100 ml water with gentle agitation for 10 min.
Repeat this step for a total of two washes.
6b. For larger 2-D gels: Incubate the gel in ∼500 ml water with gentle agitation for 15
min. Repeat this step three times for a total of four washes.
In both of the above washing steps, it is important that the gel be completely immersed in
the water in order to remove all of the methanol and acetic acid from the gel. Residual
methanol or acetic acid will interfere with Pro-Q Diamond phosphoprotein staining.
Stain the gel
7a. For minigels: Incubate the gel in the dark in 50 ml of Pro-Q Diamond phosphoprotein
gel stain with gentle agitation for 75 to 120 min.
7b. For larger 2-D gels: Incubate the gel in the dark in 500 ml of Pro-Q Diamond
phosphoprotein gel stain with gentle agitation for 3 to 4 hr.
In both of the above staining steps, if directly comparing multiple gels, it is important that
the incubation time be the same for each gel. Under no circumstances should the gel be
stained overnight, as this will result in higher background staining.
For selective detection of phosphotyrosine residues, see Support Protocol 5.
Destain the gel
8a. For minigels: Incubate the gel in the dark in 80 ml of phosphoprotein gel destain
solution at room temperature for a total of ∼3 hr with two changes of destain solution
(e.g., three incubations of 60 min each).
The first destaining step may be as short as 45 min; the last destaining step may be as long
as overnight.
8b. For larger 2-D gels: Incubate the gel in the dark in 500 ml of phosphoprotein gel
destain solution at room temperature for a total of ∼4 hr with three changes of destain
solution (e.g., four incubations of 60 min each).
Staining Proteins
in Gels
If desired, the last destaining step may be overnight.
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Image and document the gel
9. Image and document the phosphoprotein-stained gel using the appropriate instrumentation (see Support Protocol 3).
The phosphoprotein staining pattern must be viewed and documented before proceeding
with total-protein staining (steps 10 a or b), because the stain will be washed away during
the staining procedure for total protein.
Stain the gel for total protein
10a. For minigels: After obtaining results with the Pro-Q Diamond phosphoprotein gel
stain, stain the gel with a quantitative total-protein stain, such as SYPRO Ruby
protein gel stain (see Alternate Protocol 5), to ascertain the relative phosphorylation
state of proteins.
In this way, an abundant nonphosphorylated protein that exhibits low nonspecific staining
with Pro-Q Diamond stain can be distinguished from a less abundant highly phosphorylated protein. Note that nonquantitative total-protein stains, such as silver stains, are much
less useful in this application.
10b. For larger 2-D gels: After staining with Pro-Q Diamond stain, treat the gel with a
total-protein stain, such as SYPRO Ruby protein gel stain (see Alternate Protocol 5),
Coomassie blue stain (see Basic Protocol 1), or a silver stain (see Basic Protocol 2).
A quantitative stain such as Coomassie blue or SYPRO Ruby protein gel stain will be the
most useful, because it will aid in determining the relative phosphorylation state of a given
protein. Furthermore, for 2-D gels, total protein staining makes it easier to localize a
protein to a particular spot within a complex protein pattern.
IMAGING AND DOCUMENTING THE PHOSPHOPROTEIN-STAINED GEL
The phosphoprotein stain used in Alternate Protocol 5 has an excitation maximum at ∼555
nm and an emission maximum at ∼580 nm. Imaging instruments with light sources and
filters that match the excitation and emission maxima will result in the highest sensitivity.
SUPPORT
PROTOCOL 3
Visible Light–Based Scanners
Stained gels are best visualized using excitation at 532 to 560 nm, such as that provided
with a visible light laser–based or xenon arc lamp–based gel-scanning instrument. For
most instruments, a ∼580 nm long-pass or a ∼600 nm band-pass emission filter is
recommended.
Transillumination
Stained gels can be visualized on a blue-light transilluminator, such as the Visi-Blue series
of transilluminators (UVP), Dark Reader transilluminator (Clare Chemical Research), or
on a 300-nm UV transilluminator; however, the sensitivity will be lower than with a
scanning instrument. Images can be documented using either conventional or digital
photography. With a Polaroid camera and Polaroid 667 black-and-white film, use an
appropriate long-pass filter, such as the SYPRO photographic filter (S-6656), and exposure times of ∼15 to 30 sec. The red-orange filters typically used to photograph gels stained
with ethidium bromide will not work well. For digital cameras, use a filter that corresponds closely to the emission characteristics of the stain, such as a 600 nm band-pass
filter.
Analysis of
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SUPPORT
PROTOCOL 4
SELECTIVE DETECTION OF PHOSPHOTYROSINE RESIDUES
Phosphotyrosine residues can be selectively detected by removing phosphoserine and
phosphothreonine residues through a β-elimination reaction. To detect only phosphotyrosine residues, one must perform the β-elimination reaction before carrying out Pro-Q
Diamond staining as in Alternate Protocol 5. To obtain data on all phosphorylation sites,
stain with Pro-Q Diamond as in Alternate Protocol 5, then document the gel image. Next,
perform the β-elimination reaction and stain the gel with Pro-Q Diamond stain again.
All spots or bands appearing in the first staining but not appearing in the second staining
will be due to phosphoserine or phosphothreonine residues.
Additional Materials (also see Alternate Protocol 5)
Gel for phosphoprotein fluorescent staining (either before fixation/staining or after
fixation/staining/destaining; see Alternate Protocol 5)
Barium hydroxide octahydrate
Argon source
Glacial acetic acid
50°C shaking water bath
1. If the gel has not already been fixed and stained, perform fix and wash steps (see
Alternate Protocol 5, steps 5a or b and 6a or b). If the gel has already been fixed,
stained, and destained, then perform only the wash procedure (see Alternate Protocol
5, step 6a or b).
2. Prepare a saturated solution of barium hydroxide by dissolving 12.6 g barium
hydroxide octahydrate in 40 ml of degassed distilled water. Mix for 15 to 20 min and
centrifuge 10 min at 10,000 × g, room temperature, to pellet any insoluble barium
hydroxide. Store under argon gas.
3. Incubate 40 ml of the saturated barium hydroxide solution in a 50°C water bath for
30 min. At the same time, warm 40 ml of degassed distilled water to 50°C in the water
bath.
All solutions should be treated with argon gas to remove atmospheric carbon dioxide and
prevent the formation of insoluble barium carbonate.
4. Mix 40 ml of the warmed barium hydroxide solution with 40 ml of the warmed
degassed water; and incubate the gel in the diluted solution at 50°C for 30 min with
gentle agitation.
5. Stop the reaction by lowering the pH to 4.0 with addition of ∼6 ml glacial acetic acid.
6. Wash and stain the gel (see Alternate Protocol 6, steps 6a or b and 7 a or b).
7. Document the results (see Support Protocol 3).
ALTERNATE
PROTOCOL 6
Staining Proteins
in Gels
FLUORESCENT STAINING FOR DIFFERENTIALLY STAINING
GLYCOSYLATED AND NONGLYCOSYLATED PROTEINS IN THE SAME GEL
Fluorescent staining provides a powerful method for differentially staining glycosylated
and nonglycosylated proteins in the same gel. The technique combines a highly sensitive
glycoprotein stain with ultrasensitive SYPRO Ruby protein gel stain. Both stains provide
simple, sensitive, and robust detection. The Pro-Q Emerald 300 glycoprotein stain reacts
with periodate-oxidized carbohydrate groups, creating a bright green fluorescent signal
on glycoproteins. Using this stain, it is possible to detect as little as 0.5 ng of glycoprotein
per band, depending upon the nature and the degree of glycosylation, making it about
50-fold more sensitive than the standard periodic acid–Schiff base method using acidic
fuchsin dye. The green fluorescent signal can be visualized with 300-nm UV illumination.
The second staining uses SYPRO Ruby protein gel stain (see Basic Protocol 4) to detect
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total protein. This easy-to-use fluorescent stain provides the same sensitivity as silver
staining, but has the advantage that it does not require glutaraldehyde, which can produce
false positive responses when glycoproteins are stained. The use of SYPRO Ruby stain
makes it possible to detect contaminating proteins and to easily compare the sample with
molecular weight standards. For 2-D gels, total-protein staining makes it easier to localize
a protein to a particular spot in the complex protein pattern. Proteins show orange-fluorescent staining when illuminated with a 300-nm UV transilluminator or a laser scanner
with a 473-nm, 488-nm, or 532-nm light source. Glycosylated molecular weight standards
are also highlighted containing a mixture of glycosylated and nonglycosylated proteins,
which, when separated by electrophoresis, provide alternating positive and negative
controls.
Materials
Protein-containing sample of interest
8-cm × 10-cm SDS- polyacrylamide minigel (UNIT 10.2A)
Sample buffer (UNIT 10.2A)
Pro-Q Emerald glycoprotein gel staining kit (Molecular Probes) including:
Pro-Q Emerald 300 staining reagent (component A), 50× concentrate in DMF
(store at −20°C up to 6 months, protected from light)
Pro-Q Emerald 300 staining buffer (component B; store at room temperature up
to 6 months)
Oxidizing reagent (component C): 2.5 g of periodic acid (add 250 ml of 3% v/v
acetic acid and store at room temperature up to 6 months)
CandyCane glycoprotein molecular weight standards (store at −20°C up to 6
months)
Fixing solution: 50% (v/v) methanol in H2O
Wash solution: 3% (v/v) glacial acetic acid in H2O
Polystyrene staining dish (e.g., large weighing dish)
Orbital shaker
Additional reagents and equipment for SDS-PAGE (UNIT 10.2A), viewing and
documenting glycoprotein-stained gels (see Support Protocol 5), and SYPRO
Ruby staining (see Basic Protocol 4)
NOTE: The following procedure is optimized for staining 0.5- to 0.75-mm thick, 8-cm ×
10-cm minigels. Large 2-D gels (20 cm × 20 cm) require much larger volumes and longer
fixation and staining times, as indicated in the annotations to the respective steps.
1. Dilute protein sample to ∼10 to 100 µg/ml with sample buffer and load 5 to 10 µl of
the diluted sample per lane of an 8-cm × 10-cm polyacrylamide gel. Also dilute 0.5
µl of the CandyCane standard mixture (from the Pro-Q Emerald glycoprotein gel
staining kit) with 7.5 µl of sample buffer, vortex, and load in a lane of the gel. Perform
standard SDS-PAGE as described in UNIT 10.2A.
In the standard lane there will be ∼250 ng of each standard protein included in the
CandyCane standard mix; for larger gels increase the amount of standard and buffer used.
Larger gels typically require twice as much material for sample and standards.
2. Transfer gel to a polystyrene staining dish and immerse the gel in 100 ml of fixing
solution (50% methanol) and incubate at room temperature with gentle agitation (e.g.,
on an orbital shaker at 50 rpm) for 45 min. Repeat this wash step to ensure that all of
the SDS is washed out of the gel.
For large (20 × 20–cm) 2-D gels, use 700 ml of fixing solution and incubate at room
temperature overnight.
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3. Wash the gel by incubating in 50 ml of wash solution (3% acetic acid) with gentle
agitation for 10 min. Repeat this step once.
Use 700 ml of wash solution for large 2-D gels.
4. Oxidize the carbohydrates by incubating the gel in 25 ml of oxidizing solution
(component C in the Pro-Q Emerald 300 kit; 2.5 g periodic acid in 250 ml of 3%
acetic acid) with gentle agitation for 30 min.
Large 2-D gels require 500 ml of oxidizing solution and should be incubated for 1 hr. The
250-ml volume of oxidizing solution can be diluted with 250 ml of 3% acetic acid in order
to have an adequate volume for large 2-D gels.
The Pro-Q Emerald glycoprotein gel staining kit provides sufficient materials to stain ten
8-cm × 10-cm, 0.5 to 0.75–mm thick gels.
5. Wash the gel by incubating in 50 ml (700 ml for large 2-D gels) of wash solution (3%
acetic acid) with gentle agitation for 5 to 10 min. Repeat this step two more times for
a total of three washes.
For large 2-D gels, 700 ml wash solution should be used for each wash and three additional
washes (for a total of four washes) should be performed.
6. Just before use, dilute the 50× concentrate of Pro-Q Emerald 300 staining reagent
(component A in the kit) 50-fold into Pro-Q Emerald 300 staining buffer (component
B in the kit); e.g., dilute 500 µl of component A into 25 ml component B. Stain the
gel by placing it in the dark in 25 ml of the mixed staining solution while gently
agitating for 90 to 120 min.
The signal can be seen after about 20 min and maximum sensitivity is reached at about
120 min. The authors do not recommend staining overnight. A 200-ml volume of staining
solution and staining period of 2.5 hr are required for large 2-D gels.
7. Wash the gel in 50 ml wash solution (3% acetic acid) at room temperature for 15 min.
Repeat this wash once for a total of two washes. Do not leave the gel in wash solution
>2 hr, as the staining will start to decrease.
Use 700 ml of wash solution for large 2-D gels.
8. View and photograph the gel to document the staining pattern (see Support Protocol 5).
For best results, the Pro-Q Emerald 300 stain should be used first and the glycoprotein
staining pattern documented before proceeding with SYPRO Ruby staining. After SYPRO
Ruby staining, the fluorescent signal from the Pro-Q Emerald 300 glycoprotein stain can
still be seen, but the sensitivity will be somewhat decreased.
9. Stain the gel with SYPRO Ruby (see Basic Protocol 4).
10. View and photograph the gel (see Support Protocol 5).
SUPPORT
PROTOCOL 5
Staining Proteins
in Gels
VIEWING AND PHOTOGRAPHING THE GLYCOPROTEIN-STAINED GEL
The green-fluorescent Pro-Q Emerald 300 staining should be viewed and documented
before staining total proteins with SYPRO Ruby protein gel stain. The Pro-Q Emerald
300 stain has an excitation maximum at ∼280 nm and an emission maximum near 530
nm. Stained glycoproteins can be visualized using a 300-nm UV transilluminator (UVP).
The use of a photographic camera or CCD camera and the appropriate filters is essential
to obtain the greatest sensitivity. The instrument’s integrating capability can make bands
visible that cannot be detected by eye. It is important to clean the surface of the
transilluminator after each use with deionized water and a soft cloth (e.g., cheesecloth);
otherwise fluorescent dyes can accumulate on the glass surface and cause a high back-
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ground fluorescence. Some fluorescent speckling may occur, especially near the edges of
the gel. This speckling is an intrinsic property of the stain and does not affect sensitivity.
When analyzing amounts of glycoprotein near the limit of detection, the authors advise
that samples be run in the middle lanes of the gel. The authors use a 300-nm transilluminator with six 15-W bulbs. Excitation with different light sources may not give the same
sensitivity. Using a Polaroid camera and Polaroid 667 black-and-white print film, the
highest sensitivity is achieved with a 490-nm long-pass filter, such as the SYPRO protein
gel stain photographic filter, available from Molecular Probes. The authors typically
photograph minigels using an f-stop of 4.5 for 2 to 4 sec, using multiple 1-sec exposures.
Using a CCD camera, images are best obtained by digitizing at ∼1024 × 1024 pixels
resolution with 12-, 14-, or 16-bit grayscale levels per pixel. The camera manufacturer
should be consulted for recommendations on filters to use. A CCD camera–based
image-analysis system can gather quantitative information that will allow comparison of
fluorescence intensities between different bands or spots. The polyester backing on some
precast gels is highly fluorescent. For maximum sensitivity using a UV transilluminator,
the gel should be placed polyacrylamide-side-down and an emission filter should be used
to screen out the blue fluorescence of the plastic.
SYPRO Ruby staining for total protein is described in Basic Protocol 4. Viewing and
photographing SYPRO Ruby–stained protein gels is described in Support Protocol 2. One
should keep in mind that SYPRO Ruby protein gel stain has two excitation peaks and can
be viewed using either UV illumination or blue-light illumination with a laser scanner.
With UV illumination, both stains can be visualized simultaneously, although the signal
from the green fluorescent glycoprotein stain may be somewhat reduced, compared to
what it was before SYPRO Ruby staining. For documentation, the orange-red fluorescent
SYPRO Ruby staining can be separated from the green-fluorescent Pro-Q Emerald 300
staining in one of two ways. If using UV illumination, use either a long-pass filter with
a cutoff between 620 and 650 nm, or a band-pass filter with a center wavelength at about
645 nm, to document the SYPRO Ruby stain alone. Filters with cutoffs at wavelengths
shorter than 620 nm may show some bleed-through of the Pro-Q Emerald 300 signal.
Alternatively, the gel can be imaged using visible-light excitation, such as used in a laser
scanner. Visible light will excite the SYPRO Ruby stain, but not the Pro-Q Emerald 300
stain. The fluorescent signal from the SYPRO Ruby stain can then be documented as
described.
REAGENTS AND SOLUTIONS
Use high-quality deionized, distilled water (≥18 MΩ) in all recipes and protocol steps. For common stock
solutions, see APPENDIX 2; for suppliers, see APPENDIX 4.
Carbonate developing solution
0.5 ml 37% formaldehyde per liter solution
3% (w/v) sodium carbonate
Prepare fresh before use
Coomassie blue staining solution
50% (v/v) methanol
0.05% (w/v) Coomassie brilliant blue R-250 (Bio-Rad or Pierce)
10% (v/v) acetic acid
40% H2O
Dissolve Coomassie brilliant blue R in methanol before adding acetic acid and water.
Store for up to 6 months at room temperature. If precipitate is observed following
prolonged storage, filter to obtain a homogeneous solution.
Analysis of
Proteins
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Developing solution
0.5 g sodium citrate
0.5 ml 37% formaldehyde solution (Kodak)
H2O to 100 ml
Store up to ∼1 month at room temperature
Drying solution
10% (v/v) ethanol
4% (v/v) glycerol
86% H2O
Store up to ∼1 month at room temperature
Fixing solution for Coomassie blue and silver staining
50% (v/v) methanol
10% (v/v) acetic acid
40% H2O
Store up to ∼1 month at room temperature
Fixing solution for phosphoprotein gels
For minigels:
50% (v/v) methanol
10% (v/v) acetic acid
For 2-D gels:
50% (v/v) methanol
10% (w/v) trichloroacetic acid
One 6-cm × 9-cm × 0.75-mm minigel will require ∼200 ml of fix solution; one 20-cm × 20-cm
× 1-mm 2-D gel will require ∼1 liter of fix solution. In order to improve the specificity of
phosphoprotein staining in 2-D gels, the authors recommend fixing them in 10% trichloroacetic acid/50% methanol instead of 10% acetic acid/50% methanol.
Store fixing solutions up to 1 month at room temperature
Fixing solution for SYPRO Ruby staining of 2-D polyacrylamide gels
Fix with any of the following solutions (all prepared in H2O):
10% (v/v) methanol
7% (v/v) acetic acid/25% (v/v) ethanol
12.5% (w/v) trichloroacetic acid/10% (v/v) ethanol
7% (v/v) acetic acid/50% (v/v) ethanol
3% (v/v) acetic acid/40% (v/v) ethanol
10% (v/v) acetic acid
Store up to 1 month at room temperature
These fixative solutions are used in 5- to 10-fold excess of the gel volume. Thin (<0.75-mm)
gels will equilibrate with fixative much more quickly than thick gels. Larger-format gels
require more fixative to effectively exchange the gel buffers for the fixative.
Formaldehyde fixing solution
40% (v/v) methanol
0.5 ml 37% formaldehyde per liter solution
60% H2O
Store up to ∼1 month at room temperature
Staining Proteins
in Gels
Isopropanol fixing solution
25% (v/v) isopropanol
10% (v/v) acetic acid
65% H2O
Store indefinitely at room temperature
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Methanol/acetic acid destaining solution
5% (v/v) methanol
7% (v/v) acetic acid
88% H2O
Store up to ∼1 month at room temperature
Phosphoprotein gel destain solution
15% (v/v) 1,2-propanediol (propylene glycol)
50 mM sodium acetate, pH 4.0
For each liter of destain solution to be prepared, add 50 ml of 1 M sodium acetate,
pH 4.0, to 800 ml of distilled water. Add 150 ml of 1,2-propanediol (or 40 ml of
acetonitrile; see below). Bring the volume to 1 liter with distilled water and mix
thoroughly. Alternatively, this destain Solution is available from Molecular Probes
as a separate product. Store up to 1 month at room temperature.
If 1,2-propanediol (propylene glycol) is not available, 4% acetonitrile may be substituted.
However, note that acetonitrile is a hazardous compound and its use will involve waste
disposal restrictions.
The destaining step is very important for maximizing detection sensitivity for phosphoproteins while minimizing nonspecific staining of nonphosphorylated proteins. One 6-cm × 9-cm
× 0.75-mm minigel will require ∼250 ml of destain solution; one 20-cm × 20-cm × 1-mm 2-D
gel will require ∼2 liters of destain solution.
Pro-Q Diamond phosphoprotein gel stain
Purchase Pro-Q Diamond phosphoprotein gel stain from Molecular Probes (available in 200-ml, 1-liter, and 5-liter quantities). 200 ml provides sufficient material to
stain ∼4 minigels; 1 liter provides sufficient material to stain ∼20 minigels or two
large-format 2-D gels; 5 liters provide sufficient material to stain ∼100 minigels or
10 large-format gels. Upon receipt, store the stain at room temperature, protected
from light. For long-term storage, store the stain at 2° to 6°C, protected from light.
When stored properly, the stain should be stable for at least 6 months.
Rapid Coomassie blue staining solution
10% (v/v) acetic acid
0.006% (w/v) Coomassie brilliant blue G-250 (Bio-Rad)
90% H2O
Store indefinitely at room temperature
Silver nitrate solution (ammoniacal)
Add 3.5 ml concentrated NH4OH (∼30%) to 42 ml 0.36% NaOH and bring the
volume to 200 ml with H2O. Mix with a magnetic stirrer and slowly add 8 ml of
19.4% (1.6 g/8 ml) silver nitrate. Use within 20 min.
If the solution is cloudy, carefully add NH4OH until it clears. Alternatively, use NH4OH that
is <3 months old.
CAUTION: This solution is potentially explosive when dry and therefore should be precipitated by the addition of an equal volume of 1 M HCl. The resultant silver chloride can be
washed down a drain with a large volume of cold water.
SYPRO Orange or Red fluorescent staining solution
Allow stock vial of SYPRO Orange or Red protein gel stain (Molecular Probes) to
warm to room temperature and then briefly microcentrifuge to deposit the dimethyl
sulfoxide (DMSO) solution at the bottom of the vial. If particles of dye are present,
dissolve by briefly sonicating the tube or vortexing it vigorously after warming.
Dilute stock 1:5000 (v/v) in 7.5% (v/v) acetic acid and mix vigorously. Store in very
continued
Analysis of
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clean, detergent-free glass or plastic bottles, protected from light, at 4°C (stable ≥3
months).
SYPRO Orange: 300 and 470 nm excitation, 570 nm emission; SYPRO Red: 300 and 550
nm excitation, 630 nm emission.
The stock solutions should be stored protected from light at room temperature, 4°C, or
20°C.When stored properly, they are stable for 6 months to 1 year.
SYPRO Ruby protein gel stain
Purchase SYPRO Ruby protein gel stain from Molecular Probes (available in
200-ml, 1-liter, and 5-liter quantities; 200 ml provides sufficient material to stain ∼4
minigels). Store at room temperature protected from light (stable for at least 9
months). For convenient storage and dispensing, the 5-liter unit size is packaged in
a cubical box with a spigot. Once opened, the box can be stored on its side with the
top flap closed to protect the stain from light.
Thiosulfate developing solution
3% (w/v) sodium carbonate
0.0004% (w/v) sodium thiosulfate
0.5 ml 37% formaldehyde per liter solution (add immediately before use)
Store indefinitely without formaldehyde at room temperature
COMMENTARY
Background Information
Staining Proteins
in Gels
Coomassie brilliant blue (Basic Protocol 1)
binds nonspecifically to proteins (Wilson,
1983). Because the dye does not bind to the
polyacrylamide gel, proteins will be detected
as blue bands surrounded by clear gel zones.
Silver staining relies on differential reduction of silver ions, which is the basis for photographic processes. A highly sensitive photochemical silver staining technique (Switzer et
al., 1979; Merril et al., 1984) permits the detection of polypeptides in gels at more than 100×
lower concentrations than Coomassie brilliant
blue (i.e., femtomole levels of protein). The
basic silver staining protocol described here
(Basic Protocol 2) is derived from a modified
technique developed by Oakley et al. (1980),
which is simpler and less expensive than the
original procedures. The first alternate silver
staining protocol presents a very popular
method described by Morrissey (1981). The
second alternate silver staining protocol is a
very rapid method described by Bloom et al.
(1987).
Fluorescent protein gel stains provide a
number of advantages over conventional colorimetric stains. The SYPRO Orange and Red
protein gel stains described here can detect 1 to
2 ng protein per minigel band, more sensitive
than Coomassie brilliant blue staining and as
sensitive as many silver staining techniques. In
addition, staining is complete in <1 hr. After
electrophoresis, the gel is simply stained,
rinsed, and photographed; no separate fixation
or destaining step is required and there is no
fear of overstaining the gel (Steinberg et al.,
1996a,b, 1997). In addition, stained proteins
can be visualized using a standard 300-nm UV
transilluminator or a laser scanner (Fig. 10.6.2).
Because the dyes interact with the SDS coat
around proteins in the gel, they give more consistent staining between different types of proteins compared to Coomassie or silver staining
and do not exhibit negative staining. Furthermore, the dyes detect a variety of proteins down
to ∼6500 Da without staining nucleic acid or
lipopolysaccharide contaminants that are
sometimes found in protein preparations derived from cell or tissue extracts.
Critical Parameters
The high sensitivity of the silver staining
technique renders it susceptible to impurities
and staining artifacts. It is mandatory that the
polyacrylamide gels and all staining solutions
be prepared from high-quality reagents in order
to avoid staining artifacts. Especially important
is the use of high-quality water (glass-distilled
or deionized, carbon-filtered). The glassware
used for gel polymerization and plastic containers should be cleaned thoroughly, and gels
should be handled with vinyl, powder-free
gloves. To avoid uneven staining of the gel
surface, the polyacrylamide gel should be covered with a sheet of Parafilm in order to uniformly wet the gel surface during staining, and
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A
B
C
D
Figure 10.6.2 Identical polyacrylamide minigels stained with (A) SYPRO Orange gel stain, (B)
SYPRO Red gel stain, (C) silver stain, and (D) Coomassie brilliant blue stain according to standard
protocols. The SYPRO-stained gels were photographed using 300-nm transillumination, a SYPRO
Orange/Red protein gel stain photographic filter, and Polaroid 667 black-and-white print film. The
Coomassie- and silver-stained gels were photographed using transmitted white light and Polaroid
667 black-and-white print film; no optical filter was used.
should be touched only very gently with gloved
hands. If silver staining is performed infrequently, commercial silver staining kits should
be used; those distributed by Bio-Rad and
Pierce have been tested and found to be reliable
and sensitive.
For immunoblotting and other blotting techniques, fluorescent stains can be diluted in
standard transfer buffer. However, staining the
gel in transfer buffer will result in lower sensitivity. Therefore, for blotting techniques, staining the gel with SYPRO Tangerine protein gel
stain, which does not require acetic acid fixation, or staining the blot directly with SYPRO
Ruby protein blot stain is recommended.
Diluting fluorescent stain below the recommended concentration will result in reduced
staining sensitivity. Using higher staining concentrations than recommended will not result
in better detection, but will instead result in
increased background and quenching of the
fluorescence from dye molecules crowded
around the proteins.
SYPRO Red and Orange stains cannot be
used to prestain protein samples for SDS gels.
Loading solutions contain so much SDS that
the dye simply localizes in the free SDS and
binds very little to the proteins.
The SDS front at the bottom of the gel stains
very heavily with SYPRO stains. Unless the
proteins of interest co-migrate with the SDS
front, it will be advantageous to run the SDS
front off the gel. Colored stains and marker
dyes, as well as commercially prestained protein markers, interfere with SYPRO dye staining and quench fluorescence.
Highly colored prosthetic groups (e.g.,
heme) that remain bound in native gels will
quench fluorescence of the SYPRO Orange and
Red stains. Odd marks on stained gels can be
caused by several factors. If the gel is squeezed,
a mark appears that stains heavily with the
SYPRO dyes. This is probably due to a localized high concentration of SDS that has difficulty diffusing out. Glove powder can also give
background markings, so rinsing or washing
gloves is recommended prior to handling gels.
Staining with the SYPRO Orange dye occasionally results in gels with scattered fluorescent speckles.
Due to different staining properties of proteins, dual staining procedures can reveal pro-
Analysis of
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10.6.23
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teins with one procedure that the other has not
visualized. SYPRO-stained gels can be restained with either Coomassie brilliant blue or
silver stain procedures. In fact, for some silver
staining methods, the authors have found that
prestaining with SYPRO dyes actually increases the rate of staining and the sensitivity
for detection. To fluorescently stain gels that
have previously been stained with Coomassie,
the Coomassie stain must be completely removed, as it will quench the fluorescence of
SYPRO dyes. Soaking the gel in either 30%
methanol or 7.5% acetic acid with several
changes of destaining solution is effective at
removing Coomassie. Once the Coomassie has
been removed, the gel should be incubated in
0.05% SDS for 30 min before staining with the
SYPRO stain as usual.
Triton X-100 at ≥0.1% interferes with
SYPRO dye staining. If Triton X-100 is used
in the gel, the authors recommend soaking the
gel in two to three changes of buffer to be sure
the Triton X-100 is diluted out, and then incubating the gel in 0.05% SDS for 30 min before
staining as usual.
Staining for glycoproteins
The overall specificity of glycoprotein detection by the Pro-Q Emerald 300 reagent
method depends greatly upon the specificity of
the oxidation reaction, which is governed in
turn by the reaction conditions used (e.g., periodic acid concentration, pH, temperature, and
exposure to light). Careful attention to the protocol is required to avoid oxidation of serine,
threonine, and hydroxylysine residues to form
aldehyde groups, which result in false-positive
signals. Residual SDS in the gel will also lead
to nonspecific staining. This can be avoided by
adhering strictly to the fixation and wash volumes and times indicated in Alternate Protocol
6. In addition, it is advisable to stain a duplicate
gel, eliminating the oxidation step (step 4), as
a negative control.
Anticipated Results
Staining Proteins
in Gels
The sensitivity of Coomassie blue gel staining is 0.3 to 1 µg/protein band; the sensitivity
of silver staining is 2 to 5 ng/protein band. The
sensitivity of both stains varies in an unpredictable manner with the protein being stained.
For fluorescent dyes, detection limits are
typically ∼500 ng protein/band in room light,
∼50 ng protein/band with 300-nm transillumination, and ∼1 to 2 ng protein/band in a photograph taken with Polaroid 667 black-and-white
print film. The authors achieve detection limits
of 1 to 2 ng/band using a Fotodyne Foto/UV
450 ultraviolet transilluminator, which has six
15-watt bulbs that provide peak illumination at
312 nm. When using weaker illumination
sources, exposures must be correspondingly
longer. Although the authors’ detection limits
are 1 to 2 ng/band for most proteins, it should
be emphasized that bands containing 5 to 10
ng/protein are more readily detected. Bands
containing less than 5 to 10 ng protein require
longer exposures and sharp bands for good
visualization. Longer exposures can result in
higher background.
Time Considerations
Coomassie blue staining requires 8 to 12 hr.
Silver staining requires ∼5 hr. Fixation may be
extended for several days before Coomassie
blue staining. Fixation may be extended for
longer periods—up to several weeks—before
silver staining.
Use of either of the rapid staining protocols
considerably reduces the time required to visualize proteins. Detection of protein bands by
rapid Coomassie blue staining requires ≤90 min
from the time a minigel is run (30 to 60 min)
until the gel is fixed (10 min) and placed in
staining solution (5 to 10 min); however, additional time may be necessary for larger gels.
Separated proteins stained with the rapid silver
stain method can be visualized in ∼35 min.
The staining time for SYPRO dyes is 10 to
60 min, depending on the thickness and percentage of the gel. For 1-mm-thick 15%
polyacrylamide gels, the signal is typically optimal at 40 to 60 min of staining.
Literature Cited
Anderson, N.L. 1988. Two Dimensional Electrophoresis Operation of the ISO-DALT (R) System; Large Scale Biology Press, Washington,
D.C.
Bloom, H., Beier, H., and Gross, H.S. 1987. Improved silver staining of plant proteins, RNA and
DNA in polyacrylamide gels. Electrophoresis
8:93-99.
Merril, C.R., Goldman, D., and Van Keuren, M.L.
1984. Gel protein stains: Silver stain. Methods
Enzymol. 104:441-447.
Morrissey, J.H. 1981. Silver stain for proteins in
polyacrylamide gels: A modified procedure with
enhanced uniform sensitivity. Anal. Biochem.
117:307-310.
Oakley, B.R., Kirsch, D.R., and Morris, N.R. 1980.
A simplified ultrasensitive silver stain for detecting proteins in polyacrylamide gels. Anal. Biochem. 105:361-363.
Steinberg, T.H., Haugland, R.P., and Singer, V.L.
1996a. Applications of SYPRO orange and
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SYPRO red protein gel stains. Anal. Biochem.
239:238-245.
Steinberg, T.H., Jones, L.J., Haugland, R.P., and
Singer, V.L. 1996b. SYPRO orange and SYPRO
red protein gel stains: One-step fluorescent staining of denaturing gels for detection of nanogram
levels of protein. Anal. Biochem. 239:223-237.
Steinberg, T.H., White, H.M., and Singer, V.L. 1997.
Optimal filter combinations for photographing
SYPRO orange or SYPRO red dye-stained gels.
Anal. Biochem. 248:168-172.
Switzer, R.C., Merril, C.R., and Shifrin, S. 1979. A
highly sensitive silver stain for detecting proteins
and peptides in polyacrylamide gels. Anal. Biochem. 98:231-237.
Wilson, C.M. 1983. Staining of proteins on gels:
Comparison of dyes and procedures. Methods
Enzymol. 91:236-247.
Contributed by Joachim Sasse
Shriners Hospital for Crippled Children
Tampa, Florida
Sean R. Gallagher
UVP, Inc.
Upland, California
Analysis of
Proteins
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