Westermeier R., Naven T., H?pker H.-R. Proteomics in Practice: A Guide to Successful Experimental Design

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3.6 Staining of IPG Strips 321

3.6

Staining of IPG Strips

3.6.1

Acid Violet 17 Staining of IPG Strips

3% (v/v) phosphoric acid 21 mL 85% H

Make up to 1 L with

deion

11% (v/v) phosphoric acid 76.1 mL 85% H

Make up to 1 L with

deion

1% (w/v) stock dye solution 1 g Acid violet 17 In 100 mL H

deion

Heat to 50 C on

magnetic stirrer

0.1% (w/v) staining solution 1 mL 1% (w/v) stock

solution

Plus 9 mL of 11%

phosphoric acid

Staining procedure

Fix for 20 minutes in 20% (w/v) TCA.

Wash for 1 minutes in 3% (v/v) phosphoric acid.

Stain for 10 minutes in 0.1% (w/v) Acid violet 17

staining solution.

Destain with 3% (v/v) phosphoric acid until

background is clear.

Wash three times (1 minute each time) with

deion

Impregnate with 5% (v/v) glycerol, air dry.

& Note: This staining is not

reversible!

323

Step 4: SDS Polyacrylamide Gel Electrophoresis

This chapter describes casting and running of large vertical gels.

Small vertical gels and horizontal flatbed gels on film supports are

described in detail in “Electrophoresis in Practice”.

Wear disposable gloves during the entire procedure.

4.1

Casting of SDS Polyacrylamide Gels

Laboratory-made gels are cast in glass cassettes using relatively sim-

ply designed equipment. Casting gels on support films requires more

sophisticated equipment. Film-backed gels are therefore only pre-

pared in industrial scale. The preparation of gels with two different

gel casters is described, and how to cast gradient gels without a

pump. The following protocols are limited to the preparation of

1 mm thick gels, for 1.5 mm gels one-third more monomer volume

is used; there is space for a lower number of cassettes in the casters.

The large cassettes are built from 3 mm thin glass plates. It is

important to prepare such gels in multicasters, because this prevents

the cassettes from bulging, which would lead to uneven gel thickness.

However, as the polymerization process is an exothermal reaction,

care must be taken to prevent overheating in multicasters.

4.1.1

Stock Solutions

Use a ready-made acrylamide, Bis stock solution containing 40% T,

3% C.

Or prepare:

Proteomics in Practice. A Guide to Successful Experimental Design 2

Ed.

Reiner Westermeier, Tom Naven, and Hans-Rudolf Hçpker

ISBN: 978-3-527-31941-1

Westermeier R. Electrophoresis

in Practice. Fourth Edition.

WILEY-VCH, Weinheim

(2004).

1 mm gels are anyhow advan-

tageous for downstream

analysis with mass spectro-

metry, because the yield for the

tryptic digestion is higher.

Casting single gels requires a

support by thick glass or plastic

plates.

Much safer than weighing acry-

lamide and Bis powder.

Step 4: SDS Polyacrylamide Gel Electrophoresis324

Acrylamide, Bis solution (40% T,3%C):

40% Acrylamide PAGE solution 40% T 1L

Bis 3% C 12 g

Mix thoroughly in the supplied bottle

Store in the refrigerator.

& Caution! Acrylamide and Bis are toxic in the

monomer form. Avoid skin contact and dispose

of the remains ecologically. Polymerize the

remains with an excess of ammonium persulfate.

1.5 mol/L gel buffer Tris-Cl pH 8.8 (4 concentrated):

Tris base 1.5 mol/L 181.8 g

SDS 0.4% (w/v) 4 g

Water, deionized dissolve 800 mL

4 mol/L HCl titrate to pH 8.8

Water, deionized make up to 1 L

Store in the refrigerator.

Ammonium persulfate solution (APS):

Ammonium persulfate 10% (w/v) 1 g

Water, deionized Dissolve 10 mL

Prepare fresh.

TEMED is used undiluted as 100%, because it is added to the

monomer solution at the beginning.

1 % (w/v) Bromophenol blue solution:

Bromophenol blue 1% (w/v) 100 mg

Tris base 60 mg

Water, deionized dissolve 10 mL

Displacing solution:

Tris-Cl (1.5 mol/L, pH 8.8), SDS 0.375 mol/L 60 mL

85%Glycerol (v/v) 50% (v/v) 142 mL

1% Bromophenol blue (w/v)) 0.002% 480 mL

Water, deionized Make up to 240 mL

Prepare fresh.

Acrylamide powder could also

be used, but is not recom-

mended because of health

hazards during weighing.

A 10% solution is mixing more

homogeneously with the

monomer solution than a 40%

solution.

1% Bromophenol blue does not

go completely into solution

without Tris

4.1 Casting of SDS Polyacrylamide Gels 325

Overlay spraying solution:

SDS 0.1% (w/v) 1 g

Water, deionized Dissolve 1 L

Gel storage solution:

Tris-Cl (1.5 mol/L, pH 8.8), SDS 0.375 mol/L 500 mL

Water, deionized Make up to 2 L

Store in the refrigerator.

4.1.2

Cassettes for Laboratory-made Gels

It is very important to have absolutely clean glass plates and cassettes

at hand. Therefore the tips for cleaning the glass plates at the end of

this chapter are very important. To be on the safe side and to remove

any dust particles, clean glass plates again before use with distilled

water and lint-free tissue.

Hinged cassettes The standard cassettes are hinged on one side for

easy handling. Note that one of the class plates are offset to make

application of the IPG strip easier.

Separate and low fluorescence glass plates For spot picking, the gels

need to be covalently bound to one glass plate. In this case hinged

cassettes are not useful, and separate glass plates are used – one with

spacers and one without.

For pre-labeling fluorescence detection methods like DIGE, special

borosilicate glass plates are needed, which are usually not hinged.

Treating glass plates with Bind-Silane The gel will be bound to the

glass plate without spacers; this one has to be treated with Bind-

Silane.

& Note: It is important that glass plates are

properly clean.

Before re-use, the plates are placed in a 5% (v/v) Decon 90 solution

overnight. Do not leave plates standing in this solution for a longer

time, because this will cause etching due to the alkali nature of

Decon.

Thoroughly wash the plate to be treated. Any gel

fragments from previous gels must be removed.

Inspect the cassettes carefully

for any dirt or dust particles.

If Decon 90 is not available,

just use 5% (w/v) potassium

hydroxide.

Step 4: SDS Polyacrylamide Gel Electrophoresis326

The careful cleaning of the glass plates before

casting is important, to ensure a uniform coat-

ing with the Bind-Silane and to avoid keratin

contamination.

Thoroughly rinse the plates with deionized water

to remove Decon.

Dry the plate using a lint-free tissue or leave

them to air dry.

Apply two internal reference (IR) markers (self-

adhesive, fluorescent) onto the middle of the

glass plate, each approximately 3.5 cm away

from the lateral edges.

Bind-Silane working solution:

Ethanol 8 mL

Acetic acid 200 mL

Bind-Silane 10 mL

Water, deionized 1.8 mL

Pipette 4 mL of the Bind-Silane solution onto

the plate and distribute equally over the plate

with a lint-free tissue. Cover the plate to prevent

dust contamination and leave to air dry on the

bench for 1 hour.

Polish the plate with a lint-free tissue, moistened

with a small amount of deionized water or etha-

nol.

The gels will stay attached to the glass during

electrophoresis, staining procedures, scanning,

spot picking, and storage.

Prepare the cassettes with gel labels cut from printed paper or over-

head film placed at the corner on the opposite of the filling side (see

Figure 4.1).

The IR markers must also be

coated with Bind-Silane.

Note that a too thick layer of

Bind-Silane can create a high

background in fluorescent

detection.

This is not necessary for DIGE

runs, when IPG strips with

printed numbers are used: the

strip is scanned together with

the SDS gel in the cassette.

4.1 Casting of SDS Polyacrylamide Gels 327

Fig. 4.1: Hinged cassette with gel label made from paper

or overhead film (the label is not needed when IPG strips

with printed numbers and the DIGE method are employed).

4.1.3

Multiple Gel Caster (Up to 14 Gels)

4.1.3.1 Preparation of the Gel Caster

The gel caster should be set up in a tray, just in case any liquid over-

flows. Never forget to place the plastic separator sheets between the

cassettes; they would firmly stick to each other after gel polymeriza-

tion. When less than 14 gels are being prepared, block the non-used

space with blank cassettes, placing separator sheets between each

plate.

Be sure that gel caster, separator sheets, and cassettes are perfectly

clean and dry.

Place the gel caster in a tray. Tip it back, so it

rests on the support legs.

According to the instruction manual the hinged cassettes should be

placed into the caster with the opening edges to the filling side, some

proteomics teams have experienced to get a straighter gel surface,

when they are inserted with the hinged edge to the filling side.

Start with a separator sheet.

Assemble non-hinged cassettes outside of the

caster before you place them into the caster box.

Dirty cassettes cause vertical

streaking in silver-stained gels.

For non-hinged cassettes the

orientation does not matter.

Step 4: SDS Polyacrylamide Gel Electrophoresis328

Place the cassettes alternating with separator

sheets into the caster as shown in Figure 4.2,

with the offset edges of the cassettes up.

Fig. 4.2: A. Placing cassettes and separator sheets into the

caster. Here the hinged edges are oriented towards the filling side.

B. Closing the gel caster.

Press strongly on the spacers areas on both sides

to ensure that the spacers and the glass plates

stick well together.

Finally a separator sheet and a filling plate are inserted to complete

the stack up to the edge of the caster. Do not overfill it. This would

cause pressure on the gels during polymerization, when the solution

starts to get warm.

Place the foam gasket into the groove of the

front plate. Do not cover it with grease.

Turn the first four screws into the bottom holes

and place the front plate on the caster with the

bottom slots on the screws. Apply the rest of the

screws and tighten them evenly. Do not use too

much force; the sealing gasket should be com-

pressed evenly to prevent leakage.

Tip the caster to the front.

Level the gel caster horizontally.

Prepare 120 mL of displacing solution.

When the room temperature is higher than 20 C, the caster should –

like the monomer solution(s) – be pre-cooled in a cold room or a

refrigerator.

4.1.3.2 Homogeneous Gels

Very good results are usually obtained with homogeneous gels (e.g.

monomer solutions). Gradient gels need to be prepared in special

cases.

Always take the foam gasket

out of the front plate after gel

casting.

4.1 Casting of SDS Polyacrylamide Gels 329

For monomer solutions the ammonium persulfate should not be

added before everything is in place, because there are only about 10

minutes left to pour the gel until it starts to gel.

Monomer solutions with selected % T, and 3% C for 14 gels with

1 mm thickness, 900 mL:

7.5%

10%

12.5%

15%

Acrylamide, Bis solution (40% T,3%C ) 169 mL 225 mL 281 mL 338 mL

1.5 mol/L Tris-Cl, pH 8.8, 0.4% SDS 225 mL 225 mL 225 mL 225 mL

TEMED (100%) 450 mL 450 mL 450 mL 450 mL

Water, deionized, make up to 900 mL 900 mL 900 mL 900 mL

Mix and degas with a water jet pump

10% ammonium persulfate 3.6 mL 3.6 mL 3.6 mL 3.6 mL

The amounts of TEMED and ammonium persulfate are based on

the author’s experience. In order to get reproducible gels with a

straight edge, the concentration of TEMED is higher and the concen-

tration of ammonium persulfate is lower than in the instrument

instruction. Furthermore, at basic pH, ammonium persulfate can

react with the Tris; this effect is minimized by adding more TEMED

and reducing the ammonium persulfate content.

Degas and precool the monomer solution.

Add the ammonium persulfate to the monomer

solution and mix.

Pour the gel solution directly into the balance

chamber, avoiding air bubbles (see Figure 4.3A).

Stop pouring when the level reaches 3 cm below

the upper edges of the casting cassettes.

Pour the dense displacing solution to fill the

V-chamber and the sloped bottom of the caster.

The gel solution level will rise to 1 cm below the

cassette edges. A thin blue layer should be visi-

ble at the bottom of the cassettes.

Should the volume of the displacing solution in the balance chamber

not be sufficient to produce the thin blue layer in the entire area of

the caster bottom, top more displacing solution into the balance

chamber until the blue layer has distributed over the entire bottom

area. This saves you from cutting off protruding gel pieces at the bot-

tom edge of each cassette.

Immediately spray 0.1% SDS solution overlay

solution over the cassettes (see Figure 4.3C).

Cover the caster with a cling film.

Add ammonium persulfate

shortly before gel casting.

It is important to use only high-

quality TEMED and ammo-

nium persulfate.

And it prevents a too strong

contraction of the gel during

polymerization, which can

cause a gap between gel and

spacer on the side of the gels

where the solution is introduced

from the bottom.

Spraying with SDS solution is

better than pipetting water-

saturated butanol on the edge.

Step 4: SDS Polyacrylamide Gel Electrophoresis330

Fig. 4.3: Casting homogeneous gels in a 14-gel caster.

A. Pouring the monomer solution into the balance chamber.

B. Pouring the displacement solution into the balance chamber.

C. Overlaying the gel solution by spraying 0.1% SDS solution.

About 50 mL monomer solution should be left over. Let it polymerize

in a small beaker before you discard it.

Let the gel polymerize overnight at room tem-

perature.

During unloading rinse the cassettes to remove

excess polyacrylamide.

& Note: When enough displacing solution had

been used, there is no need to cut off protruding

gel pieces at the bottom edge of each cassette.

Inspect each gel cassette for eventual air bub-

bles. Gels with air bubbles should not be used.

Store the usable gels in an airtight container

with about 100 mL gel storage solution. When

Because the silent polymeriza-

tion must be completed.

4.1 Casting of SDS Polyacrylamide Gels 331

the gels are not used during the next day, place

the container with the gels into a cold room or a

refrigerator.

Take the foam gasket out of the front plate and

rinse it with deionized water.

Rinse the gel caster and the separator sheets

with mild detergent and then with deionized

water.

Let them dry in the air.

4.1.3.3 Gradient Gels

Gradient gels are more complicated to prepare than homogeneous

gels. When homogeneous gels are run for a short time only, they

achieve comparable resolution. However, for some samples, gradient

gels may show a better spot distribution. Most instructions describe

how to cast gradient gels with a peristaltic pump. The author has

experienced that very reproducible gels are achieved without a pump,

which safes a lot of work with calibrating, setting up and cleaning.

Instead of a pump, gravitation is used; the flow rate is controled by

the level of the laboratory platform carrying the gradient maker and

with a pinchcock clamp on the tubing. A pinchcock clamp can close

the tubing completely or reduce the opening of the tubing in several

steps, thus reducing the flow rate.

Gradient maker set up

Place the clean and dry gradient maker with a

magnetic stirrer on an adjustable laboratory plat-

form or on a shelf board: the level must be at

least 10 cm above the edge of the caster.

& Note: For reproducible gels, use always exactly

the same level for the gradient maker position.

Use always the same shelf board or measure and

note the position height of the gradient maker.

Insert the plastic tube into the hole in the hydro-

static balance chamber.

Place a pinchcock clamp on the flexible connec-

tion tubing, and close it completely.

Connect the plastic tube with the flexible tubing

to the gradient maker.

Fill the hydrostatic balance chamber with

100 mL of the displacing solution.

Place a magnetic stirring bar – 2 cm to 3 cm

long – into the mixing chamber.

Do not leave it in the groove,

because it would lose its sealing

property.

Pinchcock clamps are available

in different sizes.