**ARGONNE NATIONAL LABORATORY**

**PROCEDURES FOR TWO-DIMENSIONAL**

**ELECTROPHORESIS OF PROTEINS**

# Carol S. Giometti and Sandra L. Tollaksen

[INTRODUCTION](#Introduction)

1 [SAMPLE PREPARATION](#SamplePreparation)

2 [FIRST DIMENSION SEPARATION](#Chapter2)

-   [ISO](#Chapter2ISO)

-   [IPG](#IPGMethods)

-   [BASO (NEPHGE)](#Chapter2_5BASO)

-   [ACIDO](#Chapter2_6Acido)

3 [SECOND DIMENSION SEPARATION](#Chapter3Dalt)

4 [PROTEIN DETECTION](#Chapter4ProteinDetection)

-   [COOMASSIE BLUE](#Chapter4_1CoomassieStaining)

-   [SILVER STAIN](#Chapter4_2SilverStaining)

-   [AUTORADIOGRAPHY](#Chapter4Autoradiography)

5 [PROTEIN TRANSFERS](#Chapter5Transfers)

6 [WESTERN BLOTS](#Chapter6WesternBlots)

-   [HRP COLOR DEVELOPMENT](#Chapter6_1HRPColorDevelopment)

-   [CHEMILUMINESCENCE](#Chapter6_2Chemiluminescence)

7 [REFERENCES](#Chapter7References)

8.  [RECIPES](#Chapter8Recipes)

For technical questions contact: Sandra Tollaksen
(<tollaksen@anl.gov>)[]{#SamplePreparation .anchor}

[]{#Introduction .anchor}**INTRODUCTION**

The ISO-DALT system is a high-throughput approach to analysis of
proteins by two-dimensional gel electrophoresis (2DE). Originally built
at Argonne National Laboratory under the direction of Norman and Leigh
Anderson ([1978a](#ReferenceAndersons),
[1978b](#ReferenceAndersonsUrine)), the ISO-DALT system provides the
capability to generate and run first-and-second-dimension gels in
multiples of 20.

Modifications of the chemistry used for first-dimension separation in
the ISO apparatus have been developed to allow the resolution of very
[acidic](#Chapter2_6Acido) and very [basic](#Chapter2_5BASO) proteins,
in addition to the classical use of [isoelectric focusing](#Chapter2ISO)
for the separation of neutral proteins. Modifications of the
polyacrylamide gel composition for the second-dimension separation in
the DALT apparatus can be made to adjust the molecular weight range of
proteins resolved. Thus, the spectrum of proteins that can be surveyed
by 2DE has been broadened beyond that originally described by [Patrick
O'Farrell](#ReferenceOFarrellFirstPaper) in 1975.

The Argonne ISO-DALT system has stood the test of time, still producing
2DE patterns of superior quality. The procedures described in this
Web-based manual have evolved over almost three decades of use and
include the most current recipes and electrophoresis procedures used by
the Argonne Protein Mapping Group. The details of the procedures are
specific to the Argonne ISO-DALT system and may require some
modification for use with commercially available electrophoresis
systems. The recipes are generally applicable independent of the
equipment used.

Although the vendors and products described pertain to the work done by
the Argonne Protein Mapping Group, they are not specifically endorsed or
recommended by Argonne National Laboratory. The authors of this manual
have 50 combined years of work with 2DE and have run over 55,000 gels.
Thus, the procedures outlined in this document have been well tested and
fine-tuned.

**1.** **SAMPLE PREPARATION**

**1.1 GENERAL SAMPLE PREPARATION**

The solutions for sample preparation can be made in fairly large
quantities (up to 1 L) and stored frozen at −70 °C in small aliquots.
The following four solutions have been optimized for the samples
specified. Experimentation with these methods and chemicals is advised
to find the best conditions for other types of samples.

For all of the solubilization conditions described below, samples should
be centrifuged at 22 °C for 1 h at approximately 100,000 × *g* in an
ultracentrifuge or for 10 min at 100,000 rpm (435,000 × *g*) in a
Beckman TL‑100 centrifuge. For mixes containing 9 M urea, use 1.5 ml
polyallomer centrifuge tubes, as polycarbonate tubes may crack. The
supernatant is recovered for analysis by 2DE. Samples may be analyzed
immediately or stored frozen (−70 °C) for future analysis. The small
translucent pellet that contains DNA and cell debris is discarded.

1.  [SDS mix](#Recipe1BlueMix) (body fluids such as serum, plasma, or
    amniotic fluid). To a 10 μl sample, add 20−30 μl of SDS mix. Next,
    heat the sample on a 95 °C heating block for 5 min to achieve
    maximum solubilization and to inactivate any proteolytic enzymes.

2.  [NP-40/urea mix](#Recipe2PinkMix) (solid tissue samples, isolated
    cells, and pure proteins). For a wet tissue sample, use a volume of
    mix that is eight times the blotted wet weight of the sample (e.g.,
    for a 100 mg sample, use 800 μl of mix). For a frozen tissue sample,
    pulverize the sample on a platform chilled on dry ice and use a
    volume of mix that is four times the weight of the powdered sample.
    Homogenize (or sonicate) before centrifuging the sample. For
    solubilizing isolated cells, use 5 × 10^6^ cells per 50 µl of mix
    when Coomassie blue detection will be used. When silver‑stain will
    be used, use 5 × 10^5^ cells per 50 µl of mix. Disperse the cell
    pellet by tapping the bottom of the tube gently prior to addition of
    the solubilization mix. After the mix is added, mix the cell lysate
    well by drawing the solution up and down in a pipette tip. For pure
    protein samples, mix with solubilization mix to obtain a final
    protein concentration of 1 mg/mL. **Do not heat** any protein sample
    in the presence of urea, because carbamylation of the proteins will
    occur.

3.  [Urea mix without NP-40](#Recipe3YellowMix) (urine). Mix 10 mg of
    lyophilized urinary proteins with 100 μl of the mix. Load 10 μl (or
    less) of the solution onto the ISO gel.

4.  [NP-40/urea/DTE mix](#Recipe4GreenMix) (muscle). For a wet tissue
    sample, use a volume of mix that is eight times the blotted wet
    weight of the sample (e.g., for a 100 mg sample, use 800 μl of mix).
    For a frozen tissue sample, pulverize the sample on a platform
    chilled on dry ice and use a volume of mix that is four times the
    weight of the powdered sample. Homogenize (or sonicate) before
    centrifuging the sample.

**1.2 RADIOACTIVE SAMPLE PREPARATION**

1\. Samples (cells or tissue pieces) are incubated with either
[l]{.smallcaps}‑\[^35^S\]‑methionine (approximately 50 μCi per sample)
or ^32^P‑orthophosphate (200 μCi per sample) in methionine- or
phosphate-free tissue culture media for 1-18 hours.

> 2\. Cells or tissue pieces are collected from the labeling media by
> centrifugation, and the media is discarded. Samples are washed at
> least twice with phosphate-buffered saline.

3.  The radioactively labeled cells or tissues are solubilized in
    approximately 50 μl of [NP40-urea mix](#Recipe2PinkMix) and
    centrifuged in a Beckman microfuge for 8 min at 100,000 rpm. After
    centrifugation, the supernatants are stored at −70 °C.

**1.3 PROTEIN ANALYSIS**

The protein concentrations of samples in solubilization mixtures
containing NP-40/urea and mercaptoethanol can be determined using a
modified Bradford protein assay ([Ramagli and Rodriguez
1985](#ReferenceRamagliModifiedProtein)). The optimal protein load for
Coomassie blue staining of whole cell lysates is 150-300 μg protein,
while 20-60 μg of protein is recommended for silver-stained gels.

**CHAPTER 2: THE ISO SYSTEM FOR FIRST-DIMENSION SEPARATION**

[]{#Chapter2ISO .anchor}**2.1 CASTING GELS IN THE ISO APPARATUS**

There are two ISO formats, the 7-in. (18 cm) and the 10-in. (25 cm). The
selection of ISO format should be based on optimal resolution of the
proteins being analyzed. At Argonne, the 10-in. format is used for
isoelectric focusing of mouse liver and tissue culture cell proteins,
and the 7-in. format is used for BASO-DALTs and isoelectric focusing of
microbial proteins. The entire system must be [clean and
dry](#CleaningUp).

1.  Place a metal retainer on the bottom of the gel tubes.

2.  Fill the bottom chamber of the ISO apparatus with tap-distilled
    water (2 l for 7‑in. systems and 3 l for 10-in. systems). Place the
    Lucite trough on the base of the support stand, and position the
    rack containing the upper chamber and the gel tubes into the acrylic
    trough.

3.  Prepare the polyacrylamide gel solution by mixing the following in a
    150‑ml lyophilizer flask. (Note: acrylamide monomer must be handled
    as a suspected human carcinogen.)

  -----------------------------------------------------------------------
  Compound                                                 Amount
  -------------------------------------------------------- --------------
  Urea                                                     8.25 g

  [Acrylamide (30% solution)](#Recipe5_30Acrylamide)       2.0 mL

  Ampholytes (the majority of ANL isoelectric focusing     0.8 mL
  gels are 50:50 pH 5-7:pH 3-10; *P. furiosus* and *S.     
  oneidensis* gels are an exception)                       

  Double-distilled water                                   6.0 mL
  -----------------------------------------------------------------------

> Dissolution, an endothermic process for urea, is aided by warming the
> flask and its contents in a water bath, but do not heat the urea
> solution beyond room temperature.
>
> 4\. Degas the solution briefly using a vacuum pump dedicated for
> degassing acrylamide solutions. (If the degassing is too long, the
> urea will come out of solution; if this happens, warm the flask
> slightly until the urea goes back into solution.)
>
> 5\. Add 0.3 ml NP-40 detergent (except with gels for urine proteins,
> where only two drops should be added) and swirl gently (vigorous
> swirling introduces bubbles). Then carefully add:

+----------------------------------------------------+-----------------+
| Compound                                           | Amount          |
+====================================================+=================+
| [Ammonium persulfate (catalyst; 10%                | 50 μL           |
| solution)](#Recipe6AmmoniumPersulfate)             |                 |
+----------------------------------------------------+-----------------+
| TEMED (*N,N,N',N'*                                 | 5 μL            |
|                                                    |                 |
| -tetramethylethylenediamine; an accelerator)       |                 |
+----------------------------------------------------+-----------------+

> 6\. Pipette the acrylamide solution (approximately 15 mL) into the
> trough. Carefully layer 3−4 ml of double-distilled water over the
> acrylamide solution to bring the fluid level up to the top of the
> trough.
>
> 7\. Slowly lower the upper chamber/tube assembly with the acrylamide
> trough into the bottom chamber containing the appropriate volume of
> water. Allow the acrylamide to rise evenly by displacement in all
> tubes. Examine the tubes for bubbles. If there are any, use a 1 ml
> syringe with a cut off yellow pipette tip attached (the wide top end
> cut off) to suck acrylamide out the top of the tube until the bubble
> is removed. Allow the fluid to fall back to the proper level.
>
> 8\. Allow the gels to polymerize for at least 2 hours.

**2.2 PREFOCUSING**

> 1\. After the gels have polymerized, remove the upper chamber/gel tube
> assembly with the support stand from the bottom chamber. Empty the
> water from the bottom chamber and fill with 10 mM phosphoric acid.
>
> 2\. []{#Chapter2_2CleaningSetupforPrefocus .anchor}Carefully place the
> tube stand on its side and hold on to the sides of the metal retainer.
> Pull off the trough by wiggling it free of the polymerized acrylamide
> and retainer. Do not bend the metal retainer or stretch the gels in
> the tubes. Cut with a razor blade between the end of the tubes and the
> metal retainer. Remove the retainer and rinse the outside of the
> tubes. Reinsert the upper chamber with attached tubes into the lower
> chamber containing the acid solution.

4.  Degas 200 ml of double-distilled water. Add 0.4 ml of 10 N NaOH and
    pour the solution into the upper chamber. Use a 100 μl Hamilton
    syringe containing the NaOH solution to displace the air pocket that
    forms between the top of the gels and the NaOH in the upper chamber.
    Be careful not to disturb the tops of the gels.

> 5\. []{#Chapter2_2Prefocus1hr .anchor}Prefocus the 7-in. ISO for 1 h
> at 200 V, or the 10‑in. ISO setup at 300 V.

**2.3 ISOELECTRIC FOCUSING WITH CARRIER AMPHOLYTES \[ISO\]**

Using a Hamilton syringe, underlay 5 to 25 μl of sample on top of each
isoelectric focusing gel. Larger volumes result in poor resolution and
can cause the ISO gel to break. The optimum protein loading level is
100−300 μg for Coomassie blue detection or 20−60 μg for silver stain. If
isoelectric focusing standards (e.g., Carbamalyte Calibration Kit,
Amersham Pharmacia Biotech, Catalogue 17‑0582‑01) are to be run with the
samples, add 2−4 μl to each tube above the sample.

Although the optimal separation time depends on the sample type, most
cellular proteins are well separated in the 7-in. ISO apparatus with a
run time of approximately 14,000 Vhe.g., 800 V for 17.5 h), while
plasma or amniotic fluid proteins require approximately 12,000 Vh. When
using the 10-inch ISO apparatus, the optimum run for tissue and cellular
protein is 30,000 Vh.

4.  []{#IPGMethods .anchor} **ISOELECTRIC FOCUSING WITH IPG STRIPS
    \[IPG\]**

IPGs are immobilized pH gradient strips that are fixed on a solid
plastic support. They must be rehydrated before isoelectric focusing.
The apparatus with corollary equipment is purchased commercially (e.g.,
Bio-Rad), and instructions for setup are included.

Add 125-250 μl (7 cm IPG strip) or 300-600 μl (17 cm IPG strip) of the
[rehydration buffer](#Recipe24RehydrationBuffer) to each channel of the
isoelectric focusing tray, making sure that each strip is completely wet
to prevent uneven rehydration. Volume of the sample should not exceed 30
μl (5−100 μg protein for silver staining, and up to 1 mg for Coomassie
blue) per IPG strip. Apply mineral oil to each channel containing a
strip, covering the entire strip. Put the lid on the tray and place the
entire assembly on the Peltier platform. Be sure to align the electrodes
of the focusing tray with the color-coded Peltier platform's electrode
connections. Program or select the desired method, and start the run.

Isoelectric focusing is immediately done after rehydration. IPG strips
may be stored indefinitely at −70 °C after isoelectric focusing or may
be loaded on to second dimension gels. Before running in the second
dimension, IPG strips are equilibrated first with [DTT-equilibration
buffer](#Recipe25EquilibrationBuffer) for 10 min and then with
[iodoacetamide-equilibration buffer](#Recipe25EquilibrationBuffer) for
an additional 10 min. After equilibration, IPG strips are placed on top
of the second dimension slab gels.

[]{#Chapter2_5BASO .anchor}**2.5 NON-EQUILIBRIUM pH GRADIENT
ELECTROPHORESIS**

**(NEPHGE/BASO)**

Proteins with pI values higher than 8.0 do not focus well or fail to
enter isofocusing gels. To study such basic proteins, the technique of
running a BASO under nonequilibrium conditions was devised as a
modification of the NEPHGE (non-equilibrium pH gradient electrophoresis)
system ([O'Farrell et al., 1977](#ReferenceOFarrell)).

> 1\. The BASO gels are cast in the 7-in. ISO apparatus using the same
> procedure as for [ISOs](#Chapter2ISO). However, the BASO gel
> separation requires the use of the wide range pH 3−10 ampholyte (e.g.,
> Biolyte 3/10).

2.  After the gels have polymerized, remove the upper chamber/gel tube
    assembly with the support stand from the bottom chamber. Empty the
    bottom chamber and fill with 2 l of 20 mM NaOH prepared using
    degassed double-distilled water.

3\. Clean the gel tubes as described for
[prefocusing](#Chapter2_2CleaningSetupforPrefocus) ISO gels.

> 4\. Samples are applied to BASO gels without prefocusing. Using a
> Hamilton syringe, samples are loaded directly to the upper gel surface
> in each tube. Samples are then overlaid with 4 M urea, filling the
> remainder of the gel tube. Fill the upper chamber with 200 ml of 10 mM
> H~3~PO~4.~ The 4 M urea cushion serves to protect the sample proteins
> from the acid in the upper chamber.

5.  Reverse the electrodes relative to their use for isofocusing, i.e.,
    so that the positive electrode is attached to the top chamber and
    the negative electrode is attached to the bottom chamber. To run the
    BASO gels in one day, start the electrophoresis at 400 V for 1 h and
    then turn up the power to 800 V for 4−8 h, stopping the run at
    approximately 3000−8000 Vh. An overnight run is done at 300 V for
    20 h. Use shorter run times to ensure that proteins with isoelectric
    points greater than pH 10 are captured in the gels.

[]{#Chapter2_6Acido .anchor}**2.6 ACIDO ISOELECTRIC FOCUSING**

Urine contains very acidic proteins (most acid urinary protein \[MAUP\]
and α-1-acid glycoprotein) that will not focus following the ISO
protocol for isoelectric focusing gels ([Edwards, Tollaksen, and
Anderson 1982](#ReferenceEdwardsUrine)). The conditions for ACIDO gels,
designed to resolve such acidic proteins, that differ from an ISO run
are shown below.

1\. Acido gels are cast in the 7-in ISO apparatus using the following
recipe:

  -----------------------------------------------------------------------
  Compound                                             Amount
  ---------------------------------------------------- ------------------
  Urea                                                 8.25 g

  [Acrylamide (30% stock solution)](#Recipe5)          2.0 mL

  Ampholyte (pH 2.5−4)                                 1 mL

  Ampholyte (pH 3−10)                                  0.3 mL

  Water                                                5.5 mL
  -----------------------------------------------------------------------

Degas the solution and then add:

  -----------------------------------------------------------------------
  Compound                                             Amount
  ---------------------------------------------------- ------------------
  NP-40                                                0.3 mL

  [Ammonium persulfate (10%                            90 μL
  solution)](#Recipe5_30Acrylamide)                    

  TEMED                                                10 μL
  -----------------------------------------------------------------------

2.  After the gels have polymerized, remove the upper chamber/gel tube
    assembly with the support stand from the bottom chamber.

3.  Clean the gel tubes as described for
    [prefocusing](#Chapter2_2CleaningSetupforPrefocus) ISO gels. Empty
    the bottom chamber and fill with 2 l of water and 3 ml of
    concentrated H~2~SO~4.~

4.  Fill the upper chamber solution with 40 ml of degassed
    double-distilled water and mix with 1 ml of the pH 3−10 ampholyte.
    Add this solution to the upper chamber after the gels have
    polymerized. Debubble each tube carefully using a Hamilton syringe
    containing the ampholyte solution to displace the air pocket that
    forms between the top of the gels and the upper chamber solution.

5.  [Prefocus](#Chapter2_2Prefocus1hr) the setup for 1 h as described.

6.  Underlay approximately 20 μl of sample using a Hamilton syringe and
    run for 3600−4000 Vh. Electrophoresis can be finished in one day
    using 700−800 V or overnight using 200 V (18−19 h). After
    electrophoresis is complete, white precipitate (protein that did not
    enter gels) will be visible at the top of each gel.

**2.7 ISO/BASO/ACIDO GEL RECOVERY AND EQUILIBRATION**

> 1\. If stored refrigerated, stir the [equilibration
> buffer](#Recipe8EquilibrationBuffer) at room temperature until the
> precipitated SDS goes back into solution, and then dispense 4−5 ml
> into 5-ml glass screw-top vials.

2\. At the power supply, turn down the voltage and then turn off the
power. Remove the safety lid and take out the gel tube holder. Drain the
upper buffer solution into the sink (or into a radioactive waste
container if protein sample was radiolabeled) and place the gel tube
holder on the unloading rack. Use a 1 ml syringe attached to a cut-off
pipette tip and filled with double-distilled water to expel the gels
slowly into vials containing the equilibration buffer. When ACIDO gels
are unloaded into the [equilibration
buffer](#Recipe8EquilibrationBuffer) containing bromophenol blue, an
indicator dye, the liquid will change color to yellow or yellow-green.
Rinse the gels with equilibration buffer until the equilibration dye
stays blue.

> 3\. Rock the vials gently at room temperature for a maximum of 5 min
> to minimize protein diffusion. Place the gels on the DALT slabs
> immediately or freeze them at −70 °C and thaw them as needed. Frozen
> first-dimension tube gels produce higher resolution spots than fresh
> gels, but they are thinner and somewhat more fragile. Gels may be
> frozen indefinitely at −70 °C.

[]{#CleaningUp .anchor}

**2.8 CLEANING UP**

> The glass ISO tubes are cleaned by soaking them overnight in a tank of
> room temperature chromic acid cleaning solution (e.g., Fisher
> Catalogue No. SC88-212). Use extreme caution when handling chromic
> acid, as it can cause severe burns. Flush the ISO tubes thoroughly
> with double-distilled water and dry by air aspiration.

[]{#Chapter3Dalt .anchor}**3 THE DALT SYSTEM SECOND-DIMENSION
SEPARATION**

**3.1 CASTING DALT GELS**

Be sure that the entire gel casting system (comprised of a 2 l
double-chamber gradient maker, stirring box, DALT casting box,
peristaltic pump, and vacuum pump) is clean, dry, and free of
polymerized acrylamide.

> 1\. Place the bottom plate with an angle edge in the bottom of the
> casting box so that the angle is up and toward the solution entry port
> on the right side of the box.

2.  Place a Teflon sheet to serve as a spacer at the back of the box.
    Stack gel plates into the casting box with the red hinges to the
    left and vertical, interspersing three "bubble wrap" packing squares
    cut the same size as the plates between every fifth or sixth plate.
    Screw on the front cover of the box.

> 3\. Cut and place gel serial numbers (printed on #1 filter paper) in
> order in front of the gel-casting box.
>
> 4\. Turn on the aspirator. Run the [blue underlay
> solution](#Recipe13SucroseUnderlay) to the T in the feed line and
> clamp with a hemostat. Shut off the vacuum line and make sure that the
> clamp to the vacuum line is closed. Be sure that the gradient-maker
> lines are clamped off, with one clamp on the heavy (18%) line just as
> it leaves the gradient maker and the other on the main line just
> beyond the mixer. Set the gel-casting chamber to a 45° angle from
> vertical.
>
> **NOTE:** There will be about 15 min available to pour the plates
> between the addition of the ammonium persulfate and TEMED and when the
> gels begin to polymerize, so work fast!

6.  Make the appropriate acrylamide working solutions (a 9% solution and
    an 18% solution) from the stock mixes according to the following
    table:

+-----------+-------------+------+------+--------+-----------+-------+
|           | [Acryl      | [Buf | [Buf | 10%    | [10%](    | TEMED |
|           | amide/bis]( | fer] | fer] | SDS    | #Recipe6A |       |
|           | #Recipe9_27 | (#Re | (#Re |        | mmoniumPe | (μL)  |
|           | Acrylamide) | cipe | cipe | (mL)   | rsulfate) |       |
|           |             | 11Bu | 12Bu |        |           |       |
|           | (mL)        | ffer | ffer |        | [(N       |       |
|           |             | L10) | L20) |        | H~4~)~2~S |       |
|           |             |      |      |        | ~2~O~8~]( |       |
|           |             | [    | [    |        | #Recipe6A |       |
|           |             | L10] | L20] |        | mmoniumPe |       |
|           |             | (#Re | (#Re |        | rsulfate) |       |
|           |             | cipe | cipe |        |           |       |
|           |             | 11Bu | 12Bu |        | (mL)      |       |
|           |             | ffer | ffer |        |           |       |
|           |             | L10) | L20) |        |           |       |
|           |             |      |      |        |           |       |
|           |             | (mL) | (mL) |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
| For 11    |             |      |      |        |           |       |
| plates:   |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
| 9%        | 80          | 160  |      | 2.4    | 4         | 70    |
| solution  |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
| 18%       | 240         |      | 120  | 3.6    | 2         | 10    |
| solution  |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
| For 22    |             |      |      |        |           |       |
| plates:   |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
| 9%        | 192         | 384  |      | 5.7    | 6         | 70    |
| solution  |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
| 18%       | 440         |      | 220  | 6.6    | 3.5       | 10    |
| solution  |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+
|           |             |      |      |        |           |       |
+-----------+-------------+------+------+--------+-----------+-------+

Degas after mixing the acrylamide and buffer L10 or L20. Then swirl the
container gently to mix while adding the ammonium persulfate and TEMED.
The amounts of persulfate and TEMED required may vary slightly with
different brands of reagents or the temperature of the laboratory.

> 6\. Pour all the light (9%) acrylamide solution into the right side of
> the gradient maker. Open the line beyond the mixer to fill the mixer
> and lines up to the gel-casting box. Add the heavy (18%) acrylamide
> solution to the left side of the gradient maker to the same level as
> the light acrylamide. Bleed the air bubbles out of the line.
>
> 7\. Remove the clamp on the 18% line and the clamp above the mixer to
> allow the solution to run into the "funnel" formed by the V in the
> casting box.
>
> 8\. Watch the level of fluid rising in the casting box, and slowly
> rotate the casting box to vertical as filling is completed. Then drop
> serial numbers into each gel at the right side. Use a level to check
> that the plates in the box are completely level.
>
> 9\. Clamp the line between the vacuum and blue sucrose underlay
> inlets. Open the line for the [blue underlay
> solution](#Recipe13SucroseUnderlay).
>
> 10\. While the underlay solution is flowing in, open the vacuum line,
> being sure that the water suction is on. Add water to both chambers of
> the gradient maker, flushing about 2 l through the system.
>
> 11\. After underlay solution has reached the appropriate level (i.e.,
> almost to the bottom of the glass plates), immediately (and very
> gently) overlay approximately 1 ml of [Photoflo
> solution](#Recipe14Photoflo) onto each gel.

12\. Adjust the underlay solution container up or down so that the blue
level remains just below the bottom of the plates without having to
clamp the line. This placement allows slightly more displacing fluid to
be drawn in as the gels polymerize.

> 13\. Cover the box with plastic wrap. Flush two more liters of water
> through the gradient maker and tubing. Drain the system and then drain
> the mechanical mixer by inverting it (the outlet tubes are at the top
> of the mixer). Allow the gels to polymerize for at least 4 hours.
>
> 14\. After the gels have polymerized, carefully disassemble the
> apparatus. Wash the plates carefully with warm water, removing excess
> acrylamide from the sides and edges of the plates. Rinse the top
> surface of each gel with tap-distilled water, and set the plates
> sideways in a dish rack (with the red hinges up) to drain the surface
> of the gels.

**3.2 ADDITION OF A STACKING GEL**

Since the ISO gel serves the stacking function in the second dimension
of 2DE, a stacking gel is not necessary. However, when the DALT gels are
used for 1DE, a stacking gel improves resolution. Prepare the [stacking
gel recipe](#Recipe15StackingGel) and quickly pipette the mix on top of
each slab gel.

**3.3 LOADING FIRST-DIMENSION GELS ONTO DALT GELS**

> 1\. If the first-dimension gels (ISO, BASO, ACIDO) gels are frozen,
> they should be moved from the −70 °C freezer to a −20 °C freezer about
> an hour before they are to be loaded. To minimize diffusion of
> proteins in the gels, thaw out 4 or 5 vials at a time in a beaker of
> warm water.
>
> 2\. Pour the first-dimension tube gel from the vial into a plastic tea
> strainer, letting the equilibration fluid drain into a beaker. Rinse
> the gel with [Buffer L](#Recipe10BufferL) diluted 1:4. Transfer the
> tube gel to the top of the DALT loading platform, and position the gel
> so that the acidic end (bottom of ISO and ACIDO gels or top of BASO
> gel) is to the left (acidic) side. Note that the razor rough-cut end
> corresponds to the bottom and the smooth tapered end corresponds to
> the top of each tube gel.
>
> 3\. Place a DALT gel plate on the front of the lectern with the red
> hinge to the right and the label in the lower right corner. Add a few
> drops of the diluted buffer L to the top of the DALT gel and then roll
> the ISO gel onto the top of the DALT gel. Smooth the ISO gel into
> position with a spatula, eliminating any air bubbles. Turn the DALT
> plate upside down on a towel to drain off any excess fluid before
> overlaying the gel with agarose.

3.  Overlay each gel with approximately 0.5 ml of [melted
    agarose](#Recipe16AgaroseOverlay), removing any air bubbles that
    form, Allow the agarose to completely solidify before placing the
    plates in the DALT tank. Molecular weight standards (e.g., BioRad or
    Pharmacia)can be incorporated into the agarose overlay.

**3.4 RUNNING THE DALT ELECTROPHORESIS SYSTEM**

Each set of gels is run in a clean DALT tank with fresh buffer. Prepare
the buffer in the tank several hours prior to the run to assure adequate
cooling (about 4 °C) prior to use.

> 1\. Weigh out the [buffer components](#Recipe17DaltTankBuffer) and mix
> in approximately 4 l of water. Rinse the tank well, and then fill the
> tank half full with water. Pour the concentrated buffer solution into
> the tank, and add water to the fill mark. Turn on the circulation
> pump. Just before the DALT gels are to be run, put the plate spacers
> back in the tank.

2\. After first-dimension gels are loaded onto the second-dimension DALT
gels, insert the DALT plates carefully between the rubber tank spacers
such that the first-dimension gel is along the left side and the red
rubber hinge is along the bottom of the tank. Use a slight left-wise
sliding motion so that the tank gasket flaps open to the left. Slide the
plates firmly to the bottom. Dipping the plates in the tank buffer first
simplifies their installation.

> **NOTE**: After the plates are in position, the buffer level should be
> even with the spacers of the plate and *not above the top of the
> plates*. Submersion will cause an electrical short circuit.
>
> 3\. Close the lid on the tank and attach the electrodes. Run the gels
> at 150 V (limited voltage) with the current limited to 0.6 A overnight
> until the blue tracking dye runs completely off the bottom of the DALT
> gels (about 16 h). These conditions minimize heat generation in the
> DALT gels.

4.  Turn off the power supply and carefully remove the DALT plates from
    the tank one at a time. Wear gloves when handling the gels.

5.  Place the plates one at a time on the unloading lectern, pry them
    open with a screwdriver, and use a razor blade or spatula to free
    the gel along the spacers. Carefully peel the gel away from the
    glass and place it in [Coomassie Blue stain
    solution](#Recipe18CoomassieBlue), [silver-stain fixing
    solution](#SilverStainFix), or [transfer buffer](#TransferBuffer).

```{=html}
<!-- -->
```
7.  Place the box of gels (10 per box for Coomassie blue, four per box
    for silver stain) on a shaker for staining or proceed with the
    protein transfer procedure.

**3.5 CLEANING THE DALT PLATES**

Soak the used plates in distilled water with a small amount of SDS. When
cleaning, go over all surfaces (including the edges) with a Teflon
scrubber. Finally, rinse the plates with water (and, if desired,
possibly ethanol). Air-dry them in the open position in a drying rack or
stand them on a clean surface in an inverted "v" (**Λ**) configuration.
Do not open the plate books beyond a right angle or the bindings will
break.

**3.6 DRAINING THE DALT TANKS**

Pump the buffer down the laboratory drain or into liquid waste
containers (e.g. after analysis of radioactive samples). Rinse the tank
with distilled water, drain, and leave to dry until the next use. This
procedure minimizes bacterial growth in the tanks, tubes, and
circulation pumps.

[]{#Chapter4ProteinDetection .anchor}**4 PROTEIN DETECTION METHODS**

[]{#Chapter4_1CoomassieStaining .anchor}**4.1 COOMASSIE BLUE STAIN**

> 1\. After removing DALT gels from their plates, place them into
> [Coomassie blue stain](#Recipe18CoomassieBlue) (100 ml per gel,
> maximum of 10 gels per box) at room temperature and shake overnight.
>
> 2\. Destain the gels in 20% ethanol (100 ml per gel, maximum of 10
> gels per box) for at least 1 h five times, shaking the gels in the
> final destain overnight.
>
> 3\. One hour before scanning or photography, place the gels in
> distilled water (100 ml per gel, maximum of 10 gels per box) at room
> temperature with shaking. Wipe off any stain residue from both
> surfaces of the gels before scanning or photographing.
>
> 4\. Scan or photograph using a green filter (e.g., Ratten #58).

[]{#Chapter4_2SilverStaining .anchor}**4.2 SILVER STAINING**

The method described below is a modification of the techniques described
in [Guevara et al. 1982](#ReferenceGuevara) and [Giometti et al.
1991](#ReferenceGiomettiSilver). Optimal results are achieved when gels
are stained directly with silver, although gels previously stained with
Coomassie blue can be subsequently silver-stained (backgrounds are
darker). Note that "ethanol" refers to 95% 190 proof ethanol.

> []{#SilverStainFix .anchor}1. After removing DALT gels from their
> plates, place them (maximum of four gels per box) in 50% ethanol
> containing 1% (v/v) acetic acid and 0.1% (v/v) formaldehyde (250 ml
> per gel), and shake at room temperature for a minimum of 6 h.
>
> 2\. Drain the solution and replace with 50% ethanol (250 ml per gel,
> maximum of 4 gels per box) and continue to shake at room temperature
> overnight.
>
> 3\. Drain the gels and rinse in 20% ethanol (250 ml per gel, maximum
> of 4 gels per box) for 30 min. Rinse an additional 30 min in 20%
> ethanol containing 5 mg/l of dithiothreitol (250 ml per gel, maximum
> of 4 gels per box).
>
> 4\. For a box containing four gels, mix 940 ml of 20% ethanol with 1.4
> ml of 10 N NaOH. In another beaker, dissolve 4.0 g AgNO~3~ in 50 ml
> water. Immediately before use, mix 10.5 ml of NH~4~OH in the ethanol
> solution. Then with the ethanol solution stirring, add the silver
> solution. A brown precipitate will form, but should dissolve quickly
> as the solution is stirred. If the precipitate does not clear, discard
> the solution and start over. Drain the gels and add the silver
> solution. Shake at room temperature for 1 h.
>
> 5\. Drain the silver solution and wash the gels three times with 20%
> ethanol (250 ml per gel, maximum of 4 gels per box), 20 min per wash.
>
> 6\. After draining the final ethanol wash, develop the gels for 10 min
> at room temperature with shaking by adding a solution containing 50 mg
> citric acid and 0.5 ml for­maldehyde in 1 l of 20% ethanol (for four
> gels). Gently rearrange the gels in the box to ensure even
> development.
>
> 7\. Drain the developer solution and add 0.5% acetic acid in water
> (250 ml per gel) to stop the development. Shake at room temperature
> for five minutes.
>
> 8\. Drain the stop solution and wash the gels three times at room
> temperature with tap-distilled water (250 ml per gel), 20 min per
> wash.

9.  Scan or photograph the gels after the final 20 min wash.

[]{#Chapter4Autoradiography .anchor}**4.3 AUTORADIOGRAPHY**

1\. Stain gels with [Coomassie Blue](#Chapter4_1CoomassieStaining) and
destain with two 1 h washes in 20% ethanol (100 ml per gel) at room
temperature.

2.  Shake destained gels in 2% glycerol for 30 min, changing the
    glycerol solution once after 15 min.

3.  Dry the gels onto filter paper using commercially available gel
    driers (e.g., BioRad).

4.  Label each dried gel with the appropriate number using labeling
    products such as Radtape Plus phosphorescent adhesive labels
    (Diversified Biotech, Boston, MA; Catalogue No. ADP-200).

5.  Place the dried gel on film (e.g., Kodak XAR-2 film \[Catalogue No.
    165 1579\], which gives a light gray background, or Fuji RX film
    \[Fisher Catalogue No. 04-441-95\], which gives a clear light blue
    background. Store the gels on film, flattened by weights or in
    cassettes, for 1-10 days, depending on the activity on the gels and
    the isotope used.

6.  Develop the autoradiographs (e.g., Kodak's XOMAT Model M35A
    automated film processor).

[]{#Chapter5Transfers .anchor}**5 TRANSFER OF PROTEINS FROM 2DE GELS TO
MEMBRANES**

Proteins can be transferred out of 2DE gels onto membrane supports for
further analysis such as immunoblotting (Western blots) or
amino-terminal amino acid sequencing. The membranes of choice are
nitrocellulose or polyvinylidene difluoride (PVDF). The following
methods are used for PVDF membranes ([Matsudeira
1987](#ReferenceMatsudeira)) (e.g., Immobilon-P \[Millipore Corporation,
Bedford, Massachusetts; Catalogue No. IPVH 151 50; 15 ×15 cm\] for
Western blots; Immobilon-P^SQ^ \[Catalogue No. ISEQ 151 50; 15×15 cm\]
for protein sequencing). The proteins bound to PVDF are stable for an
indefinite time on dry membranes stored at −20 °C.

> []{#TransferBuffer .anchor}1. Remove DALT gels from plates and
> equilibrate in a solution containing 10 mM
> 3‑(cyclohexylamino)-1-propanesulfonic acid [(CAPS) transfer
> buffer](#Recipe26CAPS) (pH 11.0) and 10% methanol (100 ml per gel,
> maximum of 6 gels per box) for 5−10 min.
>
> 2\. Using a fine-tip felt pen, number the dry PVDF membranes to
> correspond to gel numbers, wet the membranes in 100% methanol for a
> few seconds, and place them in CAPS transfer buffer. Wet several
> squares of filter paper (3 pieces per gel) (e.g., Gel Blot paper
> \[Midwest Scientific, Valley Park, Missouri; Catalogue No.
> 3MWO-1616\]) in another box containing the CAPS transfer buffer.
>
> 3\. Trim each gel to the size of the membrane, and place the gels in
> fresh CAPS buffer, with the proper orientation, as the identification
> number will be cut off the gel.
>
> 4\. Using a semi-dry electroblotting apparatus, place the Mylar^^
> mask (with an opening the size of the gels) over the anode electrode
> surface. Then place three pieces of the wet filter paper squares on
> the bottom of the electroblotting apparatus. Next put a PVDF membrane
> (with the number in the lower right corner) on the filter paper.
> Finally, place the gel, properly oriented, on top of the membrane. To
> transfer proteins from two gels (the maximum allowed per apparatus for
> optimal results), add another piece of filter paper, then PVDF
> membrane, and, finally, another gel.
>
> 5\. Place two more pieces of wet filter paper on the top of the last
> gel, place the lid on the apparatus, and turn on the power supply and
> run for 1.5−2.5 h at 7−8 V.
>
> 6\. For Western Blots, place the membranes in a non-reactive blocking
> solution (e.g., [3% BSA](#Recipe20BSA) or [3% nonfat dry
> milk](#Recipe21MilkBlockSoln) or [1% chemiluminescence blocking
> solution](#Recipe29ChemilumBlockSolution)) and leave them overnight in
> the refrigerator.

8.  For detection of proteins by Coomassie Blue, (e.g., Immobilon-P^SQ^
    membranes for sequencing),

    -   Rinse the membranes briefly in double-distilled water and then
        saturate the membranes in 100% methanol for a few seconds.

    -   Place the membranes in a solution of stain containing 0.1%
        Coomassie Blue R-250, 40% methanol, and 1% acetic acid
        (approximately 100 ml per two membranes) for 1-2 min.

    -   Destain the membranes in two to three changes of 50% methanol.

    -   Rinse the membranes with water, and hang to dry.

[]{#Chapter6WesternBlots .anchor}**6. WESTERN BLOTS**

[]{#Chapter6_1HRPColorDevelopment .anchor}**6.1 HRP COLOR DEVELOPMENT**

> 1\. If the membrane has been dried prior to staining, rewet in
> 100% methanol, rinse in water, and equilibrate in
> [Tris-saline](#Recipe23TrisSaline) for about 20 min. Otherwise, pour
> off the BSA blocking solution and use it to dilute the primary
> antibody (1:100 or greater). Shake the membrane in 50 ml of diluted
> primary antibody at room temperature for 1 hour.
>
> 2\. Pour off the antibody (can store frozen at −20 °C) and rinse the
> transfer membrane with [Tris-saline](#Recipe23TrisSaline) solution
> five times to wash off unbound antibody.
>
> 3\. Shake the membrane in 50 ml of the appropriate
> peroxidase-conjugated secondary antibody (e.g., if the primary
> antibody was rabbit anti-human, then use goat anti-rabbit
> IgG-peroxidase) at room temperature for 1 hour.
>
> 4\. Discard the secondary antibody solution and rinse the membrane
> five times with Tris-saline solution to remove excess secondary
> antibody.
>
> 5\. Immediately before use, prepare the following two solutions
> (recipe for two to three transfers):

-   60 mg of 4-chloro-1-naphthol (HRP color development reagent) (e.g.,
    BioRad, Catalogue No. 170-6534 or Sigma, Catalogue No. C6788) in 20
    ml of methanol.

-   60 μl of 30% H~2~O~2~ in 100 ml of cold 20 mM Tris/saline.

> Mix the hydrogen peroxide (H~2~O~2~) solution with the color
> development reagent at room temperature and immediately pour the
> mixture over the membrane. Agitate until staining is adequate (blue
> spots with white or light blue background).
>
> 6\. Rinse the stained membranes well with tap-distilled water to stop
> the reaction.

[]{#Chapter6_2Chemiluminescence .anchor}

**6.2. CHEMILUMINESCENCE**

1.  If the membrane has been dried prior to staining, rewet in
    100% methanol, rinse in water, and equilibrate in [1% blocking
    solution](#Recipe29ChemilumBlockSolution). Rinse the blocked
    membrane twice with [Tris-buffered saline
    (TBS)](#Recipe27TBSChemilum). Shake the membrane for 1 hour with 50
    ml of primary antibody diluted 1:100 or greater in 0.5 % blocking
    solution or to about 4 μg/mL.

2.  Pour off the antibody and wash the membrane twice in
    [TBS-Tween20](#Recipe28TBSTween), 10 min each, and then wash twice
    with 0.5 % blocking solution for 10 min each.

3.  Shake the membrane for 30 min at room temperature with
    peroxidase-conjugated secondary antibody diluted 1:1000 in 0.5 %
    blocking solution.

4.  Pour off the antibody and wash the membrane 4 times with large
    volumes of TBST for 15 min each.

5.  Drain off the last TBST wash, and add premixed [detection
    reagent](#Recipe30DetectionSolutionChemilum). Incubate at room
    temperature for \~60 seconds. Drain off excess detection reagent and
    wrap the blot in plastic wrap.

6.  Insert the membrane, protein side up, into a film cassette. In the
    dark, add a sheet of X-ray film on top of the blot and close the
    cassette. Expose for 10−60 seconds.

7.  Immediately replace the exposed film in the cassette with a new one
    to expose for a shorter or longer time, and develop the exposed
    film. An optimized exposure can be done based on the signal
    intensity of the first two films.

[]{#Chapter7References .anchor}**7. REFERENCES**

[]{#ReferenceAndersons .anchor}Anderson, N.G., and N.L. Anderson, 1978a.
Analytical techniques for cell fractions. XXI. Two-dimensional analysis
of serum and tissue proteins: multiple isoelectric focusing, *Analytical
Biochemistry* 85: 331−340.

[]{#ReferenceAndersonsUrine .anchor}

Anderson, N.L., and N.G. Anderson, 1978b. Analytical techniques for cell
fractions. XXII. Two-dimensional analysis of serum and tissue proteins:
multiple gradient-slab electrophoresis, *Analytical Biochemistry* 85:
341−354.

Anderson, N.G., N.L. Anderson, and S.L. Tollaksen, 1979. Proteins of
human urine. I. Concentration and analysis of two-dimensional
electrophoresis, *Clinical Chemistry* 25: 1199−1210.

[]{#ReferenceEdwardsUrine .anchor}Edwards, J.J., S.L. Tollaksen, and
N.G. Anderson, 1982. Proteins of human urine. III. Identification and
two-dimensional electrophoretic map positions of some major urinary
proteins, *Clinical Chemistry* 28: 941-948.

[]{#ReferenceGiomettiSilver .anchor}Giometti, C.S., M.A. Gemmell, S.L.
Tollaksen, J. Taylor, 1991. Quantitation of human leukocyte proteins
after silver staining: A study with two-dimensional electrophoresis,
*Electrophoresis* 12: 536-543.

[]{#ReferenceGuevara .anchor}Guevara, J., S. Capetillo, D.A. Johnston,
B.A. Martin, L.S. Ramagli, and L.V. Rodriguez, 1982. Quantitative
aspects of silver deposition in proteins resolved in complex
polyacrylamide gels, *Electrophoresis* 3: 197−205.

[]{#ReferenceMatsudeira .anchor}Matsudeira, P., 1987. Sequence from
picomole quantities of proteins electroblotted onto polyvinylidene
difluoride membranes, *Journal of Biological Chemistry* 262:
10035−10038.

[]{#ReferenceOFarrellFirstPaper .anchor}O'Farrell, P.H., 1975. High
resolution of two-dimensional electrophoresis of proteins, *Journal of
Biological Chemistry* 250: 4007-4021.

[]{#ReferenceOFarrell .anchor}O'Farrell, P.Z., Goodman, H.M., and
O'Farrell, P.H., 1977. High resolution two-dimensional electrophoresis
of basic as well as acidic proteins, *Cell* 12: 1133-1142.

[]{#ReferenceRamagliModifiedProtein .anchor}Ramagli, L.S. and L.V.
Rodriguez, 1985. Quantitation of microgram amounts of protein in
two-dimensional polyacrylamide gel electrophoresis sample buffer,
*Electrophoresis* 6: 559−563.

[]{#Chapter8Recipes .anchor}**8. RECIPES**

[]{#Recipe1BlueMix .anchor}**Recipe 1**. SDS Mix (solubilizing agent)

  ----------------------------------------- -------------- --------------
  Compound                                  Amount         Final Conc.

  CHES (2-\[*N*-cyclohexylamino\]ethane     1 g            0.5 M
  sulfonic acid)                                           

  SDS (sodium dodecyl sulfate)              2 g            2%

  DTT (dithiothreitol)                      1 g            1%

  Glycerol                                  10 mL          10%
  ----------------------------------------- -------------- --------------

Adjust pH to 9.5 with NaOH and add water to a final volume of 100 mL.

[]{#Recipe2PinkMix .anchor}**Recipe 2.** NP-40/Urea Mix (solubilizing
agent)

+----------------------------------------+--------------+--------------+
| Compound                               | Amount       | Final Conc.  |
+========================================+==============+==============+
| Urea                                   | 54 g         | 9 M          |
+----------------------------------------+--------------+--------------+
| Nonidet P-40                           | 4 mL         | 4%           |
+----------------------------------------+--------------+--------------+
| Ampholyte, 8−10 range                  | 10 ml        | 2%           |
|                                        |              |              |
| (20% w/v stock)                        |              |              |
+----------------------------------------+--------------+--------------+
| 2-mercaptoethanol                      | 2 mL         | 2%           |
+----------------------------------------+--------------+--------------+

Adjust pH to 9.5 with NaOH and add water to a final volume of
100 mL.[]{#Recipe3YellowMix .anchor}

**Recipe 3.** Urea Mix without NP-40 (solubilizing agent)

+----------------------------------------+--------------+--------------+
| Compound                               | Amount       | Final Conc.  |
+========================================+==============+==============+
| Urea                                   | 54 g         | 9 M          |
+----------------------------------------+--------------+--------------+
| Ampholyte, pH 3.5−10 range             | 5 mL         | 2%           |
|                                        |              |              |
| (20% w/v stock)                        |              |              |
+----------------------------------------+--------------+--------------+
| 2-mercaptoethanol                      | 5 mL         | 5%           |
+----------------------------------------+--------------+--------------+

Adjust pH to 9.5 with NaOH and add water to a final volume of 100 mL.

[]{#Recipe4GreenMix .anchor}**Recipe 4.** NP-40/Urea/DTE Mix
(solubilizing agent for muscle samples)

+----------------------------------------+--------------+--------------+
| Compound                               | Amount       | Final Conc.  |
+========================================+==============+==============+
| Urea                                   | 54 g         | 9 M          |
+----------------------------------------+--------------+--------------+
| Nonidet P-40                           | 4 mL         | 4%           |
+----------------------------------------+--------------+--------------+
| Ampholyte, 3.5−10 range                | 5 mL         | 2%           |
|                                        |              |              |
| (40% w/v stock)                        |              |              |
+----------------------------------------+--------------+--------------+
| DTE (dithioerythritol)                 | 1 g          | 1%           |
+----------------------------------------+--------------+--------------+

Adjust pH to 9.5 with NaOH and add water to a final volume of 100 mL.

[]{#Recipe5_30Acrylamide .anchor}**Recipe 5.** 30% Acrylamide/1.8% bis
(for first-dimension and stacking gels)

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Acrylamide (Bio-Rad)                      30 g           30%

  *N,N,′′*-methylene-bis-acrylamide      1.8 g          1.8%
  -----------------------------------------------------------------------

> Add water to 100 mL. Filter with a 115-ml side-arm filter unit
> attached to the vacuum pump and store the solution in the refrigerator
> in a bottle marked "Danger: Cancer Hazard".

[]{#Recipe6AmmoniumPersulfate .anchor}

**Recipe 6.** 10% Ammonium persulfate

-   Dissolve 10 g of ammonium persulfate in 100 ml of water.

```{=html}
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```
-   Store in the refrigerator in a brown bottle, and use the solution
    within one week.

[]{#Recipe7BufferII .anchor}

**Recipe 7.** Buffer II (recipe for 375 mL)

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Trizma base                               11.25 g        3%

  SDS                                       0.75 g         0.2%
  -----------------------------------------------------------------------

Add 300 ml water. Stir chemicals and add concentrated HCl until pH is
6.8.

Bring volume up to 375 mL.

[]{#Recipe8EquilibrationBuffer .anchor}**Recipe 8.** Equilibration
buffer (recipe for 750 mL)

+----------------------------------------+--------------+--------------+
| Compound                               | Amount       | Final Conc.  |
+========================================+==============+==============+
| Glycerol                               | 75 mL        | 10%          |
+----------------------------------------+--------------+--------------+
| [Buffer II](#Recipe7BufferII)          | 375 mL       | 50%          |
+----------------------------------------+--------------+--------------+
| SDS                                    | 15 g         | 2%           |
+----------------------------------------+--------------+--------------+
| Dithioerythritol                       | 1.0 g        | 5%           |
|                                        |              |              |
| or                                     | or           |              |
|                                        |              |              |
| Dithiothreitol                         | 1.0 g        |              |
|                                        |              |              |
| or                                     | or           |              |
|                                        |              |              |
| 2-mercaptoethanol                      | 37.5 mL      |              |
+----------------------------------------+--------------+--------------+
| Bromophenol blue                       | Trace        | Trace        |
+----------------------------------------+--------------+--------------+

Add 300 ml water. Adjust pH to 6.8. Mix the above ingredients in a
convenient dispenser, such as the Repipet Jr. (Fisher Catalogue
No. 13-687-59A), and stir for approximately 1 hour.

[]{#Recipe9_27Acrylamide .anchor}**Recipe 9.** 27% acrylamide/0.8% bis
(for DALT gels)

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Acrylamide                                300 g          27%

  Bis                                       8 g            0.8%
  -----------------------------------------------------------------------

Mix on a magnetic stirrer and bring up to 1.1 l with water. Filter.

If you prefer to use liquid acrylamide instead of working with the
powder, use:

Compound Amount

  ----------------------------------------- ---------- ------------------
  Acryl/bis 29:1 ratio                      500 mL     

  Acryl/bis 37.5:1 ratio                    500 mL     
  ----------------------------------------- ---------- ------------------

Final concentration will be 33.25% acrylamide and 1.0% bis.

Note: Amresco, BioRad, and Sigma all sell liquid acrylamide and bis.

[]{#Recipe10BufferL .anchor}**Recipe 10.** Buffer L (recipe for 3 L)

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Trizma base                               400 g          13.3%

  Trizma HCl                                200 g          6.7%
  -----------------------------------------------------------------------

> Combine and add about 2 l of water. Add HCl until pH 8.5−8.6 is
> reached
>
> (approximately 120 ml of 12N HCl). Add water up to 3 L.

[]{#Recipe11BufferL10 .anchor}**Recipe 11.** Buffer L10

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  [Buffer L](#Recipe10BufferL)              3 parts        37.5%

  Water                                     5 parts        62.5%
  -----------------------------------------------------------------------

[]{#Recipe12BufferL20 .anchor}

**Recipe 12.** Buffer L20

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  [Buffer L](#Recipe10BufferL)              3 parts        75%

  Glycerol                                  1 part         25%
  -----------------------------------------------------------------------

[]{#Recipe13SucroseUnderlay .anchor}**Recipe 13.** Sucrose underlay
solution

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Sucrose                                   35 g           35%

  Water                                     100 mL         

  Methylene blue                            Trace          Trace
  -----------------------------------------------------------------------

[]{#Recipe14Photoflo .anchor}**Recipe 14.** Photoflo overlay solution

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Buffer L10                                67 mL          66.7%

  Water                                     33 mL          33.3%

  Photoflo (or similar surfactant)          0.1 mL         0.1%
  -----------------------------------------------------------------------

[]{#Recipe15StackingGel .anchor}**Recipe 15.** Stacking gel mix

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  [Buffer II](#Recipe7BufferII)             480 mL         48%

  Water                                     380 mL         \-

  [30% acrylamide/1.8% bis                  140 mL         14%
  solution](#Recipe5_30Acrylamide),                        
  -----------------------------------------------------------------------

[]{#Recipe16AgaroseOverlay .anchor}**Recipe 16.** Running buffer Agarose

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Trizma base                               3 g            0.3%

  Glycine                                   14.4 g         1.44%

  SDS                                       1 g            0.1%

  Agarose                                   5 g            0.5%
  -----------------------------------------------------------------------

> Add water to 1 L. Microwave to dissolve. Freeze in 100 ml portions.

**NOTE**: Agarose solutions often superheat when they are microwaved,
boiling over when disturbed, e.g., when a pipette is introduced, and
serious injury can result. Use extreme caution when handling heated
agarose.

[]{#Recipe17DaltTankBuffer .anchor}**Recipe 17.** DALT buffer (26 L)

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Tris                                      78 g           24 mM

  Glycine                                   374 g          0.2 M

  SDS                                       26 g           3.5 mM
  -----------------------------------------------------------------------

> Add approximately 4 l distilled water and stir with a magnetic stirrer
> until dissolved. Add to the clean DALT tank and add distilled water to
> the fill mark.

[]{#Recipe18CoomassieBlue .anchor}**Recipe 18.** Coomassie Blue Stain

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  Coomassie Blue                            20 g           0.2%

  Double-distilled water                    4.5 L          \-

  Phosphoric acid                           250 mL         2.5%

  Ethanol (95%)                             5 L            47.5%
  -----------------------------------------------------------------------

> Mix the stain and water. Add 250 ml phosphoric acid. Add 5 l 95%
> ethanol and mix thoroughly for at least an hour. Let the solution
> stand overnight before use.

[]{#Recipe19Saline .anchor}**Recipe 19.** Saline solution

  -----------------------------------------------------------------------
  Compound                                  Amount         Final Conc.
  ----------------------------------------- -------------- --------------
  NaCl                                      90 g           150 mM

  Water                                     10 L           \-
  -----------------------------------------------------------------------

[]{#Recipe20BSA .anchor}**Recipe 20.** Bovine serum albumin blocking
solution

Compound Amount Final Conc.

  ----------------------------------------- -------------- --------------
  [Tris buffer (1 M, pH                     0.5 mL         10 mM
  7.5)](#Recipe22_1MTris)                                  

  30% Bovine serum albumin solution         5 mL           3 %

  [Saline](#Recipe19Saline)                 44.5mL         150 mM
  ----------------------------------------- -------------- --------------

**NOTE:** Use sterile procedures when handling bovine serum albumin
(BSA) to minimize bacterial contamination.

[]{#Recipe21MilkBlockSoln .anchor}

**Recipe 21.** Nonfat dry milk blocking solution

Compound Amount Final Conc.

  --------------------------------------- ---------------- ---------------
  Powdered milk                           3 g              3%

  [Tris buffer (1 M, pH                   1 mL             10 mM
  7.5)](#Recipe22_1MTris)                                  

  Saline                                  99 mL            150 mM
  --------------------------------------- ---------------- ---------------

[]{#Recipe22_1MTris .anchor}**Recipe 22.** 1 M Tris buffer

> Dissolve 12.1 g of Tris in water; adjust pH to 7.5, and make a final
> volume of 100 mL.

[]{#Recipe23TrisSaline .anchor}**Recipe 23.** Tris-saline solution

Add 1 ml of [1 M Tris, pH 7.5](#Recipe22_1MTris) to 100 ml of
[saline](#Recipe19Saline) solution.

[]{#Recipe24RehydrationBuffer .anchor}**Recipe 24.** Rehydration buffer
(for IPGs)

Compound Amount Final Conc.

  --------------------------------------- ------------ -------------------
  Urea                                    48.04 g      8 M

  SDS                                     2.0 g        2%

  DTT                                     772 mg       50 mM

  Ampholyte                               0.5 mL       0.2%

  Bromophenol blue                        Trace        0.001%

  H~2~O                                   To 100 mL    
  --------------------------------------- ------------ -------------------

[]{#Recipe25EquilibrationBuffer .anchor}

**Recipe 25.** Equilibration Buffer

Compound Amount Final Conc.

  --------------------------------------- ------------ -------------------
  Urea                                    36.036 g     6 M

  SDS                                     2.0 g        2%

  Tris Base                               4.54 g       0.0375 M

  Glycerol                                20 mL        20 %

  HCl                                                  To pH 8.8

  H~2~O                                   To 100 ml    
  --------------------------------------- ------------ -------------------

-   For DTT Equilibration Buffer, add 200 mg DTT/10 ml Equilibration
    Buffer just before use.

-   For Iodoacetamide Equilibration Buffer, add 250 mg Iodoacetamide/10
    ml Equilibration Buffer just before use.

[]{#Recipe26CAPS .anchor}**Recipe 26.**
3-(cyclohexylamino)-1-propanesulfonic acid (CAPS) buffer in 10 %
methanol,

pH 11.0

-   Stir 2.2 g CAPS in 800 ml double-distilled water.

-   Add 100 ml methanol.

> The pH will be \~ 5.5. Adjust pH to11.0 with 10 N NaCl, and add water
> to 1 L.

[]{#Recipe27TBSChemilum .anchor}**Recipe 27.** Tris buffered saline
(TBS), pH 7.5, for chemiluminescence detection

> Add 5 ml of [1 M Tris](#Recipe22_1MTris), pH7.5, to 995 ml
> [saline](#Recipe19Saline).

[]{#Recipe28TBSTween .anchor}**Recipe 28.** TBS-Tween^^ 20 (TBST)

Dilute 1 ml of Tween^^ 20 in 1 l TBS (final concentration 0.1%
\[w/v\]).

[]{#Recipe29ChemilumBlockSolution .anchor}

**Recipe 29.** 1% chemiluminescence blocking solution

Add 10 ml of blocking reagent concentrated solution to 90 ml of TBS.

[]{#Recipe30DetectionSolutionChemilum .anchor}**Recipe 30.** Detection
solution for chemiluminescence (as per directions included in the kit)

Mix substrate solution A and starting solution B in a ratio of 100:1.