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, 1978b), 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 and very basic proteins, in addition to the classical use of isoelectric focusing 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 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 (body fluids such as serum, plasma, or amniotic fluid). To a 10 ml sample, add 20-30 ml 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 (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 ml 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 ´ 10⁶ cells per 50 µl of mix when Coomassie blue detection will be used. When silver‑stain will be used, use 5 ´ 10⁵ 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 (urine). Mix 10 mg of lyophilized urinary proteins with 100 ml of the mix. Load 10 ml (or less) of the solution onto the ISO gel.

4. NP-40/urea/DTE mix (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 ml 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‑[³⁵S]‑methionine (approximately 50 mCi per sample) or ³²P‑orthophosphate (200 mCi 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 ml of NP40-urea mix 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). The optimal protein load for Coomassie blue staining of whole cell lysates is 150-300 mg protein, while 20-60 mg of protein is recommended for silver-stained gels.

CHAPTER 2: THE ISO SYSTEM FOR FIRST-DIMENSION SEPARATION

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.

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
Ampholytes (the majority of ANL isoelectric focusing gels are 50:50 pH 5-7:pH 3-10; P. furiosus and S. oneidensis gels are an exception)	0.8 mL
Double-distilled water	6.0 mL

Compound

Amount

Urea

8.25 g

Acrylamide (30% solution)

2.0 mL

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

0.8 mL

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% solution)

50 mL

TEMED (N,N,N’,N’

-tetramethylethylenediamine; an accelerator)

5 mL

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. 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 ml 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. 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 ml 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 mg for Coomassie blue detection or 20-60 mg 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 ml 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.

2.4 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 ml (7 cm IPG strip) or 300-600 ml (17 cm IPG strip) of the rehydration buffer 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 ml (5-100 mg 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 for 10 min and then with iodoacetamide-equilibration buffer for an additional 10 min. After equilibration, IPG strips are placed on top of the second dimension slab gels.

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).

1. The BASO gels are cast in the 7-in. ISO apparatus using the same procedure as for ISOs. 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 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 H₃PO_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.

2.6 ACIDO ISOELECTRIC FOCUSING

Urine contains very acidic proteins (most acid urinary protein [MAUP] and a-1-acid glycoprotein) that will not focus following the ISO protocol for isoelectric focusing gels (Edwards, Tollaksen, and Anderson 1982). 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
Ampholyte (pH 3-10)	0.3 mL
Water	5.5 mL

Compound

Amount

Urea

8.25 g

Acrylamide (30% stock solution)

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% solution)

90 mL

TEMED

10 mL

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

Clean the gel tubes as described for prefocusing ISO gels. Empty the bottom chamber and fill with 2 l of water and 3 ml of concentrated H₂SO_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.

Prefocus the setup for 1 h as described.

Underlay approximately 20 ml 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 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 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.

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.

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 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:

	Acrylamide/bis (mL)	Buffer L10 (mL)	Buffer L20 (mL)	10% SDS (mL)	10% (NH₄)₂S₂O₈ (mL)	TEMED (mL)
For 11 plates:
9% solution	80	160		2.4	4	70
18% solution	240		120	3.6	2	10
For 22 plates:
9% solution	192	384		5.7	6	70
18% solution	440		220	6.6	3.5	10

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.

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 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 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 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, 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 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, silver-stain fixing solution, or transfer buffer.

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” (L) 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.

4 PROTEIN DETECTION METHODS

4.1 COOMASSIE BLUE STAIN

1. After removing DALT gels from their plates, place them into Coomassie blue stain (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).

4.2 SILVER STAINING

The method described below is a modification of the techniques described in Guevara et al. 1982 and Giometti et al. 1991. 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.

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 AgNO₃ in 50 ml water. Immediately before use, mix 10.5 ml of NH₄OH 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.

4.3 AUTORADIOGRAPHY

1. Stain gels with Coomassie Blue 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).

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) (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.

1. Remove DALT gels from plates and equilibrate in a solution containing 10 mM 3‑(cyclohexylamino)-1-propanesulfonic acid (CAPS) transfer buffer (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^Ò 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 or 3% nonfat dry milk or 1% chemiluminescence blocking solution) and leave them overnight in the refrigerator.

8. For detection of proteins by Coomassie Blue, (e.g., Immobilon-P^SQmembranes 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.

6. WESTERN BLOTS

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 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 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 ml of 30% H₂O₂ in 100 ml of cold 20 mM Tris/saline.

Mix the hydrogen peroxide (H₂O₂) 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.

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. Rinse the blocked membrane twice with Tris-buffered saline (TBS). 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 mg/mL.

2. Pour off the antibody and wash the membrane twice in TBS-Tween20, 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. 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.

7. REFERENCES

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.

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.

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.

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.

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.

Matsudeira, P., 1987. Sequence from picomole quantities of proteins electroblotted onto polyvinylidene difluoride membranes, Journal of Biological Chemistry 262: 10035-10038.

O’Farrell, P.H., 1975. High resolution of two-dimensional electrophoresis of proteins, Journal of Biological Chemistry 250: 4007-4021.

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.

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.

8. RECIPES

Recipe 1. SDS Mix (solubilizing agent)

Compound

Amount

Final Conc.

CHES (2-[N-cyclohexylamino]ethane sulfonic acid)

1 g

0.5 M

SDS (sodium dodecyl sulfate)

2 g

DTT (dithiothreitol)

1 g

Glycerol

10 mL

10%

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

Recipe 2. NP-40/Urea Mix (solubilizing agent)

Compound	Amount	Final Conc.
2-mercaptoethanol	2 mL	2%

Compound

Amount

Final Conc.

Urea

54 g

9 M

Nonidet P-40

4 mL

Ampholyte, 8-10 range

(20% w/v stock)

10 ml

2-mercaptoethanol

2 mL

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

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

Compound

Amount

Final Conc.

Urea

54 g

9 M

Ampholyte, pH 3.5-10 range

(20% w/v stock)

5 mL

2-mercaptoethanol

5 mL

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

Recipe 4. NP-40/Urea/DTE Mix (solubilizing agent for muscle samples)

Compound	Amount	Final Conc.
DTE (dithioerythritol)	1 g	1%

Compound

Amount

Final Conc.

Urea

54 g

9 M

Nonidet P-40

4 mL

Ampholyte, 3.5-10 range

(40% w/v stock)

5 mL

DTE (dithioerythritol)

1 g

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

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,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”.

Recipe 6. 10% Ammonium persulfate

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

· Store in the refrigerator in a brown bottle, and use the solution within one week.

Recipe 7. Buffer II (recipe for 375 mL)

Compound

Amount

Final Conc.

Trizma base

11.25 g

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.

Recipe 8. Equilibration buffer (recipe for 750 mL)

Compound	Amount	Final Conc.
SDS	15 g	2%
Dithioerythritol or Dithiothreitol or 2-mercaptoethanol	1.0 g or 1.0 g or 37.5 mL	5%
Bromophenol blue	Trace	Trace

Compound

Amount

Final Conc.

Glycerol

75 mL

10%

Buffer II

375 mL

50%

SDS

15 g

Dithioerythritol

Dithiothreitol

2-mercaptoethanol

1.0 g

37.5 mL

Bromophenol blue

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.

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.

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.

Recipe 11. Buffer L10

Compound

Amount

Final Conc.

Buffer L

3 parts

37.5%

Water

5 parts

62.5%

Recipe 12. Buffer L20

Compound

Amount

Final Conc.

Buffer L

3 parts

75%

Glycerol

1 part

25%

Recipe 13. Sucrose underlay solution

Compound

Amount

Final Conc.

Sucrose

35 g

35%

Water

100 mL

Methylene blue

Trace

Recipe 16. Running buffer Agarose

Compound	Amount	Final Conc.
Agarose	5 g	0.5%

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.

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

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.

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.

Recipe 19. Saline solution

Compound

Amount

Final Conc.

NaCl

90 g

150 mM

Water

10 L

Recipe 20. Bovine serum albumin blocking solution

Compound Amount Final Conc.

Tris buffer (1 M, pH 7.5)

0.5 mL

10 mM

30% Bovine serum albumin solution

5 mL

3 %

Saline

44.5mL

150 mM

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

Recipe 21. Nonfat dry milk blocking solution

Compound Amount Final Conc.

Powdered milk

3 g

Tris buffer (1 M, pH 7.5)

1 mL

10 mM

Saline

99 mL

150 mM

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.

Recipe 23. Tris-saline solution

Add 1 ml of 1 M Tris, pH 7.5 to 100 ml of saline solution.

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%
H₂O	To 100 mL

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
H₂O	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.

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.

Recipe 27. Tris buffered saline (TBS), pH 7.5, for chemiluminescence detection

Add 5 ml of 1 M Tris, pH7.5, to 995 ml saline.

Recipe 28. TBS-Tween^Ò 20 (TBST)

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

Recipe 29. 1% chemiluminescence blocking solution

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

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.