2 D Electrophoresis

2nd Dimension


Contact: Hyuk-Kyu Seoh
Rm (lab): PSC 537

Rm (office): PSC 521
Tel: (404) 413-5379

Location: PSC 537


2D electrophoresis manuals:
Sample Prep
2D Cleanup
2D Quant


2nd Dimension topics:
Supply List
Plate Treatment
Ettan gel casting - 6 gels
SE600 Ruby gel casting
Equilibration

GE Amersham Manuals:
Amersham 2-D electrophoresis Manual
Ettan DIGE System User Manual


Ettan 2nd Dimension run
SE600 Ruby 2nd Dimension run
Cleaning 2nd Dimension
Buffers and Solutions

Table/Rollin - amounts for 6 gels
Table/Anupama - amounts for 1 - 6 gels


Equipment for 2nd dimension:

Ettan gel caster
Ettan DALT six system: separation unit (can hold up to 6 gels); power supply; gel casting cassettes
SE600 Ruby Vertical Unit


2nd Dimension SDS-PAGE

SDS-PAGE (SDS-polyacrylamide gel electrophoresis is a method for separating polypeptides according to their molecular weight. Instructions for pouring 2nd dimension gels follows along with information about precast gels in the Supply section.

Notes:
• We have the Ettan DALT six electrophoresis system and the SE600 Ruby electrophorsis system. 
• SE600 Ruby is very sensitive to chips on corners.  Chips on the bottom – gel leaks in the gel caster; chips on top – gel leaks at top buffer chamber because it doesn’t form a good seal.
• DALT II system can tolerate chips.  When casting, the plates are inside a contained system.  When the gels are run in the DALT II electrophoresis system, the plate sandwich slides into a rubber gasket to form the seal for the top chamber.  It would have to be a Big chip for the top chamber to leak.
• Run the gel immediately after Equilibration steps. 
• The SDS Equilibration buffer takes a while to go into solution.  Make it at least the day before you need to use it.
• Place the buffer at 4oC the night before.  Rollin and Shelby use 1X buffer for lower and upper chambers of the DALT system. The Ettan DALT six electrophoresis system manual, GE manual and Anupama uses 1X buffer for the lower chamber and 2X buffer for the upper chamber.
• You need 5 L 1X for SE600 Ruby system; 6 L 1X for DALT system using 1X in both chambers; 5 L 1X and 1 L 2X if using 1X in the bottom chamber and 2X in the top chamber.

Read the Amersham manual for details about:
• SDS –PAGE background information.
• Equilibration solution components and why
• Setting up Equilibration
• Setting up gel apparatus
• Using pre-cast gels and pouring your own gel
• Electrophoresis conditions

Key:
LB     Lysis Buffer
SB    2X Sample Buffer
RB    Rehydration Buffer
EB    Equilibration Buffer

Pour 2nd Dimension gels
• Use Ettan DALT six system and a 6 gel caster
• Only a few differences in gel pouring between 2 labs – Rollin uses a funnel to pour gel into caster. 
ALWAYS cast gels at least the day before.  There are many reasons that a gel will split when the plates are separated, but top of our list of reasons is that the gels were not cast the night before. 
• Cast the gels and let sit for 1 – 2 hours.  Separate gel sandwich, rinse with dH2O.  Rinse wells and outside of plate.  Do not use tap H2O – the 2nd D will be wavy.  Leave all gels RT o/n.  Store unused gels at 4oC for up to 1 week.  See below for how to store gels. 

Plate Treatment – SE600 Ruby and DALT
1. See below for instructions about cleaning the glass plates.  This protocol is in the Cleaning section.
2.
Treat plates with Binding solution (see Buffer and Solutions section) if you plan to use the Spot Picker.  You want the gel bound tight to the plate so the picker won’t pull up the gel. 
3.
Lay plates without vinyl spacer (Ettan six apparatus) or low fluorescent plate (SE600 Ruby apparatus) on a CrewWipe – only treat 1 side of a plate.
4. Use 4 ml of Binding Solution/Ettan plate and 3 ml for SE600 Ruby.  Spread Solution with a Crew Wipe (can use 1 Crew Wipe for all the plates).  Using a pipet, place the Binding solution in the middle of the glass plate.  Spread evenly and thoroughly and lightly with a top to bottom and then right to left motion. Let sit for a minute or so.  When you see the liquid start to pull in, spread very lightly again with Crew Wipe.  Repeat until no drops left.  If drops are allowed to dry, they can show up when you image your plate.  Cover the plate with another wipe to keep dirt or dust from falling on plate. 

5.
Let Binding solution dry on plate.  Time depends on whether the Spot Picker will be used or not.
a. If not using the Spot Picker, let plates sit for 1 1/2 hrs before casting gel. 
b. If you are using the Spot picker, you will need reference markers.  After 45 min., attach reference markers.  Be sure to place markers so they fall within the 2 white lines of the Spot Picker tray.  SE600 Ruby – place a spacer on the plate; use tweezers to place markers and press marker hard to plate; place the marker half way down the spacer and a 4 – 5 mm away from the spacer.  EttanDALT plates -  place reference markers 10.5 cm up from plate bottoms (vinyl spacers go all the way to the bottom edge at the of the plates); measure spacer width; use tweezers to place markers and press marker hard to plate 4 – 5 mm away from spacer.  Be careful not to put them too close to the edge – leave space for the spacers if not using Ettan plates.  Let plate sit another 45 min. before casting.
6.
Once dry, buff plates with Crew wipe.

Gel Casting

Ettan DALT six system Gel Casting – 6 gels
Anupama
1. 2 kinds of plastic spacers, thin ones (separator sheets) to go in between the glass plates and/or blank cassettes and thick ones (filler sheets). These sheets have square corners on 1 end and rounded corners on the other. Be sure to have the square corners on the bottom of the caster (next to the black wedge). The blank cassettes are slightly thicker than the 2 glass plates and are used to fill the caster and also fill the 2nd Dimension DALT six apparatus.
2. Lay caster bottom flat on bench. Make sure black wedge is in place.
3. Start making sandwich (following is for 2 gels). As you work, push plates to one side of the caster: 1 separator sheet (square corners down), 1 blank cassette; 1 separator sheet (square corners down), 1 blank cassette; 1 separator sheet (square corners down), 1 blank cassette; 1 separator sheet (square corners down), 1 blank cassette,1 separator sheet (square corners down), 1 plate w/ spacer, 1 top plate (treated if needed) ***, 1 separator sheet (square corners down), 1 plate w/ spacer, 1 top plate (treated if needed) ***, 1 separator sheet (square corners down). Fill the rest of the Caster with the Filler sheets. You need to fill the castor so that the plates, sheets and cassettes fit very tight when the top of the caster is on. Check sides and bottom of sandwich to make sure everything is even and same level. Make sure all plates are to 1 side of caster. Be very careful that the glass plates are pushed down and even on all 4 sides.
***When you place the 2nd plate on top of the bottom one, it tends to stick.  If you      need to even things up or move the top plate, you will need to pop the plate apart using the BioRad gel releaser tool.  Make absolutely sure that the 2 plates areflush on all 4 sides.  If the 2 plates are uneven on the sides or bottom, you can’t use it.
4. Be sure to keep track of any untreated plates if doing both treated and untreated.
5. Screw 2 black screws in, but not all the way.
6. Top of Caster has a long gray washer that was removed to rinse the Caster when done. Push back into groove. Put gel seal/grease on washer, a thin layer all the way around.
7. Turn lid over, place down onto Caster, slots at bottom that fit around the screws,push all the way down (through), make sure washer still in. Check all the way around to make sure gray washer isn’t coming out anywhere. This is a common cause of leaking.
8. Use clamps – 3 grooves on either side of caster for clamps. Place first clamp on top. Second clamp place opposite side on bottom. Place third clamp on bottom and fourth clamp on opposite side on top. Place last 2 clamps in middle. Make sure the clamps are not crooked. Small or large side on top doesn't matter.
9. Tighten screws and stand Caster up.
10. Level caster. Important – you don’t want the IPG stripto be slanted.
11. Check your plate order to make sure it is set up correctly, all to one side and a separator sheet between all plate sandwiches and/or cassette blanks.
12. See below for gel recipe tables. You don’t need to degas if using Protogel.
13. Pour acrylamide mix in groove, rest flask against plastic and pour down groove slowly.
14. Pour to level of lower notch (or to top of notch in groove) about 1 cm from top – the gel will shrink about 1 mm when polymerized.  You do not want to go to top of plate (you need room for the 1st D strips.  You also don’t want the level too low – this just makes it harder to push the 1st D strip down to the gel surface.
15. Rollin and Shelby have a large funnel setup that they feel delivers a steadier stream of acrylamide mix so there is less chance for bubbles.
16. Layer with water-saturated butyl alcohol.  Cover top of each gel.  Only need to layer gels not blanks. 
17. Leaks – if the caster starts to leak, it will be from the bottom opposite side that you are pouring, so watch for it. If it leaks, you have to take everything apart, clean again and start over. Common causes of leaking – gray gasket is pinched or slipped out of the groove somewhere, check the black clamps to make sure they’re tight, check the black clamps to make sure they’re not crooked.
18. Polymerization should take 1 - 2. Check leftover acrylamide mix in flask to see of solidified.
19. Put whole caster in a large tray, so that as you start taking it apart, the alcohol and unpolymerized acrylamide will just stay in the tray.
20. Unscrew screws and take apart.
21. Pour off alcohol and smooth away any acrylamide.
22. Rinse plates in dH2O at the sink.
23. With a razor blade, remove any excess gel from the sides of the gel.
24. Lay out a large piece of Saran Wrap and lay 1 gel on top.
25. Pipet 1X running buffer or gel storage buffer on top of gel and some on bottom to keep gel surfaces from drying out – 15 ml.
26. Wrap gel up. If you have done a mixture of treated and untreated gels, label saran wrap if gel is untreated.
27. Put a piece of foil in bottom of a tray. Lay gel flat on top of foil.
28. Place all wrapped gels in tray. Place another piece of foil on top so that the gels are covered. Put lid on tray
29. Put tray in RT o/n or 4oC if not using the next day.

SE 600 Ruby gel casting
Shelby, Suganthi, Debby and Kyu
1. Prepare the caster and clamps. Place the spirit level into the caster center and adjust the leveling feet.
2. Loosen all clamp screws and make space for the sandwich by sliding the pressure plates toward the screws.
3. Treat plates with Binding solution that you will use on the Spot Picker. You want the gel bound tight to 1 plate. We have 3 types of plates. High fluorescent (blue-green edges of plate) plates that will not be used on the Typhoon. Low Fluorescent (regular glass look to edges) plate that will be used on the Typhoon. Notched plate (gel is never bound to this plate) that is used for Club sandwich.
4. Construct gel sandwiches. For each sandwich choose two perfectly clean, unchipped glass plates and two spacers. Lay one plate on a flat surface, lay the Spacer-Mate assembly template onto the plate (wide side at the top of the plate), place a spacer along each edge, and lay the second glass plate on top. There are 2 sets of spacers available. The white ones are 1 mm and the gray ones are 1.5 mm. If you use the white spacers, you can’t use the Spacer-Mate, the white spacers are too narrow.
5. Secure the sandwich with clamps. Slide one clamp at a time along the sandwich sides. Finger-tighten one screw on each clamp, set the sandwich upright on a flat surface, and loosen the screw to align the stack. Taking great care in alignment will ensure a good seal. The glass plates and spacers must be flush with the clamp ridges at both top and bottom for a good seal. Fingertighten all screws. Remove the Spacer-Mate.
Tip: Use the casting cradle to hold the sandwich during alignment. Remove the laminated gasket from the cradle and, instead of setting the sandwich upright on a flat surface, set it into the casting cradle.
6. Club sandwich (pour 2 gels on each side of caster for a total of 4 gels). A 16-cm-long, notched center-divider plate (ordered separately and looks like other plates expect has notch) pairs two sandwiches to double the number of gels that can be cast and run. Start with a treated plate, place 2 spacers on the plate, lay the divider/notched plate on top, a second set of spacers on the stack and lastly a second treated plate. Place the notch so that it will be at the top of the gels. It is essential that the spacers and plates align perfectly in order to seal. Make sure the spacers are at the edge of the glass. This is harder to do if you can’t use the Spacer-Mate. You might have to use a thin ruler or another spacer to push a spacer out that is too far in. The gels will be run in this sandwich.
7. Secure the sandwich with clamps. Slide one clamp at a time along the sandwich sides. Finger-tighten one screw on each clamp, set the sandwich upright on a flat surface, and loosen the screw to align the stack. Taking great care in alignment will ensure a good seal. The glass plates and spacers must be flush with the clamp ridges at both top and bottom for a good seal. Fingertighten all screws.
8. Remove the sandwich from the casting cradle and inspect the bottom to make sure that edges are aligned flush to ensure a complete seal. Adjust if necessary.
9. Apply a light film of GelSeal compound only on the bottom corner surfaces created by the spacers and plates if the sandwiches tend to leak.
10. Place the gray laminated gasket into the casting cradle with the foam side down. Place the clamp assembly in the casting cradle, screw side facing out.
11. Level the casting cradle
Note: Do not use silicone grease or petroleum jelly to seal the sandwich. These substances are difficult to remove and ultimately cause artifacts.
12. Insert a black cam into the hole on each side of the casting tray with the ridge (short end) pointing up. Seal the gel sandwich against the casting gasket by turning both cams as far as needed, usually 90°–150°, up to 180°. The cam action presses the plates down into the gasket to seal the bottom of the sandwich. The seal is complete once the glass edge appears darker and nearly transparent against the gasket. Do not turn past this point. If the cam seems loose (as compared to the other cams) exchange the clamp screw.
13. Prepare the acrylamide monomer solution (12.5% or 10%; see Buffers and Solutions)and pour the gel. Prepare the required amount of monomer solution (30 mls/gel; this allows for some extra). Add the APS and TEMED just prior to pouring the gel. Using the 50 ml syringe, pipette the solution into one corner of thesandwich, taking care not to introduce any air bubbles. Fill until about 1 inch from top – allows space for the IPG strip. For a club sandwich, pipette the solution into both sandwiches, filling each to the same level below the notched edge.
14. Overlay each gel with a thin layer of water-saturated butanol, to prevent gel exposure to oxygen. This also forms a nice flat surface for the strip. Slowly deliver the overlay solution with a Pasteur pipette. Apply the solution near the spacer at one side of the sandwich and allow it to flow across the surface unaided. You only need 4-5 mm, just enough to cover.
15. After a couple of hours, pour off butanol and add diluted gel buffer to cover gel surface.
16. Allow the gel to polymerize at RT o/n or 4oC if not using the next day.

Equilibration
Roberto Notes:
• Optimize multiple components. DTT can be between 1 - 2.5 %; Iodoacetamide can be between 2.5 - 4 %
• Time critical. If you equilibrate too long, the proteins start to ppt.
Once you are done equilibrating - load on 2nd D gel.
• Usually do 15 min. incubation for each. Use shorter time for hi MW proteins and longer time for low MW proteins.

Prep
Day Before
1. Remove SDS equilibration buffer (SDS EB), 1 tube per gel from -20oC and place at 4 oC.
2. Dilute 10X electrophoresis buffer (1X: 0.5L into 5L ddH2O; 2X: 0.2L into 1L ddH2O or 1X: 1L into 9L ddH2O)) and prechill.
Run Day
1. If gels were stored at 4 oC, bring to RT.
2. Put 1 tube of agarose sealing solution (1 ml) at 70oC.
3. Make sure round mesh pad is in bottom of gel box (sometimes falls out when washed.
4. Attach 2 tubes in back of gel box (DALT and Ruby)to heat exchanger.
5. Fill lower buffer chamber about 2/3 full with 1X electrophoresis buffer. Don’t put too much buffer in lower chamber, the gels and blanks will displace some of buffer.
6. Put big gray-blue piece in gel box (only goes in 1 way).
7. Put spacers in slots that won’t have any gels.
8. Plug in gel box.  This will start the pump.
9. Turn on MultiTemp III, heat exchanger (just turn power on).  Check H2O level.  This keeps gels from going over 20oC during the run.  Heat exchanger should be set at 9.  If not, press Mode to select the S mode.  Use arrows to set to 9.  Press Enter.
10. Remove EB from 4 oC …1 tube (40ml) per gel and bring to room temp.  If you need to use heat to dissolve SDS – DO NOT go over 30oC.
11. Split SDS EB between 2 - 50ml tubes.

Rollin. Add 0.5% DTT to one and 4.5% iodoacetamide to the other:

_____ ml EB x 0.005 = ____g DTT
_____ ml EB x 0.045 = ____g iodo
50 ul 1% bromophenol blue/20 ml

Anupama. Add 1% DTT to one and 4.5% iodoacetamide to the other:

_____ ml EB x 0.010 = ____g DTT
_____ ml EB x 0.045 = ____g iodo
50 ul 1% bromophenol blue/20 ml

12. Petri dishes – 1 for each strip. Label top and bottom. Remove lids.
13. Or you can use the equilibration tubes for the equilibration steps.
14. Fill 100ml cylinder with 1X electrophoresis buffer for rinsing strips before laying on gel.
15. Do all of above before starting equilibration.  You do not want the strips to sit in any of the equilibration solutions longer than the indicated time.

Equilibration
1. Either remove boats from IPGphor or remove strip from -70 oC.
2. Remove lid off of a boat and place lid in a tray containing dH2O (Place all parts in dH2O when done just until you have time to wash parts with nonionic detergent.  DO NOT allow any buffer to dry onto boat).
3. With forceps, lift strip out of boat.  Touch 1 end to boat and let oil drain.  Touch 1 end to a piece of filter paper (Whatman 1) to blot oil.  You can set it down, just be sure gel side is up (much easier to see now that it is rehydrated.
4. Pick strip up on each end with forceps or pull strip out of Equilibration tube.  Curl into Petri dish along inner side.  Make very sure that the plastic backing is next to the Petri dish.
5. Alternatively, if you are not starting the 2nd Dimension.  Slide the strip into an equilibration tube, gel side up.  Store at –80oC.
6. Western blotting: if you did anything special for your 1D gels, you need to do the same (ex. if your proteins need to be derivitized, treat with DNPH). Equilibrate for 20 min. Wash these with 2M Tris/30% Glycerol for 15 min.  Pour off liquid.
7. Incubate strips for 15 min at 85 rpm in 20ml EB with DTT.
8. Pour off EB with DTT and discard into waste.
9. Add 20ml EB with iodoacetamide at 85 rpm for 10 min.
10. Remove gel sandwiches from box.  Rinse with ddH20 and rinse out well.  Invert to drain off well.  Tip gel sandwich on a corner to drain off H2O.  Use the corner of a Crew Wipe to carefully remove last of the water from the well.  Be careful not to touch the gel surface.  Place on rack.
11. Shelby lays the plate flat on a Crew Wipe to do the initial positioning of the strip.  Then she holds the gel up with her left and uses the ruler to push the strip down onto the gel.  Anupama and I place the plate in the rack, position the strip the same way and push the strip down on the gel the same.  Either method is fine – just pick one that you are comfortable doing. 
12. Drain off EB with iodoacetamide and discard into waste.
13. Grab the no barcode side of the strip with forceps with your right hand.  Dip 2 –3 times into 100 ml grad. Cylinder containing 1X running buffer.
14. Ettan DALT system:  Lay on edge of gel with the acidic end (bar code end ) to the left, gel side towards you and plastic to back of gel.  SE600 Ruby:  Lay on edge of gel with the acidic end to the left, gel side towards you and plastic to spacer plate (if 4 gels were poured) or to back of gel sandwich if only 2 poured.  Use clean thin plastic ruler to position strip on top of gel (touch only the strip backing with the ruler, not the IEF gel itself).  Move the strip so that the acidic end is centered. 
15. Be very careful not to push the ruler through the gel.  With the shorter strips it is pretty easy to push the strip down using only the ends of the strip.
16. Adding MW marker.  We haven’t found an effective way to add a MW marker.  I am going to give you 3 methods to try.
a. Spot 15 ul of ProtoMarkers onto a small piece of Whatman 1 (~0.5 cm2).  Slip piece on 1 end of gel.  It is thin enough that it just drops in.  This method starts out OK, but is extremely difficult to see as the gel runs the full time.  The marker did show up when scanned on the Typhoon.
b. Rehydrate an IPG strip in the reswelling tray o/n with RB + a marker.  Suggestion from GE discussion group.  I tried this twice and it didn’t work while it was running and I couldn’t see it on the scanned image.
c. Cut a long narrow piece of plastic (same thickness as gel).  When you add the agarose sealing solution, hold the piece of plastic in place until the agarose solidifies.  Carefully remove.  This should form a well that you can use to load a marker. *We are seeing if this works now – 11/21/07.
17. Pipet about 1.5 ml of agarose sealing soln. On top of strip.  Using ruler, try to remove any bubbles.  Wait a few minutes for it to solidify.
18. Repeat for remaining gels.

Setting up 2nd Dimension Run
Ettan DALT system.
1. Rollin used 1X buffer for lower and upper chambers.  GE Manual and Anupama uses 1X buffer for the lower chamber and 2X buffer for the upper chamber.  The Ettan DALT six electrophoresis system manual recommends 1X buffer for the lower chamber and 2X buffer for the upper chamber.
2. Place finished gels in gel box slots, 3 in front and 3 in back.  Keep track of any gels that are untreated.  + (anode) on left.  Short plate of sandwich towards front.
3. Fill Lower chamber to LBC line with 1X buffer. 
4. Upper chamber with rubber gaskets – wet so that it is easier to slide over glass.  Push hard until you hear a click; there shouldn’t be a gap and the lid should go to about the LBC line.  The part with the rubber gaskets forms a tray for the 2X buffer, so push really hard.  It should rest on the plastic shoulder right and left of plate.
5. Add 2X  or 1X running buffer to MAX fill line in the upper chamber.
6. 2 liquid levels (1X and 2X) are separated at first, over about 5 min. the lines come together and should be between the Min and Max fill line.  Add more 2X to the upper chamber and 1X to the lower chamber if not.
7. Until 2 liquid levels come together between the MIN and MAX line, you can’t start the power supply.  You will get an error message.
8. Plug in lid with electrodes – only 1 way to go on and then plug electrodes into power supply.
9. Run takes 4 –5 hrs. depends on how many gels (3 gels – 4 hrs.).  Usually run until dye front at bottom.  If your proteins are near the top, you might want to run it a little longer.  You can also run o/n – just run at 2 watts/gel o/n.  Don’t need prerun step. 
10. The slower you run your gels, the better the resolution.
11. Turn power supply on.
12. Protocol has 2 steps, but depending on your power supply, you might not be able to set up both at the same time.
•1st step:  time: 30’, 600V, 400mA, 2.5 W/gel
•2nd step:   time: 4-5 hours, 600V, 400mA, 100W
•even though V,  mA and W are set at maximum, V will be rate limiting and never get to 600 V.
13. Run gel o/n.  2 watts/gel; ran 5:00 pm – noon to get dye at end of gel; increased voltage when you come in if the dye front is not at the bottom. 

Setting up 2nd Dimension Run SE600 Ruby
1. Prechill the buffer or place about 3 L buffer in lower chamber and start water circulating.
2. Turn on MultiTemp III, heat exchanger (just turn power on). Check H2O level. This keeps gels from going over 20oC during the run. Heat exchanger should be set at 9. If not, press Mode to select the S mode. Use arrows to set to 9. Press Enter.
3. Rinse both buffer chambers with water and distilled water thoroughly before each use.
4. Clean away any gel adhering to the exterior of the gel sandwiches.
5. If running only one gel: Block the second upper buffer chamber slot by installing the acrylic buffer dam included with the unit. Fit clamps onto the dam, taking care to align the clamp ends and dam edges. Install the “dummy” gel, screws facing out, in the second cradle in the dual gel caster.
6. Attach the gel sandwich to the upper buffer chamber. Turn the upper buffer chamber upside down. Spread a small amount of grease on the dark gray gasket – ends only. This will help the gasket stick to the upper chamber better. Place a slotted gasket into both sandwich holder recesses. Both the slot in the gasket and the slot in the recess must align – the gasket has raised edges along the slot on only 1 side (side with grease), these raised edges must fit into the upper buffer chamber slot. Both slotted gaskets must be used even if running only one gel sandwich. Grooves along each slot help keep the gasket in place. Release the sandwiches from the caster by removing all bottom cams (if present). Lower the upper buffer chamber onto the gel sandwiches while still sitting in the casting stand. Install the cams on the top holes of the chamber and gel sandwich. Cams should be positioned ridge (short end) pointing down. Clamp the sandwich in place by simultaneously turning one cam clockwise and the other counterclockwise a full 180°. The final cam position must be vertical so that the assembly fits into the lower buffer chamber. If one of the cams seem loose (as compared to the other cams), exchange the clamp screw.
Note: Do not force the cams. If you encounter unusual resistance, disassemble and inspect clamp and glass alignment along the top of the sandwich. Align and reinstall.
Important! A smooth fit between the sandwich and gasket is essential to a good seal.
7. Pour 100 ml of buffer into the chamber, directing the buffer stream toward the side wall. Check that no buffer leaks around the gasket. This is a very important step – if there is any leaking you can’t proceed. If there is leaking, check that the gasket is inserted properly and completely. If any of the cams are loose, the upper chamber will leak. Leaking can also be caused by chips on glass plate at the top or not enough grease on gray gasket.
8. If the assembly leaks, take it to a sink and partially release the cams to allow buffer to drain out of the upper chamber. Disassemble, check alignment of all sandwich components, and adjust if necessary.
9. Important assembly notes:
• IEF runs: The buffer level in the lower buffer chamber must never reach the upper buffer chamber; maintain at least 2 cm of clearance.
• Do not fill the upper or lower chamber above the recommended levels.
• Remove buffer in contact with the electrode posts.
• Pour buffer slowly and away from the slots in the upper buffer chamber to avoid disturbing the samples.
• Use only water or 50/50 water/ethylene glycol as coolant. Never use a commercial antifreeze or any alcohol-based mixture, or irreparable damage to the heat exchanger will result.
• Do not connect the heat exchanger to a water tap or any other source where the water pressure is unregulated.
10. Place a magnetic spin bar into the lower buffer chamber (LBC) and place the unit on a magnetic stirrer. Fill the lower chamber with up to 4 litres of buffer. Turn magnetic stirrer on at a setting that mixes the buffer well.
11. Lower the heat exchanger (part that has tubes that will be connected to MultiTemp III) into the lower chamber, fitting the ports into thenotches in the rim. (The heat exchanger must be in place for all runs because the lower electrode is integrated into the heat exchanger).
12. Fit the upper buffer chamber assembly into the lower buffer chamber. Grasp the assembly in the casting stand by the upper buffer chamber and carefully lower it into the lower chamber.
13. Inspect the installation and check the buffer levels.
• Upper buffer chamber (UBC). The electrode along the upper chamber ridge must be submerged about 1 cm. This level requires 450–600 ml of buffer— just enough to cover the upper chamber ribs, but not high enough to contact the banana plug. Do not fill above UBC MAX fill line.
• Lower buffer chamber (LBC). Fill to LBC MAX fill line.
14. Place the safety lid on the unit by engaging the safety interlock pins before lowering the electrode connections on to the banana plugs. The safety lid has black and red pins that fit onto the electrode connections. The black pin fits over a red electrode connection. This is OK.
15. Plug the color-coded leads into the jacks of an approved power supply, such as the EPS 601. Plug the red lead into the red output jack and the black lead into the black output jack. In most systems the red lead, which is connected to the bottom electrode, is the anode (+), and the black lead, connected to the top electrode, is the cathode (–).
16. Separating the sample. Gels may be run at either constant current or constant voltage settings. A constant current mode is traditionally used with a discontinuous buffer system so that the rate of electrophoretic migration remains unchanged throughout the run. Under these conditions voltage increases as the run proceeds. A lower current setting is recommended for higher resolution. The optimal current level must be determined empirically; the main factors that must be balanced include the gel % and migration speed, and the resulting Joule heating and band distortion. Table 2 in the SE600 Ruby manual lists starting-point guidelines and adjustments for gel thickness, number ofgels, and migration rate.
Debby 0406: Application Note.80-6445-94 – Amersham 2-D electrophoresis. Start at 15 mA/gel for an initial migration and stacking period of 15 min., then increase to 30 mA/gel for a period of ~3 hrs.
Shelby 0506: Run gel at 20 mA/gel. Set V at 600 and W at 100. Run takes ~5 hrs.
SE600 Ruby manual:
• Current. Current acts on the total cross-section area of all the gels because the gels are connected in parallel in the electrical circuit. Thus the current setting for one gel must be multiplied by the number of gels of the same crosssection run simultaneously. For a gel 1.5 mm thick, we suggest a starting current setting of 25 mA. (Two 1.5 mm gels = 50 mA.). For a gel 1.0 mm thick, we suggest a starting current setting of 16.7 mA. (Two 1.0 mm gels = 33.4 mA.). Note: Cooling may be required to control Joule heating. The current can be increased for faster runs if active cooling is used and it can be decreased for slower overnight runs.
• Voltage. The starting voltage for a 1.5 mm slab gel connected to a power supply set to 25 mA is usually 80–90 V (using the SE 600 Ruby unit with a Laemmli discontinuous buffer system for SDS gels). The final voltage is typically 250–400 V, depending on the length of the gel. (See Table 2 in manual.)
• Time. A run is usually complete when the tracking dye reaches the bottom of the gel. In a 16 cm gel (SE 600 Ruby), a 1.5-mm-thick Laemmli SDS gel, run at 25 mA/gel without cooling, usually requires 5 h.
17. Record each run. Keep a record of the current or voltage setting, number and thickness of gels, buffer system, and the starting and final current or voltage readings for each run so that results can be compared. Inconsistent results for the same system and settings indicate potential problems such as leaking current, incorrect buffer concentrations, high salt concentrations, or inconsistent chemical quality. Check band progress after 5 min, and again after 1 h, keeping an eye on the migration rate of the tracking dye. If migration is slow, make sure stir bar is stirring the buffer well. The run is complete when the tracking dye reaches the bottom of the gel. Watch the buffer level and, if necessary, replenish it as required to keep the top electrode submerged. (A small volume of buffer may leak past a nicked plate or gasket, or buffer may pass through the gel.)

Cleaning
1. 2nd dimension gel box. Rinse several times with dH2O until no longer soapy.
2. 2nd dimension glass plates. Scrape off gel using BioRad gel prying tool.  Soak o/n (different soaking depending on what plate will be used for).  Use a net bath ball to clean plate.  Different treatments depending on whether or Bind Silane was used on a plate.  Also Rollin includes an HCl treatment for DALT plate without spacer or SE600 Ruby plate that has Bind Silane.

a. Spot picker.  Acid plate treatment
1) This is a royal pain to do, but if you don’t do a good job the next run could have the gel splitting between the 2 plates.
2) Soak plates top and bottom in 1% Contrad 70.  Do not soak o/n.  Plates can turn cloudy. Soak up to 2 hours.
3) Rinse plates in ddH2O.  Place in drying rack and let dry some.  Check plate to make sure ALL of gel has been removed.  Hold up to the light.
4) If there is still gel stuck to plate, wet the gel prying tool with Contrad 70 and scrape plate.
5) Repeat steps 2 and 3 until the plate is COMPLETELY CLEAN. This will take a # of tries to get clean. Check again after dry to make sure all of gel has been removed. 
6) Prepare 2L of 1% HCL
7) Pour 1.5L  1% HCL into staining tray.
8) Insert top plates (the ones without spacers)…stack carefully on top of each other
9) Incubate at 30 rpms for 2h or o/n.
10) Wash bottom plates (with spacers) with 1% Contrad, rinse with dH2O, then ddH2O, then HPLC grade H2O, shake off excess water and leave to dry protected from duct contamination.
11) Repeat (10) with acid-treated bottom plates.
12) Store between Crewwipes in a staining tray.
b. No spot picker.  Use regular soap.  Scrub and look.  Rinse with dH2O.  Dry on gel rack.  Check again after dry to make sure all of gel has been removed. 

Buffers and Solutions
Read the Amersham manual for basic solutions
•Appendix 1 has solutions and buffers used to run the gels.

Binding Solution – 10 ml
8 ml 100 % EtOH
200 ul glacial acetic acid
20 ul Bind Silane (Rollin uses 12.5 ul)
1.8 ml H2O
Use 4 ml/Ettan DALT plate. Use 3 ml/SE600 Ruby plate

Binding Solution – 15 ml
12 ml 100 % EtOH
300 ul glacial acetic acid
30 ul Bind Silane (Rollin uses 12.5 ul)
2.7 ml H2O
Use 4 ml/Ettan DALT plate. Use 3 ml/SE600 Ruby plate

H2O saturated n-Butanol – 110 ml
10 ml ddH2O
100 ml n-Butanol

Gel Storage Buffer
375 mM Tris-HCl pH 8.8, 0.1 % SDS

375 ml 1M Tris – HCl pH 8.8
10 ml 10% SDS

Bring volume up to 1L

SDS Equilibration solution
50 mM Tris-Cl pH 8.8, 6 M urea, 30 % glycerol, 2% SDS

To 60 ml ddH2O:
6.7 ml 1.5M Tris-Cl, pH 8.8
72.07 gm Urea
69 ml glycerol
4.0 gm SDS
Dissolve and bring up to 200 ml. Aliquot 40 ml/tube. Store at RT. Or –20C, 4C night before use and RT the day of use. Stable for 6 months.

Tris/Glycerol used for Westerns
2M Tris, 30 % glycerol

12.11 gm Tris
15 ml glycerol
Dissolve and bring up to 50 ml with water.

Agarose Sealing Solution
0.125 gm agarose
25 ml 1 X running buffer
Heat carefully in microwave to dissolve. Let remaining solution solidify in flask until next time. Remelt in microwave just before using.

10 X Tris/Glycine Running Buffer
250 mM Tris, 1.92 M Glycine, 1% SDS

151.25 gm Tris
720.5 gm Glycine
50 gm SDS
Dissolve and bring up to 5 L.
OR
30.25 gm Tris
144.1 gm Glycine
10 gm SDS
Dissolve and bring up to 1 L.

Note: Rollin and Suganthi do NOT pH their buffer; Anupama ph’ed her buffer to 5.3. When I tried to pH the buffer, it ran REALLY slow.

SDS-PAGE (Debby’s, SE600 Ruby)
Recipe from Red Book; Table 18.3

Gel solution (12.5%) – 30ml; 30 ml is more than enough for 1 gel – you don’t need to do extra

12 ml 40% National Diagnostics Protogel
7.5 ml 1.5M TrisCl pH8.8
9.9 ml ddH2O
0.3 ml 10% SDS
0.3 ml fresh APS
12 ul TEMED (80% of recommended amount)

Gel solution (10%) – 30ml; 30 ml is more than enough for 1 gel – you don’t need to do extra

10 ml 40% National Diagnostics Protogel
7.5 ml 1.5M TrisCl pH8.8
11.9 ml ddH2O
0.3 ml 10% SDS
0.3 ml fresh APS
12 ul TEMED (80% of recommended amount)

10% APS
0.5 gm APS
4.5 ml ddH2O

SDS-PAGE (Rollin’s)

Gel solution (12.5%) – 940ml
294ml 40% monomer solution
235ml 1.5M TrisCl pH8.8
392ml ddH2O
9.4ml 10% SDS
376ul TEMED (80% of recommended amount)

Displacing solution
0.375M TrisCl pH 8.8, 50% glycerol, 0.002% bromophenol blue – 120ml

20ml ddH20
30ml 1.5M TrisCl, pH 8.8
70ml glycerol
240ul 1% bromophenol blue

10% APS
1gm APS
9.6ml ddH2O

TEMED concentration can be reduced in increments of 10 or 15% if the casting results in bumpy tops.

Rollin's table for 1 - 6 gels using Ettan DALT 6 gel caster
Components
volume units
1 gel
2 gels
3 gels
4 gels
40% acrylamide/0.8% bisacrylamide
ml
31.3
59.5
84.5
106.4
4X resolving gel buffer (1.5 M Tris, pH 8.8)
ml
25
47.5
67.5
85.0
10% SDS
ml
1.0
1.9 
2.7
3.4
ddH2O
ml
41.7
79.2
112.6
141.8
10% Ammonium persulphate
ml
1.0
1.9 
2.7
3.4
100% TEMED
ul
50
95 
133 
170
Total Volume
100
190
270
370
           
Components
volume units
5 gel
6 gels
40% acrylamide/0.8% bisacrylamide
ml
128.3
150.2
4X resolving gel buffer (1.5 M Tris, pH 8.8)
ml
102.5
120.0
10% SDS
ml
4.1
4.8 
ddH2O
ml
171.0
200.0
10% Ammonium persulphate
ml
4.1
4.8 
100% TEMED
ul
205.0
240.0 
Total Volume
410
480


Anupama's table for Ettan 6 gel Caster.
Please note that she used Protogel and a different % of SDS and % of TEMED. Use 450 ml even in only pouring 4 gels. I have used 1/2 recipe when pouring 3 gels or fewer and it was fine. You do not need to degas Protogel.
Components
volume units
6 gels
National Diagnostics Protogel
ml
188.0
4X resolving gel buffer (1.5 M Tris, pH 8.8)
ml
113.0
20% SDS
ml
2.25
ddH2O
ml
141.63
10% Ammonium persulphate
ml
4.5
10% TEMED
ul
620.0
Total Volume
 
450


 


This information is given as a guide to the facilities and instrumentation available in the DNA/Protein Core facility at Georgia State University. If you have any concerns or thoughts about the content of this website please contact: John Houghton (404) 413-5390


 


 

 

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