2-D Electrophoresis
using Immobilized PH Gradients
IPGphor and Dalt-system
24 maart 2004
1 Sample preparation
Rehydration stock solution with IPG Buffer * (lysis buffer)
Component
Final concentration Amount 10 ml
___________________________________________________________________
Utrapure Urea
8M
4.8 g
CHAPS
4 % (w/v)
0.4 g
Tris
10 mM
0.012 g
DTT
50 mM
77 mg
IPG Buffer
0.5% (v/v)
50 µl
(same pH range as the IPG strip)
Bromophenol blue
trace
(a few grains)
Deionised water
to 10 ml
Store in 1.0 ml aliquots at –80 °C. NB. If you want to do a Bradford, do not include the IPG
buffer in the lysis buffer.
Brain tissue – Total Protein Extraction
1. Usually ~ 10-20 mg wet weight homogenized in 400 l lysis buffer (18 cm) or 500 l
(24 cm) is a good amount for silver staining (check it with Bradford)
2. Homogenize the tissue with a plastic pestle in a eppendorf tube or for slightly tougher
tissue (eg. DRG or sciatic nerve), use a glass-glass homogenizer.
3. Lyse for one hour while mildly shaking
NB. When you solubilize in lysis buffer, if it gets sticky (a lot of DNA), it usually
means you have too many tissue. Stickyness should be prevented because basic
proteins bind to the DNA and you will lose them.
Brief (a few seconds) sonication with a probe sonicator can break DNA and
therefore make sample soluble again. But it is best to omit stickines altogether by
having a optimal ratio between cell quantity and lysis buffer.
4. After lysis centrifuge 5 min. to get rid of particles/dell debris
5. Estimate the amound of protein with Bradford:
6.
7.
8.
9.
…. l sample
Together 10 l
…. l lysis buffer
10 l H2O
10 l 0,1M HCl
1000 l Bradford 1:4
Dilute the sample with lysis buffer to the desired concentration
Add IPG buffer (2 l in 400 l lysis buffer), mix well
Centrifuge samples prior to loading onto the IPG strip holder.
Load ~200 ug protein in 450 ul (24 cm) or in 370 ul (18 cm) into the IPG strip holder
2
Cell culture – Total Protein Extraction
1. Lysis Buffer (for total cell lysates)
8M Urea, 4% Chaps, 50 mM DTT
NB. Leave out Tris from lysis buffer because salt is often a problem with cell
culture samples, at least when the IPGphor system is used for the first dimension.
2. Quantity
2 million cells per 2D gel is usually a good protein load (check with bradford prior to
running). Sometimes a little more or less cells are needed, it depends on the cell type and
should be tested before running a full set.
NB. Add IPG buffer to cells in lysis buffer AFTER bradford.
3. Washing
Wash cells adherent to plate (cell pellets in case of non-adherent cells) twice with PBS to
get rid of serum and twice with 10 mM Tris to get rid of salts.
NB. Make sure to pipet off all residual liquid (if you do on-plate washing, keep the
plate at an angle for a minute o so to collect residual liquid in the edge and pipet off
again and repeat the several times especially with the last washes).
4. Lysis
1. Add 400-500 ul 2D lysis buffer (8M Urea, 4% Chaps, 50 mM DTT) to plate (~ 10
cm, with the right amount of cells), scrape cells QUICKLY into lysis buffer and lyse
cells further by QUICKLY pipetting up and down (avoid foaming).
NB. 450 ul is the sample volume for a 24 cm IPG strip and 370 ul for a 18 cm IPG
strip.
NB. When you solubilize in lysis buffer, if it gets sticky (a lot of DNA), it usually
means you have too many cells. Stickyness should be prevented because basic
proteins bind to the DNA and you will lose them.
Brief (a few seconds) sonication with a probe sonicator can break DNA and
therefore make sample soluble again. But it is best to omit stickines altogether by
having a optimal ratio between cell quantity and lysis buffer.
2.
3.
4.
5.
6.
7.
Lyse during 1 hr at RT while mildy shaking (avoid foaming)
After lysis centrifuge 5 min. to get rid of particles/dell debris
Estimate the amound of protein with Bradford (see page 2), use ~5 ul sample
Add IPG buffer to samples, mix well.
Centrifuge samples prior to loading onto the IPG strip holder.
Load ~200 ug protein in 450 ul (24 cm) or in 370 ul (18 cm) into the IPG strip
holder
5. Storage
Lysed cells can be stored at –80 oC.
3
2
First-dimension isoelectric focusing
IPGphor Isoelectric Focusing System
With the IPGphor Isoelectric Focusing System, both rehydration of the IPG strip and
IEF occur in individual strip holders. The sample can be loaded by simply including it
in the rehydration solution
1. Select the stripholder(s) corresponding to the IPG strip length chosen for the
experiment.
Important: Handle the ceramic holders with care, as they are brittle
2. Pipette the appropriate volume of sample-containing rehydration solution into each
holder. For 24 cm IPG strip 450 l and for 18 cm IPG strip 370 µl. Deliver the solution
slowly in the strip holder channel while moving from left to right, so that the sample is
equally divided over the strip holder. Remove any larger bubbles.
3. Mark the IPG strips with a number on the cathodic (-) end of the strip. Use the blue
Fineliner from Staedtler.
4. Remove the protective cover from the IPG strip. Position the IPG strip with the gel
side down and the anodic (+) end of the strip directed toward the pointed end of the
strip holder. Plus end first, lower the IPG strip onto the solution. To help coat the
entire strip gently lift and lower the strip and slide it back and forth along the surface on
the solution, tilting the strip holder slightly as needed to ensure complete and even
wetting. Finally, lower the cathodic (-) end of the IPG strip into the channel, making
sure that the gel contacts the strip holder electrodes at each end. Be careful not to trap
bubbles under the IPG strip.
5. Apply IPG Cover Fluid to minimize evaporation and urea crystallisation. Pipette the
fluid drop wise in the strip holder, until the entire IPG strip is covered. 600 l for a 18
cm holder and 700 l for a 24 cm holder. Make sure there is no solution outside the
stripholder (clean with tissue).
6. Place the cover on the stripholder
7. Allow the IPG strip to rehydrate. Rehydration can proceed on the IPGphor unit
platform. Ensure that the holder is on a level surface. A minimum of 10 hours is
required for rehydration. The rehydration period can be programmed as the first step of
an IPGphor protocol.
4
Run the first dimension
Protocol-IPGphor (protocol 1 in the new machine and protocol 3 in the old
machine(RA-lab))
50 µA per IPG strip
20 °C for both rehydration and IEF
Immobiline DryStrip 18 cm, pH gradient 3-10 NL
Immobiline DryStrip 24 cm, pH gradient 3-10 NL
Step
Rehydration
1
2
3
Total
Voltage
Step duration
Volt-hours
Gradient type
30
500
1000
8000
12:00
1:00
1:00
8:00
360
500
1000
64000
step-n-hold
step-n-hold
step-n-hold
step-n-hold
22.00
65860
9. Ensure that the stripholders are properly positioned on the IPGphor platform.
Close the safety lid and begin IEF.
10. After IEF proceed to the second-dimension separation immediately or store the
IPG strips at -40 to -80° C covered with saran wrap.
Gels may be stored at least 2 to 3 months at -80° C. Do not place in the equilibration buffers required for the second dimension
prior to storage.
5
4 Equilibration of the IPG gel strips
The IPG gel strips are equilibrated twice, each time for 15 min in 2 x 5 ml equilibration
buffer. Equilibration buffer 1 contains DTT. Equilibration buffer 2 contains 260 mM
iodoacetamide to remove excess DTT, responsible for the "point streaking" in silver
stained patterns.
Buffer solutions:
1.5 M Tris-HCl buffer, pH 8.8
To make 1000 ml, dissolve 182 g of Tris in about 800 ml of deionised water. Adjust to
pH 8.8 with HCl and fill up to 1000 ml with deionised water.
Equilibration buffer
50 mM Tris-Cl pH 8.8, 6 M urea, 30% glycerol, 2% SDS, Bromophenol blue, 200 ml)
72,07 g urea
60,6 ml 99% glycerol
40 ml 10% SDS
10 ml 1.5 M Tris-HCl, pH 8.8
a few grains Bromophenol blue
Dissolve in deionised water and fill up to 200 ml.
Store in 50 ml portions at – 20 °C
Procedure: for 10 strips
1. Dissolve 250 mg of DTT in 25 ml of equilibration buffer,
(=Equilibration buffer I).
Pipette 2.4 ml equilibration buffer I (DTT) in each well of the equilibration holder.
Take out the focused IPG gel strips one by one and remove the coverfluid, but try not
to damage the gel! Then place them individually in each well of the equilibration holder
with the gel side faced up. Place the equilibration holder on a rocker. Equilibrate for a
maximum of 15 minutes.
2. Dissolve 625 mg of iodoacetamide in 25 ml of equilibration buffer
(=Equilibration buffer II).
Pipette 2.4 ml equilibration buffer II (iodoacetamide) in each well of the equilibration
holder.
Take out the focused IPG gel strips and place them individually in each well of the
equilibration holder with the gel side faced up. Place the equilibration holder on a
rocker. Equilibrate for a maximum of 15 minutes.
6
5
SDS-PAGE as second dimension
Preparing SDS DALT slab gels
Gel size:
200 x 250 x 1.5 mm
Buffer composition:
Gel Buffer: 1.5 M Tris-HCl buffer, pH 8.8
Tris
182 g
Adjust to pH 8.8 with HCl
Deionised water
to 1000 ml
Adjust to pH 8.8, store at 4 °C.
10% SDS
Sodium dodecylsulfate 10 g (weight in fume hood)
Deionised water
up to 100 ml
Store at room temperature.
Overlay Buffer
n-butanol
50ml
deionised water.
5 ml
Combine in a bottle and shake. Use the top phase to overlay gels.
Store at room temperature indefinitely.
Acrylamide Stock Solution (30.8% T, 2.6% C):
Biorad 37.5 : 1 2.6% C catalog No. 161-0158
Or use Duracryl van Genomic solutions catalog No. 80-0085
10% Ammonium persulfate
Ammonium persulfate
1.1 g
Deionised water.
11 ml
This solution should be prepared freshly just before use.
Displacing Solution
( 375 mM Tris-Cl, pH 8.8, 50 % glycerol, bromopenol blue, 200 ml )
Tris-Cl (1.5M, pH 8.8)
50 ml
Glycerol
100 ml
Bromophenol blue
2 mg
Water
50 ml
This solution should be prepared freshly just before use.
7
Recipe for casting 11 vertical DALT slab gels (1100 ml)
Component
Duracryl 30%
Acrylamide 37.5 : 1
Tris stock pH 8.8
Water
10% SDS
10% APS
TEMED
11% gel (ml)
201.5
201.5
275
400
11
11
0.5
12% gel (ml)
220
220
275
363
11
11
0.5
13% gel (ml)
238.5
238.5
275
326
11
11
0.5
Casting Homogeneous SDS- Gels
1. Be sure the entire gel casting system is clean, dry and free from any polymerised
acrylamide.
2. Prepare a sufficient volume of gel overlay solution. You need 1.0 ml of overlay for
each cassette.
3. Make up 200 ml of displacing solution.
4. Make up the gel acrylamide stock solution, without adding the 10% APS or TEMED.
Mix well.
5. Load the gel caster with cassettes, separator sheets and filler blocks, if necessary.
Place a gel label in each cassette. (for 11 gels, 4 big and 1 small filler block)
Don’t forget the rubber and the sponch to trap air bubbles!
6. Connect the feed tube to a funnel held in a ring-stand at a level about 12 inches
above the top of the gel caster. Insert the other end of the feed tube in the grommet
in the bottom of the balance chamber.
7. Load the balance chamber with 200 ml heavy displacing solution.
8. Add the appropriate volumes of APS and TEMED only when you are ready to
pour the gel, not before. Mix well.
9. Slowly pour the gel solution into the funnel, taking care to avoid introducing any
bubbles into the feed tube line.
10. Once the pouring is complete, remove the feed tube from the balance chamber
grommet. If the V-well is not completely filled and the level of gel in the cassettes is
more than 1 cm below the top of the cassettes, you may add up to 50 ml more
displacing solution to the balance chamber.
8
11. Immediately after removing the feed tube from the caster, slowly deliver 1.0 ml of
overlay buffer to the surface of each gel. The overlay should spread evenly across
the cassette with a minimum of mixing resulting in a smooth, flat gel top surface.
Apply equal volumes of overlay to each gel to produce gels of consistent heights.
12. Allow non-gradient gels to polymerise for at least 1 hour. Put some saran wrap over
the gelcaster.
Loading the IPG Strips onto the DALT Slab Gels
1. Dip the IPG gel strips in SDS electrophoresis buffer to lubricate it, then place the
IPG gel strip onto the DALT gel cassette.
Position the IPG strip between the plates, touching the surface of the second
dimension gel, with the plastic backing against one of the glass plates. For a
convenient reference, place the pointed, acidic, end of the IPG strip on the same
side as the gel label. Use a the thin plastic backing from the package of de IPG
strips to push against the plastic backing of the strips, not the gel itself, and move
the strip down into contact with the surface of the second dimension gel. The edge
of the strip should just rest on the surface of the slab gel. Avoid trapping air
bubbles between the plastic backing and the glass plate or cutting into the SDS gel
with the strip. The gel face of the strip should not touch the opposite glass plate.
2. Biorad 161-0373 standard. For silver gels dilute standard 5 times and pipette 10 µl
of the diluted marker on a pre-cut strip. For coomassie gels pipette 7 l of the
undiluted standard on a pre-cut strip.
Biorad 161-0374 dual color standard (for blotting ). Pipette 7 l of this undiluted
marker on a pre-cut strip.
Position the Standard strip next to square end of the IPG-strip.
3. Melt the agarose (0,5% in 1x TGS) at 100°C for approximately 10 minutes. Allow
the agarose to cool to 40 to 50°C and then deliver agarose sealing solution onto the
IPG strip to seal it into place. Carefully avoid bubbles when sealing with agarose.
Wait 2-5 minutes to allow the agarose to fully solidify before proceeding.
SDS Electroforese buffer
10x TGS (Tris/Glycine/SDS buffer) van Bio-Rad cat. Nr. 161-0772
Dilute 1:10 with demi-water (1.8 liter buffer and fill to 18 liters), precool the buffer
for a few hours (16°C)
9
Loading Cassettes into the DALT tank
1. Carefully load the cassettes after the agarose overlay has fully solidified. The
cassettes are correctly loaded in running orientation in the DALT Tank slot with the
IPG strips vertical along the left side, or cathode (-), side and the rubber cassette
hinge the bottom. Dip the hinge side of the cassette into the tank buffer first to
lubricate it before inserting into the flap seals. Use both hands slide cassettes firmly
to the bottom.
Be careful! The plates slip easily once your hands are immersed in tank buffer!
2. Adjust the buffer level after al cassettes are loaded in position.
Important; The tank buffer level should be even with the uppermost spacer of the cassette and
neither above the top of the cassette nor below the level of the top edge of the gel.
3. Close the lid and attach the electrical leads to make proper electrical contact with
the power supply. Migration proceeds towards the red (+), or right chamber.
4. Turn on the power supply
5. Set the power supply to 85 V constant voltage for overnight runs (it takes about 16
hours for a 12% gel)
10
Protocol Silver Staining
(compatible with MS)*
Incubations times and volumes are optimised for 'large scale' silver staining of ten 2D gels (20X25 cm, 1.5
mm) in 2 boxes (5 gels/box). All incubation volumes are 1.5 lit per box with 5 gels. All incubation steps
are performed on a circular shaker. We use a home-made 'peddle' with holes, to press down the gels to the
bottom of the container while decanting fluids.
1. Fixation: 45-60 min in 5% HAc/50% MeOH.
2. Wash steps: 15 min in 50% MeOH, followed by 15 min in distilled (or demi) water.
In the mean time, make the developer solution (2 x 3L for 10 gels), because it takes a long time for the
sodium carbonate to dissolve
3. Sensitisation: 3 min in 0.02% sodiumthiosulphate (600 mg/3 Liter)
The time for the sensitisation depends on the temperature in the lab. This means when the temperature is
more than 20°C the sensitisation time must be shortened
4. Wash steps: 1x 2 min followed by 1x1 min in distilled (or demi) water. Stick to time!
5. Silver staining: 30-45 min in 0.15% silver nitrate (4,5 gr/3 L)
Gels become yellow
Put the silver waste is the special silver waste container
6. Wash steps: 1x 2 min followed by 1x1 min in distilled (or demi) water. Stick to time!
7. Develop: Refresh the developer solution (0.04% formalin or 37% formaldehyde in 2%
sodium carbonate) when the liquid turns yellow, followed staining until the desired intensity
is achieved (usually not more than 2 min).
Make developer fresh. Dissolve 60 gr sodium carbonate in 3 liter and add 1.2 ml formalin just before use.
If duracryl (Genomics Solutions) gels are used, 5 gels can be developed at once in a box, while vigorously
shaking (I do this by hand, while watching). Although it may be safer to develop not more than 3 or 4 gels
per box. If regular acrylamide gels are used (especially when they are of low percentage), I recommend to
develop not more than 2 gels at a time.
8. Stop: Replace developer with 5% HAc. For 20-60 min.
Put the gels in a clean box with the stop solution
9. Store: Store the gels in 0,2 % HAc/10% MeOH.
*Modified from Shevchenko et al (1996) Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Electrophoresis 68,
850-858.
11
Protocol Coomassie Staining (G-250)
Coomassie stain G-250: 34% methanol
340 ml
3% phosphoric acid
35,3 ml (85%)
150 gram ammonium sulfate
150 gram
1 gram coomassie brillant blue G-250
1 gram
Add water to 1000 ml
Store at room temperature
1. Fix gel overnight in 50% ethanol/3% phosphoric acid
2. Wash 3x15 min in water
3. Stain in coomassie brillant blue G-250 solution for at least several hours to days.
No destaining is necessary
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