2-D Electrophoresis
using Immobilized PH Gradients
IPGphor and Ettan DALTtwelve System
6 april 2004
1 Sample preparation
Rehydration stock solution with IPG Buffer * (lysis buffer)
Component
Final concentration Amount 10 ml
___________________________________________________________________
Ultra pure 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 an eppendorf tube or for slightly tougher
tissue (e.g. 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 stickyness altogether by
having an optimal ratio between cell quantity and lysis buffer.
4. After lysis centrifuge 5 min. to get rid of particles/dell debris
5. Estimate the amount of protein with Bradford:
6.
7.
8.
9.
10.
…. l sample
Together 10 l
…. l lysis buffer
10 l H2O
10 l 0,1M HCl
1000 l Bradford 1:4
Measure the concentration on the spectrofotometer
Dilute the sample with lysis buffer to the desired concentration
Add IPG buffer (2,5 l in 500 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 an 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 stickyness altogether by
having an optimal ratio between cell quantity and lysis buffer.
2.
3.
4.
5.
6.
7.
Lyse during 1 hr at RT while mildly shaking (avoid foaming)
After lysis centrifuge 5 min. to get rid of particles/dell debris
Estimate the amount 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.
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 the sample in lysis buffer 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 an 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
3 Equilibration of the IPG gel strips
The IPG gel strips are equilibrated twice, each time for 15 min in equilibration buffer.
Equilibration buffer 1 contains DTT. Equilibration buffer 2 contains 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 40 ml portions at – 20 °C
Procedure: for 12 strips
1. Dissolve 400 mg of DTT in 40 ml of equilibration buffer,
(=Equilibration buffer I).
Pipette 3.2 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 1000 mg of iodoacetamide in 40 ml of equilibration buffer
(=Equilibration buffer II).
Pipette 3.2 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
4
SDS-PAGE as second dimension
Preparing SDS Ettan DALT slab gels
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
Filter through 0,45 um filter. 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% Acrylamide/Bis, 37.5 : 1 Biorad catalog No. 161-0158
Or
Duracryl High Tensile Strenght 30.65%T (30% Acrylamide/0,65% BIS) van
Genomic solutions catalog No. 80-0085
10% Ammonium persulfate
Ammonium persulfate
1.0 g
Deionised water.
10 ml
This solution should be prepared freshly just before use. Fresh ammonium persulfate “crackles”when
water is added. If does not, replace it with fresh stock.
Displacing Solution
( 375 mM Tris-Cl, pH 8.8, 50 % glycerol, bromopenol blue, 100 ml )
Tris-Cl (1.5M, pH 8.8)
25 ml
Glycerol
50 ml
Bromophenol blue
trace
Water
25 ml
This solution should be prepared freshly just before use.
7
Recipe for casting 13 vertical EttanDALT slab gels (1200 ml)
Component
Duracryl 30%
30% Acrylamide
37.5 : 1
Tris stock pH 8.8
Water
10% SDS
10% APS
TEMED
11% gel (ml)
220
12% gel (ml)
240
13% gel (ml)
260
220
300
438
12
10
0,5
240
300
398
12
10
0,5
260
300
358
12
10
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 1.5 ml of overlay for
each cassette.
3. Make up 100 ml of displacing solution.
4. Place 2 triangular sponge in de base of the V-shaped feed channel. Start filling the
gel caster by placing a separator sheet against the back wall. Fill the caster by
alternating glass plates with separator sheets, end with a separator sheet.
- For 13 gels: 13 glass plates and 1 fake glass plate, all separated by separator sheets
Place a gel label in each cassette. Don’t forget the rubber!
5. 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.
6. Load the balance chamber with 100 ml heavy displacing solution.
7. Make up the gel acrylamide stock solution, without adding the 10% APS or TEMED.
Mix well.
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.
8
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.
11. Immediately after removing the feed tube from the caster, slowly deliver 1.5 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.
9
SDS Electroforese buffer and filling the tank
10x TGS (Tris/Glycine/SDS buffer) van Bio-Rad cat. Nr. 161-0772
1.
2.
3.
4.
5.
Turn the pump valve to circulate
Fill the tank with water
Put the pump on
Turn the pump valve to drain and let some of the water out, not all of it!
Add 750 ml 10x TGS and fill to 7,5 liter with water, mix the buffer for a few hours
at the correct temperature
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 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 with some bromofenol blue) 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.
10
Loading Cassettes into the DALT tank
1. Once the electroforesis tank has reached the desired temperature and the gels are
ready, carefully slide the gels, one-by-one, into the tank. Until the buffer reaches the
bottom of the rubber sealing tubes, the cassettes should be lubricated with buffer or
water to prevent the rubber tubing from sticking to the cassettes. Be careful! The
plates slip easily once your hands are immersed in tank buffer!
NB. Forcing cassettes through the rubber tubes of the buffer seal without
sufficient lubrication can damage the buffer seal.
2. Fill any unused slots with the blank cassette inserts. When the last cassette is put
into place, buffer will be pushed out of the lower tank into the upper tank via the
two air vents at the corners of the sealing assembly.
NB. For electroforesis runs of six or fewer gels, it is helpful to alternate gel
cassettes with blank cassette inserts. Alternating cassettes will make it
considerably easier to remove the cassettes from the unit following the run.
The blank cassette inserts are easily removed first, leaving a larger gap that
makes it easier to grasp and remove the gel cassettes.
3. Adjust the buffer level after al cassettes are loaded in position.
4. 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.
5. Turn on the power supply
6. Recommended running conditions
Day run:
- Step 1
- Step 2
Set temperature to 25°C
2,5 W/gel
17 W/gel (max 180)
Overnight run: - 1,5 W/gel
Set temperature to 30°C
30 min
6 uur
17 hours
This counts for a 12,5% gel
11
Unloading and cleaning the separation unit
1. After the run has been completed, remove the blank cassette inserts or gels.
2. Open the cassettes using a wonder wedge and carefully transfer the gels to the
staining tray. Take care to ensure that the gel does not adhere to the spacers.
3. When all of the gels or blank inserts have been removed, drain the buffer by turning
the pump valve to drain with the pump on. Emptying will take about 1 min.
4. After the buffer has been removed, pour 3-4 l of distilled or deionized water into
the unit and allow it to drain. Rinse the unit with 5-7 l of distilled or deionized water
in circulate mode, empty again, and repeat if necessary.
12
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.
13
Protocol Coomassie Staining (G-250)
During all the washings, stainings etc. use continuous gentle agitation
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. Fixation: overnight in 50% ethanol/3% phosphoric acid
2. Wash steps: 3x15 min in water
3. Staining: in coomassie brillant blue G-250 solution for at least several hours to
days. No destaining is necessary
14
Protocol SYPRO Ruby protein gel stain
During all the washings, stainings etc. use continuous gentle agitation
1. Fixation: 3 hours in 10% Methanol/7% acetic acid
2. Wash steps: 3x 10 min. in water
3. Staining: Stain the gel for at least 3 hours for maximal sensitivity. 30-90 min. gives
specific staining. For convenience, gels may be left in the stain solution overnight
(16-18 hours) without overstaing.
4. Wash steps: 30-60 min. in 10% Methanol/7% acetic acid. This rinse step decreases
background fluorescence.
5. Wash gel in water before imaging
6. Scan gel on the FUJIFILM FLA-5000 (4th floor)
7. blue laser 473 and green filter
15