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EMSA using ds Oligonucleotides

时间:2003-11-20 00:00来源:本站原创 作者:admin 点击: 864次

Solutions
10X Annealing Buffer
200 mM Tris 8.0 200 ml 1M Tris pH 8.0
10 mM EDTA 8.0 20 ml .5M EDTA pH 8.0
500 mM NaCl 100 ml 5M NaCl
280 ml Q
store at room temperature
10X Klenow Buffer
500 mM Tris 7.5 500 ml 1M Tris pH 7.5
100 mM MgCl2 100 ml 1M MgCl2
10 mM DTT 20 ml 0.5 M DTT
0.5 mg/ml BSA 50 ml 10 mg/ml BSA (NEB)
330 ml Q
store at -80° C in 50 ml aliq.
2X Binding Buffer (ref: Gutman et al. GENE 110, 1992 pg 197)
20% glycerol 400 ml 50% glycerol
20 mM Tris 7.5 20 ml 1M Tris pH 7.5
100 mM KCl 100 ml 1M KCl
1 mM DTT 2 ml 0.5 M DTT
478 ml Q
make fresh as needed
Poly dI/dC
Make a 1 mg/ml stock and store in 100 ul aliq. at -80° C
Procedure
? Design complementary oligonucleotides with compatible half sites on the ends (I use BamHI and BglII). Dry down 300 ng of the two purified oligos together and resuspend in 9 ml Q with 1 ml 10X Annealing buffer. Place in a 65° water bath for 2 min and cool in 50 ml of the 65° water in a beaker on ice. This takes 15-20 minutes.
? Mix the following and incubate at room temperature for 30 min:
1 ml of the annealed oligo mixture
5 ml 10X Klenow buffer
25 ml dNTP mix (21 ml Q, 3 ml 10 mM dATP/dTTP dGTP)
5 ml a32P dCTP
14 ml Q
1 ml Klenow
Bring up to 200 microliters with TE, phenol/chloroform extract and ethanol precipitate with 0.1 volume of 3 M NaOAc pH 5.2 as well as tRNA. Dry and resuspend in 100 ml TE and count. I usually get 200,000-400,000 cpm per ml.
? Generate a table of binding reaction parameters and competitor concentrations. Prepare the gel and running buffer.
EMSA Gel:
8 ml 30% acrylamide (0.8% BIS)
6 ml 50% glycerol
6 ml 10X TBE
40 ml Q
180 ml 10% APS, 180 ml TEMED
Cool to 4° C along with the appropriate amount of 1X TBE.
? Mix the binding reagents in the following order:
i) 10 ml 2X Binding Buffer
ii) 3 ml dI/dC
iii) Q to 20 ml (see table)
iv) competitor at 10-20X excess
v) 10-50 fold dilution of the probe
vi) NE usually 2-4 mg is sufficient
? Incubate at room temperature for 30 minutes and load directly onto gel with no loading dye. Run the gel at 200V for 4 hours. Note: it is best to pre-run the gel during the 30 min binding reaction.
 
Nuclear Extract Preparation
This is the standard protocol with only minor modifications. The volumes indicated are for 6 liters of cells at 1-2 x106 cells per ml.
Solutions
Solution A (Equilibration Buffer: 100 ml)
10 mM HEPES 7.9 (pH with KOH) 1 ml 1M HEPES pH 7.9
1.5 mM MgCl2 150 ml 1M MgCl2
10 mM KCl 1 ml 1M KCl
up to 100 ml with Q
for every 100 ml add:
100 ml .5 M DTT
500 ml PMSF (Sigma #P7626)
100 ml pepstatin (2.0 mg/ml)
100 ml leupeptin (1.0 mg/ml)
100 ml aprotinin (5.0 mg/ml)
(Protease Inhibitor Cocktail Sigma #P8340)
1.0 ml 0.1 M EGTA
1.0 ml 10% NP40
Solution B (Low Salt Buffer: 100 ml)
20 mM HEPES 7.9 2.0 ml 1M HEPES 7.9
1.5 mM MgCl2 150 ml 1M MgCl2
20 mM KCl 2.0 ml 1M KCl
0.2 mM EDTA 40 ml .5 M EDTA
up to 100 ml with Q
for every 100 ml add:
100 ml .5 M DTT
500 ml PMSF
Protease inhibitors (see Solution A)
1.0 ml 0.1 M EGTA
1.0 ml 10% NP40
Solution C (High Salt Buffer: 10 ml)
20 mM HEPES 7.9 200 ml 1M HEPES 7.9
1.5 mM MgCl2 15 ml 1M MgCl2
1.2 M KCl 3.0 ml 4M KCl
0.2 mM EDTA 4 ml .5 M EDTA
up to 10 ml with Q
for every 10 ml add:
10 ml .5 M DTT
50 ml PMSF
Protease Inhibitors (see Solution A)
100 ml 0.1 M EGTA
100 ml 10% NP40
Solution D (Dialysis Buffer: 2 liter)
20 mM HEPES pH 7.9 40 ml 1M HEPES 7.9
100 mM KCl 14.9 g KCl
0.2 mM EDTA 800 ml 0.5 M EDTA
20% glycerol 400 ml glycerol
up to 2 liter with Q
for every 2 liters add:
4.0 ml 100 mM PMSF
2.0 ml 0.5 M DTT
Procedure
GENERAL NOTES: I try to gently dislodge the pellet after every step to avoid pipeting up and down to resuspend the cells or nuclei. If a given spin is not sufficient to pellet the cells or nuclei I have found a quick spin at higher rpms is useful (such as 5’ at 1.5-2K).
? Cool the SA600 rotor with the small adaptors to 4° and cool the 50 ml tube adaptor for the J6B.
? Harvest cells in 1 liter bottles by spinning in J6B at 1.2K for 30' @ 4o. Pour off most of the medium and as the cells come loose, stop. Pool the remaining medium and cells into 2-4 250 ml conical bottles and spin at 1.2K for 10 minutes. Gently resuspend in 4x 50 ml HBSS, transfer to a 50 ml tube, and spin in the J6B for 5 minutes at 1.2K. Aspirate the HBSS and gently flick the tube with your finger to dislodge the pellet.
? Resuspend the pellet up to 2x the pellet volume of Solution A and pool into 2x 50 ml tubes. (I generally get 15-25 ml cells total and resuspend up to 30 to 50 ml).
? Spin in the J6B for 5' at 1.2K and gently flick the tube to dislodge the pellet, then resuspend in the remaining Solution A (there should be about 70-80 ml left). Equilibrate on ice for 20'. The Solution A plus the cells should be about 100 ml.
? Take a look at the cells with Trypan Blue to see if they have already lysed. If the cells still appear to be intact, dounce approx. 30 ml 10 times and recheck with the Trypan Blue. Repeat this until >90% of the nuclei take up the dye.
? Spin in the J6B for 5' at 2K and aspirate the supernatant. Flick the tube with your finger to gently dislodge the pellet (it may be a little sticky at this point).
? Gently resuspend the pellet up to 40 ml final volume with Solution B and spin in the J6B for 5' at 2K. You may lose some nuclei here, but it is better than packing them too tightly.
? Remove the supernatant and flick the tube with you finger to gently dislodge the pellet.
? Resuspend the nuclei in 0.5 volumes of Solution B and slowly add 0.5 volumes of Solution C (Add it dropwise with a pasteur pipet while vortexing on 3 or 4).
? Mix on the tiltboard for 30' @ 4o.
? Transfer the nuclei into a 15 ml glass Corex tube and spin in the SA600 for 30' at 10K.
? Dialyze the supernatant against 2x 1 liter Solution D.
? Quick freeze in liquid Nitrogen and store at -80o.
HELA CELL NUCLEI PREP
I. Solutions:
a. Chelsky Buffer (200ml)
0.01M Tris-HCl
0.01M NaCl
0.003M MgCl2
0.03M Sucrose (mw. 342.3 g/mole)
pH to 7.0 and bring up to 200ml.
b. NP-40 Buffer
100ml of Chelsky Buffer
500ul of NP-40 detergent
c. Cacl2 Buffer
100ml of Chelsky Buffer
1.0ml 100x stock CaCl2 sol'n (10mM)
d. Buffer C
20mM Tris-HCl pH 7.9
20% glycerol
0.1M KCl
0.2mM EDTA
pH to 7.9
II. Procedure:
a. Cell collection:
Plate Helas on 150mm plates on day 1. (1 x107 cells yields about 1 mg of total cellular protein) Approximately 32-150mm plates.
Trypsinize every plate on day 2-3 (am).
Bring each plate up in 5ml of media, and count the total number of cells\ml. Record the volume of cell suspension.
b. Cell treatment:
Note: Everything (buffers, cells, centrifuge) must be kept on ice (4oC) throughout this entire procedure, to reduce protein degradation.
Note: Save the supernatant from each wash to keep track of protein if lost.
Wash the Hela cells 2x with 25 ml of cold PBS per tube in clinical centrifuge setting #3 in cold room(4oC).
Resuspend cell pellets in 2.0 ml of NP-40 Buffer, spin in HB-40 rotor at 3000rpm (1500xg) for 10mins,remove supernatant and resuspend as above.
Repeat NP-40 wash step above.
Remove supernatant and resuspend pellet into 2.0ml of CaCl2 Buffer.
Spin in HB-40 rotor at 3000 rpm (1500xg) for 10mins, remove supernanat and resuspend as above. Repeat CaCl2 wash step.
Resuspend pellet from second CaCl2 wash step into 2.0 ml of Buffer C, depending on how much protein was collected (use 1ml/number of cells)
Spin balanced tubes in SS34 rotor at 14,500 rpm(25,000xg) for 30mins.
Collect supernatant (nucleoplasm), resuspend pellet (nuclear envelope) in Buffer C and homogenize to get into solution.
Aliquot into cryovials and freeze at -70oC.
Oligonucleotide Purification
Diluent 5X Buffer 25% Acrylamide
209 g Urea 209 g Urea 209 g Urea
up to 500 ml Q 250 ml 10X TBE 120.8 g Acrylamide
up to 500 ml Q 4.1 g BIS
up to 500 ml Q
2.5 M NH4OAc
19.2 g NH4OAc
up to 100 ml Q
Formamide Dye
9 ml deionized formamide
500 ml 10X TBE
500 ml 0.2% bromophenyl blue and 0.2% xylene cyanol
Other Reagents Needed:
0.22 mM disposable syringe filters, short wave UV source, intensifying screen for UV shadowing
Procedure
? Pour a 20% denaturing gel: 12 ml 5X Buffer, 48 ml 25% acrylamide, 500 ml 10% APS (Gibco #15523-012), 20 ml
TEMED (Sigma #T8133).
? Dry down 100 mg oligonucleotide and resuspend in 50 ml Formamide Dye.
? Boil for 2 minutes and hold on ice until ready to load gel.
? Run the gel at 15 W constant power until the fast dye is at the bottom of the gel.
? To visualize the bands, UV shadow on the intensifying screen. Excise the band and crush and soak in 1 ml 2.5 M
Ammonium Acetate at 37°C for several hours to overnight.
? Spin for 10 minutes and remove the supernatant. Add another 1 ml of 2.5 M ammonium acetate, pellet and pool with the previous eluate.
? Filter the supernatant through a 0.22 mM filter, add 0.1 volumes of 3M NaOAc 5.2 and 2 volumes EtOH. I usually add 1 ml glycogen also.
? Spin for 30 minutes, wash and dry. Resuspend in 50 ml TE and quantitate 5 ml.
Gel Mobility Shift Assay Conditions -Mg/EDTA in Gel and Buffer
Protein Dilution Buffer
5ml 20 mM Tris pH7.9
100 microliters 1M Tris 7.9
150 mM KCl 0.75 ml 1 M KCl
1 mM DTT 50 microliters 0.1 M DTT
10% glycerol 1 ml 50% glycerol
50 micrograms/ml BSA 2.5 microliters 100 mg/ml BSA
3.1 ml H2O
Optional: add Brij 58 to 0.1%.
Store dilution buffer at -70 degrees.
Proteins are diluted in dilution buffer and quick frozen on dry ice. Thaw proteins on ice. Proteins are typically stable to multiple repeated freeze thaw.
5x binding buffer
1 ml 20% glycerol 400 microliters 50% glycerol
100 mM Tris-HCl pH8 @ 25 degrees 100 microliters 1 M Tris pH8
300 mM KCl 300 microliters 1 M KCl
25 mM MgCl2 25 microliters 1 M MgCl2
500 micrograms/ml BSA 5 microliters 100 mg/ml BSA
170 microliters H2O
optional: add 25 microliters saturated bromophenyl blue [BioRad] (~0.1% in H2O) per ml of 5X buffer (this may
inhibit the binding of some proteins)
Store buffer at -70 degrees.
Gel shift reactions are performed as follows:
20 microliter binding reaction:
4 microliters 5X binding buffer
0.2 microliters 0.1 M DTT
2000-5000 cpm labeled DNA
0.125 micrograms p[dG-dC]
H2O to 20 microliters final volume
Add proteins to reaction last. Incubate protein and DNA at room temperature for ~30-40 min and load to native gels which are run in the cold room at 4 degrees. Gels are not pre cooled but are set in cold room 5-10 minutes before loading and pre run at 160 V. The wells of the gels are rinsed out several times before prerunning and again before loading. Samples are applied to the gel while the gel is running. For best results, use a fine tip pipetman tip to load the gels. We run gels at 160V (12 cm long) for ~45 min.
For a typical gel shift reaction (20 microliter reaction), I use 1-2 ng TBPc (the conserved region of yeast TBP from the Sigler lab) and and 5-10 ng of wild-type or truncated yeast TFIIB. The amount of proteins will have to be titrated for your specific conditions.
Gels:
10.5 ml (20%/0.33%) acrylamide/bis acrylamide
3.5 ml 10X TGOE buffer
1.75 ml 50% glycerol
35 microliters 0.5 M DTT
20.9 ml H20
0.3 ml 10% ammonium persulfate
30 microliters TEMED
10X TG0E 500 ml
0.25 M Tris 15.1 g
1.9 M glycine 71.3 g
pH 8.3 with acetic acid at room temp. Adjust volume to 500 ml.
Running buffer is 1X TGOE buffer
[Note that there is no Mg or EDTA in both the gel and in the running buffer.]
Kinasing oligonucleotides with 32P ATP
Wear gloves throughout and work in radiation area. Monitor area before and after use.
Mix the following in an eppendorf tube:
1. 0.5 microgram oligonucleotide dissolved in H2O.
2. 3 microliters 10x kinase buffer.
3. 2 microliters 32P ATP from ICN (>5000 ci/mmole).
4. H20 so that the final volume is 30 microliters.
Add 25 units T4 polynucleotide kinase and incubate 60 min at 37 deg.
Purify labeled oligo using MERMAID (read manufacturers instructions):
Add to kinase reaction:
1. 3 microliters 5M Ammonium Acetate
2. 90 microliters high Salt Mermaid Buffer
3. 6 microliters Mermaid Beads
Bind oligo to beads for 10 min with constant mixing by shaking at low speed.
Spin in microfuge for 10 sec and remove supernatant. (Dispose of supernatant in radioactive liquid waste bottle)
Wash beads with 100 microliters high salt buffer. Vortex, spin and remove sup.
Wash beads 3 times with 300 microliters Mermaid ETOH wash.
Elute labeled oligo from beads two times with 50 microliters TE at 55 deg. Store oligo frozen at -20 deg.
Quantitate radioactive incorporation by counting 2 microliters of a 1/100 diluted sample. Expect between 20 -100 million cpm total.
10x Kinase Buffer
0.5 M Tris pH 7.6
0.1 M MgCl2
50 mM DTT
Ethanol Precipitation
1. Measure the volume of nucleic acid containing sample. If the volume is less than 450 ul, place it in a 1.5 ml eppendorf tube. If the volume is between 450 ul and 4.5 ml, place it in a 15 ml baked siliconized Corex tube. If the volume is between 4.5 ml and 9 ml, place it in a 30 ml Corex tube.
2. Add 0.1 times the volume of 3M NaAc pH 5.2 *. Mix. Calculate the new volume, then add 2 volumes of cold ethanol. Seal the tube tightly with parafilm. Mix.
* It is usually preferable to use ammonium acetate(1/2 volume of 7.5M) instead of NaOAc as the salt. It is more volatile, and leaves behind more protein. However, the larger volume may be inconvenient.
3. Place the sample at -20deg. (If there is a lot of DNA, you can proceed almost immediately. If there is only a few hundred nanograms, it is best to wait at least one hour or overnight) or at -70deg. for 1 hr. For dilute samples, better recovery is obtained at -20deg. O/N.
4. If the precipitation is being done in a microfuge tube, spin at least 15 minutes in the TOMY at 4deg. (For 15 ml or 30ml Conical tubes, spin 20 minutes in the Sorvall in swinging bucket rotor). IT IS IMPERATIVE THAT YOU ARE READY TO TAKE THE TUBES IMMEDIATELY AFTER THE SPIN STOPS. If the machine has stopped for even a minute by the time you get to it, turn it back on for an additional 5 minutes. Otherwise, you run a great risk of the pellet becoming dislodged and being dumped out. Don't try to precipitate too many tubes at once. It will take too long to deal with them all, and you may lose the last few pellets. 5. Dump off the ethanol and tap the tube (upside down) on a paper towel to remove as much EtOH as possible. Keep the tube upside down. Quickly dry off the inside of the tube with a Q-tip, but don't touch the pellet. Put the tube into an upright position, and dry in Speed-Vac for 5 minutes.

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