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Cleavage of fUSE55 with BglI and dephosphorylation with CIP
This protocol serves as an examplar for preparing large amounts of cut vector ready for ligation with an
insert.
MATERIALS
fUSE55 RF; 424 µg/ml; 9206 bp (see RFmaxiprep2.doc for large-scale RF preparation)
BglI 10,000 units in 1 ml (New England Biolabs)
10× NEBuffer 3 (New England Biolabs)
100× (10 mg/ml) BSA (New England Biolabs)
CIP (alkaline phosphatase from calf intestine) 1 U/µl (Roche)
8 µM test insert insert (two hybridized phophorylated synthetic oligos with ends compatible with the cut
vector)
CLEAVAGE WITH BglI AND DEPHOSPHORYLATION WITH CIP
1. In a 15-ml disposable screw-cap conical centrifuge tube pipette





6.85 ml water
1.2 ml 10× NEBuffer 2
120 µl 100× BSA
3.17 ml (1.344 mg) fUSE55 RF
675 µl (6750 units) BglI
Vortex gently; centrifuge briefly in clinical centrifuge; incubate at 37° overnight.
NOTE: During this incubation set up a 55º water bath.
2. Centrifuge briefly in the clinical centrifuge to drive solution to the bottom. Add 270 µl (270
units) of calf intestine alkaline phosphatase; vortex gently; incubate 30 min in 37º water bath.
3. Centrifuge briefly in the clinical centrifuge to drive solution to the bottom. Add an additional
405 µl (405 units) of alkaline phosphatase; vortex gently; incubate 30 min in 55º water bath.
4. Centrifuge briefly in the clinical centrifuge to drive solution to the bottom; add 600 µl 250mM EDTA pH 8; vortex gently; centrifuge briefly in the clinical centrifuge to drive solution to
the bottom; transfer half the solution (theoretically 6.69 ml) to a second conical disposable 15-ml
screw-cap tube.
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5. Extract each tube twice with 6 ml freshly neutralized phenol (you’ll need to neutralize about
25 ml altogether) and once with chloroform/isoamyl alcohol, using the “double-spin” method:
 vortex vigorously for ~1 min
 centrifuge to separate phases
 take upper (aqueous) phase greedily (including just enough of the interphase to ensure
that you have all the aqueous phase) to a second tube
 centrifuge again
 take the upper phase carefully (avoiding the interphase, which should be close to the
pointy bottom of the tube) to a third tube
Pipette the final aqueous phases into a single fresh 50-ml tube.
6. Dialyze in a 12-ml 10-KDa MWCO Slide-a-Lyzer (Pierce) against three changes of TE in the
cold.
7. Remove the dialyzed DNA into a tared 50-ml tube and determine net weight = net volume =
18.02 ml.
8. Using TE as diluent and reference solution, scan 100 µl of a 1/10 dilution from 320 to 220
nm. Data are graphed below. The DNA concentration is 61.8 µg/ml, and the total amount of
DNA recovered is 1.11 mg, which is 83% of the starting DNA at step 1.
0.15
0.125
Absorbance
0.1
0.075
Blank TE 1:10 Dialysis Buffer
Remove Blank Rescan
Remove Blank Rescan
fUSE55BglCIP
0.05
0.025
0
-0.025
220
240
260
280
300
320
Wavelength (nm)
NOTE: Since the total amount of recovered DNA is about 1 mg, we used two QIAGEN-tip 500s
(theoretical capacity 500 µg each) next step.
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PURIFICATION ON QIAGEN MAXIPREP
9. Pipette half the DNA (9 ml) step 7 (in a 50-ml tube) into a second 50-ml tube; to each tube
add QIAGEN buffer QBT to the 50-ml mark; invert many times to mix; centrifuge the tubes
briefly in the clinical centrifuge to drive the solutions to the bottom.
10. Mount two QIAGEN-tip 500s over a waste container in a suitable rack or ring-stand.
Equilibrate each tip with 10 ml QIAGEN buffer QBT, allowing the effluent to drain to the waste.
The column will stop flowing automatically when the meniscus reaches the upper frit.
11. Still collecting to waste, apply one tube of diluted DNA step 9 to each column and allow the
entire sample to flow through.
12. Still collecting to waste, wash each column twice with 30 ml QIAGEN buffer QC, allowing
the entire wash buffer to flow through each time.
13. Position each QIAGEN-tip column to collect into a fresh 50-ml tube; elute each column with
25 ml QIAGEN buffer QF; again, allow the buffer to flow completely through the column.
14. Transfer half (12.5 ml) of each eluate to a fresh 50-ml tube (there will be four 50-ml tubes in
all); to each 50-ml tube (with 12.5 ml eluate) add 9 ml (0.72 vol) isopropanol; mix thoroughly by
vortexing; mark one edge of each tube to designate the centrifugal wall (where the pellet will end
up); orienting the marked edges outward (centrifugally), centrifuge at 14 Krpm at 4º for 30 min
in FiberLite F15S-8×50C rotor1; pour off supernatant; re-centrifuge briefly to drive residual
supernatant to bottom, orienting the tube as before (the glassy pellet may not be easily visible);
suction off residual supernatant; air-dry briefly (no need to get rid of last drops of liquid).
15. Dissolve each of the four pellets in 750 µl TE; centrifuge the 50-ml tubes to drive the
solution to the bottom; pool all the solutions into a single one of the 50-ml tubes; dialyze the
pooled solutions in a single 3-ml, 10-KDa-cutoff Slide-A-Lyzer (Pierce) against three changes of
900 ml of 1/10 × TE; save some of the last outside fluid to use a diluent and reference at step 17.
16. Remove dialysate from the Slide-A-Lyzer into a tared 4-ml vial, recording the net weight =
volume in the table next step; label the vial and store it in the refrigerator or freezer.
17. In a 500-µl Ep tube make 100 µl of 1/20 dilution of the maxiprep and scan from 220 to 320,
using the saved final outside dialyis fluid step 15 as diluent and reference. The data are graphed
and tabulated below:
1
These excellent rotors allow you to spin standard disposable 50-ml conical screw-cap centrifuge tubes at 15,000
rpm! The pellet collects at the angle rather than the tip, which makes it particularly easy to free of residual
supernatant. If you don’t have this rotor, you can use OakRidge tubes and the Sorvall SS34 rotor (or equivalent
Beckman rotor) instead.
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Undiluted nucleic acid concentration
Volume (previous step)
Linear DNA size (without 14-bp stuffer)
Total yield
Percent recovery from 1.344 mg starting DNA
124.2 µg/ml (20.47 nM)
3.911
9192 bp
485.8 µg
36.14%
0.14
0.12
Absorbance
0.1
Blank 1:10 TE
Remove Blank Rescan
Remove Blank Rescan
fUSE55BglCIP1step17 1:20
0.08
0.06
0.04
0.02
0
-0.02
220
240
260
280
Wavelength (nm)
300
320
LIGATION AND ELECTROPORATION TEST
18. In a 500 µl Ep tube make 1× ligase buffer by mixing:
 180 µl water
 20 µl 10× ligase buffer with ATP (New England Biolabs)
19. Into a 500-µl Ep tube labeled Vector make 25.05 µl 2.5 nM cut vector in 1× ligase buffer:
 19.5 µl water
 2.5 µl 10× ligase buffer (previous step)
 3.05 µl cut vector step 16 (4-ml vial labeled top and side with orange tape in RB130)
Vortex; microfuge briefly to drive solution to bottom; pipette 4 µl from tube Vector into five
additional 500-µl Ep tubes labeled 0–4. Into the remaining solution in tube Vector (theoretically
5.05 µl left) pipette:
 7.6 µl 1× ligase buffer step 18
 3.1 µl lysis mix
Vortex tube Vector; save temporarily in the deepfreeze.
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20. In a 500-µl Ep tube labeled 525 make 525-nM test insert by mixing
 63.5 µl water
 7.6 µl 10× ligase buffer with ATP
 5 µl 8-µM test insert (see Materials)
Vortex; microfuge briefly to drive solution to the bottom. Label four additional 500-µl Ep tubes
26, 15, 9 and 5. Into tube 26 pipette 95 µl 1× ligase buffer step 18; into tubes 15, 9 and 5 pipette
15 µl 1× ligase buffer step 18. In tube 26 make a 1/20 dilution of tube 525 by passing 5 µl from
tube 525. In tubes 15, 9 and 5 (in that order) make serial 1.75-fold dilutions of tube 26 by
passing 20 µl. Pipette 2-µl portions into tubes 0–4 from previous step as follows:





2 µl from tube 26 this step into tube 4 previous step
2 µl from tube 15 this step into tube 3 previous step
2 µl from tube 9 this step into tube 2 previous step
2 µl from tube 5 this step into tube 1 previous step
2 µl of 1× ligase buffer step 18 into tube 0 previous step
21. In a 500-µl Ep tube labeled Enzyme pipette
 39 µl 1× ligase buffer step 18
 1 µl T4 DNA ligase
Vortex gently; microfuge briefly; pipette 4 µl into tubes 0–4 previous step; vortex tubes 0–4
gently; microfuge them briefly; incubate the tubes in the cold room overnight.
22. Pour an 8-lane 0.7% agarose/4×GBB minigel
23. In a 1.5-ml Ep tube premix
 1.15 ml water
 15 µl 10× TE
 35 µl 7.8-mg/ml yeast RNA (acts as carrier for low concentrations of DNA)
Vortex; dispense 200 µl into five 500-µl Ep tubes labeled E0–E4; save these 500-µl
electroporation Ep tubes in the refrigerator for use in next step.
24. Next day, microfuge the ligation tubes 0–4 step 21 briefly; pipette 2 µl from ligation tubes
0–4 into electroporation tubes E0–E4 previous step, respectively; into the remaining 8 µl in
ligation tubes 0–4 pipette 2 µl lysis mix; vortex tubes 0–4 and E0–E4 and microfuge them briefly.
25. Thaw 500-µl Ep tube Vector step 19 (80-hole rack on the bottom shelf of the door of
deepfreeze); vortex it and microfuge it briefly. Load 10-µl samples of tube Vector and tubes 0–4
onto the minigel step 22 along with marker as follows:
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10 µl .BstEII
10 µl tube Vector
All 10 µl of tube 0
All 10 µl of tube 1
All 10 µl of tube 2
All 10 µl of tube 3
All 10 µl of tube 4
×
Electrophorese at 30 V until BPB has run almost to the end; stain with sybr green for 2 hr;
photodocument as usual; here is the image:
There seems to be evidence of ligation only in Tube 4, with a 5.25:1 molar ratio of insert to
vector. In that tube, it looks like perhaps half the linear vector was converted to open circles.
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26. Set up the electroporator at 1250 V and following supplies:
 Rack with five 15-ml tubes labeled 0–4 containing 1.5 ml SOC with 0.2 µg/ml
tetracycline
 Sterile transfer pipettes in the 15-ml tubes
 Space in the S-I for strapping the rack in
 Ice bucket with electroporation tubes E0–E4 step 24
 Five 1-mm cuvettes on ice containing 55 µl 20% glycerol underlay (6.9 ml water, 2 ml
glycerol, 1 ml TE pH 8, 100 µl 10-µM phenol red)
 15 NZY/Tet plates (plates with NZY medium supplemented with 40 µg/ml tetracycline)
labeled E0-neat, E0-10-1, E0-10-2, E1-neat, E1-10-1, E1-10-2, E2-neat, E2-10-1, E2-10-2,
E3-neat, E3-10-1, E3-10-2, E4-neat, E4-10-1 and E0-10-2
27. Put five tubes with 22-µl aliquots of frozen electrocompetent MC1061 cells (see
ElectrocompetentCells.doc) in a bucket of powdered dry ice.
28. Immediately do five electroporations as follows
 Thaw one of the tubes of electrocompetent between finger tips
 Pipette 1 µl (60 pg) of one of the tubes E0–E4 into the electrocompetent cells and stir
with pipette tip; leave on on ice 30 sec.
 Carefully layer 18 µl (use regular yellow tip) on top of the red 20% glycerol underlay in
one of the cuvettes, being very careful to avoid bubbles.
 Zap
 Using the transfer pipette, draw up the SOC in the correspondingly labeled 15-ml tube,
use it to resuspend the zapped cells by vigorously pumping up and down a few times, and
transfer the resuspended cells back into the 15-ml tube (don’t worry about the small
volume that remains inaccessible in the cuvette).
29. Shake the 15-ml tubes at 37º for 45 min.
30. Meanwhile, label 15 sterile dilution tubes to correspond to the labeled NZY/Tet plates step
26. Pipette 450 µl SOC into all but the five neat tubes.
31. When the 45-min incubation step 29 is finished, pour each electroporation culture step 29
into the corresponding neat dilution tube. In the remaining two dilution tubes in each series
make serial 1/10 dilutions by passing 50 µl (e.g., in dilution tubes E1-10-1 and E1-10-2 make
serial 10-fold dilutions of dilution tube E1-neat). Spread 200-µl portions of all 15 dilution tubes
(including the neat tubes) on the 15 corresponding NZY/Tet plates; each neat plate represents
5.97 pg vector DNA. Incubate plates overnight at 37º. Enter the colony counts below.
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Electroporation
Concentration of vector during ligation (nM)
Concentration of insert during ligation (nM)
Neat
Number of colonies at following
10-1
dilutions
10-2
Electroporation efficiency (colonies/µg)
E0
1
0
0
0
0
<1.67e5
E1
1
0.98
5
1
0
8.37e5
E2
1
1.71
19
2
0
3.18e6
E3
1
3
25
1
0
4.19e6
E4
1
5.25
353
21
1
5.91e7
These results are satisfactory. The vector with a 5-fold molar excess of insert gives at least ~300
times more colonies than the vector without insert, and the yield per µg is OK given that the cells
are pretty old and even when new gave an efficiency of only ~1.5 × 108 colonies/µg with intact
RF.