533583453 Page 1 3/9/2016
Construction of a large random peptide library (RPL) in fUSE55
PRELIMINARY COMMENTS
This protocol serves as an exemplar for constructing a large random peptide library (RPL). In
this particular RPL the displayed peptide has a fixed hexapeptide sequence KCCYSL with
flanking randomized amino acids (the Xs):
xxxxxKCCYSLxxxx
The N-terminus of the mature pIII protein (after signal peptidase cleavage) will be:
ADGAxxxxxKCCYSLxxxxGAAGAETVE
Here in outline is the scheme for preparing vector-ready insert:
CTCACTCGGCCGACGGGGCCNNKNNKNNKNNKNNKAAATGCTGTTATAGCCTG
CAGTTTCGGCCCCAGCGGCCCCMNNMNNMNNMNNCAGGCTATAACAGCATTT
Klenow
QIAGEN midiprep
Concentrate on Centricon
CTCACTCGGCCGACGGGGCCNNKNNKNNKNNKNNKAAATGCTGTTATAGCCTGNNKNNKNNKNNKGGGGCCGCTGGGGCCGAAACTG
CAGTTTCGGCCCCAGCGGCCCCMNNMNNMNNMNNCAGGCTATAACAGCATTTMNNMNNMNNMNNMNNGGCCCCGTCGGCCGAGTGAG
A
D
G
A
X
X
X
X
X
K
C
C
Y
S
L
X
X
X
X
G
A
A
G
A
E
T..
G
A
E
T..
SfiI
QIAGEN midiprep
Concentrate on Centricon
GGGCCNNKNNKNNKNNKNNKAAATGCTGTTATAGCCTGNNKNNKNNKNNKGGGGCCGCTG
CGGCCCCMNNMNNMNNMNNCAGGCTATAACAGCATTTMNNMNNMNNMNNMNNGGCCCCGT
A
D
G
A
X
X
X
X
X
K
C
C
Y
S
L
X
X
X
X
G
A
A
The foregoing scheme exploits the complementary non-degenerate sequences at the 3´ ends of
the starting oligos to allow the two non-contiguous degenerate segments to be synthesized in two
separate oligonucleotides. When you are making a fully degenerate RPL, in contrast, all the
degenerate nucleotides will have to be synthesized in a single, longer oligonucleotide, which will
serve as the template for a non-degenerate primer base-paired to its non-degenerate 3´ end:
degenerate segment
5’
5’
Klenow extension
5’
5’
533583453 Page 2 3/9/2016
The procedure for preparing vector-ready insert is not affected by this change in oligonucleotide
structure.
Electroporation is key to library construction. General comments on electroporation, as well as a
small-scale procedure used to check several stages of the RPL construction below, can be found
in electroporation.doc. The main electroporation at steps 53–57 is a 133-fold scale-up of the
small-scale procedure.
SPECIAL SOLUTIONS FOR QIAGEN ION EXCHANGE CHROMATOGRAPHY OF
SMALL DNAs
NOTE: Standard solutions and procedures are italicized and are found in stdpreps.doc. Here we
give recipes for special solutions required for ion exchange chromatography of small DNA
inserts on QIAGEN columns.
Buffer QB (750 mM NaCl, 50 mM MOPS, 15% ethanol, pH 7.0):





Dissolve 42.70 grams NaCl and 11.56 grams MOPS (sodium salt) in 800 ml water
Adjust pH to 7.0 with HCl
Add 150 ml 100% ethanol
Adjust volume to 1000 ml in a 1-liter graduated cylinder
Store at room temperature
Buffer QP (400 mM NaCl, 50 mM MOPS, 15% ethanol, pH 7.0)





Dissolve 22.247 grams NaCl and 11.56 grams MOPS (sodium salt) in 800 ml water
Adjust pH to 7.0 with HCl
Add 150 ml 100% ethanol
Adjust volume to 1000 ml in a 1-liter graduated cylinder
Store at room temperature
QIAelution buffer (1.25 M NaCl, 50 mM Tris, 15% ethanol, pH 8.5)





Dissolve 73.05 grams NaCl and 6.055 grams Tris in 800 ml water
Adjust pH to 8.5 with HCl
Add 150 ml 100% ethanol
Adjust volume to 1000 ml in a 1-liter graduated cylinder
Store at room temperature
PREPARATION OF VECTOR-READY DEGENERATE INSERT
1. In a 500-µl Ep tube mix 1 nmol of each oligonucleotide and 10 µl 10× NEBuffer 2 (New
England Biolabs) in a total volume of 88 µl; vortex; microfuge briefly; incubate 15 min at 50ºC
and 15 min at 37ºC; allow to cool to room temperature.
533583453 Page 3 3/9/2016
NOTE: The scale of the synthesis was dictated by the amount of the least abundant synthetic
oligonucleotide available—1 nmol in this case. If we’d had more of the limiting oligonucleotide,
we’d have increased the scale of synthesis accordingly. You need to consider that scale in
determining the capacity of QIAGEN column to use at step 7 (see note under step 4).
2. Microfuge the 500-µl Ep tube briefly; add 1 µl 25-mM dNTP mixture; vortex gently;
microfuge briefly.
3. Label a 500-µl Ep tube “hybridized oligos step 3”; into it pipette:
 11 µl water
 2 µl 70/75/BPB
 1 µl hybridized oligos previous step
Store the tube in the deepfreeze.
4. Meanwhile, to the bulk of the primed template in the 500-µl Ep tube step 2 add 2 µl (10 units)
exo- Klenow fragment (Fermentas); vortex gently; microfuge briefly; incubate at 37º for 1 hr.
NOTE: Theoretically this Klenow extension makes 1 nmol of an 87-bp double-stranded product;
that corresponds to a theoretical mass of 57 µg—well within the capacity of the QIAGEN-tip 100
midiprep column (nominal capacity 100 µg) used at step 7.
5. Microfuge the 500-µl Ep tube previous step briefly; add 5 µl 250-mM EDTA; vortex gently;
microfuge briefly.
6. Label a 500-µl Ep tube “87 bp primer extension”; into it pipette
 11 µl water
 2 µl 70/75/BPB
 1 µl step 5
Store the tube in the deepfreeze.
NOTE: QIAGEN purification (next six steps) is intended to free the 87-bp primer extension
product of enzyme and other high- and low-molecular weight components and contaminants in
preparation for SfiI digestion at step 14.
7. Set up a QIAGEN plasmid midi kit QIAGEN-tip 100 in a suitable rack over a waste
container; equilibrate it by passing 5 ml QIAGEN QBT then 10 ml buffer QP (see Materials)
through it, collecting into waste.
8. Transfer the remainder of the 500-µl Ep tube step 5 to a 15-ml tube containing 5 ml buffer QP
(see Materials); vortex to mix; load the DNA into the column previous step, continuing to collect
into waste.
533583453 Page 4 3/9/2016
9. Flow an additional 10 ml QP through the column, continuing to collect to waste.
10. Wash the column with four 5-ml portions of buffer QB, continuing to collect to waste.
11. Elute the column with two 3.6-ml portions of QIAelution buffer, collecting into a fresh 50ml tube.
12. Concentrate in two passes (1.8 ml per centricon per pass) on two 10-KDa Centricons
(Millipore); wash three times with 1/10 × TE, concentrating all the way each time. Collect
retentates as usual by back-centrifugation into conical collection tubes (see Millipore
instructions); transfer to a single 500-µl Ep tube with a 100-µl pipetter, measuring volumes as
you do so; the combined volume was 188 µl. Store in freezer.
13. Scan 100 µl of a 1/50 dilution from 220 to 320 nm, using 1/10 × TE as diluent and reference
(save the 1/50 dilution for step 22 below). The data are graphed below. The calculated
concentration of the 87-bp purified primer extension product is 228 µg/ml = 3.97 µM; the total
yield in the 188 µl is therefore 0.746 nmol (74.6% of theoretical).
0.1
Absorbance
0.08
0.06
Blank TE 1:10
Remove Rescan Blank
Remove Rescan Blank
Erb1.docstep12
0.04
0.02
0
-0.02
220
240
260
280
300
Wavelength (nm)
14. To a 1.5-ml Ep tube add:





604 μl water
100 μl of 10× NEBbuffer 2 (New England Biolabs)
10 μl of 100× (10-mg/ml) BSA (New England Biolabs)
The remaining 186 μl (56 µg) of the extension reaction step 12
100 μl (2000 units) SfiI (New England Biolabs)
Incubate overnight at 50ºC.
320
533583453 Page 5 3/9/2016
NOTE: QIAGEN column purification (next six steps) is intended to free the 60-bp vector-ready
insert of enzyme, cut-off end pieces (13–14 bp) and other high- and low-molecular weight
components and contaminants in preparation for ligation. Special buffers for chromatography of
small DNAs like the insert here are given in Solutions above.
15. Set up a QIAGEN plasmid midi kit QIAGEN-tip 100 in a suitable rack over a waste
container; equilibrate it by passing 5 ml QIAGEN buffer QBT then 10 ml buffer QP (see
Solutions) through it, collecting into waste.
16. Microfuge the restriction digest step 14 briefly to drive solution to the bottom; transfer the
solution to a 50-ml tube containing 15 m buffer QP (see Solutions); vortex to mix; load the DNA
into the column from the previous step, continuing to collect into waste.
17. Wash the column with an additional 10 ml QP, continuing to collect to waste.
18. Wash the column with four 5-ml portions of buffer QB (see Solutions), continuing to collect
to waste.
19. Elute the column with two 3.6-ml portions of QIAelution buffer (see Solutions), collecting
into a fresh 50-ml tube labeled “eluate.”
20. Concentrate in two passes (1.8 ml per centricon per pass) on two 10-KDa Centricons; wash
each Centricon three times with 1/10 × TE, concentrating all the way each time. Collect
retentates as usual by back-centrifugation into conical collection tubes (see Millipore
instructions); transfer both retentates to a single 500-µl Ep tube with a 100-µl pipetter, measuring
volumes as you do so; in this case the combined volume was 81 µl. Store in deepfreeze. In the
next step the nominal concentration is measured at 6.49 µM, but the actual concentration of
vector-ready insert is probably more like 3 µM.
21. Make 100 µl of a 1/50 dilution and scan from 220 to 320 nm, using of 1/10 × TE as diluent
and reference (save the 1/50 dilution for next step). The results are graphed below:
0.12
Absorbance
0.1
Blank 1:100 TE
Remove Blank Rescan
Remove Blank Rescan
Erb2.docstep8 1:50 dil
0.08
0.06
0.04
0.02
0
-0.02
220
240
260
280
Wavelength (nm)
300
320
533583453 Page 6 3/9/2016
The DNA concentration in the undiluted concentrate is 257 μg/ml; assuming that all the DNA is
in the form of the intended 60-bp doubly-cut vector-ready insert, this corresponds to a molar
concentration of 6.49 µM and a total molar yield of 0.526 nmol (52.6% of the starting
oligonucleotides). In fact, as shown at step 23 below, only about half the DNA mass is in the
form of this 60-bp insert, so the actual yield is probably ~0.25 nmol.
22. In two 500-µl Ep tubes make electrophoresis samples as follows:
 Purified primer extension:
o 12 µl (54.7 ng) of the 1/50 dilution step 13
o 2 µl 70/75/BPB
 Vector-ready insert:
o 12 µl (61.7 ng) of the 1/50 dilution previous step
o 2 µl 70/75/BPB
 LMW DNA ladder:
o 550 µL of water
o 100 µL 70/75/BPB
o 50 µl low molecular weight DNA ladder (New England Biolabs); 357 µg/ml
23. Set up a BioRad 10% acrylamide 1× TBE Ready Gel in the BioRad Ready Gel Cell, using a
356:44 vol/vol mixture of 1×TBE and water as the running buffer (BioRad’s TBE has 89 mM
Tris and borate; our TBE had 100 mM Tris and borate). Into the middle six lanes load 14-µl
electrophoresis samples as indicated below. Electrophorese at 70 V until BPB has reached to  1
cm from the bottom. Stain with sybr green and photodocument as usual. Here is the image (sizes
of the LMW DNA ladder markers are shown at the left):
LMW DNA
ladder step 22
Vector-ready
insert step 22
LMW DNA
ladder step 22
Purified primer
extension step 22
Primer extension step 6
Hybridized
oligos step 3
533583453 Page 7 3/9/2016
Evidently SfiI cleavage is incomplete: in addition to the desired 60-bp doubly-cut insert, there’s a
little uncleaved 87-bp primer extension product and one or both of the two singly-cut products
(73–74 bp). Despite these imperfections, this preparation is acceptable as “vector-ready” insert.
TEST LIGATIONS
24. 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)
25. Into a 500-µl Ep tube labeled Vector make 25 µl 2.5 nM cut vector in 1× ligase buffer:
 2.5 µl 10× ligase buffer (previous step)
 62.4 pmol (378.8 ng) cut fUSE55 vector (see CutVector.DOC:17); this vector has been
cleaved with BglI, 5´-dephosphorylated with calf intestine alkaline phosphatase, and
purified by ion exchange chromatography on a QIAGEN column to remove enzymes, the
14-bp stuffer (released by BglI digestion) and other components and contaminants.
 water to bring the total volume to 25 µl
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 µl left) pipette:
 7.6 µl 1× ligase buffer step 18
 3.1 µl lysis mix (5× electrophoresis sample buffer)
533583453 Page 8 3/9/2016
Vortex tube Vector; save temporarily in the deepfreeze.
26. In a 500-µl Ep tube labeled 260 make 30 µl of 260-nM insert by mixing
 25.8 µl water
 3 µl 10× ligase buffer with ATP (see step 24)
 1.2 µl 6.49-µM insert step 20
Vortex; microfuge briefly to drive solution to the bottom. Label four additional 500-µl Ep tubes
52, 26, 13, and 6.5. Into tube 52 pipette 8 µl water and 2 µl 260-nM insert; into tubes 26, 13, and
6.5 pipette 5 µl 1× ligase buffer step 24. In tubes 26, 13, and 6.5 make serial 2-fold dilutions of
tube 52 by passing 5 µl and vortexing. Pipette 2-µl portions of these insert tubes and of 1× ligase
buffer into vector tubes 0–4 previous step as follows:





2 µl from tube 52 this step into tube 4 previous step
2 µl from tube 26 this step into tube 3 previous step
2 µl from tube 13 this step into tube 2 previous step
2 µl from tube 6.5 this step into tube 1 previous step
2 µl of 1× ligase buffer step 24 into tube 0 previous step
27. 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 (~8ºC) overnight.
28. Pour an 8-lane 0.7% agarose/4×GBB minigel
29. 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.
30. Next day, microfuge the ligation tubes 0–4 step 27 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.
533583453 Page 9 3/9/2016
31. Thaw 500-µl Ep tube Vector step 25; vortex it and microfuge it briefly. Load 10-µl samples
of tube Vector and tubes 0–4 previous step (not tubes E0–E4) onto the minigel step 28 along
with marker as follows:
10 µl (400 ng) .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
×
Tube 4
Tube 3
Tube 2
Tube 1
Tube 0
Vector
.BstEII
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 in Tubes 1 and 2, with 1.3 and 2.6 molar ratios of insert to
vector, respetively. In Tube 2 (molar ratio 2.6) it looks like perhaps 20% of the linear vector was
converted to open circles. This is about as good as we achieve with a library ligation.
533583453 Page 10 3/9/2016
32. Set up the electroporator at 1250 V and a time constant of 10 ms (for example, 400 Ω × 25
µF = 10,000 µs = 10 ms in our electroporator design electroporator.doc), 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
33. Put five tubes with 22-µl aliquots of frozen electrocompetent MC1061 cells (see
ElectrocompetentCells.doc) in a bucket of powdered dry ice.
34. 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).
35. Shake the 15-ml tubes at 37º for 45 min.
36. Meanwhile, label 15 sterile dilution tubes (capless 2.2-ml polypropylene tubes from
Sarstedt) to correspond to the labeled NZY/Tet plates step 32. Pipette 450 µl SOC into all but
the five neat tubes.
37. When the 45-min incubation step 35 is finished, pour each electroporation culture step 35
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.
533583453 Page 11 3/9/2016
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
0
E1
1
1.3
60
9
0
1.0×107
E2
1
2.6
195
18
1
3.3×107
E3
1
5.2
36
51
0
7.2×106
E4
1
10.4
50
1
0
8.4×106
The optimal transfection efficiency was observed from tube E2; it is acceptable for a large
library, but about 1/30 of the transfection efficiency (109 colonies/µg) obtained by transfecting
good electrocompetent cells with purified RF DNA. Since ligation Tube 2 also gave the
strongest evidence of ligation by gel electrophoretic analysis step 31, we used the conditions of
Tube 2 (1 nM vector, 2.6 nM insert) for the large-scale (80-ml) ligation.
LARGE-SCALE LIGATION
38. In a 15-ml tube make 10 ml of 10× ligase buffer:






2.75 ml water (sufficient to bring total volume to 10 ml after adding other components)
500 mM Tris.HCl pH 7.5 (5 ml 1 M stock pH 7.5)
100 mM MgCl2 (1 ml 1 M stock)
10 mM diothreitol (15.4 mg powder)
10 mM ATP (1 ml 100-mM stock)
250 µg/ml BSA (250 µl 10-mg/ml BSA from New England Biolabs)
39. Into a 50-ml tube pipette
 3.9 ml (79.8 pmol) cut fUSE55 vector (see CutVector.DOC:17; see step 25)
 32 µl (207.7 pmol) vector-ready insert step 20
 Water to bring the total volume to 20 ml
Measure half (10 ml) into a second 50-ml tube, so that there are now two 50-ml tubes. Into each
50-ml tube pipette:
 4 ml 10× ligase buffer previous step
 250 µl (250 units) T4 DNA ligase (Roche)
 Water to a total volume of 40 ml
Incubate overnight in a cold room (~8ºC). The final vector concentration is 1 nM; the final
nominal insert concentration is 2.6 nM. The total mass of DNA in both tubes combined is 492
µg.
40. To each 50-ml tube add:
 2 ml 250 mM EDTA pH 8 (1.25 molar ratio to Mg2+)
533583453 Page 12 3/9/2016
 3.6 ml of 5 M NaCl (increasing total NaCl concentration to ~400 mM)
 1.77 ml 0.5 M MOPS sodium salt adjusted to pH 5.7 with HCl (this had been determined
in advance to reduce the pH to 7.0 as required for QIAGEN ion exchange
chromatography)
41. Set up a QIAGEN plasmid maxi kit QIAGEN-tip 500 in a suitable rack over a waste
container; equilibrate it by passing 5 ml QIAGEN buffer QBT then 10 ml buffer QP (see
Solutions) through it, collecting into waste.
42. Transfer 12-ml portions of the ligations step 40 (total volume in both tubes combined ~95
ml) into the QIAGEN-tip 500 until the entire volume of the reaction has been applied to the
column, continuing to collect to waste.
43. Wash the column with 10 ml QP and four 5-ml portions of buffer QB, continuing to collect
to waste.
44. Elute the column with two 12-ml portions of QIAelution buffer (see Solutions), collecting
into a fresh 50-ml tube.
45. Dialyze in two 12-ml 10-KDa MWCO Slide-a-lyzer dialysis cassettes against two changes of
1 liter 10 mM Tris.HCl pH 8.5 and 1 change of 1 liter 1 mM Tris.HCl pH 8.5 in the cold room.
46. Withdraw the contents of the dialysis cassettes into a 50-ml tube; apply the dialyzed DNA in
2-ml portions to two 10-KDa 30-KDa MWCO Centricons (Millipore), concentrating all the way
each time; collect final retentates into conical retentate cups as per Millipore instructions;
transfer the retentates to a single 500-µl Ep tube, measuring volumes as you do; the combined
volume turned out to be 180 µl.
47. Scan 100 µl of a 1/100 dilution from 220 to 320 nm, using 1 mM Tris.HCl pH 8.5 as diluent
and reference (save the dilution for next step). The results are graphed below; the calculated
DNA concentration is 1.077 mg/ml and the total yield of DNA is therefore 194 µg—39.4% of the
input to the ligation at step 39.
533583453 Page 13 3/9/2016
Absorbance
0.2
0.15
0.1
0.05
0
-0.05
220
240
260
280
300
320
Wavelength
48. In two 1.5-ml Ep tubes make electrophoresis samples:
 Purified ligation product:
o 12 µl (129 ng) of the 1/100 dilution previous step
o 2 µl 70/75/BPB
 Cut vector
o 1 µl (124 ng) cut vector
o 11 µl water
o 2 µl 70/75/BPB
Load these 14-µl samples next to 10 µl (400 ng) .BstEII marker in the middle lanes of a 0.7%
agarose/4×GBB minigel; electrophorese at 30 V until the BPB is close to the end of the gel; stain
with sybr green and photodocument as usual. Here is the image:
Ligation
Cut vector
.BstEII marker
533583453 Page 14 3/9/2016
This pattern is reasonably consistent with the test ligation step 31.
PILOT ELECTROPORATION
NOTE: In order to construct the library, we planned 12 electroporations, each with 200 µl frozen
electrocompetent cells and 1/12 (14.8 µl) of the purified ligation product step 46. In order to test
in advance the efficiency of these transfections, we did a 1/10 scale pilot electroporation in which
1.5 µl was transfected into 20 µl of frozen electrocompetent cells from the same batch.
533583453 Page 15 3/9/2016
49. Set up the electroporator at 1250 V and a time constant of 10 ms (for example, 400 Ω × 25
µF = 10,000 µs = 10 ms in our electroporator design electroporator.doc), and the following
supplies:





A 15-ml tube containing 1.5 ml SOC with 0.2 µg/ml tetracycline
Sterile transfer pipette in the 15-ml tube
Space in the 37º shaker-incubator for strapping the rack in
Ice bucket
One 2-mm cuvette 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)
 The electroporation product step 46 on ice
 Six NZY/Tet plates (containing NZY nutrient agar medium supplemented with 40 µg/ml
tetracycline) labeled neat, 10-1, 10-2, 10-3 , 10-4 , and 10-5
 A 250-ml culture flask containing 50 ml NZY supplemented with 20 µg/ml tetracycline
50. Remove one tube with a 22-µl aliquot of frozen electrocompetent MC1061 cells (see
ElectrocompetentCells.doc; same batch as will be used for the full-scale electroporation) from
the –80º freezer and immediately do the electroporation as follows:
 Thaw the tube of frozen electrocompetent cells between finger tips
 Pipette 1.48 µl (1.59 µg) of purified ligation product step into the electrocompetent cells
and stir with pipette tip; leave on ice 30 sec
 Carefully layer as much of the cell/DNA mixture as possible on top of the red 20%
glycerol underlay in the cuvette, being very careful to avoid bubbles
 Zap
 Using the transfer pipette, draw up the SOC in the 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).
 Shake the 15-ml tube at 37º for 45 min.
51. When the 45-min incubation previous step is finished, pipette the entire electroporation
culture from the 15-ml tube to the 250-ml flask (containing 50 ml NZY + tetracycline; step 49).
After removing a 250-µl portion next step, shake the 250-ml flask vigorously overnight at 37º.
52. Label six sterile dilution tubes neat, 10-1, 10-2, 10-3 , 10-4 , and 10-5 to correspond to the
labeled NZY/Tet dilution plates step 49; into all but the neat tube pipette 225 µl NZY (no need
for tetracycline); pipette 250 µl from the 250-ml culture flask previous step into the neat dilution
tube; in the remaining five dilution tubes make serial 10-fold dilutions by passing 50 µl and
vortexing; spread a 200-µl portion of each dilution tube on the corresponding NZY/Tet plate step
49; incubate plates overnight at 37º. Next day, enter the colony counts in table below.
533583453 Page 16 3/9/2016
Dilution
neat
10-1
10-2
10-3
10-4
10-5
Efficiency (clones/µg)
Colonies
Too many to count
~1300
135
16
0
0
2.19 × 106
The total number of clones can be calculated from the colony count on the 10-2 plate: (135
clones/0.2 ml spread on plate) × 100 (dilution factor) × 51.5 ml (total volume of culture in the
250-ml flask) = 3.48 × 106. Since the amount of input ligation product was 1.59 µg, the
efficiency is 2.2 × 106 clones/µg. This is disappointingly low (more than 10 times lower than at
step 37, for example), but that may reflect the fact that the student carrying out this experiment
was a novice working pretty much on his own. At this efficiency, the total number of primary
clones expected from the remaining ligation product would be only ~4 × 108. Nevertheless we
decided to press ahead. If the size of the library had been extremely important to our particular
needs in this experiment, however, we would have repeated the pilot electroporation, including
untreated vector RF as a positive control to confirm the quality of the frozen electrocompetent
cells. If necessary, we would have made and tested a new batch of frozen electrocompetent cells.
LARGE-SCALE ELECTROPORATION
53. Set up the electroporator at 2500 V and a time constant of 10 ms (for example, 400 Ω × 25
µF = 10,000 µs = 10 ms in our electroporator design electroporator.doc), and the following
supplies:
 Twelve 50-ml tubes containing 15 ml SOC supplemented with 0.2 µg/ml tetracycline
 Sterile transfer pipette in each 50-ml tube
 In the 37º shaker-incubator mount six clips for 125-ml culture flasks and six clips for 2.8liter Fernbach flasks
 Six sterile 125-ml culture flasks (plugged with absorbent cotton and capped with a metal
cap)
 Six sterile 2.8-liter Fernbach flasks numbered 1–6, containing 1 liter NZY supplemented
with 20 µg/ml tetracycline, plugged with absorbent cotton, and capped with aluminum
foil
 Ice bucket with twelve 2-mm electroporation cuvettes and the remainder of the purified
electroporation product step 46
 A bucket of powdered dry ice
 Thirty NZY/Tet plates labeled 1-neat, 1-10-1, 1-10-2, 1-10-3, 1-10-4, 2-neat, 2-10-1, 2-10-2,
2-10-3, 2-10-4, 3-neat, 3-10-1, 3-10-2, 3-10-3, 3-10-4, 4-neat, 4-10-1, 4-10-2, 4-10-3, 4-10-4, 5neat, 5-10-1, 5-10-2, 5-10-3, 5-10-4, 6-neat, 6-10-1, 6-10-2, 6-10-3, and 6-10-4
533583453 Page 17 3/9/2016
54. Remove twelve tubes containing 200-µl aliquots of frozen electrocompetent MC1061 cells
(see ElectrocompetentCells.doc) and put them in the bucket of powdered dry ice. Immediately
do twelve electroporation as follows:
 Thaw one of the tubes of frozen electrocompetent cells between finger tips
 Pipette 14.8 µl (15.94 µg) of purified ligation product step 46 (in ice bucket previous
step) into the electrocompetent cells and stir with pipette tip; leave on ice 30 sec
 Carefully pipette the entire cell/DNA mixture into the bottom of one of the 2-mm
cuvettes (in ice bucket previous step, being very careful to avoid bubbles
 Zap
 Using the transfer pipette, draw up the SOC in one of the 50-ml tubes, use it to resuspend
the zapped cells by vigorously pumping up and down a few times, and transfer the
resuspended cells into one of the 125-ml flasks
 When you’ve filled a 125-ml flask with two 15-ml electroporation cultures, put it in the
shaker incubator and shake vigorously at 37º for 45 min
NOTE: Two of the twelve electroporations arced and were therefore lost; so two of the six flasks
got only one electroporation not two. The reason for arcing is inadvertently including air bubbles
in the cell/DNA mixture; experienced electroporators learn to avoid this problem.
55. Meanwhile, label six sets of five sterile dilution tubes to correspond to the labeled NZY/Tet
dilution plates step 49. In all tubes except the six neat tubes pipette 225 µl NZY.
56. When the 45-min incubations step 54 are finished add one of the 30-ml electroporation
cultures (in the 125-ml culture flasks) to each of the six numbered sterile 2.8-liter Fernbach
flasks, each containing 1 liter sterile NZY/Tet (20 µg/ml). Mix by swirling. Remove a 250-µl
aliqout from each of the numbered cultures into the corresponding neat dilution tube previous
step. Shake the six Fernbach flasks vigorously overnight at 37º.
57. In the four 10-1–10-4 dilution tubes in each series step 55 make serial 10-fold dilutions of the
corresponding neat tube by passing 25 µl and vortexing. Spread 200-µl portions of each dilution
tube on the corresponding NZY/Tet plate step 49. Incubate plates overnight at 37º. Enter the
colony counts below (TNTC = too numerous to count):
Dilution
Neat
10-1
10-2
10-3
10-4
Total clones (theoretically)
Culture 1
TNTC
TNTC
225
158
34
8.14e8
Culture 2
TNTC
TNTC
253
62
25
3.19e8
Culture 3
TNTC
TNTC
TNTC
140
35
7.21e8
Culture 4
TNTC
TNTC
400
136
30
7.00e8
Culture 5
TNTC
TNTC
TNTC
164
60
8.45e8
Culture 6
TNTC
TNTC
90
9
1
0.46e8
The total number of primary clones in the library is nominally 3.45 × 109, using the counts on the
10-3 plates; this corresponds to an overall transfection efficiency of 1.8 × 107 clones/µg.
Obviously, though, these data show inconsistencies in that the count doesn’t drop by a factor of
~10 with successive 10-fold dilutions in each series. This probably reflects the fact that the
533583453 Page 18 3/9/2016
student carrying out this experiment was a novice working largely on his own. An experienced
electroporator would generate more consistent data, and probably would achieve higher
transfection efficiencies with such a highly purified ligation product. For our purposes we were
entirely satisfied with the library. In any case, there’s no way to repeat these titers once the cells
in the Fernbach flasks have started to divide. After overnight growth, all shake cultures look the
same (fully saturated stationary phase), whether the number of primary clones in them is 1 or 10
billion.
PURIFYING VIRIONS FROM PRIMARY LIBRARY
NOTE: Although over 6 liters of library culture were propagated at step 56, we only processed
350 ml from each of the six cultures, since we didn’t need any more. The reason for propagating
6 liters of library culture when we needed only 2 liters was to ensure that the volumes of the
cultures were sufficient to allow the successfully transfected cells to grow and secrete virions.
58. Pour 350 ml from each of the six Fernbach flasks step 56 into a sterile 500-ml centrifuge
bottle; centrifuge at 5 Krpm at 4ºC for 10 min in Sorvall GS3 rotor (or equivalent).
59. Pour the supernatants into six fresh sterile 500 ml bottles. Re-centrifuge the supernatants at
8 Krpm at 4ºC for 10 min. Pour the supernatants into six fresh 500-ml centrifuge bottles; to each
bottle add 52.5 ml PEG/NaCl solution (see stdpreps.doc); mix by 100 inversions; allow
precipitate to develop overnight at 4ºC.
60. Centrifuge the bottles at 8 Krpm at 4ºC for 30 min; RRR (see stdpreps.doc); dissolve each
pellet in 10 mL TBS (see stdpreps.doc) by shaking vigorously at 4ºC (preferably, but room
temperature OK); once the pellets are dissolved, centrifuge the bottles briefly to drive the
solution to the bottom; pool all six solutions in a single bottle and transfer half (30 ml) to each of
two disposable conical 50-ml tubes (assuming a FiberLite or equivalent rotor is available;
otherwise use OakRidge tubes); centrifuge the tubes at 10 Krpm 10 min at 4ºC to clear insoluble
material; carefully pour the cleared supernatants into fresh 50-ml (or OakRidge) tubes; to each
tube add 0.15 vol (4.5 ml) PEG/NaCl solution (see stdpreps.doc); mix by 100 inversions. If a
copious turbidity appears almost immediately as the PEG solution is mixed in, indicating a high
concentration of virions, go directly to the next step; otherwise, incubate the tubes for several
hours or overnight to allow the precipitate to develop.
61. Centrifuge the tubes at 10 Krpm at 4ºC for 10 min; RRR (see stdpreps.doc); dissolve the
pellet in each tube in 1 ml TBS (see stdpreps.doc) by vigorous vortexing; allowing the pellets to
soften and re-vortexing as necessary; centrifuge the tubes briefly to drive solution to the bottom;
transfer each 1-ml solution to a 1.5-ml Ep tube; microcentrifuge 5 min at top speed to clear
insoluble material; carefully transfer the supernatants into a single 4-ml glass vial (or other
suitable vessel); store in the refrigerator; the total volume is nominally 2 ml.
62. Make 1 ml of a 1/100 dilution and scan from 220 to 320 nm, using TBS as diluent and
reference. The results are graphed and tabulated below:
533583453 Page 19 3/9/2016
0.2
Absorbance
0.15
0.1
0.05
0
-0.05
220
240
260
280
Wavelength (nm)
Measure
Reference
A269
0.0026
A320
-0.00085
corrected net A269
undiluted corrected net A269
Bases per viral genome
Virion concentration in undiluted stock
300
320
Sample
0.1485
0.0263
0.11875
11.875
9252
7.70 × 1013 virions/ml
The overall yield corresponds to ~7.5 × 1010 virions/ml in the unprocessed culture supernatant;
although that yield is only 15% of the virion yield from an fd-tet-derived clone, it’s a perfectly
acceptable yield for a primary library.
63. Infectious units are titered as tetracycline transducing units (TU) as described in TUtiter.doc;
results are tabulated below:
Phage
fd-tet control
Library
Diluent only
Colonies at following dilutions:
-5
10
10-6
10-7
10-8
188
TNTC ~2000 259
18
0
Titer
(TU/ml)
9.588 × 1011
1.321 × 1012
Physical particle
concentration
(virions/ml)
Infectivity
13
1.816 × 10
5.28%
7.70 × 1013
1.72%
The infectivity is lower than a pure fd-tet clone, but perfectly acceptable for a primary library.
533583453 Page 20 3/9/2016
AMPLIFYING THE PRIMARY LIBRARY
The primary library step 61 is amplified, taking care to avoid substantially reducing its diversity.
Amplification is accomplished by infecting fresh cells with a portion of the library, growing the infected
cells in large cultures, and isolating the phage secreted by the infected cells into the medium. They key
to maintaining diversity is to ensure that the number of cells infected (before the cells have had a chance
to replicate) is much larger than the number of clones in the primary library (nominally 3.45 × 109
primary clones in this case; see step 57).
64. Inoculate six 1-liter culture flasks containing 100 ml terrific broth (see stdpreps.doc) with 1 ml of an
overnight culture of K91BK (= K91BluKan; see Strains.doc). Shake vigorously at 37º until the OD600 of
a 1/10 dilution reaches ~0.2 (late log phase = ~5 × 109 cells/ml; only one culture is measured, the other
five being assumed to be comparable; we use a Spectronic 20 colorimeter, but a regular
spectrophotometer with a 1-cm cell could be substituted):
64. Slow the shaking way down for 5 min to allow sheared F pili to regenerate.
65. To each flask add 166.7 µl (1.28 × 1013 virions; see step 62) of the library to be amplified; this is
about 30.6 physical particles = 0.38 TU per cell (since infectivity = 1.245%; see step 63). Continue slow
shaking for 15 min to allow virions to infect cells.
66. Pour each of the six 100-ml cultures sterilely into a pre-warmed 2.8-liter Fernbach flask containing
1 liter of NZY (see stdpreps.doc) supplemented with 0.22 g/ml tetracycline; shake vigorously for 35
min at 37º.
67. To each flask add 1 ml 20 mg/ml tetracycline, bringing the concentration up to 18 g/ml.
68. Remove a 7-µl sample from two of the flasks for the dilutions (next step), then continue shaking the
flasks vigorously overnight.
69. Spread 200 µl of 10-4 and 10-5 dilutions of the two 7-µl samples from the previous step (diluent =
NZY) on NZY plates containing 40 g/ml tetracycline and 100 g/ml kanamycin. Count the colonies the
next day. Here are the results for the 10-4 plates:
Plate A = 216 colonies
Plate B = 338 colonies
Average = 277 colonies
This means that the six cultures combined contain:
277 clones
× 104 dilution factor × 1100 ml/culture × 6 cultures = 9.14 × 1010 infected cells
0.2 ml plated
which corresponds to 26.5 infected cells per primary clone in the library. This is number is an
acceptable overrepresentation of the primary library, even though it’s only 9.5% of the anticipated
533583453 Page 21 3/9/2016
number of infected cells given the number and infectivity of the input virions at step 65. This result
points up the advisability of including a wide margin of error in planning the scale of library
amplification.
PURIFICATION OF VIRIONS FROM THE AMPLIFIED LIBRARY
NOTE: The virions from the amplified library will be purified by CsCl density equilibrium
centrifugation to improve their long-term storage stability. As for the primary library, we processed only
2 of the total of 6 liters of culture since we didn’t need more of this highly specialized library. For a
general-use random peptide library, all 6 liters would be processed.
70. Pour 350 ml from each of the six Fernbach flasks step 68 into a sterile 500-ml centrifuge
bottle; centrifuge at 5 Krpm at 4ºC for 10 min in Sorvall GS3 rotor (or equivalent).
71. Pour the supernatants into six fresh sterile 500 ml bottles. Re-centrifuge the supernatants at
8 Krpm at 4ºC for 10 min. Pour the supernatants into six fresh 500-ml centrifuge bottles; to each
bottle add 52.5 ml PEG/NaCl solution (see stdpreps.doc); mix by 100 inversions; allow
precipitate to develop overnight at 4ºC.
72. Centrifuge the bottles at 8 Krpm at 4ºC for 30 min; RRR (see stdpreps.doc); dissolve each
pellet in 12.5 mL TBS (see stdpreps.doc) by shaking vigorously at 4ºC (preferably, but room
temperature OK); once the pellets are dissolved, centrifuge the bottles briefly to drive the
solution to the bottom; pool all six solutions in a single bottle and transfer half (nominally 37.5
ml) to each of two disposable conical 50-ml tubes (assuming a FiberLite or equivalent rotor is
available; otherwise use OakRidge tubes); centrifuge the tubes at 10 Krpm 10 min at 4ºC to clear
insoluble material; carefully pour the cleared supernatants into fresh 50-ml (or OakRidge) tubes;
to each tube add 0.15 vol (5.625 ml) PEG/NaCl solution (see stdpreps.doc); mix by 100
inversions. If a copious turbidity appears almost immediately as the PEG solution is mixed in,
indicating a high concentration of virions, go directly to the next step; otherwise, incubate the
tubes for several hours or overnight to allow the precipitate to develop.
73. Centrifuge the tubes at 10 Krpm at 4ºC for 10 min; RRR (see stdpreps.doc); dissolve the pellet in
each tube in 10 ml TBS (see stdpreps.doc) by vigorous vortexing; allowing the pellets to soften and revortex as necessary; centrifuge the tubes at 10 Krpm for 10 min to clear insoluble matter; pool the
supernatants in a single TARED 50-ml tube; add additional TBS to the tube to bring the net weight in
grams (= volume in ml) to 21.82 ml.
74. Add 9.803 g pure CsCl to the 50-ml tube previous step and vortex until the salt is completely
dissolved, giving 24.33 ml of a 31% w/w CsCl solution, with a density of 1.3 g/ml. Divide the
solution evenly into two SW41 tubes (by eye is sufficiently accurate); centrifuge in SW41 rotor
at 37 Krpm at 4º for ~36 hr.
75. Illuminating each tube from above against a dark background, a sharp, thin flocculent band
was visible above the phage band, which was a translucent, non-flocculent light-scattering band
about 1 cm long with a sharp upper boundary (easily visible as a refractive index discontinuity)
533583453 Page 22 3/9/2016
and a diffuse lower boundary; another much larger flocculent band was visible just below the
phage band. The flocculent bands were relatively opaque, and are therefore much more apparent
at first glance than the very translucent phage band. We tried to avoid the lower flocculent band
to the extent possible, but there would have been little harm if some of this band had
contaminated the phage band. Using sterile transfer pipettes, the solution above the phage band,
including the flocculent band, was removed from each tube. Then the phage band was removed
with a clean transfer pipette, taking care to aspirate slowly and to avoid stirring the gradient.
Both phage bands were pooled in a single 50-ml tube. An experienced technician can generally
obtain both bands in a combined volume of ~3.5 ml—a volume that will fit into a single 3-ml
Slide-A-Lyzer dialysis cassette (next step). In this case, however, the inexperienced student
obtained the bands in a combined volume of ~6 ml.
76. Dialyze the phage in two 3-ml, 10-KDa MWCO Slide-A-Lyzers (Pierce) against three
changes of TBS (see STDPREPS.DOC); as mentioned in the previous step, an experienced
worker will be able to dialyze in a single Slide-A-Lyzer.
77. Remove the dialysate(s) into a single TARED vessel and record net weight (= net volume)
(the student failed to do this, but the total volume was ~6 ml). Store the vessel in the
refrigerator; the virions are usually stable for decades in this state.
78. In a 500-µl Ep tube make 250 µl of a 1/50 dilution and scan from 220 to 320 nm, using TBS
as diluent and reference solution. Results are graphed and tabulated below:
0.4
0.35
Absorbance
0.3
0.25
0.2
0.15
0.1
0.05
0
-0.05
-0.1
220
240
260
280
Wavelength (nm)
300
320
533583453 Page 23 3/9/2016
Corrected net A269
Dilution factor
Genome size
Undiluted virion concentation
(virions/ml)
Volume (ml)
Total yield (virions)
0.365
50
9252
1.184 × 1014
~6 ml
~7.1 × 1014
The final yield of purified virions corresponds to ~3.6 × 1011 virions per ml of unprocessed culture
supernatant—pretty close to the nominal yield of ~5 × 1011 virions/ml for fd-tet-derived phage.
NOTE: The library’s infectivity should be quantified by titering 10-7 and 10-8 dilutions (along with a
positive control phage and diluent negative control) as in step 63. The amplified library should have an
infectivity comparable to that of the positive control (typically 5–10%). In this case, the student didn’t
do this step. Typically about 0.1 ml (~1013 virions = ~5 × 1011 TU) would be used in the first round of
affinity selection with a target receptor or other selector, corresponding to ~100 TU per primary clone in
the library. This ensures that a high-affinity clone, with a typical final yield of ~1% of input during
affinity selection, has a good chance of being captured during the first round and therefore of ultimately
being identified after subsequent rounds of affinity selection.
SEQUENCE ANALYSIS
79. Viral single-stranded DNA (ssDNA) was extracted from both the amplified virions step 77
and from the primary library step 61 as described in ssDNA.doc and sequenced through the
degenerate insert region. A section of the sequence profile that includes the degnerate positions
is shown below:
The N degenerate residues showed up as a roughly equal mixture of all four nucleotides; the M
degenerate residues (in the minus strands, complementary to mRNA) showed up as equal
mixtures of A and C. These data confirmed that both versions of the library had the expected
533583453 Page 24 3/9/2016
degeneracy, though they had essentially nothing to say quantitatively about the true complexity of
the library (i.e., how many different coding sequences were present).