construction of a large random peptide library (rpl) in

document.doc 4/13/2023

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

CTCACTCGGCCGACGGGGCCNNKNNKNNKNNKNNKAAATGCTGTTATAGCCTGNNKNNKNNKNNKGGGGCCGCTGGGGCCGAAACTGCAGTTTCGGCCCCAGCGGCCCCMNNMNNMNNMNNCAGGCTATAACAGCATTTMNNMNNMNNMNNMNNGGCCCCGTCGGCCGAGTGAG

A D G A X X X X X X X X X G A A G A E T..K C C Y S L

GGGCCNNKNNKNNKNNKNNKAAATGCTGTTATAGCCTGNNKNNKNNKNNKGGGGCCGCTGCGGCCCCMNNMNNMNNMNNCAGGCTATAACAGCATTTMNNMNNMNNMNNMNNGGCCCCGT

A D G A X X X X X X X X X G A A G A E T..K C C Y S L

K le n o wQ IA G E N m id ip re pC o n c e n tra te o n C e n tr ic o n

S fiIQ IA G E N m id ip re pC o n c e n tra te o n C e n tr ic o n

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:

5’

5’

d e g e n e ra te s e g m e n t

5’

5’

K le n o w e x te n s io n


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.


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.

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 50-ml 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).

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220 240 260 280 300 320

Wavelength (nm)

Ab

sorb

ance

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.

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:


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0.12

220 240 260 280 300 320

Wavelength (nm)

Ab

sorb

ance

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 13o 2 µl 70/75/BPB

Vector-ready insert:o 12 µl (61.7 ng) of the 1/50 dilution previous stepo 2 µl 70/75/BPB

LMW DNA ladder:o 550 µL of watero 100 µL 70/75/BPBo 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):


Hyb

ridi

zed

olig

os s

tep

3

Pri

mer

ex-

tens

ion

step

6

Pur

ifie

d pr

imer

exte

nsio

n st

ep 2

2

Vec

tor-

read

yin

sert

ste

p 22

LM

W D

NA

ladd

er s

tep

22

LM

W D

NA

ladd

er s

tep

22

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)


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.


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) .BstEII10 µl tube VectorAll 10 µl of tube 0All 10 µl of tube 1All 10 µl of tube 2All 10 µl of tube 3All 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:

.E

IIB

st

Vec

tor

Tub

e 0

Tub

e 1

Tub

e 2

Tub

e 3

Tub

e 4

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.


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.


E0 E1 E2 E3 E4Concentration of vector during ligation (nM)

1 1 1 1 1

Concentration of insert during ligation (nM) 0 1.3 2.6 5.2 10.4

Number of colonies at following dilutions

Neat 0 60 195 36 5010-1 0 9 18 51 110-2 0 0 1 0 0

Electroporation efficiency (colonies/µg) 0 1.0×107 3.3×107 7.2×106 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+) 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.


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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 stepo 2 µl 70/75/BPB

Cut vectoro 1 µl (124 ng) cut vectoro 11 µl watero 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:


.E

II m

arke

rB

st

Cut

vec

tor

Lig

atio

n

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.


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.


Dilution Coloniesneat Too many to count10-1 ~130010-2 13510-3 1610-4 010-5 0Efficiency (clones/µg) 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

2.8-liter 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, 5-neat, 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


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 Culture 1 Culture 2 Culture 3 Culture 4 Culture 5 Culture 6Neat TNTC TNTC TNTC TNTC TNTC TNTC10-1 TNTC TNTC TNTC TNTC TNTC TNTC10-2 225 253 TNTC 400 TNTC 9010-3 158 62 140 136 164 910-4 34 25 35 30 60 1Total clones (theoretically)

8.14e8 3.19e8 7.21e8 7.00e8 8.45e8 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 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:


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0

0.05

0.1

0.15

0.2

220 240 260 280 300 320Wavelength (nm)

Abs

orba

nce

Measure Reference SampleA269 0.0026 0.1485A320 -0.00085 0.0263corrected net A269 0.11875undiluted corrected net A269 11.875Bases per viral genome 9252Virion concentration in undiluted stock 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

Colonies at following dilutions:Titer (TU/ml)

Physical particle concentration (virions/ml) Infectivity10-5 10-6 10-7 10-8

fd-tet control 188 9.588 × 1011 1.816 × 1013 5.28%Library TNTC ~2000 259 18 1.321 × 1012 7.70 × 1013 1.72%Diluent only 0

The infectivity is lower than a pure fd-tet clone, but perfectly acceptable for a primary library.


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 coloniesPlate B = 338 coloniesAverage = 277 colonies

This means that the six cultures combined contain:

× 104 dilution factor × 1100 ml/culture × 6 cultures = 9.14 × 1010 infected cells

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 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 re-vortex 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) 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:

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0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

220 240 260 280 300 320Wavelength (nm)

Abs

orba

nce

Corrected net A269 0.365Dilution factor 50Genome size 9252Undiluted virion concentation (virions/ml) 1.184 × 1014

Volume (ml) ~6 ml Total yield (virions) ~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 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).

construction of a large random peptide library (rpl) in

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