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Sequencing Phage
NOTE: Nowadays, we do all our sequencing through the standard “automatic” sequencing
services available in most academic and industry research settings. It must be admitted,
however, that preparing template for these services is a very inefficient way of meeting the
routine sequencing task that confronts the typical phage-display lab: determining 18–60
unknown nucleotides in each of, say, 50 or 100 templates. The manual procedure outlined here,
which used to be routine but is no longer used in our lab, incorporates several shortcuts that
greatly simplify this task.1 These include: (1) release of phage DNA template by disassembly in
alkali rather than extraction and ethanol precipitation; (2) use of a single 32P-labelled primer
rather than individual labeling reactions for each individual template; (3) a rapid method for
pouring bubble-free sequencing gels. The protocol below is for our standard four-lane
sequencing using combinatorial W, M, K and S termination mixes and 49-lane sequencing gels.
This gives clear reads for ~150 base-pairs from the primer, and thus is suitable for all our random
peptide libraries. Additional shortcuts for high-volume sequencing—~800 clones in a week—
have been described [Haas, S.J. and Smith, G.P.: Rapid sequencing of viral DNA from
filamentous bacteriophage. Biotechniques 15 (1993) 422-424, 426-428, 431]. These include: (1)
two-lane sequencing with Q and R termination mixes, especially when the unknown coding
sequence is very short (for example, the fUSE5/6-mer library, with only 18 bases of unknown
coding sequence; library.doc); piggy-backing two sets of sequences on top of each other—again,
suitable when the unknown sequence is short; and 97-lane gels.
TEMPLATE (PHAGE)
1. The four-lane sequencing protocol uses 6 µl of phage particles at a physical particle
concentration of ~2.5 × 1013 virions/ml (exact concentration not important; 2-fold errors in either
direction make little difference). The buffer should not be strong enough to prevent alkaline
denaturation; typically the phage are in water or 1/10 × TE. The phage need not be highly
purified; crude PEG precipitate prepared as described in SmallScaleVirions.doc or in steps 1–3
and 7 of Propagation_1ml.doc is perfectly suitable. We will call these intact phage “templates,”
even though it would be more accurate to call the single-stranded viral DNA (ssDNA) inside the
particles the template.
PRIMER PREPARATION
NOTE: The primers for the fUSE vector clones (5´-TGAATTTTCTGTATGAGG-3´) and the
f88-4 vector clones (5´-AGTAGCAGAAGCCTGAAGA-3´) are aligned with the sequences of our
random peptide libraries in libeseq.doc. (We no longer supply these primers.)
1. In a 1.5-ml Ep tube mix
water in sufficient volume to give a final volume of 10 µl
2 µl 22.5 µM primer in water (45 pmol)
1
On the other hand, abandoning manual sequencing (as efficient as it is) has allowed us to abandon radioactivity
altogether in our work, thus freeing us of the extremely burdensome (and completely unnecessary) demands of
radiation “safety.”
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200 µCi crude [-32P]ATP (~7 µCi/pmol) in  6 µl
NOTE: Crude label at ~166 µCi/µl is available from Dupont (Cat. No. NEG035C)
and ICI; concentrating purified label from much more dilute solution doesn’t
work in our experience, probably because the concentrated buffer components
poison the subsequent reactions. Label is still usable up to ~4 weeks (2 half-lives)
after its reference date; be sure to use larger volumes as needed to compensate for
decay.
1 µl 10 × kinase buffer
1 µl (~8 units) T4 DNA kinase
Incubate in a 37º water bath for 15 min.
2. Add 140 µl TE; incubate 15 min at 65–70º to inactivate enzyme.
3. Meanwhile, tap a NENSORB 20 cartridge (Dupont Cat. No. NLP-022) to shake the resin into
the narrow part of the column; remove the shipping cap and insert the cartridge into an 18 × 150
mm test tube in a rack; fill the column with 6 ml methanol; attach a 30-ml syringe filled with air
to the syringe adapter, secure the adapter onto the top of the column, and push the methanol
through the column (this will not require much pressure); it’s OK to expel all the methanol with
air. Remove the adapter and syringe and add 5 ml NENSORB reagent A to the cartridge; fill the
syringe with air and reattach it to the cartridge; push the reagent A through the column; this will
require considerable pressure, and it usually is necessary to detach the syringe, fill it with air,
reattach it, and continue pushing in order to expel all the reagent A. Move the cartridge to a
second 18 × 150 mm tube.
4. To the primer from step 2 add 400 µl NENSORB reagent A; open the NENSORB cartridge
and apply the mixture. Fill the syringe with air, reattach to the cartridge, and push the mixture
through the column. Open the cartridge and add 3 ml NENSORB reagent A; fill the syringe with
air, reattach to the cartridge, and push the reagent A through the cartridge (it’s OK to expel the
reagent with air). Open the cartridge and add 3 ml water; fill the syringe with air, reattach to the
cartridge, and push the water through the cartridge (it’s OK to expel the reagent with air). This
completes elution of unincorporated label. The liquid in the 18 × 150 mm tube is discarded in
liquid radioactive waste.
5. Mount the column on a ringstand over a rack containing seven 2.2-ml capless microtubes
(Sarstedt Cat. No. 72708) in a rack. Open the cartridge and add 3 ml of a 1:1 v/v water-ethanol
mixture (the eluent); fill the syringe with air; push part of the eluent through the cartridge,
collecting five 2-drop fractions and two 5-drop fractions. Discard the remaining eluent into
liquid radioactive waste, and the cartridge and syringe adapter into solid radioactive waste (the
syringe is saved in the radioactive work area for re-use). Use a radioactivity survey meter to
identify the two most ratioactive fractions (usually fractions 1 and 2). Pool these fractions in a
1.5-ml Ep tube; measure the volume with a 200-µl pipetter (typically ~80 µl) and dilute with
water if necessary to bring the total volume to 150 µl. Store at –20º in a lead pig. The primer
concentration, assuming losses are negligible, is ~300 nM; however, the exact primer
concentration is not important. The labeled primer is usable for ~4 weeks. Unused labeled
primer is discarded into solid radioactive waste.
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NOTE: The ethanol in the eluate is sufficiently diluted during the chain-termination reactions
that it doesn’t interfere. Don’t attempt to evaporate the ethanol; we have documented almost
complete breakdown of the labeled primer during this process.
6. To quantify incorporation, dilute 1 µl in 500 µl TE, spot 5 µl of the dilution on a 25-mm disk
of DE81 paper (Whatman), and count in a scintillation counter. The counts are typically 4,000–
15,000 cpm, corresponding to ~15–50% incorporation.
The exact amount of incorporation is not important experimentally, but it can aid in NRC
record-keeping. All unincorporated label ends up in liquid radioactive waste (step 4 above),
while essentially all the incorporated radioactivity ends up in solid waste, as follows: (1)
Radioactivity that is actually loaded onto the sequencing gel ends up in the neutralized strippedoff gel, which is disposed of at step 29. Radioactivity that is used for sequencing reactions but
not loaded onto gels ends up in solid waste when the 48-well GeNunc modules are discarded
after loading the gel at step 21. Unused labeled primer (step 5) and unused label are discarded in
solid waste (their liquid volumes being negligible). Therefore, for the purpose of recordkeeping, we enter the incorporated label immediately as being disposed of in solid radioactive
waste and the unincorporated label in liquid radioactive waste.
ALKALINE DENATURATION AND CHAIN-TERMINATION REACTIONS
NOTE: Our sequencing gel is a BRL Model S2, which uses two rectangular plates, one long and
one short, made of ordinary plate glass and obtainable as a custom order from a glass shop. We
use a pair of BRL sharkstooth combs that form 49 ~6-mm sample compartments.
7. In preparation for sequencing, make up all the necessary components:
NaOH/primer (6 µl/template plus an extra 35 µl for pipetting errors)
0.18 N NaOH (made from accurately titrated 2 N NaOH)
45 nM end-labeled primer (step 5; do not allow for radioactive decay)
Neutralizer (i.e., add the MnCl2 to the stock without that salt; 6 µl/template plus extra
100 µl)
W, M, K and S termination mixes diluted as necessary with termination diluent
3 µl/template plus ~20 µl for pipetting errors
Pipette in 500-µl Ep tubes and keep on ice
For very short sequences (fUSE5/6-mer library) use undiluted termination mix
Otherwise, dilute 1 vol termination mix with 2 vol termination diluent
Each tube can serve up to 24 templates
If necessary, use more than one tube of each termination mix
Using a 10-µl pipetter, mix 6 µl NaOH/primer with 6 µl neutralizer; confirm that the phenol red
turns from yellow to reddish pink, indicating successful neutralization. (If the neutralized
solution is still yellow, add ~1 µl 1 N NaOH to the NaOH/primer, and re-test. If the neutralized
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solution is dark purple, try 7 rather than 6 µl neutralizer, and if that works use 7 µl rather than 6
µl neutralizer at step 10 below.)
8. Using a 10-µl pipetter, pipette 6 µl of each template (~1.5 × 1011 physical particles = 250
fmol) into a well of a 48-well GeNunc module (Nunc InterMed Cat. No. 2-32298; well volume
25 µl).
9. Using a 10-µl pipetter, pipette 6 µl NaOH/primer (step 7) into each template well (no need to
change tips). Centrifuge 1 min at 3 Krpm in a centrifuge fitted with a microtiter plate rotor; this
mixes the two solutions. Float the GeNunc module on a 37º water-bath (use finger to sweep out
bubbles from under the module) for 5 min. Re-centrifuge as above.
10. Using a 10-µl pipetter, pipette 6 µl neutralizer (with MnCl2 added; step 7) to each template
well, pumping up and down ~4 times with pipette tip with each addition to mix; there is no need
to change pipette tips, but try to avoid significant carry-over of one template into the next. The
phenol red should change from yellow to reddish pink, indicating neutralization. Examine the
module from the side to confirm that there is no well with unmixed yellow and pink layers;
pump with pipette tip to mix such wells if they exist. Centrifuge the GeNunc module as in step
9. The wells now contain primed template, the volume being nominally 18 µl.
11. Using a multichannel pipetter, pipette 3 µl of each row of primed templates into rows A–D
of a new 48-well GeNunc module; these wells will be chain-terminated with the W, M, K and S
termination mixes, respectively. All the sequencing reactions that will be loaded onto one
sequencing gel should be on a single GeNunc module. Even though a sequencing gel can
theoretically accommodate 12 templates (= 48 chain-termination reactions), we like if possible to
load only 11 templates on a single gel, leaving 2 empty lanes on one side and 3 empty lanes on
the other side.
12. Add Sequenase version 2 (USB) to the W tube (step 7) to give a final enzyme concentration
of ~86 units/ml (~1/150 dilution; exact concentration not important); vortex gently to mix.
Using a 10-µl pipetter, pipette 3 µl to each of the primed template wells in Row A of one or two
48-well GeNunc modules from the previous step; there is no need to change tips. Likewise, add
enzyme to the M tube and add 3 µl to the primed template wells in Row B; tube K to Row C; and
tube S to Row D. Centrifuge the module(s) as in step 9. Float the 48-well module(s) on a 42º
water bath (sweep out bubbles from under the module as necessary) for 10 min to allow the
chain-termination reactions to be completed. Re-centrifuge as in step 9.
13. To each well add 4 µl formamide load buffer; there is no need to stir, pump or change tips.
Centrifuge as in step 9. The 48-well module(s) can be stored at –20º for several days waiting to
be loaded.
POURING SEQUENCING GELS
NOTE: Our sequencing gel is a BRL Model S2, which uses two rectangular 32.5-cm wide
plates, one long and one short, made of ordinary plate glass and obtainable as a custom order
from a glass shop. We use a pair of BRL sharkstooth combs that form 49 ~6-mm sample
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compartments. Before attempting to pour the gel, you should familiarlize yourself with the
apparatus and read its manual. (Other brands of apparatus with appropriate changes in the
protocol below.) We purchase 6% sequencing gel mix from AMRESCO (Solon ,OH; Cat. No.
E568); the “1 × TBE” buffer in their formulation has 89 mM Tris and 89 mM H3BO3 (“1 × TBE”
as we define it has 100 mM of these two components). We coat the long glass plate with a
silanizing reagent that bonds covalently to the acrylamide gel as it polymerizes. After
electrophoresis, the gel is washed, dried and autoradiographed while still attached to the glass
surface. This way of processing gels has advantages and disadvantages compared to the usual
practice of sticking gels to filter paper. Some key advantages are that it is much easier to handle
and autoradiograph the gel; the major disadvantage is that the gel must be stripped off the glass
plate by alkaline hydrolysis—a messy (though not very time-consuming) process.
13. In a 50-ml disposable tube pipette:
20 ml 95% ethanol
20 µl -methacryloxy-propyltrimethoxysilane (Sigma)
60 µl glacial acetic acid
Coat the long glass plate three times with this solution by pouring ~10 ml onto the plate and
spreading with a Kimwipe in a gloved hand; after each coat has been applied, allow the reagent
to dry and buff the plate with Kimwipes.
NOTE: This treatment covalently bonds acryloxy groups to the glass; these groups in turn
become covalently coupled to the acrylamide gel as it polymerizes, thus firmly bonding the gel
to the glass surface. It is vital that the short glass plate, which must be lifted off the gel after
electrophoresis (step 23), not be contaminated by even traces of this reagent!! (If a short glass
plate becomes contaminated, as evidenced by part of the gel sticking to it at step 23, it must be
stripped with alkali like the long glass plates, as described in steps 27–29 below.)
14. Coat the short glass plate once (twice if this is the first time the plate has been coated) with
Rain-X (Unelko Corp., Scottsdale, AZ; available at hardware and auto supply stores) by
squirting ~10 ml on the plate, spreading with a Kimwipe in a gloved hand, drying, and buffing
with Kimwipes.
15. On a level bench place a pan that is larger than the sequencing gel plates (from a photo
supply store we bought CESCO-LITE Photoquip Co. print developing pans big enough for a 16
× 20 inch print to lie flat on the bottom). In the pan put four identical plastic beakers face-down
as supports for the long glass plate. Put the long glass plate coated-side-up on these beakers so
that the edges hang over the beakers.
16. Remove the foam sealing pads from two spacers, wet them with water, and lay them on the
edges of the long glass plate; use a Kimwipe to press down along the length of the spacers to
“glue” them lightly to the glass surface.
17. In a plastic 250-ml beaker measure ~100 ml sequencing gel solution (from AMRESCO; see
above). Add 1 ml freshly-dissolved 10% w/w ammonium persulfate in water and 70 µl TEMED.
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Swirl to mix and pour onto the upper ~1/2 of the long glass plate; use a spatula if necessary to
sweep off any bubbles and wet any unwetted area in the upper ~2/3 of the glass plate. Lay the
edge of the short glass plate across the spacers on the long glass plate so its surface makes an
angle of ~45º with the surface of the long glass plate. Carefully lower the short glass plate onto
the long glass plate, sliding it upward or downward as necessary to its bottom and side edges
flush with those of the long glass plate. Clamp the side edges of the two plates together with
bulldog clips (be sure the clips press on the spacers, not inside them; otherwise, the plates will be
bent together, significantly reducing the spacing between them. Put the flat edges of the two
sharkstooth combs between the glass plates at the top, so that the bottom of the combs are ~3
mm from the edge of the short glass plate. Use an additional bulldog clamp to clamp together
the glass plates at the upper edge of the short glass plate. Allow the gel to polymerize at least 1
hr at room temperature. The gel can be stored overnight in this state.
18. Remove the bulldog clamps. Run a spatula under the sharkstooth combs to loosen them,
then pull them out. Wash off the outer surfaces of both glass plates with a scotch bright scouring
pad under water in the sink; also clean off the inner surface of the long plate above the top of the
short plate, to facilitate re-inserting the combs. Attach the foam sealing pads on the spacers.
Following the instructions for the gel apparatus, install the gel in the apparatus and fill the lower
and upper buffer reservoirs with “1 × TBE,” which in our terminology means 0.89 × TBE.
(Don’t install the sharkstooth comb at this point.) The gel can be safely stored for several days
in this condition.
LOADING AND RUNNING THE SEQUENCING GEL
19. Still without installing the sharkstooth combs, fill a transfer pipette with the running buffer
from the upper reservoir and squirt it across the top of the gel to sweep out the urea that has
leached out of the gel. Mix 133 µl TE and 66 µl formamide load buffer in a 500-µl Ep tube and
load the mixture across the entire gel. Preelectrophorese at 1500 V until the bromphenol blue
band is within ~2 cm of the bottom. Periodically readjust the voltage as necessary to keep the
temperature of the outer glass plate at ~50º.
20. Meanwhile, heat a water bath to ~90º. When pre-electrophoresis is about through, turn off
the heat to the water bath and float the 48-well GeNunc module containing the samples on the
hot water. The plastic will bow up slightly, allowing bubbles to escape, and then flatten out on
the hot water. About 3 min after that, carefully remove the module (you’ll need to wear gloves,
and you man need to fashion a metal tray to lift up the module).
21. Working as rapidly as possible, stop the preelectrophoresis, sweep out leached urea, install
the sharkstooth combs teeth-downward, and load 4 µl from wells A–D (in that order) from each
numbered column in the 48-well GeNunc module into adjacent wells in the gel (so that the W,
M, K and S reactions for a single template are adjacent and in that order). You can discard the
48-well module into solid radioactive waste.
22. Resume electrophoresis and continue at the same voltage until the xylene cyanol FF band
(blue-green) has migrated ~19 cm (this, at least, is a good distance for our random peptide
libraries). Adjust voltage periodically as necessary to maintain a glass-plate temperature of ~50º.
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PROCESSING AND AUTORADIOGRAPHY OF GEL
23. Stop the electrophoresis, remove the glass plate sandwich, remove the sharkstooth combs,
and pry off the short glass plate. Put the long plate (with the gel bonded to it) in a large pan just
large enough for the gel to lie flat on the bottom (we use Nalgene polypropylene sterilizing pans;
Cat. No. 6900-0020); add 1.5 liter water and rock gently for ~20 min; pour off the water into the
sink (there’s no significant radioactivity in it); add another 1.5 liter of water and again rock
gently for ~20 min and discard water; lift the glass plate (with washed gel attached) out of the
pan, pat it dry with Kimwipes. (The upper and lower buffers in the gel apparatus are not
significantly radioactive and can be discarded in the sink.)
24. Air-dry the gel at room temperature until completely dry (~4 hr, but usually we let it dry
overnight; we have set up a rack for this purpose in the upper part of our fume hood).
25. Autoradiograph overnight with Kodak X-Omat XAR X-ray film (Kodak No. 165-1512) in a
light tight box. Since the gel is on a perfectly flat surface, there is no need for an
autoraidography cassette. Instead, simply stack glass plates (gel side up) and films in alternation,
weighting down the upper film with an additional 4 unused glass plates.
26. Develop the X-ray film with an automatic developer or by hand. Here is how the banding
pattern for one template is interpreted:
Nucleotide
in template
T
A
G
C
W
——
——
Lanes
M
K
——
——
——
——
S
——
——
Notice that it is the nucleotide in the template (viral) strand that is given in the table—the strand
whose 5´3´ direction points downward in the sequencing gel. One advantage of WMKS
sequencing is that the information is redundant: only three lanes are required to deduce the full
sequence, the fourth lane giving an added measure of assurance. Another advantage is that for
all possible pairs of adjacent nucleotides, it is always possible to compare bands side-by-side in
order to ascertain more accurately which is on top and which on bottom; this allows an additional
~50 nucleotides of accurate base-calling compared to standard sequencing gels, in which it is
frequently necessary to compare bands across one or two empty lanes.
STRIPPING LONG GLASS PLATES
This is admittedly a drawback of bonding the gel to the long glass plate: the procedure is messy
and requires some space behind a plastic radioactivity shield (in order to satisfy NRC
regulations). However, it doesn’t require much time investment. For this purpose, we have
constructed an enclosed plexiglas rack with flanges that allow us to slide in up to five Nalgene
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polypropylene sterilizing pans (Cat. No. 6900-0020) on top of each other. These pans are just
big enough for a long glass plate to lie flat on the bottom.
27. Lay the used long glass plate flat on the bottom of the pan. If this is a fresh pan, add 1 liter
of 1 N NaOH—just enough to cover the gel. If the pan already has stripped-off gel in it (usually
dried down), lay the glass plate flat on the bottom and add additional 1 N NaOH as necessary to
cover the surface (usually ~500 ml). Allow the gel to strip at least 24 hr.
28. Remove the glass plate from the pan, scraping off as much of the stripped-off gel as possible
back into the pan. The stripped plate is washed and put in a rack to dry. Meanwhile, store the
pan open in the rack, allowing liquid to evaporate.
29. After ~5 glass plates have been stripped in a pan, the goopy solution becomes hard to cope
with. When this happens, neutralize the accumulated stripped-off gel by adding 3.78 M
NaH2PO4 (~ 1 liter), testing for neutralization with a broad-range pH strip. We usually allow the
neutralized gel to air-dry for several weeks more into a hard gel before discarding it into solid
radioactive waste.