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Synthesis of a Degenerate Oligonucleotide Insert
NOTE: This protocol is an amalgam culled from recent experiments. The aim is to prepare a
double-stranded fragment containing a degenerate coding sequence, with ends ready to splice
into a phage-display vector (fUSE5 or f88-4 in most cases in our lab). There are many possible
routes to this goal; in this protocol, a synthetic degenerate oligonucleotide is base-paired with a
primer to make a partial duplex, which is then converted to a fully double-stranded fragment
with Klenow fragment of DNA polymerase and dNTP’s. This duplex is cleaved with
appropriate restriction enzyme(s), and the desired doubly-cleaved cleavage product purified by
acrylamide gel electrophoresis. A central difficulty of this process is purifying and concentrating
the product after each step. Of all the many procedures we’ve tried (including ethanol
precipitation with glycogen and purification on “MerMaid” from Bio101, QIAGEN-tips,
Promega Wizard Minipreps, etc.), the one that has worked best and most reliably by far is
centrifugation through a Centricon ultrafilter (Amicon, now a subsidiary of Millipore).
Furthermore, Amicon makes an electroelution device called a “Centrilutor,” which uses
Centricons in a very convenient and effective electroelution process.
The starting material is a degenerate oligonucleotide whose non-degenerate flanks have
restrictions sites in the right position for splicing the degenerate coding sequence into the vector
in the right reading frame. There should also be an ~20-base primer complementary to the 3´
end of the degenerate oligonucleotide. Here is an example of a pair of oligonculeotides for
making a degenerate insert for vector fUSE5; the degenerate oligonucleotide strand is on the top,
the primer strand in the middle, and the intended amino acid sequence at the N-terminus of
mature pIII on the bottom:
SfiI
SfiI
5´-TCACTCGGCCGACGGGGCCNNKNNKNNKNNKTGTNNKNNKNNKNNKNNKNNKNNKNNKNNKNNKNNKGGGGCCGCTG-3´
3´-CCCCGGCGACCCCGGCTTTGA-5´
A D G A X X X X C X X X X X X X X X X X G A A G A E T
The two oligonucleotides are complementary over a 10-base-pair overhanging region—enough
for Klenow fragment to extend both 3´ ends.
NOTE: The protocol below starts with large amounts of the degenerate oligonucleotide and the
primer—enough theoretically to make 70 µg of the 88-bp double-stranded product in the case of
the above oligonucleotides, for instance. This is actually more than can be loaded on the gel at
step below. However, it’s probably a good idea to work on the extra large scale in case there are
losses.
1. Into a 1.5-ml Ep tube pipette
1.21 nmol degenerate oligonucleotide
1.21 nmol primer
50 µl 1 M Tris.HCl pH 7.8
10 µl 1 M MgCl2
water to bring the total volume to 240µl
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Vortex; microfuge briefly; incubate to anneal the two oligos:
5 min 68°
10 min 37°
10 min RT
2. Put the tube on ice and add:
10 µl 5 mM dNTP’s (mix made from 100 mM commercial stocks)
677 µl water
375 units of Klenow fragment (New England Biolabs; ~75 µl)
Final concentrations in 1 ml:
dNTP’s: 50 µM each
degenerate oligo: 1.2 µM
primer: 1.2 µM
Tris.HCl pH 7.8: 50 mM
MgCl2: 10 mM
Klenow: 375 units/ml
Vortex gently; incubate at 30° 45 min (your can use a temporary water bath in a covered ice
bucket). Immediately put tube in ice-water (to cool immediately).
3. Remove 3 µl (+ 3 µl TE + 1 µl 70/75/BPB) on a 15% acrylamide/TBE minigel next to
suitable markers (we use 1 µg pGEM markers from Promega in 10 µl sample buffer).
Meanwhile, IMMEDIATELY freeze the reaction in the -80° freezer while you await the results
of the check gel.
4. (Assuming the gel results are satisfactory) Add 50 µl 250 mM EDTA to the reaction tube and
mix as you thaw the liquid between the fingers; incubate at 68° 10 min to inactivate Klenow.
5. Pipette half the reaction mixture (525 µl) to a second Ep tube; extract both tubes once with
phenol and twice with chloroform, using the double-spin method.
6. Evaporate in the SpeedVac in order to reduce traces of chloroform (the Centricon to be used
in the next step is not resistant to this solvent).
7. Load the solution from both tubes in a single Centricon 10-Kda-cutoff centrifugal ultrafilter;
fill the filter to the 2-ml mark with TE. Centrifuge at 5 Krpm in the Sorvall SS34 rotor at 4º (use
thick-walled rubber adapter) to concentrate the solution (~45 min). Wash three times with TE,
concentrating to ~100 µl each time. Back-centrifuge as in the Centricon instructions to collect
the retentate; pipette the retentate into a 1.5-ml Ep tube; wash the filter by adding 100 µl TE,
vortexing the Centricons, and back-centrifuge again, pooling the wash with the original retentate.
Measure the total volume of retentate with a pipetter (should be ~200 µl). The concentration will
be theoretically ~100–350 µg/ml, depending on the size of the oligos.
8. Run 1 µl on a check-gel (see step 3) with suitable markers at suitable concentrations to enable
you to estimate the amount of DNA from band intensity.
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9. Digest the double-stranded product with appropriate restriction enzymes—e.g., HindIII plus
PstI (2 hr at 37º in REact 2 buffer) for f88-4 inserts, SfiI or BglI for fUSE5 inserts). Keep in
mind that the density of restriction sites on these small synthetic duplexes is much higher than in
natural DNA; therefore, it is probably best to use, say, ~10 units of enzyme per µg of DNA to
ensure good digestion. In order to keep the volume of enzyme to a reasonable level, it is
advantageous to use high-concentration forms of the enzymes available from many suppliers.
After digestion, run ~100–300 ng on a check-gel (see step 3) to confirm that cleavage has
occurred; you should be able to resolve uncleaved, singly-cleaved and doubly-cleaved fragment,
the latter being the target product; cleavage is satisfactory as long as there’s a substantial amount
of the doubly-cleaved product. Meanwhile, freeze the bulk of the digest immediately after
removing the electrophoresis sample and keep it frozen until you get the check-gel results.
10. Add sufficient 250 mM EDTA to the bulk digest to bring the EDTA concentration to 1.25 ×
the Mg++ concentration. Add TE as necessary to bring the total volume to 1 ml. Extract,
concentrate and wash as in steps 5–7.
11. Pour a 15% acrylamide/TBE gel with 1-mm spacers and a 1-mm comb with a 7.2-cm-wide
preparative well and a small marker well (these are for the BioRad Mini Protean II minigel
system; other systems will have similar dimensions). Add 1/6 vol 70/75/BPB to the digest
(previous step) and load it into the preparative well (you can load up to ~50 µg DNA in one of
these wells). At the same time, load suitable DNA markers in the marker well (load a lot of
marker so bands can be seen even with light staining). Electrophorese as usual. Stain lightly
with 0.5 µg/ml ethidium bromide.
12. Have a scalpel blade and a piece of used Xray film ready to use as blades for excision. Also,
have a small spatula for manipulating the excised band and clean weigh boat on hand to receive
the excised band. When these materials are ready, put on gloves and illuminate the stained gel
on a long-wave UV transilluminator. Working as rapidly as possible, excise the doubly-cut band
and put it in the weigh boat.
13. Cut the excised band (roughly 72 × 3 mm) evenly into eight ~9×3 mm strips. Put two of
these strips into each of four Centrilutor capped inserts (these have holes in both the bottom and
top to allow current to pass through and to allow electroeluted DNA molecules to pass through
the bottom).
14. Set up the Centrilutor electroelution apparatus with four 10-KDa Centricons as in the
instructions. Fill both bottom and top reservoirs with TBE buffer. Dip each insert (with gel
fragments inside) in and out of the upper buffer a few times to ensure the air bubbles are
removed, then put it in one of the Centricons. Using a 9-inch Pasteur pipette whose end has been
bent into the shape of a shepherd’s crook, squirt the TBE buffer vigorously up under each
Centricon to expel trapped air bubbles (there’s a mirror on the bottom of the lower buffer
reservoir to aid in seeing these bubbles). (The Centrilutor instructions say to suck the bubbles
out, but we find that much more difficult than squirting them out with vigorous positive upward
pressure). Electroelute at 100 V for 2 hr.
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15. Carefully lift the upper buffer reservoir (with Centricons) off the lower reservoir; pour out
the buffer from the lower reservoir; re-install the upper buffer reservoir; by pulling one of the
stoppers from an unused hole, drain the upper buffer reservoir into the lower one. Breathe a sigh
of relief when the buffer level falls below the tops of the Cenricons. Slowly and carefully lift the
inserts out of the Centricons, allowing any buffer therein to drain back into the Centricons.
Gently pull the Centricons out of their holes in the upper buffer reservoir, being exceedingly
careful not to allow the buffer in the Centricons to spill out.
16. Concentrate the electroeluates by centrifuging the Centricons at 5 Krpm at 4º in the SS34
rotor (use thick-walled rubber adapters). Collect the retentate from three of the Centricons by
back-centrifugation; pipette all three of those retentates into the fourth Centricon; add 200 µl TE
to the three Centricons, vortex them to ensure dissolution of any further retentate, and again
back-centrifuge and pool the washes with the first retentates in the fourth Centricon, which now
contains all the electroeluted DNA. Concentrate on this fourth Centricon, wash three times with
TE, and collect the retentate as in step 7.
18. Run ~0.5 µg (nominally) along with appropriate amounts of a suitable marker DNA on a
check gel as in step 3. Estimate the concentration of insert by comparison with the known
amounts of DNA in the marker bands.