sequencing.doc 10/19/18 Page 1

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)

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.

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 stripped-off 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 record-keeping, 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 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 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. 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).