Polony PCR Amplification
This section outlines the protocol for single-molecule amplification within the acrylamide gel. The protocol given here may differ in specifics from what is presented here, but reflects what we are doing currently (Summer 2003). We have tried to include excessive detail here but let us know if anything is unclear. The general steps are:
1. Cast acrylamide gels
2. Diffuse in PCR reagents
3. PCR amplification
4. Slide clean-up
Several points to note before starting in on the protocols:
1. Slides must be treated with Bind-Silane prior to use. See relevant protocol.
2. It is strongly recommended that both a lab area and a freezer area be designated as "pre-PCR". Post-PCR slides and post-reaction PCR tubes should always be kept away from pre-PCR designated areas. We are performing single-molecule PCR, often with "common" primers, so a small amount of contamination can go a long ways towards uninterpretable or miseleading results. For the same reason, our pre-PCR lab area (where we set up all slides prior to putting them into the thermal cycler) is an AirClean hood with a UV lamp. We have had little or no problems with cross-contamination since designating pre-PCR areas and implementing use of the AirClean hood. The AirClean hood has a dedicated set of pipet-men, pipette tips, dH20, 1.5 mL tubes, ChemWipes, etc.
3. Try to always include one slide as a no-template negative control. This is a quick means of determining when cross-contamination has / is becoming a problem.
4. The below protocols assume that 8 slides are being processed in parallel, and the volumes, etc. are specified accordingly. This is a particularly convenient number of slides to process because the plastic Coplin jars can fit nine slides. We have successfully processed up to 32 slides in a single run, but suggest starting with 8 or 9 and scaling up once everything is working.
5. Try not to come into contact with the central oval-region of the Teflon-coated slides. Best to keep it clean as possible both before and after Bind-Silane treatment, and stay away from it after casting the gel as it is quite fragile.
6. An argon chamber can be as simple as a Ziplock bag with a tube fitting such that it can be filled with argon gas.
7. It is quite possible to include all PCR reagents in the gel from the get-go, such that a diffuse-in is not necessary. However, we have found that PCR amplifications are considerably more efficient when as many PCR-related reagents as possible are diffused in after the gel has polymerized (probably because the polymerization process can damage DNA and/or enzymes). In the protocol described below, template is included in the gel during polymerization. It is also quite possible to diffuse-in template (and this may lead to even better amplificaton results). However, the distribution of polonies across the space of the gel can be highly non-uniform. Consequently, we still often elect to mix the template into the polymerization mix.
CAST ACRYLAMIDE GELS
1. Remove 8 Bind-Silane treated slides from storage box in dessicator and place face up in AirClean hood.
2. Turn on UV lamp in AirClean hood for 15 minutes.
3. Label slides with numbers and date using a SHARPIE pen (other inks will wash off in hexane)
4. Almost completely cover slides with coverslips (Fisherbrand, 18mm x 30mm, #1, untreated). Keep a small area of oval exposed for subsequent gel loading.
5. Prepare fresh "ABD mix" in a 15 ml polystyrene conical:
9 mL IEF 40% Acrylamide
1 mL Acrylamide / Bis (19:1; 38%:2%)
200 mg DATD
6. Using a 3-cc syringe and a 0.22 micron filter, filter ~1 mL of the ABD mix into a 1.5 mL microcentrifuge tube.
7. Remove Acrydite-Modified Amplification Primer from pre-PCR area of -20'C freezer to thaw.
8. Prepare fresh 5% APS. Notably, the APS bottle should be stored in some sort of room-temperature dessicator. We just store it inside a large plastic screw-top container.
0.5 g APS →10 mL dH20
9. Prepare fresh 5% TEMED
2 uL TEMED → 38 uL dH20.
10. Put a few drops of 30% BSA into a 1.5 mL tube (BSA should be stored at 4'C)
11. Prepare the gel-casting mix (200 uL total volume). Do not add APS until immedietely prior to casting the gels. This recipe is for a 10% gel with sufficient mix for at least 8 (identical) slides.
50 uL A-B-D mix (FILTERED)
1.33 uL 30% BSA
2 uL JCF-AC (100 uM)
136.66 uL dH20
4 uL 5% TEMED
4 uL 5% APS
0.5 uL Template (at appropriate concentration)
12. Gel Loading:
Suck up 18 microliters of the mix and pipet it into the small exposed area of the oval, such that surface tension pulls the liquid into the space between the coverslip and glass to cover the full surface area of the oval. The liquid should distribute under the coverslip such that only a small amount (1 to 2 uL) cannot fit. Slide the coversip over such that the oval is completely covered.
We will load as many as 8 gels from a single mix (after adding APS), but if you are doing more slides than this, we suggest splitting your master mix prior to adding APS (so that polymerization doesn't proceed to far in the tube before you cast all of the gels). The master-mix can also be split prior to adding APS if you need to have a different template on each slide, for instance.
13. Place the slides on a flat tray and load the tray into the argon chamber (and fill with argon). Allow gels to polymerize for ~30 minutes.
14. Remove slides from argon chamber. Use a razor-blade cleanly remove cover-slips from polymerized gels (just stick the edge of the blad under the coverslip and gently "pop" it up)
15. To wash off the excess acrylamide monomer, place slides in a dH20 filled plastic Coplin jar and incubate for 30 minutes at RT with slow shaking.
16. Remove slides from Coplin jar and place face up in PCR hood.
DIFFUSE IN PCR REAGENTS
17. Prepare diffuse-in mix while allowing slides to dry. You don't want the slides to over-dry, though. You will observe a thin, shrinking film of liquid on the surface of each gel. Generally, we try to add the diffuse-in mix within 5 minutes or so of this shrinking film's disappearance. This generally means about 30 minutes of drying.
18. Prepare diffuse-in mix (200 uL total volume):
152.33 uL dH20
10 uL 5 mM unlabeled dNTP mix
20 uL 10x Taq Buffer (with MgCl2)
1.33 uL 30% BSA
2 uL 10% Tween-20
1 uL unmodified amplification primer (100 uM)
13.33 uL Jumpstart TAQ (2.5 units / uL)
19. Pipet 25 uL of diffuse-in mix to the center of each gel. Assuming the gel is newly dried, the liquid will ball up rather than spreading over the surface.
20. Apply 18 x 30 mm cover-slip. Donít slide it on. Drop it on. Usually we do this by resting one edge of the coverslip on the Teflon coat and allowing the coverslip to "fall" onto the gel. The liquid should spread to evenly cover the surface of the gel.
21. Apply an orange SecureSeal chamber. Seal down the edges with the blunt end of a set of tweezers.
22. Fill chamber with mineral oil. This is a little tricky but you get good with practice. Use a P1000 pipetman filled with ~550 uL of mineral oil. Use a Chemwipe to dry off the tip of the pipet tip. The goal is to get the mineral oil in without spilling any near the surface of the hole (as this makes it more of a pain when you go to seal the hole with the stickies). A good technique to avoid spilling is to hold the slide tilted at a 45-degree angle while filling. Use the lower hole (in whatever direction you are holding the slide) to fill. It is OK to leave some air near the upper hole- more important to avoid letting oil seep out of it. When sufficient oil is in, turn the slide flat (0 degrees) before withdrawing the pipetman from the hole.
23. Wipe off any excess mineral oil from the surface of the SecureSeal, if neccessary. Seal holes with stickies.
24. Place slides in thermocycler (with labels facing out).
25. To PCR, cycle slides as follows:
Both the annealing temperature and the extension time can be adjusted to optimize for the set of amplification primers being used and the length of the PCR products being formed, respectively. We've been using the above protocol for an 800 bp template.
SLIDE CLEAN UP
26. Fill a GLASS Coplin jar with Hexane
27. Fill a PLASTIC Coplin jar with Wash 1E
28. Without removing cover-slips, place the slides in the hexane-filled glass Coplin jar and leave for 5 minutes. If the cover-slip comes off, don't worry about it. It doesn't seem to matter much.
29. One-by-one, remove the slides and cleanly pop off coverslips with a razor-blade as above. Quick wave in the air is sufficient to evaporate off residual hexane. Remove all residual adhesive from the cover-chamber using a razor-blade or your finger-nail (through the glove that you are wearing), and place the slide in the plastic Coplin containing Wash 1E.
30. Wash 2 x 4' in Wash 1E (meaning, 4 minutes shaking in Wash 1E, then replace the solution in the same Coplin jar with fresh Wash 1E and allow it to go another 4 minutes on the shaker).
31. Gels are fine in Wash 1E and can remain indefinitely (at least a week or two) before proceeding. Be sure to cap plastic Coplin jar containing slides if you are planning to let them sit for more than a day or two.