Analysis of DNA Replication in Fission Yeast by Combing

Labeling Cells

Plug Preparation and Cell Wall Digestion

• For plug preparation, always use 2 mL round-bottomed microcentrifuge tubes to avoid damage to plugs.

• 7. Pellet 1 × 10⁸–2 × 10⁸ cells from a mid-log phase culture labeled with analog as in Steps 1–3 or 4–6.

• 8. Wash pellet twice with 1 mL spheroplasting buffer in a 2 mL round-bottomed microcentrifuge tube. Resuspend cells gently with a pipette, and centrifuge at 5000g for 1 min, at room temperature.

• 9. Add 150 µL of DNA combing enzyme mix to the washed pellet. Mix well and incubate for 5 min in a 37°C heat block.

• 10. Add 150 µL of 1.5% LMP agarose (cooled to 42°C) using a cut off 200 µL tip and mix well.

• 11. Immediately dispense 150 µL into each well of the plug mold (two wells per sample) and leave for 20–30 min at 4°C to solidify.

• 12. Place the two plugs in a single tube and add 1 mL of DNA combing plug solution.

• 13. Incubate the plugs for at least 6 h (or up to 9 h) in a 37°C heat block.

Proteinase K Treatment

• 14. Transfer the plugs into a new 2 mL round-bottomed tube containing 1 mL Proteinase K buffer and incubate overnight in a 50°C water bath.

• 15. Replace the Proteinase K buffer roughly every 12 h. Continue the Proteinase K treatment for 60 h. Change the Proteinase K buffer a total of five times.

  • Make fresh Proteinase K buffer for each change.

TE Washes

• 16. Remove the buffer and transfer the plugs to a 15 mL conical bottom tube.

• 17. Wash plugs twice for 2 h each time with 10 mL TE + 1 mL 0.5 m EDTA pH 8.0 with gentle rocking at room temperature.

• 18. Wash plugs twice for 2 h each time with 10 mL TE with gentle rocking at room temperature.

  • The plugs can be stored at 4°C in TE for many months until needed.

Melting Plugs

• 19. Place one-half or one plug in a 2 mL round-bottomed microcentrifuge tube.

• 20. Add 500 µL TE/YOYO, 480 µL 0.5 m MES pH 5.35 and up to 1.4 mL H₂O.

  • The pH of MES is extremely critical for combing. It determines the density of DNA that will stick to coverslips and their degree of stretching. Titrate the pH of MES using λ DNA. Prepare several combing solutions of λ DNA with MES buffer of varying pH. Add 200 ng λ DNA to 500 µL TE/YOYO and 480 µL 0.5 m MES of pH varying between 5.2 and 6.5 in a final volume of 1.4 mL. Comb the λ DNA and check for density of fibers and their stretching as described below. With 200 ng of DNA, the coverslip should be densely covered with fibers. Use very small increments of pH for standardization (e.g., 0.05). Once the pH is standardized, use the optimum pH for melting sample plugs. Combing can vary substantially between pH 5.30 and pH 5.35; a more basic pH gives fewer, but longer fibers on the coverslip and a more acidic pH allows more DNA of shorter length to stick on the coverslip but allows more stretching.

• 21. Incubate the plug for 20 min at 65°C. Confirm that the plug has fully melted. If it has not, continue incubating for another 10 min.

• 22. Transfer the plug to 42°C and incubate for 30 min.

• 23. Add 40 µL β-agarase mix (4 µL 10× NEB buffer, 4 µL β-agarase, 32 µL sterile H₂O) to the tube. Digest overnight at 42°C.

  • Do not mix after adding β-agarase. Gently swirl with a tip if needed. After digestion the samples should appear very clear. Usually, a small amount of wispy, thread-like agarose remnant is seen but there should be very few, if any, agarose clumps. If clumps are present, remelt the plug at 65°C and digest again with β-agarase mix. Improper cell wall digestion will make the plugs appear very cloudy and the β-agarase digestion will not occur optimally leading to poor DNA recovery.

• 24. Centrifuge the melted plug at 800g for 5 min at room temperature.

  • Handle the tubes with extreme care to avoid any damage to DNA fibers.

• 25. Transfer the supernatant directly into the Teflon reservoir of a DNA Combing instrument (see Step 35) or, if the sample is to be stored for latter processing, into a new round-bottomed tube using a cut off 1 mL tip.

  • This supernatant is the DNA combing solution and there should be hardly any pellet. Use extreme care while transferring the supernatant to avoid DNA fiber breakage. The combing solution can be stored at 4°C. However, with time the DNA fibers break; therefore, avoid storing for >1 mo.

Preparation of Silanized Coverslips for DNA Combing

• Coverslips are cleaned in a plasma cleaner such as a Harrick basic plasma cleaner. Liquid-based cleaning protocols can be used if a plasma cleaner is not available (Demczuk and Norio 2009; Marheineke et al. 2009).

Combing

• This protocol is based upon the DNA Combing System instrument from Genomic Vision, although other surface-coating systems should work (Marheineke et al. 2009). Over 50 coverslips can be combed using the combing solution from one-half a plug.

• 35. Transfer the DNA combing solution very carefully (to avoid DNA breakage) into a Teflon reservoir.

  • To avoid breakage of DNA, after spinning the melted plug the supernatant can be transferred using a cut off 1 mL tip directly into the Teflon reservoir, instead of transferring it into a new tube and then into the reservoir.

• 36. Attach silanized coverslips to the instrument holder and dip it into the reservoir for 5 min.

• 37. Withdraw the coverslips at a constant speed of 500 µm/sec to allow DNA molecules to stretch.

• 38. Check the density of fibers and stretching on the coverslips by epifluorescence using a GFP filter set and a 100× objective to visualize YOYO-1 stained DNA fibers. Select coverslips with well-stretched fibers of appropriate density for further analysis. Usually 5–10 coverslips will be suitable for further analysis.

  • The coverslips can be easily viewed with an inverted microscope. An upright microscope can also be used, but a coverslip holder is required to hold the coverslip DNA-side down while it is being viewed. Such a holder can be made by cutting a square hole in a thin piece of stiff plastic.

• 39. Aspirate away the immersion oil from the coverslip. Place a drop of super glue on a slide. Place the coverslip oil-side down on the glue. Label the slide using pencil.

• 40. Incubate the slide for 2 h or overnight at 65°C.

  • Baking robustly attaches DNA strands to the cover slip.

  • Proceed to Step 41 or Step 56.

Triple Staining (Two Analogs and ssDNA)

• 41. Cool slides for 5 min at room temperature. Place the slides in a Coplin jar. Denature the slides with freshly prepared 0.5 n NaOH or 2.5 n HCl for 30 min; add acid or alkali and place the jar on a rocking platform.

  • The choice of denaturing agent, either HCl or NaOH, depends on the batch of coverslips. Test the coverslip batch for optimal staining using spare labeled DNA before proceeding with test samples. Typically, CldU is best visualized using HCl denaturation. However, IdU staining does not work well with HCl denaturation. IdU is best visualized using NaOH denaturation, while CldU staining is moderate. At low concentrations, CldU staining is quite punctate, which can complicate analysis. At high CldU concentrations, NaOH gives good results.

• 42. Transfer the slides to a fresh Coplin jar and wash the slides with 0.1× PBS three times for 5 min each on a rocking platform.

• 43. Place the slides in a humid chamber (box containing wet paper towels).

• 44. Block coverslips with 1% BSA by adding 50 µL of solution per coverslip. Place a second coverslip on top of each to spread out the solution and to prevent evaporation. Incubate the box for 25 min at 37°C.

  • Apply fresh top coverslips as described in Step 44 for all subsequent antibody incubations.

• 45. Remove the top coverslip by dipping the slide horizontally in a wide chamber containing 0.1× PBST. Wash the slides with 0.1× PBST twice for 2 min each.

  • Remove top coverslips as described in Step 45 for all subsequent washes. In most cases the top coverslip will float off easily; however, it may be necessary to gently ease it off using a micropipette tip.

• 46. Add 50 µL CldU (rat) and IdU (mouse) primary antibodies. Incubate for 1 h at 37°C.

• 47. Wash the slides with 0.1× PBST twice for 3 min each.

• 48. Add anti-rat 594 and anti-mouse 488 antibodies. Incubate for 30 min at 37°C.

• 49. Wash the slides with 0.1× PBST twice for 3 min each.

• 50. Add rabbit anti-ssDNA primary antibody. Incubate for 1 h at 37°C.

• 51. Wash the slides with 0.1× PBST twice for 3 min each.

• 52. Add anti-rabbit 350. Incubate for 30 min at 37°C.

• 53. Wash the slides with 0.1× PBST twice for 3 min each.

• 54. Drain off excess 0.1× PBST. Add 18 µL of anti-fade mounting medium per coverslip. Place a top coverslip on each and seal with nail polish. Allow the nail polish to dry before visualization.

  • If using ProLong Gold Antifade, allow the slides to cure for 24 h at room temperature before sealing with nail polish.

• 55. Keep slides at −20°C for long-term storage.

Double Staining (Single Analog and ssDNA)

• 56. Follow the same protocol as for triple staining (Steps 41–55), except at Step 46 add a single primary antibody instead of both and similarly add a single secondary antibody at Step 48.

  • Addition of analog and ssDNA primary antibodies together may compromise analog staining; therefore, add them sequentially. ssDNA can be visualized better in 488 and 594 channels as opposed to 350. Therefore, depending on the secondary antibody used for the analog, either Alexa fluor 488 or 594 labeled antibodies can be used for ssDNA visualization rather than an Alexa fluor 350 labeled antibody.