Electrophoretic Analysis of Replication Through Structure-Prone DNA Repeats Within the SV40-Based Human Episome

NOTE: The plasmid designed for our outlined 2DGE analysis in mammalian cells should contain an SV40 origin of replication several kb upstream of structure-prone repeats (Figure 2). Leading and lagging synthesis should be kept in mind when choosing what orientation relative to the origin the repeats should be cloned into the plasmid.

Figure 2: Digestion of repeat-containing plasmid for 2DGE analysis. Structure-prone repeats are depicted several kb downstream from the right-moving replication fork. Digestion with unique cutters 1 and 2 will place the repeat sequence on the descending arm of the Y-arc, given the sequence beyond the halfway point of the digested fragment. Abbreviation: 2DGE = Two-dimensional neutral/neutral gel-electrophoresis. Please click here to view a larger version of this figure.

1. Plasmid transfection into mammalian cells

1. Seed 600,000 HEK293T cells in a 10 cm tissue culture plate prior to transfection. Allow the cells to recover at 37 °C overnight.
   NOTE: Many cell lines may be used for this experiment, although cells containing the SV40 Tag are recommended for optimal results. CAUTION: HEK293T cells are considered BSL-2 and all culturing work should be carried out in a biosafety cabinet using appropriate aseptic technique and proper PPE.
2. When the cells reach 60% confluency, transfect 8 μg of repeat-containing plasmid DNA into the seeded cells using appropriate transfection reagents in accordance with the manufacturer's protocol.
3. If not isolating intermediates at this time point, aspirate old media and replace with 10 mL of fresh media after 24 h.
4. Begin collecting cells 24-48 h following transfection.
   1. Aspirate media and carefully wash with 10 mL of phosphate-buffered saline (PBS). Detach and collect cells using 0.5 mL of trypsin and spin down at 340 × g for 4 min.
   2. Aspirate the supernatant and wash the cell pellets with PBS. Spin again at 340 × g for 4 min and aspirate the supernatant.
      NOTE: The experiment may be paused here by freezing cell pellets at -80 °C. We have found the best resolution isolating replication intermediates 48 h following transfection; however, 24 h has yielded viable results.

2. Isolation of replication intermediates

1. Resuspend the cells in 1.5 mL of modified Hirt lysis buffer [10 mM tris-HCl (pH 7.5), 10 mM ethylenediaminetetraacetic acid (EDTA)] in 50 mL conical tubes and begin cell lysis.
   1. Add sodium dodecyl sulfate (SDS) to a final concentration of 0.6% (approximately 650 μL of stock 2% SDS) and proteinase K to a final concentration of 100 μg/mL (approximately 10 μL of stock 20 mg/mL proteinase K) to remove nucleases.
   2. Mix gently by pipetting until homogeneous and incubate the mixture at 37 °C for at least 90 min.
2. Increase NaCl concentration to 1 M (approximately 540 μL of stock 5 M NaCl) and mix gently until homogeneous. Incubate overnight (18-24 h) at 4 °C to allow for precipitation of cell debris, RNA, and protein by salting out.
   NOTE: The mixture will be highly viscous, so take care and be patient while mixing well.
3. The following day, separate the DNA from cell debris, RNA, and protein.
   1. Centrifuge the mixture at 29,500 × g for 45 min at 4 °C.
   2. Transfer the supernatant containing DNA, add one volume of phenol:chloroform:isoamyl alcohol 25:24:1 (v/v), and mix briefly until homogeneous.
      CAUTION: Phenol:chloroform:isoamyl alcohol is a hazardous material and should be handled with appropriate PPE in a chemical fume hood.
   3. Centrifuge again at 15,000 × g for 5 min at room temperature. Transfer the aqueous layer to a new conical tube.
4. Precipitate and wash the isolated DNA.
   1. Add one volume of pure isopropanol and incubate at room temperature for at least 5 min. Spin down the DNA at 15,000 × g for 30 min at 4 °C.
   2. Decant the supernatant and wash the pellet with cold 70% ethanol to remove excess salt.
   3. Spin once more at 15,000 × g for 30 min at 4 °C, air dry, and gently resuspend the pellet in Tris-EDTA (TE) buffer (10 mM Tris-HCl, 1 mM EDTA).
      NOTE: The experiment can be paused here, and samples can be frozen at -20 °C; however, freeze/thaw cycling should be avoided as this can decrease the quality of the DNA replication intermediates.

3. Sample preparation and 2-dimensional gel electrophoresis

1. Digest the isolated intermediates of plasmid replication.
   1. Add 100 units of the appropriate restriction enzymes to the sample to digest the plasmid DNA, specifically placing the repeat-containing sequence on the origin-distal half of the linear fragment (Figure 2). In addition, add DpnI to cut the methylated DNA, thus removing any plasmid DNA that was not fully replicated in cultured human cells.
      NOTE: For best results, restriction enzymes should be unique cutters that yield a 3-5 kb fragment that places the structure-prone sequence on the descending arm of the Y-arc.
   2. Incubate the samples at 37 °C for 6-10 h to allow full plasmid digestion.
   3. Precipitate the DNA with either 2.5 volumes of cold pure ethanol and incubate at -20 °C overnight, or add one volume of isopropanol and incubate at room temperature for 5 min.
   4. Centrifuge the digested and precipitated samples at 15,000 × g for 30 min at 4 °C.
   5. Decant the supernatant and wash the sample with cold 70% ethanol. Spin again at 15,000 × g for 30 min at 4 °C.
   6. Decant the supernatant, air dry for 10 min, and resuspend the samples in 15 μL of TE buffer.
2. Prepare the first-dimension agarose gel at 0.4-0.5% in 1x Tris-borate-EDTA (TBE) (89 mM tris base, 89 mM boric acid, 2 mM EDTA). Allow the solution to solidify for at least 1 h.
3. Begin loading the samples into the first dimension.
   1. Load the ladder within the first 3 cm relative to the gel's leftmost edge. Then, load the entirety of the prepared samples, ensuring 3 cm between each pair.
   2. Run the gel in 1x TBE for 19-24 h at 0.85 V/cm to separate the intermediates with respect to their size. Ensure the chamber is covered to protect the samples from light, which may cause damage to the DNA.
4. The following day, remove the gel from the buffer and estimate the location of the digested linear fragment using a ruler.
   1. Excise the first 3 cm of the gel containing the ladder and stain the gel segment in 1x TBE containing 0.3 μg/mL ethidium bromide for 10-15 min. Visualize the ladder using a gel documentation system.
   2. Add 1.3 cm to the estimated location, yielding value a. Then, subtract 7.5 cm from value a, yielding value b. Align the ruler against the first-dimension gel and cut horizontally across at values a and b. Then, cut vertically down the 3 cm space reserved for each sample. Refer to Figure 3 for a visual schematic.
   3. In a new casting tray, rotate the segments clockwise and place them in the position of the sample wells (Figure 3).
5. Prepare the second-dimension agarose gel at a concentration of 1-1.3% in 1x TBE at 0.3 μg/mL ethidium bromide.
   1. Upon cooling to approximately 55 °C, pour the second-dimension gel over the rotated first-dimension segments and allow it to solidify for at least 1 h.
   2. Transfer the second dimension to a chamber with 1x TBE at 0.3 μg/mL ethidium bromide, and allow the gel to equilibrate for at least 30 min.
   3. Run the gel, again covered, for 9-10 h at 4.23 V/cm at 4 °C to separate the intermediates with respect to their shape.

Figure 3: Excision of first-dimension intermediates prior to second-dimension separation. Following visualization of the ladder, the mobility of unreplicated fragments can be estimated. (I) This value can then be used to determine appropriate cut sites (a and b) to excise it and its replicated counterparts (II). The section of the gel should then be rotated and placed in the position of wells for the second-dimension separation. Abbreviation: CW = clockwise. Please click here to view a larger version of this figure.

4. Southern blotting and hybridization with radiolabeled probe

1. Remove the second-dimension gel from the chamber and depurinate the DNA fragments for 10 min in a 0.24 M HCl solution with gentle rocking. Rinse the gel with deionized water and soak it in 0.4 M NaOH for 10-15 min.
   CAUTION: HCl and NaOH are corrosive and should be handled with appropriate PPE in a chemical fume hood.
2. Begin assembling the Southern blot to facilitate the transfer of the separated intermediates from the gel to the membrane. Refer to Figure 4 for a comprehensive schematic.
   1. Fill a sufficiently sized container with 1 L of 0.4 M NaOH.
   2. Align a long glass sheet across the container and fold (along the length) two long sheets of chromatography paper perpendicular across the glass sheet, extending into the container of NaOH.
   3. Wet the top of the paper with NaOH and carefully remove any air bubbles under its surface.
   4. Soak three sheets of chromatography paper with NaOH and place them over the folded paper, again removing any bubbles.
   5. Flip the second-dimension gel upside-down and transfer it over the papers.
   6. Wet a positively charged nylon membrane (0.45 μm pore size) with DI water and place it over the gel.
   7. Finally, add three more sheets of paper, wet with DI water, over the membrane.
   8. Cover any exposed NaOH in the bottom container with plastic wrap to prevent evaporation. Place a stack of either napkins or paper towels over the blot, ensuring it is 0.3-0.5 m tall. Place a weight on the top, compressing the entire blot to facilitate tight capillary action. Allow at least 2 days for DNA to transfer to the membrane.
      NOTE: The length and width of the chromatography paper and membrane depend on the size of the agarose gel used for the second dimension. For the most efficient transfer, use paper and membrane with the same dimensions as those of the gel.
3. Following transfer, crosslink the DNA to the membrane using a UV crosslinker at 120 μJ/cm² for 1 min.
   NOTE: The experiment may be paused here by placing the membrane in an airtight, dry, and clean sheet protector at room temperature.
4. Wash the membrane 2x for 5 min with 2x saline sodium citrate (SSC buffer) (0.3 M NaCl, 0.03 M sodium citrate).
5. Prehybridize the membrane with 0.18 mL/cm² of Church & Gilbert's hybridization buffer [1 mM EDTA, 1% bovine serum albumin (BSA), 0.5 M sodium phosphate, 7% SDS] at 65 °C, rotating in a hybridization incubator for at least 2 h.
   NOTE: The membrane may prehybridize for several days.
6. Prepare the radiolabeled probe using α-³²P dATP or dCTP and a DNA labeling kit in accordance with the manufacturer's protocol.
   CAUTION: Radiolabeled dNTPs are hazardous, and appropriate PPE should be worn when handling them. All radioactive work should be performed behind shielding, and trained individuals should be monitored for radiation uptake using dosimeters.
   1. Design a 400-900 bp linear DNA fragment complementary to the restriction digested sequence (Figure 2) and amplify the fragment using polymerase chain reaction (PCR).
      NOTE: We recommend having a stock PCR fragment of 50-100 ng/μL
   2. Combine 100 ng of the complementary PCR fragment with DNA Pol I, Klenow fragment (3' 5' exo-) buffer, and random decanucleotide oligos.
   3. Denature the fragment at 100 °C for 10 min.
   4. Add 5 units of DNA Pol I, Klenow fragment (3' 5' exo-), 50 μCi of α-³²P dNTP, and 30-50 μmol of dNTP mix deficient in radiolabeled dNTP type to the sample.
   5. Incubate at 37 °C for 10 min to allow for polymerization and radiolabeled dNTP incorporation.
   6. Add 30-50 μmol of previously absent dNTP type to the sample and incubate at 37 °C for 10 min.
   7. Purify the radiolabeled fragment using a spin column at 3,000 × g for 2 min.
7. Add radiolabeled probe to 50 mL of hybridization buffer and incubate with the membrane overnight at 65 °C rotating in a hybridization incubator.
8. The following day, remove the probe and wash the membrane 2x with wash buffer 1 (0.1x SSC, 0.1% SDS) at 42 °C and 2x with wash buffer 2 (2x SSC, 0.1% SDS) at 65 °C.
   ​NOTE: All washes should be performed with fast rotations for 15 min in the incubator.
9. Dry the membrane for 10 min and place it in a thin, transparent sheet protector. Store the sealed membrane in a controlled, radiation-resistant cassette with a phosphor-sensitive screen. Allow the membrane to be exposed to the screen for 1-10 days.
10. Visualize the results using a biomolecular imager set to phosphor imaging. Wash and re-expose if necessary.

Figure 4: Assembly of Southern blot. Comprehensive schematic of a typical apparatus used for Southern blot transfer of intermediates from the second dimension onto a nylon membrane. Please click here to view a larger version of this figure.