"Blotting" to Nylon membrane

always wear gloves!

• Run PCR Products to be probed on an agarose gel.
• Illuminate the gel with UV light and take a picture of the gel. Take care not to expose the gel for too long as prolonged exposure can degrade DNA
• Mark positions of bands in ladder in gel by poking holes through the gel with a paper clip.
• Depurinate (this is important to ensure efficient transfer of PCR products greater than ~ 1 kb): Place 250 ml 0.2 M HCl in glass baking dish. Add gel. Rock gently room temp, 10 min (and NO LONGER or DNA will be broken to bits). (0.2M HCl =16.7 ml concentrated HCl per liter, store less than 2 weeks). Rinse 2X briefly (and carefully) with water.
• Denature: soak gel for 15min in 250 ml of Denaturing solution in the same baking dish. Carefully pour out the old denaturing solution and replace with 250ml fresh Denaturing solution. Soak for 15 more min.
• Carefully pour out the denaturing solution and briefly rinse the gel with water. Add 250ml 5xSSC to the dish and wash for 15min, rocking.
• While gel is washing, cut two pieces of Whatman Paper and two pieces of Magna Charge nylon membrane to be a wee bit larger than the gel itself. (setting up the blot is easier if you err on the side of too large a piece, rather than too small).
• Wet the nylon membranes and Whatman Paper in 5xSSC.
• Once the gel has been washed, make a sandwich as follows from the ground up making sure to avoid bubbles...

One Way Transfer:

1. A glass plate large enough to hold the gel.
2. Two pieces of wet (5xSSC) Whatman Paper (just a bit larger than your gel).
3. The gel (avoid air bubbles by rolling with a test tube)
4. A piece of wet (5xSSC) MagnaCharge nylon membrane (again avoid air bubbles and have the membrane a bit larger than your gel).
5. A “mask” around the edge of your gel (I generally use old developed film).
6. Two pieces of wet (5xSSC) Whatman Paper (just a bit larger than your gel).
7. A 1" tall stack of paper towels

Two Way Transfer:

1. A 1" tall stack of paper towels
2. A piece of wet (5xSSC) Whatman Paper
3. A piece of wet (5xSSC) MagnaCharge nylon membrane (avoid air bubbles)
4. The gel (avoid air bubbles)
5. Another piece of Magna Charge nylon membrane (avoid air bubbles)
6. Another piece of Whatman Paper
7. Another 1" tall stack of paper towels
8. An electrophoresis box top to hold everything down.

Let the sandwich sit at least 3 hours or overnight.

Disassembling Transfer/Crosslinking:

• Remove the dry paper towels and save them. Toss the wet paper towels.
• Using a pencil, trace the well holes onto the nylon membrane. Also, make pencil marks through the gel holes indicating the MW ladder bands, so the marks appear on the nylon. It is important that the markers be placed asymmetrically on your gel so that you can determine the orientation of the gel from these marks.
• Peel the membrane off the gel and lay it on a paper towel DNA-side up.
• Crosslink the DNA onto the membrane using a UV crosslinker at the 'Optimal' setting on Stratalinker. The blot can now be air dried and stored at room temp until ready to hybridize.

Prehybridizing:

• Wet the membrane in 5xSSC briefly.
• Roll up the wet membrane and place it into a roller bottle.
• Add 10ml of Church Hybridization Buffer and incubate at 65°C for 30min.

Hybridizing:

• Pour out the pre-hyb solution and add 10ml of fresh Church Hybridization Buffer.
• Roll in the oven until the solution has warmed to 65°C.
• Place 100-200ng of probe in 200 µl TE in a 1.5 ml microfuge tube, and place in 95-100¾ heat block or boiling water bath for 5min.
• Pipet the probe into the bottom of the hybridization buffer, avoiding contact with the membrane.

If this is a reused probe that is already in hyb solution...

• Heat probe /hyb buffer to 95°C for 20min (in beaker of water).
• Pour out the pre-hyb solution and add the hybridization solution to jar with blot.
• Hybridize 2hrs-overnight, rolling, at 65°C
• Put a bottle of 0.2xSSC/0.1%SDS in the 65°C oven so that is warm for the next day.

Washing:

• Pipet out the hyb solution/probe using a disposable transfer pipet.
• Store the probe/hyb solution at -20°C, labelled with date, probe name, and your initials.
• Wash the membrane FOUR times in 0.2xSSC/0.1% SDS for 15-20min each wash. 65°C, rolling in the oven.
• After adding each fresh wash solution, agitate the bottle so that the membrane comes away from the edge of the tube and is completely immersed in solution.
  

Detecting:

• Cool the oven down to room temperature or transfer to different bottle roller at room temp.
• Rinse the membrane in the jar with 50ml FRESHLY MADE TBST for 5min. REPEAT.
• Block with 50ml TBST and 1% casein, rolling for 1hr at room temperature.
• Incubate blot with 1µl anti-DIG Antibody in 20ml casein/TBST solution. Agitate bottle to insure good contact between the blot and the antibody. Roll at RT for 30min-1hr.
• Wash with 50ml of TBST 5min. REPEAT.
• Transfer the blot to a clean, glass dish and rinse with 100ml TBST. Gently rock for 10min at room temperature.
• Do THREE RINSES with TBST like this.
• Pour TBST out of the baking pan and add 100ml of FRESH Detection Buffer. Rock 5min.
  Place membrane in a Seal-a-Meal pouch. Add 5µl CDP-Star in 0.5ml Detection Buffer to the DNA-side of the membrane. Squeeze out any excess bubbles and seal up the bag.
• In the dark room, expose the membrane in the bag to X-ray film, in light tight folder. Develop film after 5-20 min (depending on probe quality).
• Do a second exposure up to ~ 2 hours, depending upon signal in the short exposure. Aim NOT to over-expose blot, because signals saturate and you lose the ability to distinguish real signal from background. If the positive control signal is weak after 2 hours, dig probe is bad or something else is wrong.
• Rinse the blot with water prior to storage. If you don't do this, your blot will be ruined and cannot be reprobed. Store dry for a few days, in case you want to reprobe the blot.

Solutions for DIG Southerns:

***May need to be warmed in order for SDS to go into solution

***Make Fresh! Do not use if more than 3 days old.

***The MgCl2 in this solution likes to precipitate. This causes excessive background spots. Make fresh and filter through two layers of Whatman paper #4 immediately before using.

***Disolve 20g Sodium Casein in 700ml water. Casein is slow to disolve. Even with stirring and heating it can take several hours. Bring up the volume to 1L. Aliquot into 40ml batches and store in the -20° Freezer.