1 Jul 2011

Southern blot protocol

1. Genomic DNA.
Prepare using Gentra DNA prep kit, use 75ul of worms per prep (less is ok but more than 100ul will result in excess protein contamination as seen by a brownish/yellow pellet).
Wash pellet twice with cold 70% ethanol (this removes any excess salt that might alter the mobility of your sample in the gel). Note that although pellets are usually tightly bound to the eppendorf tube when first precipitated, pellets can loosen during the washes, so it is adviseable NOT to pour off the ethanol during the 70% washes but rather to remove it using a 1ml pipetman (so that loose pellets will remain in the bottom of the tube and will not end up in the sink). The same 1ml tip can be used for removing ethanol from multiple tubes, so long as you do not jab it down into the pellet, which could result in cross-contamination of your samples. I recommend stopping with about 50ul left and pipeting slowly so that if a DNA pellet is loose it will not get sucked up into the pipet tip.
After removal of the majority of ethanol from the second 70% wash, spin samples again briefly (to 8,000 rpm) to bring ethanol from the sides down to the bottom of the tube. Then use a P200 pipet tip to remove most of the remaining ethanol but not the pellet (side of tube that pellet rests against should face up, place tip against bottom of tube that does not contain pellet). Use a different tip for each tube at this step to avoid cross-contamination. Allow tubes to air dry for 10 minutes on a clean, dust-free benchtop. It is better if the pellets are still moist, as they are difficult to resuspend when dried completely.
Resuspend DNA in 60ul or so of DNA hydration solution, vortex several times at low speed (3 or 4 setting), resuspend overnight at room temperature, use a P200 to pipet up and down several times to make sure that the DNA is evenly dispersed (different tip for each tube), vortex several more times.
Expected results: There is apparently little correlation between the initial volume of worms and the yield of DNA. A cloudy DNA solution may look unsettling but usually cuts fine if white. Yellow or brown pellets or solution indicates protein contamination that will inhibit restriction enzymes and should be re-processed (see Gentra protocol) to ensure that all protein has been removed. This takes extra time but it is important to get all DNA samples right if possible.
2. HinfI digest.
As it is difficult to accurately guage the concentration of genomic DNA samples relative to one another, I recommend cutting 15ul of DNA for each sample, then adjusting the volume of each sample such that the concentration is slightly above that of the least concentrated sample. For each sample, add 2ul Neb 2 10x buffer, 1.5ul HinfI, 1.5 ul H20 for a 20ul reaction (make a master mix of these, aliquot 5 ul per tube with the same tip, then add 15ul genomic DNA and pipet up and down 3x to mix). Spin samples briefly to ensure that all DNA is in the reaction and incubate in 37C room overnight.
3. The Gel.
First, pour two large gels - one should be a 1% gel with many combs, so that you can check the DNA concentration three times; one should be a 0.6% gel for the Southern, which can use two combs for samples that are run halfway or a single comb for samples that can be run further (half way is fine to get a decent, publishable result on the Cambridge boxes). Make sure the combs have a nice visible space (1mm or so) from the bottom of the gel box, so that the wells will not crack when removing the combs. Also make sure that the combs are exactly parallel with the ends of the box, so that your samples will run straight. Make sure the gels boxes are very clean (any mineral oil will ruin the blot, and we have boxes set aside strictly for Southerns). I use 200 ml 1x TBE for the gel with 2ul EtBr solution added.
Add 2.5ul Loading Dye to the side of each tube with the same tip (a little more than 1x to ensure that the samples will sink to the bottom of the well). Spin the LD down and vortex each sample to on 3 or 4 speed to disperse the LD. Incubate rack at 65C for 15-20 minutes to denature any annealed, cut ends. During this time, prepare the gel by adding 1x TBE (roughly 1.8L), removing combs, and blowing out each well to be used twice with a 200ul pipetman using 1x TBE buffer in the gel box (this removes any high salt that may seep in from the gel and inhibit your samples getting to the bottom of a well). Load 2ul of each sample immediately (use a different tip for each sample to ensure that exactly the same volume is being loaded each time). The rack may have cooled by the time you finish, but the cut DNA ends should not have reannealed. Run gel at 100V to separate DNA, slice off the piece of interest, and take 1st picture. Most of the DNA should be below 2kb, but there may be bands at about 50kb and a few in between. Compare the DNA samples, lump the most dilute samples together, decide on the relative concentrations of the samples with more DNA (2x, 3x, 4x, etc), and dilute with 1x Neb2 buffer, 1x LD, in an attempt to equalize the concentrations, being careful not to dilute too much. Run samples again and take picture 2. Dilute again, run samples again, and take picture 3. Decide on the relative concentrations of each sample based on picture 3, and load the volume that you feel would best make for equal concentrations of samples. Again it is important not to underload, and do not to try to equal the least concentrated sample as there is almost always variation in the total amount of DNA loaded no matter how hard you try - instead try to adjust the volume such that it is equivalent amongst the majority of samples that are fairly dilute. Heat rack of samples to 65C, prepare the 0.6% gel as above, and load samples from warm rack (easiest to start at the top or bottom of your volume range and load all samples that need the same volume at once, though it is important that you load the correct lanes! Load 10ul 50ng/ul 1kb ladder at for all gels. Run gel overnight at 22V/cm or so (if midnight), try 15V/cm if 6pm.
The next day, stop the gel when BPB is still on it. Use a dedicated, clean Southern scalpel to cut the gel off at the BPB dye front (this removes everything below 1kb which is ok for telomere blots but you may want to leave it on if your fragment is small and you are using another probe). Remove the top 1cm of the gel below the wells (this gets rid of blank space that contains no DNA). Carefully measure the size of each gel piece. Use a spatula to move the gel to a clean piece of saran wrap, wipe the UV transilluminator down with water several times so that it is clean if your gel slips, dry it, take a picture of your gel, and carefully transfer the gel piece back into a bucket with ddH20 (don't dip your hand in or the Saran wrap in, if possible). Take picture of second gel piece if necessary.
4. Southern blot
Perform a downward blot as described in the Red Book (Current Protocols). It works very well (complete transfer occurs in 1hr, saving you a day).
Break large pieces of DNA by depurinating gel in 0.25M HCl for 7 minutes, shaking gently (too long and the DNA will be completely depurinated and in very tiny pieces). Rinse gel 2x with ddH2O, then denature DNA 2x 20 minutes with 1.5 M NaCl, 0.5 M NaOH. During this time, dump the gel box buffer into the liquid Ethidium Bromide waste, rinse the gel box carefully, replace lid and put it in a safe place (so that no one inappropriate can use it). Put on clean gloves (no powder), rinse off with ethanol, do not touch anything dirty, and use clean, flat forceps to handle the membrane (2 pairs is useful). Cut Hybond N membrane such that it is 0.5 cm larger than the gel dimensions, wet the Hybond in ddH20, pour off the water and incubate in 10x SSC until needed. Cut 3 pieces of Whatman to exactly the same size as the gel, cut two long pieces of Whatman that are the same width as the gel but much longer (so that they can wick buffer from a dish), cut a piece of Whatman paper a little larger than the membrane (use paper from membrane as reference), and make 4 copies of this larger piece. Place a 1" or 1.5" stack of paper towels that is larger than the gel dimensions in a pan (put two stacks together if necessary), add two pieces of the larger than the gel Whatman and top with three larger pieces soaked with 10x SSC from a large container (carefully lay these down and roll out any bubbles each time with a CLEAN pasteur pipet (that has been washed with ethanol-soaked kimwipe and then dried with a clean kimwipe).
Carefully lay down membrane on top of wet Whatman, then gel on top of membrane (use spatula), roll out bubbles each time (should not have to do for gel which is dripping wet), cut Saran wrap to lay on Whatman that lies to all sides of the gel and lay Saran Wrap down such that it just curls up each side of the gel (use pipet to roll it), add each piece of exactly the same size Whatman soaked in 10X SSC one at a time, rolling bubbles out each time, pile racks and place container with 10x SSC such that it is higher than the gel, soak long pieces of Whatman together and lay from gel into bucket, place light piece of plastic/glass on top of Southern to prevent evaporation, wait 1 hr (or more), dismantle Southern (gel should be flat and dye transferred to membrane), cut top right or left corner of membrane diagonally so you will know which side of the membrane is which, rinse membrane twice in 2x SSC to remove gel pieces, lay on newly cut Whatman paper cut to be larger than the membrane (DNA side up), autocrosslink DNA to membrane using UV crosslinker (I have forgotton this step it did not matter), place membrane in clean 2x SSC buffer. Prepare hybridization tube previously by washing well with hot soap and water, add 200ml 2x SSC to hyb tube, carefully place membrane DNA side-up in hyb tube using forceps such that there are no bubbles beneath the membrane, pour off SSC and add 20ml new hybridization buffer (prewarmed to 65C), rotate at 65C for 1hr.
Preheat 20ml hyb buffer at 65C in 50ml tube. Boil 600ng DIG probe + 7.5 ul DIG-labelled 10pg/ul 1kb ladder in boiling water bath on hot plate 5 minutes, cool in ice/NaCl bath, spin briefly, add 1ml warm hyb buffer to probe tube and transfer all of probe into 50ml tube, swirl briefly to disperse probe, pour off prehyb and pour probe/hyb buffer into hybridization tube, incubate at 65C overnight (be sure that blue side is pointing to the right).
The next morning, first prepare the DIG detection buffers, which must warm to room temperature before use. Make 500ml 2x SSC, 0.1% SDS, place half of this in a bucket, pour off probe into 50ml tube for storage at -20C, quickly move the membrane to the bucket (all incubations should be membrane-side-up which ensures contact with abundant buffer), rinse off excess probe twice with 2x SSC, 0.1% SDS using gentle shaking at room temperature. During this time I use the same 500ml bottle (dedicated SSC/SDS bottle), rinse it, make 500 ml 0.5x SSC, 0.1% SDS, place 200ml of this in well-cleaned hyb tube, carefully add membrane back to tube DNA-side-up, wash 15 min at 65C (incubate 200ml more of 0.5x SSC, 0.1% SDS in bottle at 65C at this time), pour off 0.5x SSC, 0.1% SDS and add rest of it for second 15 minute wash, pour last 100ml 0.5x SSC, 0.1% SDS (from 65C bottle) into clean bucket, pour off wash number 2 into sink and add membrane DNA-side-down to bucket with forceps to wash off any extra probe for 1 min (membrane is DNA-side down for all antibody incubations/washes but DNA side-up when performing hybridization and washes in tube with DIG probe), wash 1 min in DIG wash buffer, wash 30 min in 200ml buffer B (blocking), wash 30 min in buffer B + Ab (diluted 1:10,000), wash 2x15' in 200ml wash buffer, wash 2' in buffer C, place membrane DNA-side-up on transparancy cut larger than membrane but slightly smaller than second transparancy piece (on paper towel), add 2ml CSPD (40 drops or alter for membrane size), cover with second, large transparency piece, gently roll off bubbles, wait 5', roll off excess liquid (transfer several times to new paper towels), place transparency/membrane sandwich in plastic folder, incubate at 37C for 15', expose to film for 1 hr, develop. Always make several different exposure of every Southern to get spectrum of intensities (strong signal is sometimes good to see faint bands, weak signal is sometimes better for scanning/publication).

Depurination: 1 L 0.25 M HCl
12 ml 12N HCl (concentrated) + 988ml H20
Denaturation: 1.5 M NaCl, 0.5 M NaOH
300 ml 5M NaCl + 40 ml 12.5 N NaOH + 660 ml H20
Wash 1: 500ml 2x SSC, 0.1% SDS
50ml 20x SSC, 5ml 10% SDS, 445ml H20
Wash 2: 0.5x SSC, 0.1% SDS
12.5ml 20x SSC, 5ml 10% SDS, 482.5 ml H20
Standard hybridization solution: 200ml 5xSSC, 0.1% N-Lauroylsarcosine w/v (liquid), 0.02% SDS, 1% dry blocking reagent (Boerhinger)
50ml 20x SSC, 0.11ml 20% N-lauroyl sarcosine, 0.4 ml 10% SDS, 2g Blocking reagent, H20 to 200 ml.
Buffer to strip DIG probe off membrane: 400ml 0.2M NaOH, 0.1% SDS
4 ml 10% SDS, 6.4 ml 12.5N NaOH, 389.6 ml H20.

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