Sequencing Procedure

1. If new plates, mark the treated side of the plate with an RVS using the diamond pencil.
2. Using Alconox soap and glass sponge, clean larger (non-treated) plate with small circular motions including corners in dish sink using styrofoam box for support.
3. EtOH and wipe clean with kimwipe.
4. Lay on two green test tube racks and EtOH/kimwipe dust off.
5. Place side spacers with one foam end.
6. Repeat cleaning as in step 1 & 2 on treated glass (smaller plate).
7. Treated plate can be used 3-5X between treatings with Rain-X(see below).
8. Check treated plate for dust and clean.
9. Place plate treated side down onto of spacers.
10. Position bottom spacers so seal if formed.
11. Place one clip on each side and two clips on each corner, 5 clips on each side total.

1. Clean smaller treated plate as usually
2. Squirt a small amount of Rain-X onto kimwipes.
3. Apply to previously treated side of plate in circular motion until sol'n becomes cloudy (opaque).
4. Repeat steps 2 and 3.
5. Clean plate again and proceed with Plate Prep (above).

1. In 250 ml flask mix 38.4 g urea and 30 ml (26 ml) ddH₂O.
2. Dissolve by putting flask in pan H₂O bath on stir plate with heat on #3.
3. Add 1/2 to 1 g of Mixed Bed Resin.
4. Add 12 ml 40% 19:1 acrylamide/bisacrylamide.
5. Stir till dissolved, do not leave unattended.
6. While urea is dissolving, filter 4 ml 20X Glycerol Tol. Buffer (8 ml 10X TBE). IF using TBE for buffer, first filter 70 ml 10x TBE buffer to make buffer chamber solUn. Leave 4 ml (8 ml) in bottom of filter unit.
7. Make 10% APS by dissolving 0.1 g into 1 ml of ddH₂O in 1.5 ml bullet.
8. Once the urea sol'n is dissolved, filter it onto the 4 ml (8 ml) of filtered buffer.
9. Transfer sol'n into 125 ml flask
10. Clean filter unit by rinsing beads out and then vacuum washing 3X with distilled water.
11. Allow to air dry (filter can be used until tears are visible).

1. Use rinsed 250 ml flask from #1 above as syringe stand.
2. Add 20 l of TEMED and 600 l of 10% APS to gel mix, swirl well to mix.
3. Pour mix into 60cc syringe with ground off needle.
4. Angle plates with use of rack and hands and allow mix to gravity feed between plates.
5. Stop and remove any bubbles.
6. Place combs so bottom of oval lines up with top edge of smaller glass.
7. Pour sol'n to cover combs.
8. Remove clamps and slowly pull out bottom spacer.
9. Reposition top clip on each side inward slightly.
10. Tilt leftover sol'n in flask to indicate when plates have polymerizied.
11. Clean syringe, etc. so gel doesn't polymerize.

1. Prepare buffer while gel is polymerizing. In a grad. cylinder, dilute 25 ml of 20X Glycerol Tolerant Buffer (50 ml 10X TBE) for a FV of 500 ml in ddH₂O for upper buffer chamber.
2. Also, in a grad cylinder dilute 10 ml of 20X Glycerol Tolerant Buffer (20 ml 10X TBE) for a FV of 200 ml in ddH₂O for lower buffer chamber.
3. After polymerization, remove clips, rinse plates, remove combs and rinse excess acry away.
4. Close upper buffer chamber drain on right side of apparatus.
5. Place gel in "box", clamp into place, put buffer in upper chamber.
6. Put buffer in lower chamber making sure there are NO bubbles under gel.
7. Puff bubbles and urea from between plates.
8. Place combs so tips are at surface but not into gel.
9. Run at 65 watts for 5-15 minutes.

1. Radioactive components will be underlined from this point on.
2. Divide samples and place approx. half in 70°C heat block for 2-10 minutes.
3. Mark gel for samples (4 lanes each) and comb meeting.
4. Load 3 l of samples L to R, alphabetically (A,C,G,T). Put tips in solid radioactive waste container!
5. Run at 65 watts while other samples are in the heat block. Do this so a) makes gel asymmetrical b) gets samples into gel before diffuse.
6. Load other samples. Put tips in solid radioactive waste container!
7. Run total of 2' 45".
8. Prepare 80 g NaOAc into 200 ml (not FV) ddH₂O. Stir till dissolved on stir plate H₂O bath with some heat.
9. One hour before gel is done, add NaOAc sol'n to lower buffer chamber. Helps slow down front and compresses bands.
10. Cut a piece of Whatman paper 43 x 34 cm & roll up into pipet cylinder lid.

1. Prep: have two green test tube racks set aside for top plate (treated), rolled paper, water bottle, and spatula.
2. Drain upper chamber into holding well.
3. Disconnect plates.
4. Let plates drain into lower chamber -- bottom edge is hot!
5. Rinse off edge in sink.
6. On bench, remove combs.
7. Separate plates with spatula placing top plate gel side up on racks.
8. Squirt water around sides and top of gel on plate.
9. Position paper over the gel and place on gel from center.
10. Smooth firmly from center with hands.
11. Water assists in gel rolling up onto paper.
12. Transfer gel/paper to dryer.
13. Layer in the following manner (bottom to top): Mesh, filter paper to protect mesh, gel/paper (gel side up), saran wrap (must completely cover gel/paper), plastic cover, gasket.
14. Dry for 2 hours at 80°C.
15. Let cool 15-30 min before trying to remove from dryer.

1. Remove gel/paper from dryer.
2. Take gel, cassette and Biomax MR film to dark room.
3. Lock door and turn on outside "in use" light.
4. Place film in cassette with notch in upper right corner.
5. Place gel face down with top of gel at top of film, close the cassette.
6. Steps 3 and 4 allow gel to be read with the notch in the upper left corner.
7. Leave at room temperature in drawer for 20 hr for S³⁵, 20 hr for P³³, and 8 hr if P³³ is fresh.
8. To develop, place one edge against side of film developer.
9. Make sure write down what size film was developed on sheet.

1. Rinse clips, spacers, and combs. Allow to air dry.
2. Rinse plates well but not necessary to soap wash. Allow to air dry.
3. Put everything back in proper place when dry.