20.109(S14):Complete Western and prepare damaged DNA (Day4)

Introduction

More coming Wednesday. Thanks for your patience!

Brief reminder of where we are. Then…

Topic 1: gel purification and perhaps idea of diagnostic digestion more broadly

Topic 2: western secondary considerations and visualization options

Protocols

Today you get to experience grad student life, juggling multiple assays blah blah blah...

Part 1: Digest plasmid for NHEJ assay

• You will digest pMax-BFP-MCS at the cut site(s) that you chose to investigate last time. We will then evaluate and purify the DNA using gel electrophoresis.
• To avoid pipetting very small volumes, you will either prepare a reaction cocktail that uses no less than 1 μL of any restriction enzyme, or you will prepare an intermediate dilution of said enzyme(s).
• Note that enzyme stock concentrations can be found on the NEB product page for that enzyme.

1. By whichever approach outlined above, combine 3.5 μg of DNA with water, buffer, and enzyme in a well-labeled eppendorf tube. Whether you prepare an enzyme dilution or a master mix, the enzyme should be added last.
   • Why? What would happen if you added the enzyme directly to water?
   • Recall that you are using 2.5 U of each enzyme per μg of DNA.
2. Flick the tubes to mix the contents, touch-spin, then incubate the mixtures at 37°C for at least one hour. Write down your start time and also set a timer, in case you get distracted later on.
   • While your samples are digesting, you can finish the Western. (Or at least start finishing it!)

Part 2: Complete Western protein assay

A week ago, you prepared protein extracts from K1 and xrs6 gels, separated them by SDS-PAGE, and transferred them to a nitrocellulose membrane. On that day, blots were moved to blocking buffer. Four days later, they were moved to plain PBS. Finally, late yesterday, they were incubated with primary α-Ku80 antibody overnight at 4 °C. (See reagent list at end of lab for concentrations/compositions/etc.)

1. Obtain your blots from the front bench. Pour the antibody solution into a conical tube, writing the identity of the antibody and the date on the tube.
   • Because the antibody is in excess, sometimes the primary solution may be re-used on another blot. Worth saving until we see how our Westerns come out today, at least!
2. Add enough TBS-T to cover your membranes – between 10-15 mL should work, but you don't need to measure out this volume. Keep in mind that the washing steps work by dilution, so it is a balance between adding enough to create a sink for the primary antibody, but not so much that you make a huge mess on the shaker!
   • TBS-T stands for Tris-buffered saline with Tween 20 (a surfactant).
3. Shake your container for 5 min at 80 rpm, using the room temperature shaker in fume hood.
4. Repeat for a total of 3 washes.
5. Just before pouring off the last wash, prepare the secondary antibody. Specifically, prepare a 1:3000 dilution of GAR-AP in TBS-T at a total volume of 9 mL.
   • GAR-AP stands for goat anti-rabbit--alkaline phosphatase conjugate.
6. Shake at 80 rpm for about 45 minutes.
7. Wash the blot as before, for 3 washes.
8. Once you start the third wash, begin to prepare your development solution. Add 1 mL of development buffer stock to 24 mL H2O. Next, add 250 μL of reagent A and 250 μL of reagent B to the well-mixed buffer.
9. After removing the last TBS-T wash, add about half the development solution to your blot; save the rest for now.
10. Put your blot on the shaker. Check it about every 10 minutes. Full development should take somewhere between 20 and 60 minutes, perhaps a little longer.
    • Be sure you know what size band you are looking for! There are likely to be some cross-reactive/off-target/non-specific bands as well.
11. Once you are satisfied with the look of your bands, remove the development solution and add distilled water. Shake the blot for a final 10 minutes.
12. Finally, move the blot on top of a piece of filter paper to dry. Either you or the teaching faculty will take and post pictures of these on the wiki, depending on whether they appear ready by the end of lab.

Part 3: Gel purify digested plasmid

1. When your digest is ready – and you are, too – add 5 μL of 6x NEB loading dye to it, and then pipet 27 μL (or however much you end up with after pipetting error losses) into a 1% gel according to the scheme in the table below.
2. The gels will be run for about 30 minutes at 100 V, which should give sufficient separation between the plasmid fragment of interest and the nonsense fragment.
3. At least 10 minutes before your gel run is slated to be over, label and weigh an eppendorf tube.
   • I usually find it easiest to write the value right on the tube, especially if I am measuring multiple weights.
4. After your gel run is finished, the teaching faculty will show you how to safely cut the band out of your gel.

Three groups can fit on one gel. We are leaving space between the samples for two reasons:

• We don't want the differently digested DNA bands to bleed into each other, but rather to be well separated when they are being cut.
• We don't want to expose the bands that are cut out later to too much excess UV.

Lane	Sample (27 μL)	Lane	Sample (27 μL)
1	DNA ladder (load 10 μL)	6	BLANK
2	BLANK	7	Group 3
3	Group 1	8	BLANK
4	BLANK	9	DNA ladder (load 10 μL)
5	Group 2	10	BLANK

To purify your DNA from the agarose, you will use a kit from the Qiagen company. As we learned during Module 1, reagents in such commercial kits can have uninformative names and their contents are in part proprietary.

1. Estimate the volume of your gel slice by weighing it.
   • The easiest way to do this task is to pre-weigh an eppendorf tube (above), then weigh it again after adding the gel, and take the difference.
   • What can you assume about the density of agarose and why?
2. Add 3 volumes of QG for every 1 volume of agarose.
3. The maximum advised volume is 550 μL. If you have a greater volume, continue for now, but first read step 6 to understand how to proceed later. Feel free to ask the teaching faculty for clarification.
4. Incubate in the 50°C water bath for 10 minutes, until the agarose is completely dissolved. Every few minutes, you should remove your tube from the 50°C heat and flick or vortex it for a few seconds to help dissolve the agarose.
5. Add 1 volume — original gel volume, not current solution volume — of isopropanol to the dissolved sample and pipet well to mix.
6. Get one QIAquick columns and one collection tube from the teaching faculty. Label the spin-column (not the collection tube!) with your team color. Gently pipet the dissolved agarose mixture onto the column. Microfuge for 60 seconds at maximum speed (approx. 16,000 rcf). The maximum capacity of the QIAquick columns is 800 uL! If you have more than 800 uL in your mixture, you will need to repeat this step using the same column.
7. Discard the flow-through in a temporary waste conical tube and replace the spin-columns in their collection tubes. Add 500 μL of QG to the top of the column and spin as before.
8. Discard the flow-through as before, and then add 750 μL of PE to the top of the column and incubate for 5 min at room temperature.
9. Spin for 1 min at max speed.
10. Discard the flow-through once more and replace the spin-column in its collection tube.
11. Add nothing to the top but spin for 60 seconds more to dry the membrane.
    • This step completely removes remaining ethanol.
12. Trim the cap off of a fresh, pseudo-sterile eppendorf tube, and prepare a sticky label (in your team color) for the top: write "M2D3", your section day, and your team color. Please also add a plain sticky label to the side of each tube, so we don't lose track of whose sample is whose later on!
13. Place the labeled spin-column in its matching trimmed eppendorf tube and add 30 μL of EB to the center of the membrane.
14. Allow the columns to sit at room temperature for one minute and then spin as before. The material that collects in the bottom of the eppendorf tubes is your purified, digested DNA.
15. If you have time, join Su (one team at a time!) to observe her measure the DNA concentration on a Nanodrop in the Niles lab just down the hall.
    • Because your sample is precious and we don't have much to spare, using enough sample to get a good reading on our DU640 spectrophotometer would not be a great idea today!
    • A Nanodrop can reliably evaluate nucleic acid concentrations of very small volumes!

For next time

revising microbiota summaries, mainly

cell doubling calculation for fun

first steps of lipofection calculation? (suggestion though, not collected?)

Reagent list

write something here or not accessible to edit

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