We are currently constructing (for DNA) and validating (for DNA and cells) our system for investigating NHEJ. In fact, it's quite typical in scientific research that a long development phase is required just to get to the point where a researcher can collect the data that is actually of interest to her – much less to process and make sense of it!
You’ll begin the day by implementing the cut topology that you chose last time via restriction digest of the pMax-BFP-MCS plasmid. Introducing restriction site-based damage into DNA is just one creative way that scientists and engineers have harnessed the power of nature. In both Module 1 and this current module, you have witnessed molecules (such as ligase) and cells (such as E. coli) stripped from their natural context and applied to solve a problem. Though it now seems mundane, this practice of purifying and re-engineering biology for human ends is pretty amazing if you think about it! Several of the enzymes that will be used in class are “high fidelity” variants, designed for more specific activity than nature requires. Other enzymes have been engineered for faster kinetics.
The way that we are using restriction enzymes in Module 2, while not unprecedented, is somewhat unusual. There are two far more common laboratory practices that utilize restriction enzymes. One we have already examined in some detail, namely restriction-based cloning of distinct pieces of DNA with compatible ends. Briefly, an insert and vector are ligated together, and then amplified in bacteria. Next, individual DNA clones are miniprepped and tested for correctness. As for the second practice, it turns out that restriction enzymes are just as useful in assessing whether cloning worked as they are in creating a clone in the first place. You learned last time that KpnI was used to insert the dual-MCS construct into pMax-BFP; in other words, non-directional cloning was performed. Several miniprepped clones were immediately subjected to a so-called “diagnostic digest”: a clone is digested with (usually) two enzymes, one on the insert and one on the original vector, and the resultant banding patterns suggest which orientation the insert ended up in. One or a few clones that appear to be in the correct orientation can then be sent off for sequencing.
Now you might be wondering why you would ever go through the trouble of designing and performing diagnostic digests, when sequencing is relatively simple and yields more information. Here, the idea of scale becomes important. Sequencing costs $6-8 per reaction, which can add up if you need to examine, say, 10 or more candidates. Agarose gel electrophoresis, by comparison, costs perhaps $1 per candidate. Since both methods require DNA isolation, one is not dramatically more labor intensive than the other. Finally, banding patterns can give a quick readout of many candidate colonies at once compared to the time it takes for individual sequencing analyses (at least if performed manually).
But I digress.
After digesting plasmid DNA to create a desired topology, you will evaluate the success of the digestion on a gel, and then isolate and purify the primary DNA fragment from said gel. You already know plenty about agarose electrophoresis, and a fair bit about purifying DNA as well. In gel purification, a DNA band is melted, and then isolated on a silica (SiO2) column similar to the ones you used in Module 1. Salt concentration and pH effects, along with ethanol precipitation, will alternately allow for binding and eluting the DNA while washing away contaminants.
Besides DNA preparation and evaluation, you will complete the Ku80 Western blot that you began last week. Each blot has already been probed with primary α-Ku80 antibody. This particular antibody was raised in a rabbit. Hence, it’s only logical that we use an anti-rabbit secondary antibody! Secondary antibodies such as these are made by injecting a different species of animal, such as a goat, with rabbit antibodies and an adjuvant that stimulates an immune response. The resultant goat anti-rabbit antibodies may then be conjugated with an enzyme such as alkaline phosphatase. In the final step of our Western, a substrate for said conjugated enzyme will be added, which creates a colored precipitate on the nitrocellulose membrane. The amount of precipitate is proportional to the amount of secondary antibody bound to the blot, which in turn correlates with the amount of primary antibody and ultimately the amount of Ku80 adsorbed to the blot. So what is the utility of a secondary antibody? First, it amplifies the assay signal, especially as several secondary antibodies can bind to one primary. Second, it simplifies carrying out multiple Westerns. It is much less labor and/or cost-intensive to have one enzyme-conjugated secondary that can be used with many primaries, rather than to conjugate each primary directly. We will return to the topic of antibodies and protein assays during Module 3.
Today you get to experience grad student life, juggling multiple assays with staggered incubation times. Be sure not to forget about your digesting DNA while completing your Western, or about your developing Western while gel purifying your DNA!
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).
- See the D3 FNT and/or talk to your instructors for more details!
- Note that enzyme stock concentrations can be found on the NEB product page for that enzyme.
- 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.
- 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 cells, 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.)
- 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!
- 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).
- Shake your container for 5 min at 80 rpm, using the room temperature shaker in the fume hood.
- Repeat for a total of 3 washes.
- 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.
- Shake at 80 rpm for about 45 minutes.
- Wash the blot as before, for 3 washes.
- 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.
- After removing the last TBS-T wash, add about half the development solution to your blot; save the rest for now.
- 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.
- Add your remaining development solution if progress seems slow.
- 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.
- 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.
- 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 blots on the wiki, depending on whether they appear ready by the end of lab.
Part 3: Gel purify digested plasmid
- 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.
- 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.
- 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.
- 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.
||Sample (27 μL)
||Sample (27 μL)
|| DNA ladder (load 10 μL)
|| Group 3
|| Group 1
|| DNA ladder (load 10 μL)
|| Group 2
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.
- 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?
- Add 3 volumes of QG for every 1 volume of agarose.
- The maximum advised volume is 550 μL. If you have a greater volume, continue in one tube for now, but first read step 6 to understand how to proceed later. Feel free to ask the teaching faculty for clarification.
- 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.
- Add 1 volume — original gel volume, not current solution volume — of isopropanol to the dissolved sample and pipet well to mix.
- Get one QIAquick column 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.
- 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.
- 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.
- Spin for 1 min at max speed.
- Discard the flow-through once more and replace the spin-column in its collection tube.
- Add nothing to the top but spin for 60 seconds more to dry the membrane.
- This step completely removes remaining ethanol.
- Meanwhile, 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 the tube, so we don't lose track of whose sample is whose later on!
- Place the labeled spin-column in its matching trimmed eppendorf tube and add 30 μL of EB to the center of the membrane.
- Allow the column 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.
- 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
- Your Module 1 microbiota summary revisions will be due by 11 AM the Friday after spring break. We recommend that you read your comments from Jon ASAP, and then take your time sleeping on them, prioritizing which are most important to respond to, and ultimately implementing your revision. Cramming this entire process into two days instead of two weeks is probably not wise.
- Every cell line can be characterized by two important features: cell doubling time and cell recovery time. Doubling time is just what it sounds like, and is on the order of 24 hours for most mammalian cells. Recovery time is the delay before cells start growing; first they need to attach (which usually happens within hours) and get comfortable (which can take additional time). It’s really handy to know these values, if only empirically, to avoid coming in on the weekends to split cells. :)
- Under our incubator/media/plating/etc. conditions, if CHO-K1 cells are seeded at 1.4 million (hereafter M), about 11-12 M cells can be recovered 3 days later. Assuming the doubling time is right around 24 h, is the K1 recovery time on the order of 0 h or 24 h?
- In contrast, xrs6 plated at 1.4 M are recovered 3 days later at about 8.5 M cells. Assume that these cells have a recovery time of 24 h. About how many doublings do they therefore undergo in the remaining 2 days? Therefore, approximately how long is their doubling time?
- At this point, you have a lot of practice crafting figures and captions! One change between Module 1 and Module 2 is that your associated results text will be completely in narrative form, rather than partially in bullet points. Another change is that you will be describing protein and cell assays, in addition to the ubiquitous DNA gel. For next time, prepare a figure and caption depicting your Western blot results, as well as the associated results narrative.
- Remember not to mix your caption and results text together; they are two separate elements.
- Follow the best practices we have taught you, reviewing the WAC talks and the 20.109 scientific writing guidelines as needed.
- We recommend that you read the M2D5 transfection protocol and begin preparing an automated calculator in advance of class. (We will NOT collect this assignment.)
All digest reagents from New England Biolabs (NEB).
Agarose gels: 1% in TAE (see Module 1 for details)
QIAquick gel extraction kit (Qiagen)
Western Day 2
- Previously added by teaching faculty
- Being used today
- Goat anti-rabbit alkaline phosphatase conjugate (Bio-Rad)
- AP conjugate substrate kit (Bio-Rad)
- 50 mM Tris-Cl, to pH 7.5
- 150 mM NaCl
- Tween 20 at a final concentration of 0.1%
Next Day: Cell preparation for DNA repair assays
Previous Day: Choose system conditions and paper discussion