20.109(S13):Bacterial amplification of DNA (Day3)

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(Part 1: Agarose gel electrophoresis)
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==For next time==
==For next time==
#BL21(DE3) ''E. coli'' are often used for protein expression. In contrast, XL1-Blue ''E. coli'' are ‘workhorse’ cells useful for plasmid propagation. One gene modification in XL1-Blue is the ''hsdR'' mutation. What are the two other modified genes in XL1-Blue that make them ideal for the task of propagating a desired DNA? Briefly explain why each is important. It may help you to refer to the cell manual: [http://www.chem-agilent.com/pdf/strata/200249.pdf  (pdf download)], but be sure to answer the question in your own words. Note that this question is about propagation, not screening.
<font color=red>need to revise, with apologies</font color>
Digestion plan needs to be ready for D4, not D5! Focus on that.
==Reagent list==
==Reagent list==

Revision as of 15:41, 18 March 2013

20.109(S13): Laboratory Fundamentals of Biological Engineering

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DNA Engineering        Protein Engineering        Cell Engineering              



Assuming all went well, your reaction tubes from last time contain mutagenized DNA that encodes mutant inverse pericam. However, the desired DNA plasmid is likely present at a low concentration, and moreover it is nicked rather than in intact circular form. What we would like to do now is repair and further amplify only the mutagenized product. Thankfully, we have E. coli bacteria to do this for us quite efficiently!

Bacterial transformation
Bacterial transformation

Recall from Module 1 that bacteria can take up foreign DNA in a process called transformation. Today you will transform a cell strain engineered to be good at plasmid DNA amplification, using cells that are already competent. Next time you will transform a cell strain engineered to produce protein on demand, and prepare your own competent cells. [Bit more about that here or save all for D4? Lecture or ask them re: XL1B?] Whether prepared by a company or by you, remember that competent cells are extremely fragile and should be handled gently, i.e., kept cold and not vortexed.

Bacterial transformation is efficient enough for most lab purposes, resulting in as many as 109 transformed cells per microgram of DNA, but even with highly competent cells only 1 DNA molecule in about 10,000 is successfully transformed. The process is somewhat less efficient with a mutagenesis product rather than an intact plasmid. The vector carrying inverse pericam is called pRSET and also encodes a gene that leads to ampicillin-resistance; thus, selection can be performed on ampicillin-containing agar plates. Given the low concentration and nicked structure of your DNA to begin with, you should perform your transformations today with great care.

Before setting up transformations, you will test your mutagenized DNA for the presence and approximate concentration of product via gel electrophoresis. Because the product is several Kbp long, a standard 1% agarose gel will serve us just fine. The long mutant plasmid DNA should be separated from the short digested fragments of parental DNA and thus can be identified. Note that the parental plasmid is originally present at a concentration too low to detect on a gel, and even the amplified mutant may show up as only a faint band. If you do not see any band at the expected size of the mutant plasmid, you might increase the amount of DNA used during the transformation procedure at the end of lab.

Say something explicitly about stats?


Several changes being made for S13; stay tuned

Part 1: Agarose gel electrophoresis

Using a 1% agarose gel prepared by the teaching faculty, you will each run your digested reaction mixture. Most of you will also load either a reference lane containing standards of known molecular weight or a somewhat concentrated sample of wild-type plasmid. Divide up those responsibilities however you see fit. Recall that you should always handle all gels and gel equipment with nitrile gloves.

  1. In an eppendorf tube, combine 10 μL of of your DpnI-digested product with 2.5 μL of loading dye. Save the rest of the digested DNA, keeping it on ice.
    • Remember to flick and quick-spin your sample, or pipet up and down to mix.
    • Recall that loading dye contains xylene cyanol as a tracking dye and glycerol to help the samples sink into the well.
  2. Load the gel in the order shown in the table below, 11 μL per SDM sample and 10 μL per ladder lane.
    • We will use a 14-lane rather than a 10-lane gel, to concentrate the DNA into a smaller area.
    • To load, lower your sample-containing tip below the surface of the buffer and directly over the well. Expel your sample into the well. Do not release the pipet plunger until after you have removed the tip from the gel box.
  3. Once all the samples have been loaded, the teaching faculty will attach the gel box to the power supply and run the gel at 100 V for 45 minutes.
Lane Sample Lane Sample
1 Red group 8 Pink group
2 Orange group 9 DNA Ladder
3 DNA Ladder 10 Parent IPC
4 Parent IPC 11 Purple group
5 Yellow group 12 Platinum group
6 Green group 13 BLANK
7 Blue group 14 BLANK

While the gel runs, we will have today's pre-lab. During the remaining time, you can work on Parts 3 and (optional) 5, label the tubes you will need in Part 4, work on your notebooks, start the FNT assignment, etc. Be sure to pre-chill your 14 mL tubes on ice for at least a few min before adding competent cells to them.

Part 2: Gel analysis

  1. The teaching faculty will photograph and post a digital image of the gel.
  2. In the following analysis, you will need the information for the 1 Kbp ladder you used, which is available at this link.
  3. First, see if you got a band at the expected size of the pRSET plasmid with an inverse pericam insert, or ~ 4 Kbp.
  4. If you did not get a band, you should use 2-3x the usual recommended DNA amount in your mutant transformation.
  5. If you did get a band, estimate the approximate amount of DNA in that lane in ng (by comparing to the ladder standards), then the concentration in ng/μL (based on the sample volume that you loaded). Write this information in your notebook.

Part 3: Prepare tubes for liquid O/N cultures

Please label 2 large glass test tubes with your team color and sample name in duplicate (X#Z-1, X#Z-2). Prepare a stock of LB culture broth with ampicillin (provided at 1000x concentration) and aliquot 2.5 mL of this mixture per tube. As in Module 1, these will be used to set up liquid overnight cultures from your two colonies for next time.

Part 4: Bacterial transformation

You will transform competent cells called XL1-Blue with your X#Z mutagenesis reactions and plate them on ampicillin-containing Petri dishes. Before next time, two candidate colonies will be chosen from each group's plate. The efficiency of this mutagenesis protocol is reported to be ~80%. We will test two candidates per mutation to cover our bases, so to speak.

  1. Get an aliquot of competent cells from one of the teaching faculty. Keep these cells on ice at all times, allowing them to thaw slowly (over a few minutes).
  2. Label three 14 mL polypropylene round-bottom tubes as follows: (-) control, (+) control, X#Z.
    • The negative control will receive no DNA, but otherwise go through all of the following steps.
    • In your case, the positive control is reference mutant DNA. Normally, a control that comes with the mutagenesis kit we used on Day 2 would be used, but comparing to an IPC derivative is more meaningful. Be sure to use a fresh pipet tip when taking from the positive control stock DNA!still some left? have to prep more?
  3. Add 50 μL of competent cells to each tube, followed by 2 μL of the appropriate DNA. Gently swirl (do not vortex) to mix, then incubate on ice for 10 min.
  4. Bring the tubes over to the 42 °C water bath, and immerse them for exactly 45 seconds according to your digital timer.
  5. Immediately return the cells to ice for 2 minutes, and take an aliquot of pre-warmed LB medium.
  6. Add 0.5 mL of warm LB to each sample, then move them to the 37 °C incubator. Ask the teaching faculty to show you how to operate the roller and balance your tubes.
  7. Allow the cells to recover and begin expressing ampicillin resistance for 30 minutes. At the same time, pre-warm and dry three LB+AMP plates by placing them in the 37°C incubator, media side up with the lids ajar.
  8. Plate 250 μL of each transformation mix on LB+AMP plates. After dipping the glass spreader in the ethanol jar, you should pass it through the flame of the alcohol burner just long enough to ignite the ethanol. After letting the ethanol burn off, the spreader may still be very hot, and it is advisable to tap it gently on a portion of the agar plate without cells in order to equilibrate it with the agar (if it sizzles, it's way too hot). Once the plates are ready, wrap them together with one piece of colored tape and incubate them in the 37°C incubator overnight. One of the teaching faculty will remove them from the incubator and set up liquid cultures for you to use next time.

Part 5: Statistics practice (optional or not?)

Lecture during gel, then exercise

For next time

need to revise, with apologies

Digestion plan needs to be ready for D4, not D5! Focus on that.

Reagent list

  • QuikChange II Site-Directed Mutagenesis Kit from Stratagene
    • XL1-Blue supercompetent cells
  • LB (Luria-Bertani broth)
    • 1% Tryptone
    • 0.5% Yeast Extract
    • 1% NaCl
    • autoclaved for sterility
  • Ampicillin: 100 mg/mL, aqueous, sterile-filtered
  • LB+AMP plates
    • LB with 2% agar and 100 μg/ml Ampicillin
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