20.109(S09):Bacterial amplification of DNA (Day3)

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20.109(S09): Laboratory Fundamentals of Biological Engineering

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Bacterial transformation

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!

Bacteria can take up foreign DNA in a process called transformation, during which a single plasmid enters a bacterium and, once inside, replicates and expresses the genes it encodes. Most bacteria do not exist in a transformation-ready state, but can be made permeable to foreign DNA by chemical treatment or other means. Cells that are capable of transformation are referred to as competent. 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. Thus we need a way to identify transformed cells, which is usually accomplished with antiobiotics. For example, the plasmid carrying inverse pericam (calle pRSET) also carries a gene that leads to ampicillin-resistance. Consequently, a transformed bacterium will grow on ampicillin-containing agar medium, while untransformed cells will die before they can form a colony (see figure above right). 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 using gel electrophoresis. This technique separates large molecules by size using an applied electrical field and appropriate sieving matrix. DNA fragments are typically separated in gels composed of agarose, a seaweed-derived polymer (see figure, left). To prepare these gels, molten agarose is poured into a horizontal casting tray containing a comb. Once the agarose has solidified, the comb is removed, leaving wells into which the DNA sample can be loaded. The loaded DNA samples are then pulled through the matrix when a current is applied across it. Specifically, DNA molecules are negatively charged due to their phosphate backbones, and thus travel toward the positive charge at the far end of the gel (see figure, right).

Scanning EM image of agarose polymer

Although all DNA molecules travel in the same direction during gel electrophoresis, they do so at different rates: larger molecules get entwined in the matrix and retarded, while smaller molecules wind through the matrix more quickly and thus travel further from the well. Ultimately, fragments of similar length accumulate into “bands” in the gel. Bands of DNA are usually visualized by adding the fluorescent dye ethidium bromide to agarose gels. This dye intercalates between the bases of DNA, allowing DNA fragments to be located in the gel under UV light and photographed. The intensity of the band reflects the concentration of molecules that size, although there are upper and lower limits to the sensitivity of dyes. Because of its interaction with DNA, ethidium bromide is a powerful mutagen and will interact with the DNA in your body just as it does with any DNA on a gel. You should always handle all gels and gel equipment with nitrile gloves. Agarose gels with ethidium bromide must be disposed of as hazardous waste.

Today you will run your DpnI-digested and undigested mutagenesis reaction mixtures through an agarose gel. The long mutant plasmid DNA should be separated from the short digested fragments of parental DNA and thus can be identified. However, the bands may be very faint. If you do not see a 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. In between electrophoresis and transformation, we will have a visit from the Writing Across the Curriculum faculty.


Part 1: Agarose gel electrophoresis

Using a 1% agarose gel prepared by the teaching faculty, you will run two samples and a reference lane containing standards of known molecular weight (also called a DNA ladder).

  1. Set aside 10 μL of of your DpnI-digested product in eppendorf tubes. Save the rest of the digested DNA, keeping it on ice.
  2. Retrieve the aliquot of undigested sample that you prepared last time, and add 2 μL of loading dye to both 10 μL samples.
    • Loading dye contains xylene cyanol as a tracking dye to follow the progress of the electrophoresis (so you don’t run the smallest fragments off the end of your gel!) as well as glycerol to help the samples sink into the well.
  3. Flick the eppendorf tubes to mix the contents, then quick spin them in the microfuge to bring the contents of the tubes to the bottom.
  4. Load the gel in the order shown in the table below, where U is undigested and D is the digested sample. Three groups will share each row of wells.
    • To load your samples (see figure below), draw 10 μL into the tip of your P20. Lower the tip below the surface of the buffer and directly over the well. You risk puncturing the bottom of the well if you lower the tip too far into the well itself (puncturing well = bad!). Expel your sample into the well. Do not release the pipet plunger until after you have removed the tip from the gel box (or you'll draw your sample back into the tip!).
  5. 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.
  6. While the gels run, you will hear from the Writing Across the Curriculum teaching faculty. If you are done early, you can skip ahead to Part 4 of the protocol.
Loading a gel
Lane Sample Lane Sample
1 DNA ladder 6 Group 2, D
2 Group 1, U 7 BLANK
3 Group 1, D 8 Group 3, U
4 BLANK 9 Group 3, D
5 Group 2, U 10 DNA Ladder

Part 2: Gel analysis

  1. Photograph and save a digital image of your gel on a USB key according to the orientation day protocol. (Feel free to ask the teaching faculty for help.)
  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, consider preparing an extra transformation for your mutant sample with 2-5x the usual recommended DNA amount.
  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: Bacterial transformation

You will transform competent cells called XL1-Blue with your X#Z mutagenesis reactions and plate them on ampicillin-containing Petri dishes. Tomorrow, 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 the following steps.
    • The positive control is a mutagenized plasmid made from materials in the mutagenesis kit we used on Day 2. Be sure to use a fresh pipet tip when taking from the positive control stock DNA!
    • You can prepare 1 extra tube for the X#Z transformation if you wish to try both a standard and an elevated DNA concentration.
  3. Add 50 μL of competent cells to each tube, followed by 1 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 (or four, if needed) 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 4: Prepare tubes for liquid O/N cultures

You will make your teaching faculty very happy if you contribute to their preparatory work. Please label 2 large glass test tubes with your team color and sample name (X#Z-1, X#Z-2). Mix 10 mL LB with 10 μL of ampicillin. Aliquot 2.5 mL of LB+Amp per tube. These will be used to set up liquid overnight cultures from your two colonies for next time.

For next time

IPC plasmid map (Q1)
Mutant plasmid map (Q1)

1. The pRSET plasmid with inverse pericam insert, or pRSET-IPC, is 4169 basepairs long. According to the cutters list that you used on Day 1, restriction site PvuI occurs at ~1685 bp, and again at ~2730 bp into pRSET-IPC. Thus, digesting this parental plasmid with the PvuI enzyme should result in two linear fragments of DNA, with about 1050 and 3120 bp sizes.

A silent mutation can be introduced that results in a new PvuI site at the 341st-342nd residues of inverse pericam (ATT → ATC and TAC → GAC) , or approximately the 1020th basepair of IPC. When IPC is inserted into pRSET, its starting point is ~200bp into the pRSET plasmid. Thus, if the mutated pRSET-IPC plasmid is digested with PvuI, three linear fragments of DNA are the result: 470, 1050, and 2660 bp. To understand these calculations, see also the plasmid maps at right.

For this assignment, you should plan restriction enzyme digests that allow you to distinguish parental and mutant pRSET-IPC for S101L and for your X#Z mutation. You are probably best off doing a single enzyme digest for this particular experiment. However, in other kinds of experiments (notably cloning) using two enzymes per digest can give more information.

Use the NEB site to determine the appropriate buffer and temperature for your reactions.

Update: In addition to giving the reaction conditions for each digest, you should explicitly show how the digest distinguishes between parent and mutant. In other words, what are the expected band sizes for each upon digestion?

2. Consider a mutant pRSET-IPC with a newly introduced NotI restriction site that did not occur at all on the parental plasmid (zero cutter). After digestion, you see banding patterns that indicate an uncut plasmid (supercoiled and circular) rather than a linearized piece of DNA for both the parental and the mutant plasmid. Your lab partner tells you that the mutagenesis reaction must not have worked, and that the so-called mutant couldn't possibly contain a NotI site. What’s an equally valid interpretation of this data?

3. Write an early draft of the Materials and Methods section of your lab report, describing the work you have done up through today.

4. The first participation self-evaluation is due next time.

Reagent list

  • 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