Bryan Hernandez/20.109/Lab notebook/Module 1/Day 3

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20.109: Laboratory Fundamentals of Biological Engineering

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Ligations and Transformations

--Bryanh 17:17, 16 February 2007 (EST)


ligate linearized M13K07 backbone with the epitope insert. transform ligated plasmid into bacteria and screen with Kan.


When you have a break from the work described below, be sure to examine the plaques you plated last time. Record the number of plaques on each plate, their appearance and any observations or conclusions you can draw.

Part 0

counting PFUs:

Phage infection

Phage dilution Phage Strain PFUs Counted
none E4 lawn
none K07 lawn
10^-4 E4 too many to count
10^-4 K07 too many to count
10^-6 E4 2000
10^-6 K07 no PFUs (this is probably an error in our dilution step)

Part 1: Ligation reactions

For your ligation, you will mix the M13K07 backbone you prepared with the annealed oligonucleotides. As control reactions you will also prepare a “bkb, no ligase” reaction to control for any errant uncut plasmid that might have wandered into your solutions. Additionally you will prepare “bkb only, plus ligase” reaction to assess the frequency of backbone religation.

The contents of each ligation will be

no ligase
bkb only,
plus ligase
bkb + insert,
plus ligase
M13K07 bkb 4 μl 4 μl 4 μl
insert none none 5 μl
10X Ligation Buffer^ 1.0 μl 1.0 μl 1.0 μl
T4 DNA Ligase none 0.5 μl 0.5 μl
Water To 10 μl not including volume of enzyme

^New England Biolabs sells 10X Ligation buffer to use with their ligase. It contains ATP so must be kept on ice.

  1. Assemble the reactions in eppendorf tubes but not in the order listed. Please ask if you are unsure what order to assemble the components.
  2. When the ligation mixes are complete, flick the tubes to mix the contents, quick spin them in the microfuge to bring down any droplets, then incubate the reactions at room temperature for at least 10 minutes.
  3. Recall that the ligation of your insert into the M13K07 backbone will destroy the PstI or BamHI restriction sites originally used to linearize the genome. Consequently, you can enrich for the proper ligation products by digesting the ligation reactions with the enzyme used to open up the backbone. These are called "kill cuts" since in theory they destroy unwanted ligation products. So while your samples are ligating, you should prepare a "killcut" cocktail according to the table below. By making one mixture that contains water, buffer and enzyme, you can add the same mixture to all reactions, minimizing effects of pipet-error and possible accidental errors, like leaving one component out of one reaction that you remember in all the others.You will prepare enough cocktail to perform 4 killcut reactions, even though you have only three. This will assure you have enough volume for the three reactions.
volume in each reaction x4
= volume in cocktail
water 8 μl 32 μl
10X NEB buffer 2 μl 8 μl
Enzyme 0.25 μl PstI or BamHI 2 μl

4. Add 10 ul of the killcut cocktail to each of the ligation reactions you've prepared, pipetting up and down to mix.
5. Incubate at 37° for 15 minutes.

Part 2: Precipitation of DNA

In this step, salts and buffers are removed from the reactions. DNA is precipitated with salt and ethanol. Yeast tRNA is added to the precipitation as “carrier,” allowing you to better visualize the DNA pellets. The salts are washed from the pellets with 70% ethanol. The tRNA is not removed. Rather it enters the bacteria with the ligation DNA, but is then rapidly degraded.

  1. Add 20 μl 3M sodium acetate to each tube.
  2. Add 5 μl tRNA to each tube.
  3. Add 200 μl cold 100% ethanol to each tube and vortex.
  4. Spin in a room temperature microfuge 15 minutes. Be sure to orient your tubes in the microfuge so you know where your pellets should be and balance your tubes with those of another group or with a water-filled eppendorf tube.
  5. When the spin is done, locate the pellets in each of the eppendorf tubes. They may appear as solid white dots at the bottom corner of the tube or they may appear to be a diffuse white smear along the wall of the tube. Both are OK. Carefully remove the ethanol from the pellets with your P1000, taking care not to disturb the pellet. You do not have to remove every last drop.
  6. Wash the pellets with 500 μl cold 70% ethanol. This is done by dribbling the 70% ethanol along the wall of the eppendorf tube that is opposite your pellet and then removing the 70% ethanol with the same pipet tip. Again you should not disturb the pellet and you do not have to remove every drop of liquid in the tube. If the pellet seems to float away from the wall of the tube, you can re-spin the tubes for 2 minutes with the liquid to adhere the pellets to the wall again.
  7. Once you have washed all your pellets, give the tubes a quick spin in the microfuge to bring down any droplets of ethanol that cling to the sides of the tube then remove any remaining liquid from the tubes using your P200. Allow your tubes to dry in the hood for 10 minutes. All the ethanol must be removed or evaporated.
  8. Resuspend the pellets in 15 μl sterile water. This is done by adding water to the tubes and mixing. If the DNA does not readily go into solution, it helps to heat the DNA in the 42°C heat block, then vortex and pipet up and down several times. Bring any droplets down to the bottom of the tubes with a quick spin in the microfuge.

Part 3: Bacterial transformation

You will perform 4 bacterial transformations, one for each of the ligation mixtures as well as one transformation with 5 ng of plasmid DNA to assess transformation frequency.

  1. Prewarm and dry five LB+Kan plates by placing them in the 37°C incubator, media side up with the lids ajar.
  2. Get an aliquot of competent cells from one of the teaching faculty. Keep these cells on ice at all times. There should be at least 200 μl of cells in each tube. Aliquot 50 μl of cells into 4 clean eppendorf tubes.
  3. Add DNA to each tube of cells as shown in the table below.
  4. Flick to mix the contents and leave the tubes on ice for at least 5 minutes.
  5. Heat shock the cells at 42°C for 90 seconds exactly. Use your timer.
  6. Move the samples to a rack on your bench then use your P1000 to add 0.5 ml of LB media to each eppendorf tube. Invert each tube to mix.
  7. Incubate the tubes in the 37°C incubator for at least 30 minutes. This gives the kanamycin-resistance gene some time be expressed in the transformed bacterial cells.
  8. While you are waiting, label 4 large test tubes with your team color and numbers 1, 2, 3, 4. Mix 10 ml LB with 10 ul of the kanamycin stock. Aliquot 2.5 ml/tube. This will help the teaching faculty to set up overnight cultures for you for next time.
  9. Plate 200 μl of each transformation mix on LB+Kan plates, plating the bkb+insert transformation twice. Note: After passing the glass spreader through the alcohol burner and 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 (and if it sizzles, it's way too hot). Once the plates are done, wrap them with 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.
Tube Transformation Add
1 positive control plasmid 1 μl (5 ng) of M13K07 DNA
2 bkb, no ligase 5 μl
3 bkb, plus ligase 5 μl
4 bkb+insert, plus ligase 5 μl


For next time

Questions 1 and 2 are theoretical but they should help prepare you to interpret the results you will collect next time.

  1. You have purchased some supercompetent bacteria that are provided at a transformation efficiency of 109 colony forming units/ug of DNA. You transform the cells with 1 ng of plasmid DNA and plate 1/1000th of the cells. How many colonies do you expect? Next you transform another aliquot of cells, also at 109 colony forming units/ug of DNA, with 2 μl of plasmid DNA. You spread 1/100th of the cells and find 50 colonies growing on the plate after 24 hours at 37°C. What is the concentration of plasmid?
  2. To illustrate your understanding and the importance of the controls you performed today, please write a one-sentence interpretation for each of the following transformation outcomes.
    • Outcome 1: no colonies on any plate.
    • Outcome 2: thousands of colonies on all the plates.
    • There may be more than one valid interpretation for some of the data (only one answer for each is required for the assignment).
  3. Next time you will prepare DNA from four transformants and begin to characterize the plasmids in these bacteria. Using the plasmid map you drew last time, plan at least two restriction digests that will confirm the presence of the PCR insert. It will help to read the introduction for the next lab before you complete this part of the assignment. Be sure to predict the size of the fragments you expect when the plasmid does and doesn’t have the PCR insert. Also include reaction conditions such as buffer and temperature. Use the NEB website for details on various enzymes and reaction conditions (
Diagnostic digest 1 plasmid with insert plasmid no insert
Enzyme(s) used BbsI, HindIII BbsI, HindIII
Buffer used neb2 neb2
Temperature 37 37
Predicted fragments 6kb, 2.7kb 8.7kb
Diagnostic digest 2
Enzyme(s) used PstI, HindIII PstI, HindIII
Buffer used neb2 neb2
Temperature 37 37
Predicted fragments 8.7kb 6kb, 2.7kb

4. Based on the results of your plaque assay, what is the titer of each stock solution of phage? Please show your work. If the plaques appeared different, please consider how the phage genomes differ (M13K07 is a "helper phage" while E4 is identical the the M13 genome except four glutamic acids are presented on the N-terminus of the p8 protein) and suggest how these differences might account for the differences in plaque morphology.

5. Read the article by Chan, Kosuri and Endy. "Refactoring bacteriophage T7" Nature/EMBO Molecular Systems Biology 13 September 2005 doi:10.1038/msb4100025 and News & Views. Come prepared to discuss this paper during lab next time. To guide your reading and test your understanding, try to answer the following questions:

  • from the Introduction:
    • What is "refactoring" and what makes is T7 an attractive candidate for this approach?
    • What experimental techniques give us "compenent level" understanding, i.e. allow us to attribute particular functions to particular sequences in the genome? How completely can "component level" understanding provide "system level" understanding?
    • How predictive have computational and quantitative models for T7 behavior proven to be? What's important about predicting behavior?
  • from the Results:
    • What design principles were the authors pursuing? How well do these map to our class effort at M13 re-design?
    • Are there T7 components specified in the Registry of Standard Biological Parts? While you're looking around the Registry, please sign-up for an account if you haven't already. You'll need it for work that we do later in this experimental module.
    • Was the entire T7 genome refactored?
    • What techniques were used to verify the refactoring? What techniques were used to evaluate it?
  • from the Discussion:
    • How do the authors' findings extend knowledge of T7 biology?
    • Does T7.1 resolve disagreements between model-based behavior predictions and those that are observed though experimental approaches?
    • Could nature have produced the T7 phage that now exists in the Endy lab in Building 68?
    • What's next for this phage?

Reagents list

  • 50 ng M13K07
  • T4 DNA Ligase Buffer (1X)
    • 50 mM Tris-HCl
    • 10 mM MgCl2
    • 10 mM DTT
    • 1 mM ATP
    • 25 μg/ml BSA
  • LB
    • 1% Tryptone
    • 0.5% Yeast Extract
    • 1% NaCl
  • LB+kan plates
    • LB with 2% agar and 25 μg/ml Kanamycin