Experimental Overview

For this lab we will purify plasmid DNA from cultures of last week's transformed bacteria. Several colonies were picked from the pBS/yeast ligation plates and grown in LB/Amp liquid media. These colonies contain either the plasmid vector alone, or the recombinant subclone consisting of both the plasmid vector and yeast insert DNA. These plasmid DNAs will be purified away from the bacterial chromosomal DNA, proteins, cell membranes, carbohydrates and other cell components. The DNA will be digested with Bam HI and Eco RI enzymes and analyzed via agarose gel electrophoresis. The uncut DNA samples will also be submitted for DNA sequence analysis.

To manipulate cloned genes, it is important to be able to isolate and analyze plasmid DNA. There are many different techniques available for doing this which result in different yields and purity. Often, it is necessary to screen large numbers of bacterial colonies to determine which ones have the correct plasmid subclone. So technique rapidity and ease is important. Also, because 'mini-prep' DNA often needs to be analyzed via restriction enzyme digests or DNA sequencing, it is important that the DNA isolated is relatively ‘clean’.

1) Spin down 1.5 mls of bacterial cultures in eppendorf tubes.

2) Carry out resuspension, lysis, and precipiation steps.

3) EtOH precipitate and resuspend plasmid DNAs.

4) Set up restriction enzyme digestions.

5) Pour 0.7% agarose gel.

6) Load and run agarose gel. Take picture.

Bacteria are grown on solid or liquid media. The solid media, i.e. agar plates, is useful for obtaining individual colonies, however these colonies do not contain enough cells to isolate large amounts of plasmid DNA. Alternately, a single bacterial colony can be used to inoculate a test tube filled with liquid medium. The culture is then grown at 37°C using a shaking incubator. This results in a large amplification of bacteria. Typically, the yield of plasmid DNA from a single eppendorf tube (1.5 mls) of culture is sufficient for several restriction enzyme digests.

There are many different methods for performing plasmid DNA 'mini-preps'. In the one described in this protocol, the bacterial cells are pelleted, resuspended in a small volume of buffer, and lysed open with a combination of detergent and alkali to release their contents. The bacterial proteins and chromosomal DNA are precipitated together using a high salt solution. RNA is degraded via an enzyme, RNAse, present in the resuspension buffer. The plasmid DNA is then further purified from proteins and lipids via extraction with the organic solvents phenol and chloroform. The DNA is then precipitated in ethanol (EtOH), resuspended in dH2O, and analyzed via restriction enzyme digestions.

In order to determine whether there are differences between the plasmid DNA present in different colonies, an aliquot of DNA will be digested with Eco RI and Bam HI. The products will separated via agarose gel electrophoresis in order to determine the number and sizes of products. If a successful bi-molecular ligation between plasmid vector and yeast DNA fragment has occurred, the subclone will yield a distinct pattern of bands. Alternately, if only a uni-molecular re-ligation of the plasmid vector occurred, a different pattern will be observed. What are these patterns?

One potential complication to interpreting the data is that if the DNA is not sufficiently purified, the restriction enzymes may not completely digest the DNA, resulting in a partial digestion. This will produce bands at different sizes than expected for a complete digestion. How will you differentiate between the products of complete and partial digestions?

1) Label 4 eppendorf tubes 1-4.

2) Use a P-1000 pipetman to transfer 1.5 mls of each of 4 different bacterial cultures to the labeled eppendorf tubes.

3) Cap eppendorf tubes and place them in the micro-centrifuge with the hinge of the tube facing the outside of the rotor. Spin for 1 minute. You should see a compact brown pellet at the bottom of each tube.

4) Remove the supernatant using a P-1000 pipetman. Discard liquid into sink. Try to remove as much liquid as possible. If necessary use a P-200 pipetman.

5) Use the P-200 pipetman to add 100 µl of Resuspension Buffer (100mM NaCl, 10 mM Tris pH 7.5, 1mM EDTA pH 8.0, 5 ug/ml RNAse A) to each eppendorf tube. Vortex tube for several minutes to completely resuspend cell pellet. There should be no clumps of cells remaining.

6) Use the P-200 pipetman to add 200 µl of Lysis Buffer (0.2 M NaOH, 0.1% SDS) to each eppendorf tube. Invert the tube several times to mix solutions. The solution should clear (lose its silky appearance) and become viscous as the cells are lysed open and DNA is released.

7) Use the P-200 pipetman to add 150 µl of Precipitation Buffer (5 M KOAc pH 5.0). Invert tube several times. You should see white clumps of protein and chromosomal DNA precipitate out of solution.

8) Place samples in ice for 10 minutes.

9) Spin down samples in microcentrifuge at top speed for 5 minutes. While sample is spinning, label 4 new eppendorf tubes. You should see a large white pellet along the outside of the eppendorf tubes.

10) Use a P-1000 pipetman to transfer 400 µl of supernatant into a new labeled eppendorf tube.

11) Add an equal volume (~400 µl) of buffer-equilibrated phenol (the lower organic phase in the phenol container) to your DNA samples using a pasteur pipette.

You should see two separate solution phases in your eppendorf tube. The denser organic phase is on the bottom.

Be Careful!! Phenol is highly caustic and so must be handled carefully. Be sure to wear gloves and wipe up any drops that are spilled. Wash any phenol off your hands immediately.

12) Cap tube and vortex thoroughly to mix aqueous and organic phases. You should no longer see two distinct phases, but rather, there will be an emulsion formed between the phenol and DNA solution.

13) Place tubes in the microcentrifuge and spin down for 3 minutes at top speed. When you remove the tubes you should again see two distinct phases. The aqueous phase containing the DNA is on the top.

14) Carefully remove the upper phase using a P-200 pipetman. Try to avoid taking any of the lower phase. Transfer the upper phase into new labeled eppendorf tubes.

15) Add an equal volume of chloroform to the aqueous solution. Repeat extraction procedure and transfer aqueous phase to new labeled eppendorf tubes. At this point be sure not to transfer any of the organic phase. It is better to leave some of the aqueous phase behind.

16) Add 1.0 ml of 95% EtOH. Mix by inversion. Spin tubes in microcentrifuge at top speed for 1 minute.

17) Remove EtOH via P-1000 pipetman. You should see a significant white pellet on the side of the tube. Gently rinse this pellet with 1 ml of 70% EtOH. Be careful as the pellet may come loose with the 70% EtOH wash. If the pellet does dislodge, briefly spin the sample down in the micro-centrifuge for 1 minute. Remove the 70% EtOH using a pasteur pipette. Remove any residual liquid using a P-200 pipetman.

18) Dry the pellet in the Speed-Vac for 2 minutes or until tube is dry. Be sure that the rotor is spinning before you pull down a vacuum.

19) Resuspend pellet in 20 µl of dH2O. Vortex pellet if necessary.

20) Transfer 5µl of each DNA solution into 4 new eppendorf tubes. Set up restriction enzyme digests with Eco RI and Bam HI as follows:

DNA 5 µl

10 X enzyme buffer 2 µl

Eco RI 1 µl

Total Volume 20 µl

Flick tubes to mix and spin down in micro-centrifuge to collect liquid at bottom of eppendorf.

21) Incubate each tube at 37°C for 1 hour.

22) Set up and pour a 0.7% agarose gel in TAE to run your samples out.

23) Add 2 µl of 10X gel loading buffer to each sample. Load your DNA samples and Molecular Weight markers into adjacent lanes on the agarose gel using a P-20 pipetman.

24) Run gel at 130 volts for 20 minutes.

25) Remove gel and examine under UV trans-illuminator. Take a picture of the gel.