Biology 475

Molecular Biology

Library Screening

Restriction Enzymes

Agarose Gels

National Science Foundation

Western Oregon University

Yellowstone National Park

Now that you have isolated DNA from your hot spring library, you are ready to screen for the presence of insert. We will carry out two distinct methods to screen our clone libraries for the presence of insert: (1) cutting with restriction enzymes; and (2) hybridization with specific probes (today, you will only do the primary blotting).

DNA can be cut at specific sequences by bacterial restriction enzymes. Restriction enzymes recognize and cut different sequences. Based on probability, how many times would enzymes that recognize 6 vs. 4 base pairs cut a 1 million bp chromosome? Why might this be useful? Restriction enzymes work best in conditions defined by their parent organism's habitat. To satisfy these conditions, enzyme-matched buffers must be used.

Your clones should be either vector with no insert, or vector with insert. The vector is about 4000 bp (or 4.0 kb), the insert 900-1500 bp (0.9-1.5 kb), depending on the primer set used. To screen for insert, we will cut each sample DNA with EcoRI. EcoRI cuts the plasmid twice - at sites that flank the TA cloning site in the lacZ marker. What will cutting the plasmid and plasmid/insert with EcoRI yield?

Procedures
General Rule: Enzymes never freeze solid and MUST be kept on ice as much as humanly possible!

Typical screening restriction digests involve the following ingredients:

(1) 10 ul of DNA sample (corresponds to 1 ug)
(2) 1.5 ul 10X buffer (final concentration 1X, 15 ul total reaction)
(3) 0.3 ul EcoRI (corresponds to 3 units)
(4) 3.2 ul water

Label 10 small microfuge tubes to correspond with your samples.

Using different tips for each sample, add 10 ul DNA sample to each digest tube. Add to the bottom of the tubes and be careful not to touch the tops of the tubes.

Because we are cutting multiple samples, we will set up our digests using a 15X cocktail. Mix your cocktail for EcoRI in the following order:

(1) Water: 3.2 X 15 = 48
(2) Buffer: 1.5 X 15 = 22.5
(3) Enzyme: 0.3 X 15 = 4.5

Mix the cocktail and, using the same pipette tip, add 5 ul to EACH restriction digest tube - specifically to the upper side. Tap to pull down the cocktail.

Place the tubes in a floating rack and incubate at 37 degrees C for 45-90 minutes.

While You Wait - Electrophoresis

Electrophoresis uses an electric current to separate molecules by size and conformation. The principle of gel electrophoresis can be used in combination with many different kinds of gels. Agarose begins as a powder, is mixed with a liquid buffer, boiled, and then allowed to harden in a tray with a comb. The percent agarose will determine how quickly DNA can travel through it. Gels we pour are 1% agarose (i.e. 1 gram of solid agarose per 100 ml liquid buffer). We will be staining our gels with ethidium bromide, a mutagenic compound that, in combination with UV light, allows us to visualize the DNA following the run. Some additional reagents needed for gel electrophoresis include loading/tracking dyes and standard markers (lambda phage cut with HindIII). Dyes allow us to visually track the progress of the gel and assist with loading by weighing down the DNA. We will use a pre-cut marker standard made by cutting DNA with a known sequence into fragments of known sizes. Following staining and documentation of the gels, they must be disposed of as hazardous waste.

Procedures
Remember to run your PCR Product and Genomic DNA as well!

Prepare 50 ml of 1% agarose per gel box in TAE buffer. After boiling the solution (heat on high and watch it like a hawk), you will have to cool it to 50 degrees C or it will warp the gel trays.

Carefully tape the ends of a gel tray and place TWO 12-lane combs - one in the upper holder and one in the middle. One set will be used for your restriction digests (10 lanes plus marker), the other for your PCR product and genomic DNA (11 lanes plus marker).

Pour the cooled agarose into the tray. It will take 10-15 minutes to harden.

Place the hardened gel/tray into the gel box (there is only one easy way that the tray fits into the box). Check everything carefully to make sure the gel is oriented such that, when the power is turned on, the DNA will run to the positive end.

Cover the gel completely with 1X running TAE buffer. CAREFULLY pull the comb out of the gel. If there are "bits" or bubbles in the wells, clean them out by carefully pipetting buffer into them to "blow out" the pieces.

Mix your samples with loading dye by adding 2 ul dye to the top walls of each tube and tapping down.

I will do a quick demonstration load of the standard markers (10 ul - very expensive!).

Load your samples by setting the pipetteman to 15 ul (even though there is now 18)

Connect the cover with leads to the box and power source, respectively, and turn on the gel. Run 45 minutes at 125 V for this preparation.

Proceed to slot-blotting and macroarray analysis while you wait!

After consulting with your classmates, mark your gel in some distinguishing way (e.g. notched upper right corner) and give me your gel. I will stain gels together in the fume hood.

From this point on, you MUST wear gloves and handle the gel very carefully! We also have a dedicated ethidium spatula that works well.

Following staining, remove the gel to the UV trans-illuminator and place directly on the UV lightbox. If you spill anything, clean it up twice!!!

Open the Fotodyne software (on the desktop). Pull down the File menu option to acquire a new image. To see anything, turn on the camera, the lights, and focus. Turn off the lights and turn on the trans-illuminator/UV. You will likely have to increase the exposure on the gel image to 100. Save the image to an appropriate folder, save a copy to your disk, and print out a copy for your notebook. Clean everything, disposing of gloves, tissues, and spent gels in the hood.