Molecular Biology 480/580 LAB MANUAL

Bio 480 Laboratory Manual (8/24/99 edition)

General Procedures

(8/24/99)

1. Apparel:

Shoes are to be worn at all times. Open toed shoes or sandals are not acceptable, since they offer no protection from spills of caustic chemicals. Store coats and backpacks away from your working area. Uncluttered work areas are safer and easier to use.

2. No Eating, Drinking or Smoking in the laboratory at any time.

3. Spills:

If you spill a chemical on yourself, wash immediately with copious amounts of water and notify the instructor. In the event of a spill on the floor or a bench involving hazardous materials (such as strong acid or base or a volatile organic compound) notify the instructor immediately and receive instructions regarding cleanup before attempting to clean it up yourself.

4. Accidents:

If you injure yourself in the laboratory in any way (however minor you may think the injury is), report the injury to the instructor.

5. Broken Glass:

Everybody breaks glass occasionally. If you break something, don't rush to clean it up with your hands. Find a broom or dust brush, sweep up the glass and place it in the appropriate broken glassware container. Don't put it in the regular trash can.

6. Waste chemicals:

Do not put any waste chemicals down the sink. The instructor will provide appropriately labeled waste containers for proper disposal. If you are in doubt about whether or not something should go down the sink, ask the instructor.

7. Pipetting:

Mouth pipetting is forbidden. Use pipettors at all times.

8. Volatile Chemicals:

Use the fume hood when working with volatile chemicals. Check to make sure the hood is working before opening the volatile chemical.

9. Dirty Labware:

Disposables: Make sure an item is really disposable before disposing of it. Place dirty labware in a dish pan containing a detergent solution until you are ready to wash them. Wash labware with soap and water, rinse well with hot water, and rinse twice with deionized water. Use the deionized water sparingly. Air dry the labware by placing it inverted in a rack or on a paper towel.

Isolation of Anonymous cDNA Clones

A cDNA (copy DNA) is a fragment of double stranded DNA synthesized from a messenger RNA. cDNA libraries are collections of 10s of thousands of different cDNAs from a given tissue and are derived from mRNA extracted from that tissue. In order to study an individual cDNA one has to "grow" them to obtain enough DNA to work with. The cDNAs, however, are incapable of replicating on their own but when they are placed into plasmid vector (small circular DNA molecules), they can be maintained in an E. coli host indefinitely - whenever the host cells are grown, the plasmid grows and can be extracted from the cells for study. The cell line which contains a particular cDNA is called a clone and the library is actually maintained as a collection of clones. All of the different cDNAs in the library were inserted into the same kind of plasmid (see Appendices 2 and 3 for maps of this vector) thus each independent plasmid (maintained in a single cell line) contains a common region (the plasmid vector) and a variable region (the cDNA insert). We are interested in studying the inserts.

At the beginning of the semester the instructors prepared an Arabidopsis cDNA library such that the plasmids would be growing in independent colonies of E. coli on a petri plate. It is possible, however, that what appears to be single colony is actually 2 growing at the same place. This is why you need to prepare a new plate with only one clone streaked out on it.

Procedure

1. Label one LB/AMP (1 blue stripe, 1 green stripe) plate on the bottom with the last 4 digits of your student number, your initials and the date.

2. From one of the master plates choose one colony and mark the bottom of the plate directly underneath the chosen colony. This will prevent others from selecting the same colony as you have.

3. Using sterile technique, touch an inoculating stick to the colony and transfer it to your LB/AMP plate. Use the technique which will result in isolated colonies (see demonstration by instructor). Note: Be sure to take a large white colony (1-2 mm dia) and avoid the smaller pinpoint colonies (satellite colonies) around the large ones.

4. Place the plate in the 37 C incubator for approximately 24 hours.

5. After incubation for 24 hours, place the plates in the refrigerator.

6. Keep in mind you will have to use this plate next week to inoculate a broth culture in preparation for plasmid DNA extraction. This inoculation must be done next week on the DAY BEFORE LABORATORY.

Preparation of liquid culture for plasmid extraction

Each person must do this the day before lab meets the first time.

The two goals of this procedure are for you to 1) inoculate a broth (liquid) medium with an isolated colony of the clone you picked last week so that the cDNA-containing plasmid can be extracted the next day during lab, and 2) streak the same clone on a plate to use as an archive of the cDNA clone.

1. Label an LB/AMP (1 blue stripe, 1 green stripe) agar plate and a 15 mL conical centrifuge tube containing 5 mL LB/AMP liquid medium (both the plates and the tubes are in the refrigerator) with the last 4 digits of you student number and the date. Remember that this number is the "name" of your cDNA clone.

2. If your culture did not grow, choose a plate from the bag labeled "replacement cultures" (refrigerator) and continue with this clone as your own. Use a sterile inoculating stick to pick an isolated colony from that plate, dip the stick in the LB/AMP broth in your tube, and then streak it onto the labeled plate (in the same way you did last week). This way you will have inoculated both the new LB/AMP plate and the tube of LB/AMP broth with the same colony.

4. Incubate your plate, upside down in the same incubator you used last week. Incubate the tube in the shaking incubator in Room 333. It is in the middle of the room. Lift the lid and the shaker will stop. Place the tube in the rack marked "Bio 480". Make sure the lid of your tube is screwed down securely, or it will shake off during incubation. Close the lid and the shaker will start again.

When you come to lab the next day, you will begin the plasmid miniprep procedure using the broth culture that you inoculated.

Plasmid Miniprep

This procedure is used to extract plasmid DNA from bacterial cell suspensions and is based on the alkaline lysis procedure developed by Birnboim and Doly (Nucleic Acids Research 7:1513, 1979). The procedure takes advantage of the fact that plasmids are relatively small supercoiled DNA molecules and bacterial chromosomal DNA is much larger and less supercoiled. This difference in topology allows for selective precipitation of the chromosomal DNA and cellular proteins from plasmids and RNA molecules. The cells are lysed under alkaline conditions, which denatures both nucleic acids and proteins, and when the solution is neutralized by the addition of Potassium Acetate, chromosomal DNA and proteins precipitate because it is impossible for them to renature correctly (they are so large). Plasmids renature correctly and stay in solution, effectively separating them from chromosomal DNA and proteins. Cool, no?

Procedure

Note: The procedure below is used to make duplicate minipreps. This provides balanced tubes for the centrifuge as well as twice as much product when you are finished.

1. Shake the culture tube to resuspend the cells.

2. Label two 1.5 mL tubes and pipet 1000 uL of the cell suspension into each tube.

3. Close the caps and place the tubes in a centrifuge (remember to balance the centrifuge by putting the tubes opposite one another) and spin at maximum speed for 20 s.

This part of the procedure collects the bacterial cells which are suspended in the liquid medium into a cell pellet at the bottom of the tube.

4. Withdraw and discard the supernatant using a pipettor, being careful not to disturb the cell pellet. Discard the supernatant in a waste container.

5. Add 100 uL of Buffer 1 (50 mM Tris-HCl, 10 mM EDTA, 100 ug/mL RNase A, pH 8.0 ) to each tube and resuspend the cells by vortexing. It's very important that the cell suspension is homogenous and no clumps are visible.

The cells are now resuspended in a buffered solution with RNase. When the cells are lysed in the next step, the RNase will catalyze hydrolysis of all RNA molecules into nucleotides, but the DNA will not be affected.

6. Add 200 uL of Buffer 2 (1% SDS, 0.2 M NaOH ) to each tube. Close the caps and mix the solutions by rapidly inverting them a few times. DO NOT VORTEX since the chromosomal DNA released from the broken cells could be sheared into small fragments and contaminate your plasmid prep.

SDS is an acronym for Sodium Dodecyl Sulfate. It is an ionic detergent which disrupts cell membranes and destabilizes all hydrophobic interactions holding various macromolecules in their native conformation. The high pH of the 0.2 M NaOH also denatures macromolecules by changing the condition of ionizable groups (ionizing certain groups and deionizing others). The clearing you see is because the cells are lysing. The viscosity of the solution is increased by the increase in concentration of macromolecules in solution (a result of the cell lysis).

7. Let tubes stand on ice for 5 minutes (it's OK if they go longer).

8. Add 150 uL of ice-cold Buffer 3 (3.0 M Potassium Acetate, pH 5.5 ) to each tube. Close the caps and mix the solutions by rapidly inverting them a few times. A white precipitate will form.

This is really the key step in the alkaline lysis procedure. The low pH of the Potassium acetate solution neutralizes the NaOH and when the pH returns to near-neutrality then the macromolules renature. The proteins and large DNA molecules do not renature correctly however. They form hydrophobic, ionic and hydrogen bonds with each other nonspecifically because the correct conformation of the molecule was not maintained during denaturation. The plasmid DNA molecules, however, never really fully denatured because they are small circular molecules which are supercoiled. Even though the hydrogen bonds between base pairs were broken by the high pH, they reform correctly when the pH is lowered. The large DNA molecules (chromosomal DNA) and proteins form precipitates because they bind to each other in a large aggregate but the plasmids don't precipitate because they renature correctly and don't become part of the large multi-molecule aggregates. Thus plasmid DNA remains in solution while proteins and other DNA molecules precipitate.

9. Let tubes stand on ice for 5 minutes (it's OK if they go longer).

10. Place the tubes in a centrifuge (balanced) and spin at maximum speed for 5 minutes. Team up with some other folks on this spin. The precipitate will pellet along the side of the tube.

11. Transfer the supernatants into clean 1.5 mL tubes, being careful not to pick up any of the precipitate. Discard the tubes with the precipitate and KEEP the tubes with the supernatant.

12. Before you do this step, make sure there is a centrifuge available. To each tube of supernatant add an equal volume (about 400 uL) of isopropanol to precipitate the nucleic acids. Close the caps and mix vigorously. Let the tubes stand at room temperature for 2 minutes, place them, with their hinges pointing outward from the center, in a centrifuge (balanced) and spin at maximum speed for 5 minutes. This step pellets the nucleic acids but if you leave it around too long, proteins remaining in solution will begin to precipitate as well.

This concentration of isopropanol will cause the DNA to become insoluble. Since DNA molecules are ionic (uniform negative charge due to the phosphates) they are highly soluble in water but not soluble in organic solvents. Isopropanol and ethanol are commonly used to precipitate DNA from aqueous solution.

13. The reason the you oriented the tubes with the hinges outward is because the plasmid DNA pellet may be difficult to see. It will be at the bottom and along the hinge side of the tube. Carefully remove and discard the supernatant. The pellet is usually visible at this point. If not, do not despair. It may be too small to see but there is probably enough DNA there. Remember that we are dealing with small molecules that we can't see unless there is a whole lot of them together!

14. Add 200 uL of absolute ethanol to each tube and mix by inversion several times.

15. Spin the tubes at maximum speed in a centrifuge for 2-3 minutes (hinges out).

This "ethanol wash" will speed up the process because the next step is to evaporate the alcohol used in the precipitation step. Absolute Ethanol evaporates much faster than a solution of 50% isopropanol.

16. Carefully remove and discard the supernatant. Try to get as much out as possible without dislodging the pellet of plasmid DNA.

17. Place the tubes in the fume hood with the caps open for 15-20 minutes to dry off the last traces of ethanol.

18. When the ethanol is gone (you can check this by smelling the tube) add 20 uL of TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) to dissolve the pellet. Pipet the 20 uL in and out, up the side of the tube to ensure that all of the plasmid DNA comes into contact with the TE buffer.

TE buffer is commonly used to redissolve DNA because it contains EDTA. The EDTA will chelate magnesium ions which are a cofactor for most nucleases (enzymes which degrade nucleic acids). If your DNA prep becomes contaminated with a nuclease (like the ones produced by the cells in your skin) then the nuclease will be inactivated by the fact that the magnesium cofactor is unavailable in the solution (because it is chelated by the EDTA).

19. Pool the two 20 uL solutions into one labeled tube. You will use some of the DNA in for the restriction digest. The rest will be stored in the freezer.

Digestion of Plasmid DNA with Restriction Endonucleases

Restriction endonucleases such as EcoRI recognize specific palindromic sequences and cleave a phosphodiester bond on each strand at that sequence. After digestion with a restriction endonuclease the resulting DNA fragments can be separated by agarose gel electrophoresis and their size can be estimated. A restriction map is generated by using the fragment size data to determine the location of the specific endonuclease recognition sequences on the plasmid.

Each restriction enzyme requires specific reaction conditions for optimum activity. One of the most important reaction conditions which varies between different restriction enzymes is the salt (usually NaCl) concentration. Enzyme buffers are specifically formulated to provide the salt concentration for optimal enzyme activity. It is important, therefore, that the correct buffer solution is used for a particular restriction enzyme.

Listed below is a general procedure for conducting restriction digests. Keep in mind, however, that one should always consult the manufacturers recommendations of optimal conditions for each restriction enzyme (See Resources for more information on restriction enzymes).

General Procedure

1. For each digest combine the following solutions in a microcentrifuge tube.

Item Amount

deionized water 6 uL

10x reaction buffer 1 uL

Plasmid miniprep DNA 3 uL

Total 10 uL

2. Remove the restriction endonuclease (specific enzymes to use will be listed on the black board) from the freezer and place on ice. It is important to keep the enzyme cold. Add 0.5 uL of enzyme solution to the mixture you made in step 1. If digesting with two restriction enzymes (double digest), add 0.5 uL of the second enzyme.

Item	Amount
deionized water	6 uL
10x reaction buffer	1 uL
Plasmid miniprep DNA	3 uL
Total	10 uL

3. Tap the tube to mix the contents and then pulse down the tube in a microcentrifuge.

4. Incubate at 37 deg C for 2 h to overnight. After incubation place tubes in the freezer.

The First Digest of the Semester

Your first restriction digest will be a double digest to determine the size of your insert (the cDNA). A double digest is one where two restriction enzymes are used to digest DNA in a single reaction. In this case you will be using EcoR I and BamH I. There is only one site in the plasmid vector for each of these enzymes and they are located on either side of your insert DNA.

Digesting with both will cut the insert from the vector. Next week you will determine the size of the insert by seperating the digested DNA on an agarose gel. The insert may also contain a site for one or both of these enzymes and if so, the insert will be cut into multiple pieces. By adding up the sizes of each fragment you can still determine the size of the insert.

Today's Digest

1. In a microcentrifuge tube place 6 uL of deionized water, 1 uL of 10x EcoR I reaction buffer , 3 uL of your plasmid miniprep, 0.5 uL of EcoR I enzyme and 0.5 uL of BamH I enzyme. The enzymes and buffer are stored in the freezer. The tube with the buffer has a black top. The tube with EcoR I enzyme has an "E" with a black dot in top and the tube with the BamH I enzyme has a "B" and a brown dot on the top.

2. Tap the tube to mix the contents and then pulse down the tube in a microcentrifuge.

3. Incubate in the 37 degree incubator overnight. Be sure to stop in the next day to put your digest in the freezer. If you let it go for more than a day or so, the DNA will begin to degrade.

Agarose Gel Electrophoresis of DNA

Introduction

During this laboratory you will use agarose gel electrophoresis to separate DNA fragments which have been generated by digestion of your plasmid DNA with restriction endonucleases. You will then use a molecular size marker (a 1 kb ladder) to generate a standard curve of mobility vs. log bp and use the standard curve to estimate the size of the fragments separated on the gel (See Resources for a link to NEB's 1 kb ladder page including a picture and the sizes of the marker fragments)

A. Casting the gel:

1. Make 25 ml of a 1% (w/v) solution of agarose in TAE buffer.

2. Weigh the container with the mixture and record the mass.

3. Heat the mixture to boiling using the microwave oven. Examine the flask and continue boiling if any agarose is undissolved.

4. Weigh the container with the mixture again and add deionized water to compensate for loss of mass during boiling.

5. Allow the agarose to cool to by placing it in a water bath set to 55 deg C.

6. Make sure the wedges are in place firmly against the ends of the casting tray. Pour all of the agarose solution into the casting tray, being careful not to overflow the tray. Add the comb and leave the gel to cool and solidify.

B. Preparing the samples

1. While the gel is cooling, prepare the DNA samples by adding 1 uL of tracking dye to 5 uL of each restriction digest. The tracking dye is bromphenol blue in a 50% glycerol solution. Adding tracking dye to the sample will increase its density so it falls into the well of the gel and provides a visible marker to monitor the progress of electrophoresis. Also prepare a molecular size standard by mixing 5 uL of the 1 kb ladder with 1 uL of tracking dye.

C. Loading and running the gel

1. Remove the wedges from the casting tray and fill the buffer reservoir with TAE buffer until the buffer is 1-2 mm deep over the gel.

2. Carefully remove the comb by lifting it straight out of the gel slowly.

3. Carefully pipette each mixture (6 uL) into a well in the gel . See the demonstration by the instructor before doing this step. Load one well with the prepared 1 kb ladder.

4. After all the lanes have been loaded, connect the leads from the power supply to the gel box. Make sure the gel is oriented correctly (wells at negative [black] end, DNA will "run to the red"). Set the output level to 100 volts and turn the power on.

5. Run the gel until the tracking dye is approximately 3/4 the way across the gel.

D. Staining the DNA in the gel with ethidium bromide.

1. After turning the power off, remove the gel from the gel box and submerge it in the ethidium bromide staining solution. WARNING: ETHIDIUM BROMIDE IS A POWERFUL MUTAGEN. USE GLOVES WHEN WORKING WITH IT. Allow the gel to stain for 5 minutes.

2. Remove the gel to a tray of water and allow it to destain for 5 minutes.

E. Photography

1. Place the gel on the transilluminator. Place the clear plastic shield over the transilluminator window before turning on the UV light. WARNING: THE TRANSILLUMINATOR EMITS SHORT WAVE UV LIGHT WHICH WILL DAMAGE SKIN AND EYES, DURING PROLONGED EXPOSURE. BE SURE THAT PROPER SHIELDING IS IN PLACE BEFORE TURNING ON THE TRANSILLUMINATOR.

2. Turn on the transilluminator (BE SURE THAT PROPER SHIELDING IS IN PLACE) and look at the gel. Confirm the presence of DNA (orange bands) and turn off the transilluminator.

3. Remove the plexiglass shield and position the styrofoam shield and camera on the transilluminator. Turn on the transilluminator, wait a few seconds for the light to come on and shoot the picture by squeezing and quickly releasing the trigger on the camera.

4. Turn off the transilluminator. Remove the white tab from the film holder. Then pull out the photograph by grasping the black tab and pulling it completely out of the camera.

5. Wait 30-45 seconds and peel off the print from the backing. Don't touch the inside of the backing which has a photochemical gel on it. Dispose of the backing in the trash can.

Cycle Sequencing using Sequitherm DNA Polymerase

Introduction

Cycle sequencing is a modification of the traditional Sanger sequencing method. The principles are the same as in Sanger sequencing; Dideoxynucleotides are used in a polymerization reaction to create a nested set of DNA fragments with dideoxynucleotides at the 3' terminus of each fragment. The key difference is that cycle sequencing employs a thermostable DNA polymerase which can be heated to 95 deg C and still retain activity. The advantage of using such a polymerase is that the sequencing reaction can be repeated over and over again in the same tube by heating the mixture to denature the DNA and then allowing it to cool to anneal the primers and polymerize new strands. Thus, less template DNA is needed than for conventional sequencing reactions. Furthermore, the repeated heating and cooling can be automated using an instrument called a DNA thermal cycler.

Procedure

1. Label 4 PCR tubes A, C, G, and T.

2. To the A tube add 2 uL of "A mix". To the C tube add 2 uL of "C mix". To the G tube add 2 uL of "G mix". To the T tube add 2 uL of "T mix". Set the tubes on ice while you set up the master mix.

3. In a separate tube, prepare the master mix as follows

Item Amount

Sterile ddH2O 9 uL

Template DNA 3 uL of plasmid miniprep

10x Sequitherm reaction buffer 2.5 uL

LiCor IRD41-labeled primer 1.5 uL

Total 16 uL

Item	Amount
Sterile ddH2O	9 uL
Template DNA	3 uL of plasmid miniprep
10x Sequitherm reaction buffer	2.5 uL
LiCor IRD41-labeled primer	1.5 uL
Total	16 uL

4. Add 1 uL of Sequitherm DNA polymerase to the master mix and mix by pipetting.

5. To each PCR tube add 4 uL of the master mix and mix by pipetting.

6. Place tubes in the DNA thermal cycler (Gene Amp 2400, PE) and run the following thermal profile;

94 deg C, 2 minutes (initial denaturation step)
30 cycles of the following 3 steps

94 deg C, 30 sec (denaturation)
50 deg C, 15 sec (primer annealing)
72 deg C, 15 sec (polymerization)

94 deg C, 3minutes (final denaturation step)
6 deg C, indefinitely (refrigeration)

7. When the file is complete (2 hours), remove the PCR tubes, pulse them down and add 3 uL of sequencing stop buffer to the reaction. Store tubes in the dark, refrigerated until the sequencing gel is ready.

Preparation of a DNA Sequencing Gel

A. Preparing the Glass plates

1. Locate two LiCor plates (one with a notch and one without) and clean them as follows:

- Scrape off the polyacrylamide with a razor blade
- Scrub both sides of the plate with detergent and water using crub brush and wet paper towel.
- Rinse the plate well with tap water, running your hands over the gel to detect any residual polyacrylamide.
- Set the plates on the drying rack to drain.
- Dry the plates completely using kimwipes (not paper towels)
- Clean the plates scrupulously with di-water and kimwipes. Be super-scrupulous with the side of each plate which will contact the polyacrylamide.

2. Assemble the plates using 0.25 mm spacers on each side. Set it up so that the plate without the notch is on the bottom and the notched plate is on top with the notch facing you. Place the clamps on and add the U-shaped plexiglass spacer to the slot in the top and snug down the clamps (firm but not tight). Pick up the assembly and admire your work to see that everything is lined up correctly.

B. Preparing the Gel mix.

Caution! Avoid contact with unpolymerized acrylamide. It is a mild neurotoxin and can be absorbed through the skin. Wear gloves and safety glasses when handling solutions containing acrylamide.

1. Make the gel mix in a 50 mL polypropylene tube.

Add: 10.5 g urea (ultrapure)
3.0 mL of Long Ranger Acrylamide Mix (50% Acrylamide)
3.0 mL of 10x TBE buffer
ddH2O to 25 mL
Mix until the urea is completely dissolved.

2. Filter the gel mix using a 0.22 um pore filter unit and a 30 mL syringe.
3. Degas the gel mix. - Transfer the gel mix to a small side arm flask and place a stopper in the top. Attach the side arm to a vacuum source and apply vacuum to the flask while agitating for 1-2 min. Continue the application of a vacuum until bubbles no longer appear when flask is shaken (1-2 minutes).
4. Place the flask on ice.

C. Pouring the gel.

1. Gather the following items: Pipettor set to 100 uL, pipettor set to 10 uL, 10% ammonium persulfate solution (APS: stored in refrigerator and less than a week old), TEMED, a 30 mL syringe, and the spacer for the top of the gel.
2. Pull the stopper out of the syringe, fill the pipettors with 100 uL APS and 10 uL TEMED.
3. Add 100 uL APS and 10 uL TEMED to the gel mix, agitate gently to mix without introducing bubbles. While holding your finger over the end of the syringe pour the gel mix into the syringe and put the plunger just into the barrel of the syringe. Invert the syringe and use the plunger to expell the air.
4. Pour the gel, being careful not to introduce bubbles (bang on the plates while pouring).
5. Insert the top spacer and place three clamps on the top of the gel. 6. Allow 45 min for polymerization.

D. Pre-electrophoresis

1.Carefully pull the spacer out and use a razor and wet kimwipes to remove excess polyacrylamide and dried urea from the top of the gel.
2. Make sure the outsides of the gel plates are clean and dry, especially in the area where the scanner will be pointing.
3. Assemble the gel rig on the sequencer, being careful not to get any liquid behind the glass plates.
4. Make 1000 mL of 0.6x TBE buffer and fill the buffer chambers.
5. Use the syringe with the bent needle to get rid of bubbles clinging to the bottom of the plates.
6. Attach the top and bottom buffer chamber covers, attach the cable between the top buffer chamber and the sequencer, and close the sequencer.
7. Run the gel for 45 minutes.

E. Denature the sequencing reactions and prepare them for loading.

1. Place the reactions in a heat block at 90ûC for 5 minutes and cool them quickly on ice.
2. Transfer 10 uL of the reaction from underneath the oil (wipe the oil off the tip) to a clean, labelled tube.
3. Store on ice in dark until ready to load.

F. Loading and running the gel

1. Turn off the power to the gel and the scanner. Open the sequencer and remove the top of the upper buffer chamber.
2. Use a pasteur pipet to flush out the urea which has diffused from the gel into the top well.
3. Insert the sharkstooth comb so that the teeth rest firmly on the gel, but are not digging into the top of the gel.
4. Load the gel (1.2 - 1.5 uL per lane) by placing the end of the pipet tip in the space between the glass plates, directly over the well. Dispense the contents of the tip slowly. You should be able to see the contents run down into the bottom of the well.
5. After all the wells have been loaded, put the top back on the upper chamber, close the sequencer, and start the gel running. Let it run for 10 minutes and set the autogain and focus.

Determining DNA sequence from LI-COR sequencer output

LEGEND

** = DOUBLE CLICK

* = SINGLE CLICK

*| = CLICK AND HOLD

# = LET GO OF CLICK

1. OPEN THE IMAGE FILE

2. CHANGE VIEW TO BLACK ON WHITE

3. LANE DEFINITION (ASSUMING A, C, G, T LOADING ORDER)

** ON A

BRING LINE JUST INSIDE OF "A" LANE AND *|, MOVE TO NEAR THE RIGHT SIDE OF "A" LANE AND #.

BRING LINE JUST INSIDE OF "C" LANE AND *|, MOVE TO NEAR THE RIGHT SIDE OF "C" LANE AND #.

BRING LINE JUST INSIDE OF "G" LANE AND *|, MOVE TO NEAR THE RIGHT SIDE OF "G" LANE AND #.

* ON "OK"

4. MARK 1ST BAND

POSITITON THE GREEN X ON THE FIRST BAND YOU WISH TO USE AND *. (TO ADJUST WHERE THE X GOES *| THE RIGHT MOUSE BUTTON WHILE MOVING UP OR DOWN) 5. AUTO SEQUENCE

6. EDIT AUTO SEQUENCE

-TO MAKE AN AMBIGUOUS CALL

Position cursor over the band and ** with the right mouse button. Move the cursor to the other choice for that call and ** with right mouse button (two red marks will appear and the international code letter for the ambiguity is seen in the column on the left)

7. TO SAVE SEQUENCE: (YOU WILL SAVE TWO FILES: ONE IS A "SAMPLE FILE" WHICH CONTAINS THE IMAGE AND YOUR BASE CALLS; THE OTHER IS A REPORT FILE WHICH IS A TEXT FILE OF THE SEQUENCE that will be used for your analysis of the sequence. The latter file is saved to a disk and then the instructor will email it to you.

Using Sequence Databases and Search Tools

Sequence comparisons

Once you have obtained DNA sequence you can compare it with all known sequences in the existing databases using a program called BLAST (Basic Local Alignment Search Tool). This program is maintained by the National Center for Biotechnology Information (NCBI) and is the standard for searching large DNA sequence databases like GenBank.

BLAST searches can now be carried out via a WWW page which contains a simple form for submission of search jobs. You can simply paste your sequence into the window on the page and submit the request. Searches must be submitted in the FASTA format. BLAST is actually a set of 5 programs and the program you choose depends upon the type of search you are doing.

blastp - compares an amino acid query sequence against a protein sequence database;
blastn - compares a nucleotide query sequence against a nucleotide sequence database;
blastx - compares the six-frame conceptual ranslation products of a nucleotide query sequence (both strands) against a protein sequence database;
tblastn - compares a protein query sequence against a nucleotide sequence database dynamically translated in all six reading frames (both strands).
tblastx - compares the six-frame translations of a nucleotide query sequence against the six-frame translations of a nucleotide sequence database.

The BLAST form also has an option for choosing which database(s) you would like to search. The nonredundant (nr) option is the most thorough because it effectively searches all the databases. There are different databases which may be used for a variety of other applications. Finally, you can choose the option of having the results sent to you via email. This can be useful because often the computers at NCBI are swamped with requests and it can take a while before your request comes up in the queue.

Downloading sequences from the public databases

If you are successful in identifying your sequence you may want to search the sequence databases (using names of proteins, authors or ID numbers), download the related sequences, and align them using a program such as DNASIS. This program allows you to identify open reading frames, restriction sites, align sequences with other known sequences, translate your DNA sequence into a protein sequence, and a host of other usefull applications. You can use sequence alignments to learn where your sequence is located in the full length gene (if they are closely related), how similar the sequences are, and which amino acids within your sequence are conserved. For a more complete list of sequence databases see the Biologist's Control Panel.

DNA Isolation with GeneClean

Introduction

Geneclean is a commercially available kit that is used to purify DNA from agarose gels. Purification is based on the use of a suspension of finely ground silica (glass milk) which DNA will bind to in high ionic strength buffers. DNA is eluted (removed) from the glass milk in a low ionic strength buffer or water. Before the DNA will bind to the agarose, the agarose must be liquified. Heating to the melting point of agarose would denature the DNA so a solution of NaI is used to dissolve the agarose at a lower temperature. The procedure is rapid, simple and produces a high yield of pure DNA. It is widely used in molecular biology.

Procedure

1. Excise the DNA band from the ethidium bromide-stained gel with a razor blade. To minimize damage to the DNA sample, expose the gel to the UV light for as short a time as possible. Remember to wear eye and face protection as well as gloves.

2. Determine the approximate volume of the gel slice by measuring its mass (1 g = 1 mL) and transfer it to a microcentrifuge tube.

3. Add 3 volumes of NaI stock solution to the tube. (e.g., if your gel slice was 0.25 g then the gel volume would be 0.25 mL. You would add 3 times that volume, or 0.75 mL (or 750 uL), of NaI solution.)

4. Place the tube in the 50 deg C water bath for as long as it takes to dissolve the agarose.

5. Remove the tube from the water bath and add 5 uL of glass milk solution. Before removing the silica from the stock tube, vortex it to make sure the glass powder is suspended in the solution.

6. Incubate at room temperature for 15 minutes, mixing every 2-3 minutes to ensure that the glass milk stays suspended.

7. Pellet the silica by spinning in the microcentrifuge for 5 seconds. (When the microcentrifuge reaches maximum speed, count to five and turn off)

8. Use a micropipet to remove the NaI solution, being careful not to disturb the pellet of silica. Centrifuge the tube again for 5 seconds and remove the last trace of NaI using a micropipet.

9. Wash the pellet 3 times with NEW (salts in an ethanol solution). To do this add 1 mL of ice cold NEW to the tube and resuspend the pellet in the wash by pipetting back and forth while digging into the pellet with the pipet tip. After it is resuspended, spin for 5 seconds in the centrifuge and discard the supernatant. Repeat the wash procedure two more times.

10. After the supernatant from the third wash has been removed, spin the tube again for a few seconds and remove the last little bit of liquid with a micropipet.

11. Resuspend the washed white pellet with 5 uL of TE buffer and incubate the tube at 50 deg C for 2-3 minutes to elute the DNA from the silica.

12. Centrifuge for 30 seconds to make a solid pellet. Carefully remove the 5 uL of supernatant which contains the eluted DNA and place in a new tube.

Ligation of Plasmid DNA

The plasmid DNA that was isolated from the agarose gel was linear due to being cut with restriction enzymes. In order to propagate the DNA for later study you will need to close the circle to recreate a plasmid that can be transformed into E. coli. The process of covalently closing a plasmid is called ligation and is catalyzed by an enzyme called T4 DNA ligase. T4 DNA ligase uses either blunt or compatible overhanging ends, and ATP as substrates to form a phosphodiester bond. (See Resources for more information on T4 DNA ligase)

Procedure

1. For each ligation combine the following solutions in a labeled microcentrifuge tube.

Item Amount

Sterile ddH2O 4 uL

5x ligase buffer 2 uL

Gel purified DNA 3 uL

Ligase enzyme 1 uL

Total 10 uL

2. Vortex the contents briefly to mix, and pulse down in the microcentrifuge. Place the tube in the refrigerator for 1 day to 1 week.

Transformation of E. coli

Introduction

By mixing E. coli cells with plasmid DNA carrying an antibiotic resistance gene, one can isolate cells that have incorporated the plasmid as an autonomously replicating DNA molecule. This is a naturally occuring process, called transformation, but it occurs at such a low frequency in nature that it is not feasible as a routine method for getting recombinant plasmids into E. coli. Much higher frequency of transformation can be obtained if E. coli cells are treated with a solution of divalent cations (usually Ca) at low temperatures to make them "competent" to take up plasmid DNA. It is thought that the Ca++ ions form an ionic bridge between the negatively charged phosphate groups of both the DNA and the membrane phospholipids. The cold temperature crystallizes the membrane stabilizing this interaction. When the cells are then exposed to a "heat shock" some of the cells take up the DNA. If the cells are allowed to warm up above 4 deg C before the heat shock step, they will not be competent. Be sure to use good sterile technique during the entire procedure.

Usually the number of cells in a transformation is in excess and the limiting "reagent" is the DNA. A measure of the quality of the competent cells is the transformation efficiency. This is the number of transformants (colonies which grow on the plate) obtained per ug of plasmid DNA. In order to determine the transformation efficiency, a control transformation with a specified amount of plasmid DNA is carried out along with the experimental transformation. The number of colonies is counted and multiplied by the appropriate dilution factor to get the total number of transformants. This is divided by the amount of DNA used in the transformation and expressed as transformants per microgram of DNA. Transformation efficiencies between 10^6 and 10^9 represent the normal range for competent E. coli cells.

Making Competent Cells

E. coli cells are harvested by centrifugation when in late log phase of growth. They are chilled and treated with ice cold solutions containing relatively high concentrations (0.5 M) of divalent cations (usually Ca, but Mn also works well). To obtain the highest efficiency cells extraordinary care must be taken to keep cells cold during the preparation. The process involves multiple centrifugations and can by very tricky. Many researchers purchase aliquots of competent cells which have been prepared commercially and frozen at -80 C. You will be given an aliquot of such cells. The transformation efficiency of these cells should be approximately 10^9.

Procedure for Transforming E. coli Cells (Today we are using strain DH1-alpha cells purchased from Life Technologies)

1. Place 2 sterile 15 ml polypropylene tube on ice. Label one with your plasmid name and the other "control".

2. To each of the polypropylene tubes add 50 uL of the competent cells. Be sure to keep the cells cold during the transfer process.

3. Add 1 uL of your ligation mix to the first polypropylene tube and 1 uL of the control DNA (pUC18 at 10 pg/uL) to the control tube. Mix gently and incubate on ice for 30 minutes. Be sure to keep the cells cold during the transfer process.

4. Heat pulse the tubes in a 42 C water bath for 45 seconds exactly. The length of the heat pulse is critical for obtaining highest efficiencies.

5. Place tubes on ice for 2 min.

6. Add 0.9 mL of LB broth and incubate the tubes in a shaking incubator at 37 C for 60 minutes.

7. Label 3 LB/Amp plates with your plasmid name and 3 LB/Amp plates with "control". Number each set of plates 1-3.

8. Plate 0.1 mL of your plasmid transformation mix onto the LB/Amp plate labeled 1 and spread the broth out evenly over the plate. Also place 0.1 mL of your plasmid transformation mix into a second tube with 0.9 mL of LB broth. Mix and plate 0.1 mL of this mixture onto the LB/Amp plate labeled 2. Place 0.1 mL of the dilution from the second tube into a third tube with 0.9 mL of LB broth. Mix and plate 0.1 mL of this mixture onto the LB/Amp plate labeled 3.

9. Use the same plating procedure (step 8) to plate out the control transformation mix.

10. Incubate all plates at 37 C for 14-18 hours. Do not incubate any longer than 18 hours. After incubation is complete place plates in the refrigerator until needed.

Item	Amount
Sterile ddH2O	4 uL
5x ligase buffer	2 uL
Gel purified DNA	3 uL
Ligase enzyme	1 uL
Total	10 uL