Lab 3:  Plasmid minipreps, restriction digest, agarose gel

 

(See “Lab 4 References” handout – on reserve in library)

 

 

Plasmid Miniprep (also see “Lab 4 References” handout for an alternate protocol)

 

Plasmid DNA will be prepared using the Wizard Miniprep Kit from Promega.  See “Lab 4 References” handout for protocol – follow steps III “Sample Preparation” and V “Plasmid Purification without a Vacuum Manifold”.  Each miniprep will be prepared using 1.5 ml of DH5α (pPBH) culture.  This kit combines two techniques:  alkaline lysis and silica resin-based DNA purification.

 

Alkaline lysis is the most widely used technique for plasmid DNA preparation.  Cells are partially lysed using an alkaline solution of the detergent sodium dodecyl sulfate (SDS).  This allows small plasmid DNA molecules to escape from the cell, while genomic DNA remains within the cells.  When a concentrated potassium acetate solution is added to the cell lysate, cell debris (containing high molecular weight genomic DNA) is precipitated, while plasmids and soluble proteins remain in solution.  If cell membranes are dissolved completely, sheared genomic DNA may be released, contaminating the plasmid prep.  To avoid this, the lysis step is carried out for a limited time – just enough for the solution to clear.  Excessively vigorous mixing (e.g. vortexing) should also be avoided.  Standard alkaline lysis protocols follow the lysis steps with phenol/chloroform extractions and ethanol or isopropanol precipitation to produce a pure nucleic acid solution.

 

The Wizard Miniprep Kit purifies DNA by adhesion to a silica matrix.  This technique is based on the observation that nucleic acids adhere to silica in high-salt conditions, but not in low-salt conditions.  DNA will bind to silica in the lysis solution, and will be eluted from the silica matrix by TE.  The silica matrix may be composed of finely ground glass (e.g. Pasteur pipettes crushed with a mortar and pestle), diatomaceous earth (fossilized diatoms) or patented “resins” from various manufacturers.  The commercially-supplied products usually give better and more reproducible results, and are probably worth the cost.

 

The plasmid being prepared in this lab is pPBH.  This is pUC19 containing a HindIII fragment of P. ochraceus histone gene sequence.  The bacterial strain that the plasmid is maintained in is DH5α.  This is an E. coli K strain that allows alpha-complementation.

 

Restriction Digest (also see “Lab 4 References” handout)

 

The plasmid prep will be digested using HindIII.  The Wizard Miniprep Kit claims to produce approximately 5 µg DNA from 1.5 ml of a culture containing high-copy-number vectors.  If the yields match expectations, you will have 5 µg DNA in 50 µl TE buffer.  Try digesting 5 µl of your plasmid DNA solution.

 

 

Several points to remember when setting up a restriction enzyme digest:

 

1.         Always use a clean, fresh sterile tip every time you pipette from a restriction enzyme stock. 

 

2.         Keep enzymes cold before use – the time that the stock spends outside the freezer should be minimized, and the tube should be in an ice bucket or freezer block for the entire time it is outside the freezer.

 

3.         Normally, the volume of enzyme stock solution used should not exceed 1/10 the total volume of the restriction digest.  This is because enzyme storage buffers contain high concentrations of glycerol, which will interfere with restriction enzyme activity. 

 

In this lab, you will be provided with a HindIII aliquot which has been diluted 1/10 (in 1X reaction buffer) to a final concentration of 1 unit per µl.  Because the enzyme stock has been diluted, you do not have to worry about the factor above.

 

4.         One unit of restriction endonuclease activity is defined as the amount needed to completely digest 1 µg lambda phage DNA in 1 hour at the optimum temperature for the enzyme.  Cutting supercoiled plasmid DNA often requires excess enzyme – see the table (from the New England Biolabs Catalogue) included in the “Lab 4 References” handout  to find the amount of HindIII required to digest pUC19.

 

5.         Reaction buffers are usually supplied by manufacturers in 10X concentrations.  To get a final concentration of 1X in a restriction digest, 1/10 the volume of the digest should be the 10X reaction buffer stock.

 

            In this lab, the enzyme stock has already been diluted in 1X reaction buffer, so the amount of 10X reaction buffer stock used should be 1/10 the volume of the reaction mix before the addition of the restriction endonuclease.

 

Using the above information, calculate the amounts of enzyme, 10X reaction buffer stock, and dH2O needed to set up a 20 µl restriction digest containing 5 µl of plasmid DNA.  (Check your calculations with a TA before adding enzyme to the mix.)

 

When setting up your digest, keep the tube that contains the digest on ice.  After all the ingredients have been added (including the enzyme, which should be added last, pulse spin your tube on the microcentrifuge to concentrate all the ingredients at the bottom.  Mix briefly by flicking the bottom of the tube,  (vortexing DNA solutions is not recommended), and then pulse spin again to recollect your mix at the bottom of the tube.  

 

Digest your DNA sample for 1 hour at 37°C.  During this time, you can prepare your gel and test the DNA concentration in your genomic DNA sample from last week.  When the digest is complete, add 4 µl of 6X gel-loading buffer (see “Lab 4 References” handout for recipes), and prepare to load your samples on the gel.

Gel Electrophoresis

(also see “Lab 4 References” handout and Sambrook reference – in the lab)

 

DNA molecules can be sorted for size by electrophoresis through an agarose gel matrix.  The porous polysaccharide structure of  agarose gels separates DNA molecules by size – smaller (negatively charged) DNA molecules are able to move more easily through the agarose, and therefore travel further towards the positive electrode in the same length of time than larger molecules.  Within a specific range (depending on the specific conditions – agarose concentration, voltage, electrophoresis buffer, ethidium bromide concentration – the gel is run under) the distance travelled by a linear DNA molecule will be inversely proportional to the log10 of the molecular weight.  This allows the size of linear DNA fragments to be estimated by comparing their mobility to a standard curve prepared using linear DNA fragments of known length.

 

Plasmid DNA that has not been cut with a restriction endonuclease will migrate in 3 forms:  supercoiled closed circular DNA, single-strand nicked open circle DNA, and sheared linear DNA.  The exact mobilities of the 3 forms will vary based on conditions, but closed circular DNA will usually migrate faster than the other two because of its supercoiling.  This means that if the DNA band farthest from your well on your uncut plasmid sample is the largest, you have prepared a plasmid DNA sample without excessively damaging your DNA.  (Note that your gel may also contain a diffuse band of partially degraded RNA – which will migrate faster than your DNA bands.)

 

We will be using ethidium bromide to visualize DNA during electrophoresis (see “Lab 4 References” handout).  Ethidium bromide is often added to the agarose gel, the gel electophoresis buffer, or both.  While ethidium bromide allows the mobility of DNA fragments to be observed during electrophoresis (using a UV lamp), it also affects the mobility of the DNA by introducing positive supercoils.  For this reason, gels intended for publication are often run without ethidium bromide and stained with ethidium bromide later.

 

Preparing an agarose gel (see diagram in “Lab 4 References” handout)

 

Each agarose gel will be shared by 2 groups.  (The gels will have 8 wells, and each group will have 3 samples – HindIII cut plasmid DNA, uncut plasmid DNA, and genomic DNA.  One of the remaining 2 lanes will be used for “1 Kb DNA Ladder” molecular weight markers.

 

1.        Calculate the amount of agarose powder needed to make 50 ml of a 0.7 % agarose solution.  Weigh out this amount of agarose and add to 50 ml of 1X TBE buffer (see “Lab 4 References” handout for recipes of electrophoresis buffers).

 

2.         Melt the agarose in a microwave.  (Watch the solution while heating – if it starts to boil over, stop heating immediately.)  Mix the solution until the melted agarose is evenly distributed in solution, and then cool to 50°C.  This will stop the hot solution from warping the gel tray, and also reduce ethidium bromide breakdown.

3.         While the gel is cooling, set up the gel tray.  Seal off the ends of the gel tray with masking tape.  Make sure that the entire end of the tray is covered, no gaps or tears are present, and that the ends and sides of the tape are folded around the outside and bottom of the gel tray.  (If you fold one end of the tape back on itself before sticking it to the tray, it will be easier to remove later.)  Seams may be tested for leaks by pipetting molten agarose along them, but this is not usually necessary if the gel has been set up carefully.

 

After the tray has been taped, place it on a level surface, and position the comb about 0.5 cm from one end of the tray.  Adjust the vertical position of the comb so that it is ~ 0.5-1 mm from the bottom of the tray.  (If the comb is too high, you will have smaller wells to load in, and if the comb is too low, the bottom of the gel will tear out and samples will leak through the bottom of the gel.)  Line up the comb carefully so it is at right angles to the gel tray.  (This will mean that your samples will not run crooked.)

 

4.         Working in the fume hood, add enough 10 mg/ml ethidium bromide stock solution to produce a final concentration of 0.5 µg/ml in the gel mix.  Always wear gloves when handling ethidium bromide solutions – or gels containing ethidium bromide.  Dispose of ethidium bromide contaminated tips in the designated container in the fume hood.  Gently swirl until the dye is evenly distributed – try not to introduce bubbles by vigorous mixing.

 

5.         Pour the agarose into the gel tray.  If any bubbles are present, they can be popped with a p200 tip.  (If you do this, dispose of the tips properly.)  If the gel starts to leak, the leak can usually be plugged by holding a piece of ice against the leak to solidify it rapidly.  Let the gel cool until it is solid.

 

6.         Once the agarose is cool and solid, carefully remove the comb by gently pulling straight up.  Wash the comb, and return it to your drawer.  Remove the tape from the gel, and place in ethidium bromide waste container.  (Handle your gel with gloves.)  Place your gel (in the gel tray) inside the apparatus, with the wells near the negative electrode.  Fill the apparatus with 1X TBE until the gel is completely submerged.

 

7.         If you have never loaded a gel before, you should practise loading dummy samples on one of the demonstration gels that have been set up.  The proper technique will be demonstrated.  (Note – to save time, do this while your DNA is still digesting.)

 

8.         When your digest is finished, set up the following 3 samples:

 

20 µl digested plasmid + 4 µl loading buffer

5 µl uncut plasmid + 5 µl TE + 2 µl loading buffer

2 µg genomic DNA prep in 10 µl TE + 2 µl loading buffer

 

9.         Carefully load your samples in 3 wells on one side of the gel.  (Note that the maximum capacity of a minigel well is about 20  µl – so you will probably not be able to load all of your digested plasmid).  Load one of the centre lanes with 5 µl of 1 Kb DNA Ladder.  Do not move gel once wells have been loaded.

 

10.       Connect the positive electrodes with the red wire, and the negative electrodes with the black wire.  Run the gel at 100 volts for 45-60 minutes, or until the bromophenol blue (dark blue dye) is near the bottom of the gel.

 

11.       Remove the gel from the apparatus.  (Use gloves.)  The TAs will demonstrate photography of the gel.

 

Study Questions (due in week 4)

 

1.         Using your gel photograph, measure the distances of the bands in the 1 Kb DNA Ladder lane from the well.  Graph the distances of the bands from the well (on the x axis) against the log10 of the molecular weight of the bands (taken from the “Lab 4 References” handout).

 

2.         Using the standard curve you have prepared, work out the size of pUC19 and the HindIII fragment that has been cloned into it.

 

3.         Describe the purpose of the different ingredients in the Wizard kit Cell Resuspension Solution, Cell Lysis Solution, and Neutralization Solution.