Experiment 1: Site-directed Mutagenesis

Overview: You’ll be using Strategene’s QuickChange reagents and protocol in this experiment.  A PDF file of the manual is on the class website.  You are going to perform oligonucleotide-mediated, site-directed mutagenesis to introduce a mutation into the A. nidulans nimX gene sequence.  The change is a T to C transition at position 6232 in the pNIG6 plasmid sequence.  This results in a change of Tyr (Y) condon #306 to His (H).  The basis of this technique is the use of mutagenic oligonucleotides as primers to PCR amplify the entire plasmid with a high fidelity thermal stable polymerase.  The oligonucleotides will match the pNIG6 sequence on either side of 6232 perfectly (with one caveat) but encode a CG base pair at 6232 instead of TA.  The difference in the sequence between the primers and pNIG6 is called a “mismatch”. Since the procedure yields both mutant and wild type plasmids, one has to screen the results of mutagenesis to distinguish between WT and mutants.  Because T to C transition at 6232 is only detectable by DNA sequencing, we will use an additional trick to identify mutants.  The mutagenic oligonucleotides will contain a second mismatch, one that either destroys a NsiI site at 6228 (Y306H NsiI) or adds a SacI site at 6246 (Y306H SacI).  This will allow you to determine whether clones that result from your mutagenesis are wild type or mutant by a simple restriction digest (either gain of SacI site or loss of the NsiI site).  The next logical step would be to test whether your mutant nimX gene can function in A. nidulans.  You will not perform this test in this exercise, but you will test the function of nimX mutations created in subsequent lab exercises.

Chronological Protocol:

1.  Each group performs one QuickChange PCR reaction:

            All groups mutate pNIG6 plasmid

            Groups 1-4 use the oligo combination Y306H NsiI and Y306H NsiI Comp = “Loss of NsiI” Reaction

            Groups 5-8 use the oligo combination Y306H SacI and Y306H SacI Comp = “Gain of SacI” Reaction

            The TAs will prepare a master mix containing everything except oligonucleotides, pNIG6 DNA, and enzyme.  Each group will receive 44.5 ul of master mix.

            Master mix, all reagents, and the reaction are kept on ice.

            All groups add 1.25 ul each of  each oligonucleotide (concentration – 100 ng/ul)

                        Groups 1 and 5 add 2 ul of 2.5 ng/ul pNIG6

                        Groups 2 and 6 add 2 ul of 5 ng/ul pNIG6

                        Groups 3 and 7 add 2 ul of 10 ng/ul pNIG6

                        Groups 4 and 8 add 2 ul of 25 ng/ul pNIG6

            TAs will add 1 ul of enzyme to each reaction and start the PCR.  No oil will be added to the reactions.  The PCR reaction conditions will be as described in your QuickChange manual, with step 3 of cycle 2 being 7.5 minutes (>1 min/kb plasmid).

            [Note to TAs: make L-AMP plates and L-AMP+XGAL+IPTG plates.  Perform the control reaction according to the manual, all the way through transformation of XL1-Blue cells on L-AMP Xgal+IPTG  plates]

2.  Each group will remove 20 ul of their PCR reaction to a 1.5 ml tube, add 1 ul DpnI, and incubate at 37 degrees for 1 hour.

3.  Each group transforms 2 ul of their DpnI digest reactions into XL1-Blue cells

            a) TAs will provide you with 50 ul of XL1-Blue cells in tubes on ice, NZY+ medium in tubes in the 42 degree water baths

            b) Add 2 ul of DpnI digested DNA to the cells on ice, incubate on ice 10 minutes

            c) Heat shock cells by moving tube from ice to 42 deg water bath for 45 seconds and then returning the tube to ice for 2 minutes

            d) Add 0.5 ml of pre-heated NZY+ medium, and incubate the tubes shaking at 37 degrees for 1 hour.

            e) During the incubation, pipet 0.45 ml of NZY+ medium in a 1.5 tube and store at room temperature.  You’ll use this to dilute the transformation mixture before plating.

            f) After the hour incubation, vortex the cells, pipet 50 ul of cells into the tube containing .45 ml of NZY+ to make a 1:10 dilution of the transformation mixture.

            g) Plate 250 ul of the 1:10 dilution of the transformation mixture on one L-AMP plate.

            h) Plate 250 ul of the udiluted transformation mixture on one L-AMP plate.

i) Incubate plates at 37 degrees.  The TAs will store the plates at 4 degrees after colonies have grown to an appropriate size.  Plates will be ready for you at the start of the next lab period. [ Note to TAs: Pick 6 colonies from each group’s plates and inoculate 2 ml of L-AMP in culture tubes and incubate tubes shaking overnight at 37 degrees the night before the next lab period.  Bring the culture tubes and plates to the lab]

4.  Each group will do 6 miniprep DNA isolations on clones from their QuickChange reaction using Qiagen kit reagents (protocol from Qiagen manual inserted below).

            a) Transfer ~1.5 mls of each cell culture into labeled, 1.5 ml centrifuge tubes.

            b) Harvest the bacterial cells by centrifugation in a the microcentrifuge at full speed for 1 min at room temperature (15–25°C).

            c) 1. Resuspend pelleted bacterial cells in 250 μl Buffer P1 containing RNase.

            d) Add 250 μl Buffer P2 and mix thoroughly by inverting the tube 4–6 times. Mix gently by inverting the tube. Do not vortex, as this will result in shearing of genomic DNA. If necessary, continue inverting the tube until the solution becomes viscous and slightly clear. Do not allow the lysis reaction to proceed for more than 5 min.

            e) Add 350 μl Buffer N3 and mix immediately and thoroughly by inverting the tube 4–6 times.  To avoid localized precipitation, mix the solution thoroughly, immediately after addition of Buffer N3. Large culture volumes (e.g. ≥5 ml) may require inverting up to 10 times and vigorous shaking. The solution should become cloudy.

            f) Centrifuge for 10 min at full speed in a table-top microcentrifuge. A compact white pellet will form.

            g) Apply the supernatants from step 4 to the QIAprep spin column by pipetting.  Label the tops of the spin columns.

            h) Centrifuge for 30–60 s at full speed. Discard the flow-through.

i) Wash QIAprep spin column by adding 0.75 ml Buffer PE and centrifuging for 30–60 s at high speed.

            j) Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer. Important: Residual wash buffer will not be completely removed unless the flow-through is discarded before this additional centrifugation. Residual ethanol

from Buffer PE may inhibit subsequent enzymatic reactions.

            h) Place the QIAprep column in a clean 1.5 ml microcentrifuge tubes that have the lid cut off (use scissors). To elute DNA, add 50 μl Buffer EB (10 mM Tris·Cl, pH 8.5) to the center of each QIAprep spin column, let stand for 1 min, and centrifuge for 1 min at high speed.

            j) Transfer the DNA solution to a clean, labeled, 1.5 ml microcentrifuge tube and store on ice.  NOTE: there is no EDTA in EB, so keep the DNA cold to minimize the effects of any contaminating nucleases.

NOTE TO STUDENTS ON NAMING PLASMIDS:  Your plasmids will be named as follows:

Group number – Experiment Number – Miniscreen Number

Example: The plasmid isolated in the fourth miniscreen DNA in the step above by group 3 would be called: 3-1-4 (group 3 –Experiment 1 – plasmid 4).  That is all that needs to be put on the tube.  All plasmids should be stored in your box at -20 degrees (freezer).  Your notebook should have a list of all plasmids created and their position in your box.  I should then be able to check your notebook at ANY TIME, look up a DNA, find the tube in your box, find the data in your notebook.  Failure to have this information in your notebook by the start of the following lab period, in a manner that I can understand without much help form you, will result in a negative class participation point.  This includes all DNAs (plasmids and PCR products) created during my half of BIO 510.

5.  Each group digests each mutant clone and also does one pNIG6 control digest.  TAs will supply you with pNIG6 DNA at 0.2 ug/ul, and a tube containing a Master Mix containing buffer and enzyme for your digest.

            Groups 1-4 do NsiI plus PstI double digests using NEB Buffer 3 plus BSA

            Groups 5-8 do SacI single digests using NEB Buffer 1 plus BSA

            a) Distribute 35 ul of master mix to each of 7 1.5 ml microfuge tubes.

            b) Add 5 ul of each miniprep DNA to one tube.

            c) Control Reaction: Add 4 ul ddH₂0 to the last tube and 1 ul of pNIG6.

            d) Incubate reactions at 37 degrees for 1 hour.

6. Each group runs all seven (7) reactions along with markers on a 1% gel and gets photo of result.  Record and analyze your results in your lab notebook.  [Note to TAs: prepare gels, have 10X buffers and 10X BSA aliquots ready at the start of this lab period.]

            a) Add 4 ul of 10X loading buffer to each digest.

            b) Load 15 ul of the reaction on a gel.

            c) Load 5 ul of DNA MW Markers in one lane.

            d) Run until the blue dye is near the bottom of the gel ! 45 minutes.

7.  Record and Discuss Results

Mutagenic Primer TM Calculation: Each student should calculate the TM of the PCR primer their group will be using in the Quick Change PCR reaction and record that value in their lab notebook.  The primer sequences and the mismatches between the primers and the pNIG6 template can be determined using Vector NTI (you did this already).  The equation for calculating the TM of primers with mismatches is in the QuickChange user manual. 

            XL1-Blue Transformation: Each student should calculate their group’s transformation efficiency (= number of transformants per ug of pNIG6 DNA added to the cells) and record that information in the notebook.  To perform this calculation, you’ll have to calculate how much pNIG6 DNA was present in the 2ul of the DpnI digest you transformed into XL1-Blue cells.  Record your group’s results on the blackboard and copy down the results from all the groups, including the control transformation done by the TAs.  How do your results compare with the transformation efficiency advertised by Stratagene?  How do they compare with the other groups?  What are the factors that could contribute to differences between your calculated efficiency compared to Stratagene’s advertised efficiency?  Between your efficiency and that of other groups?

Mutagenesis Efficiency: Each student should also record the mutagenesis efficiency of the control reaction run the by TAs (efficiency = percent of E. coli transformants that contain mutant plasmid).  To do this, you will have to consult the QuickChange manual and to learn what the control reaction was and how you would use the blue/white colony color assay to determine which colonies carry mutant plasmid and which carry wild-type plasmid.  Also record the mutation efficiency obtained by your group and all other groups (record information of the blackboard).  To determine this, you should use Vector NTI to predict the results of restriction digests for pNIG6 and the mutated form of pNIG6 that you tried to create.

DNAs:

Plasmids: pNIG6 at 0.2 ug/ul; 25 ng/ul; 10 ng/ul; 5 ng/ul; 2.5 ng/ul

Oligonucleotides (all at 100 ng/ul): Y306H NsiI; Y306H NsiI Comp; Y306H SacI; Y306H SacI Comp.