Project 2 Manual: Experimental Procedures
The technique we will be using involves the application of the polymerase chain reaction (PCR) to produce large quantities of the E7 gene, which has been modified in the region of the epitope. In this technique, an oligonucleotide consisting of the modified DNA sequence is used as the primer or starting point for the reaction. This results in half the DNA created having the “normal” sequence and the other half having the modified sequence.
The possible epitope modifications which can be made are:
|Original sequence :||RAHYNIVTF|
Each pair of students will be given a sample of the E7 gene inserted into a vector, a circular length of DNA which carries the gene into the E. coli. In addition, they will each be given a oligonucleotide with the appropriate codon swapped out. When the DNA is mixed, the oligonucleotide binds to the complementary sequence on the E7 gene, with all of the base pairs matching except for the modified codon. During PCR, the polymerase enzyme manufactures a copy of the entire vector, starting off at the oligonucleotide. As each copy uses one of the original strands as a template, half of the vectors made in this way will have the original, unchanged E7 gene, and half will have the version with the modification (see Figure 4). We will use an enzyme which favourably digests the original DNA and leaves the modified version.
While PCR is a useful method for amplifying small amounts of DNA, it is complicated and time-consuming. Luckily, we can rely on nature to do some of the hard work for us. If the vector is inserted into E. coli, the bacterium will copy it as it copies its own DNA. As the bacterium multiplies, they become “plasmid factories” producing large amounts of DNA. To harvest the DNA, we will use a technique called a Mini-prep which separates the plasmid DNA from the rest of the cell contents. Finally, we will sequence the plasmid and check to make sure that we have created our mutant version of the E7 gene.
eg. substituting the Histidine at position 3 (CAT) for a Proline (CCT)
Putting the whole procedure together, we get :
The procedure for the week is summarised in the following figures :
|What is 10X Buffer ?|
Sometimes you will see a solution provided at “10X” concentration. As the name suggests, this means that the solution is ten times more concentrated than it needs to be. Whenever you add one solution to another, each solution is diluted by the presence of the other. In order to use a solution appropriately, you will need to add it to the reaction mixture so that it will be properly diluted by the addition of the other reagents. For a 10X solution, this means that the volume of this solution is one tenth of the final volume.
- Deoxyribonucleotide triphosphates (dNTPs) – adenosine triphosphate, thymidine triphosphate, cytidine triphosphate and guanosine triphosphate – these are the raw materials for making our DNA during PCR.
- Buffer – this is a solution which stabilises the reaction by ensuring that the pH remains constant.
- Oligonucleotide primer.
- Plasmid vector containing the E7 gene.
- A polymerase enzyme to make the DNA (Pfu Turbo Poly).
- Distilled water to make the volume up to 50µL.
This mixture contains all of the materials needed for PCR. Keep in mind that PCR is a notoriously sensitive procedure - it will amplify any DNA it encounters in the reaction mixture. Therefore, it is important to limit contamination in this part of the experiment. We use specially purified water and sterile tips on our micropipettes.
Many of the volumes are quite small and will need the use of the P2 micropipette. To ensure that the majority of the material is dispensed, add the largest volume (the dH2O) first and draw the liquid up and down inside the pipette tip a few times. Make sure you dispose of your tips between samples.
You will be working in pairs, with each person trying to make one mutant. In addition, for the whole class, we will also prepare a negative control (which has everything except the DNA vector) as a way of testing whether the reaction worked or not. Therefore, as a pair, you will be preparing two tubes.
One way of limiting the amount of error due to the small volumes used is to make up a master-mix. Since so many of the reagents used in PCR are the same, we can make a stock solution with the common ingredients. Then we just add the correct volume of the stock solution to the reagents which vary between samples. When calculating the volumes needed for a master-mix, make up enough for as many samples as you have, plus one spare. As well as allowing for reagents left in the tube, it gives us some space if we make a mistake. Therefore, in this step, we will make up a master-mix capable of supplying reagents for three samples.
The volumes needed for the PCR reaction are provided below :
|Tube Label||10mM dNTP||10X Buffer||Oligo. Primer||DNA Vector||Pfu Turbo Poly||dH2O|
The reagents which are needed in the same proportions for each tube are the dNTPs, the 10X buffer, distilled water and the Pfu Turbo Polymerase. Therefore we will use these to make the master-mix (multiplying each of the volumes needed by 3) :
|10mM dNTP||10X Buffer||dH2O||Pfu Turbo Poly|
We can now use the master-mix to set up our tubes :
|Tube Label||DNA Vector||Master-Mix||Oligo. Primer|
The polymerase chain reaction is a technique used to amplify small quantities of DNA into amounts which can be used in experimental investigations. It involves a series of heating and cooling stages, during which the amount of DNA doubles. As DNA is heated, the strands separate. It is then cooled slightly and the primers attach. A polymerase enzyme from a thermophilic (heat loving) bacterium then synthesises the complementary strands of DNA and the process repeats. A typical PCR procedure features 35-36 cycles of 9 minutes each. While this might not seem like much, in that time, a single molecule of DNA could potentially yield 68,719,476,740 (~236) molecules. More information on PCR can be found here.
When PCR was originally developed, researchers had to carefully time the placement of individual tubes in waterbaths set at the appropriate temperatures. Luckily for us, PCR thermal cyclers have been developed which automate the entire process. Tubes are inserted into the cyclers and run through a series of heating and cooling steps which are programmed in. At the completion of the run, samples are stored at 4°C. Programs can be “tweaked” to suit the conditions of the DNA or enzymes used.
The cycler in the SPARQed laboratory has been programmed for this experiment and the reaction will run overnight. For your own information, the details of the program are :
|60°C||30 seconds||35 cycles|
The next step is to remove the original DNA, leaving us with only the plasmids containing the mutated E7 gene. Dpn 1 is an enzyme which favourably digests methylated DNA. Since the original plasmid containing the E7 gene was methylated, addition of some Dpn 1 to the PCR mixture should remove all of this material.
- To each of your tubes, add 1µL of Dpn 1.
- Incubate the reaction mixture for 1 hour at 37°C.
DNA electrophoresis is performed using a gel made out of agarose. The gel must be made fresh for each test we perform. The running buffer, which is used in the electrophoresis tank, must also be made fresh. The working running buffer (1X) is made by diluting the stock buffer (50X) 1 in 50 using distilled water.
You will need to prepare around 500mL of 1X TAE running buffer to fill the tank and make the gel. Since you are provided with 50X TAE stock buffer, you will need to calculate how much buffer to add to water to ensure a 1 in 50 dilution. Use the following procedure :
Total volume = 500mL
1/50 of 500mL = 500 ÷ 50 = ____ mL
volume of 50X stock needed is ____ mL
Volume dH2O needed = Total volume - Volume stock needed
= 500mL – ____ mL
= ____ ml
Dilute ____ mL stock in ____ mL of dH2O
The gel we will be using is 0.8% agarose. This means that we should add 0.8g of agarose to every 100mL buffer. We do not need this much gel, however, so instead we will work on a total volume of 40mL. To adjust the masses involved, use the following procedure :
|New Mass Agarose||=||New Volume|
|Old Mass Agarose||Old Volume|
|New Mass Agarose||=||New Volume||x Old Mass Agarose|
|=||40 x 0.8|
- Microwave the solution on HIGH for 2 minutes (for a small gel). Make sure that the agarose is completely dissolved by swirling the heated mixture. Allow it to cool for 3 minutes.
- Wipe a plastic gel tray and comb with 70% ethanol and use tape to seal up the gel tray.
- Add 4µL of SYBR-Safe into the melted agarose and swirl to mix.
- Pour the melted agarose into the gel tray. Place the comb into the right position and allow it to set for approximately one hour (this can be done faster by placing the gel tray in the refrigerator.
- Carefully remove the comb and tape from the gel. Place the tray containing the gel into the electophoresis tank with the wells at the black (negative) electrode end. Cover the gel with 1X TAE running buffer.
- Samples are prepared by adding the appropriate amount of 6X loading dye to make it to 1X (i.e. a 1 in 6 dilution). The combined volume of the digest solution and 6X stock dye must be 6 times the volume of the dye added.
- We are going to use 5µL of our digest solution
- if the volume of dye added is “x” and the volume of digest solution 5µL
- Use this calculation to determine how much dye to add to your sample and prepare loading solutions for each of your samples
- Load all of the loading solutions into separate wells in the gel (loading 10µL of the molecular markers last into a separate well on the left or right hand side of your gel). Each well holds a maximum 30µL. Make sure you keep track of where you load each sample
- Run the gel at 80V. There must be small bubbles rising from both ends of the electrophoresis chamber. Check after 5 minutes to make sure the gel is running (i.e. the dye front has moved, is relatively straight and has run the correct direction). Then allow the gel to run for the necessary amount of time (about 1 hour however, check that the dye front has almost run through the gel).
- Switch off the power pack and take the gel to the illuminator. Take a photograph, print off and glue into your workbook. Annotate the photograph, indicating bands of interest.
- Pour away the buffer from the electrophoresis tank and rinse well with water. Rinse the gel tray and comb as well.
Interpreting Your Gel
Whenever we run a gel, we should always include a DNA “Ladder” which features fragments of DNA of known size. This ladder serves as a reference point to indicate the size of the DNA fragments in our sample. A map of the ladder we are using in this exercise is provided below in the figure to the right.
There are a number of different ways that bacteria can be encouraged to take up plasmids. In the wild, they do this at their own pace using extensions of their bodies called “pilli”. Natural transformation results in a success rate of around 1%. In laboratory transformations, we need to get as much of the plasmids inside the cells as possible, so we use a number of techniques to hurry the process along.
If we are inserting large plasmids, a process called electroporation is used. In this method, an electrical field is applied to the cells. This opens up pores in the membrane which allows the plasmids to enter.
In our procedure, we will use a method of heat shock. The bacterial cells are chilled and then rapidly heated to around 42-45°C. The change in temperature creates pores in the membrane through which the plasmids can enter.
In our experiment we will be using competent cells – E. coli which have been treated in such a way to ensure maximum efficiency in transformation. While these can be prepared in the lab, the process is long, laborious and prone to error. Instead, we will be using commercially available Sure 2 cells.
The vector containing the gene for E7 also contains a gene which confers resistance to the antibiotic Ampicillin to any bacteria which are successfully transformed. We can use this as a way of determining which bacteria have been transformed by trying to grow the bacteria in the presence of Ampicillin – the only bacteria which should grow are those which have taken up the vector (and the Ampicillin resistance gene).
- Warm a set of LB-Ampicillin culture plates (two for each sample) in the bacterial incubator. Label the plates with the name of the sample you have been working on (one set as “1X”, the other as “5X”). You will need a positive transformation control containing pcDNA3-16E7 and a negative transformation control in which distilled water is added instead of the PCR product.
- Thaw the Sure 2 cells from the -80°C freezer on ice. Pre-chill a transformation tube for each sample on ice.
- Add 40µL of Sure 2 cells to each tube.
- Add 0.8µL of β Mercaptoethanol to each tube. Cap, and mix gently.
- Incubate on ice for 10 minutes, mixing gently every 2 minutes.
- Add 4µL of the PCR reaction mixture, pcDNA3-16E7 or distilled water to the appropriate tubes.
- Incubate on ice for 30 minutes.
- Heat shock the cells for 30 seconds in a 42°C waterbath and quickly add to ice.
- Incubate on ice for 2 minutes.
- Add 500µL of LB (broth form) to the cells.
- 6 Incubate in the bacterial shaker at 37°C for 1 hour.
- Spread 100µL of each suspension onto the “1X” plates.
- Transfer the remaining suspension to a 1.5mL centrifuge tube and spin at 8,000rpm for 5 minutes.
- Remove the supernatant and resuspend in 100µL of LB.
- Spread the remaining 100µL onto the “5X” plates.
- Incubate the plates (lid side down) overnight in the bacterial incubator at 37°C.
- Prepare your LB-Ampicillin broth by adding 10µL of Ampicillin for every 10mL of broth (e.g. if you are going to need 60mL, add 60µL).
- Prepare 5 x 30mL tubes for each sample (not the controls). Into each tube, aliquot 3mL of the LB-Ampicillin broth you have prepared.
- Select a suitable colony from each plate using a sterile yellow micropipette tip and subculture by dropping the tip into the LB-Ampicillin tubes prepared above (Select 5 colonies per plate and subculture one colony into each tube).
- Incubate the tubes for at least 6 hours at 37°C (overnight if possible).
Once we have our culture of transformed E. coli we need to demonstrate that these bacteria are carrying the plasmid we hope to have inserted. A Mini-Prep is a procedure in which cells are lysed in an alkaline environment and larger chromosomal DNA is separated from the smaller plasmids using a chromatography method. The presence of the plasmids is then demonstrated using electophoresis.
Production of Cleared Lysate
- Divide the entire culture in each tube into two equal volumes in Eppendorf Tubes and centrifuge at 8,000rpm for 5 minutes.
- Remove supernatant from one of the tubes and resuspend in 250µL Buffer P1. Ensure that there are no cell clumps visible after resuspension of the pellet.
- Add 250µL Buffer P2 to each sample and invert 4-6 times to mix.
- Add 350µL Buffer N3 and invert 4-6 times to mix. The solution should become cloudy as proteins precipitate.
- Centrifuge at 13,000rpm for 10 minutes at room temperature.
Binding of Plasmid DNA
- Insert a QIAprep spin column into a centrifuge tube for each sample.
- Transfer the supernatant from the lysis stage into the spin column.
- Centrifuge at 13,000rpm for 1 minute at room temperature.
- Discard the flowthrough in the centrifuge tube and reinsert the column into the centrifuge tube.
- Add 500µL Buffer PB to the Spin Column.
- Centrifuge at 13,000rpm for 1 minute.
- Discard the flow through in the centrifuge tube and reinsert the column into the centrifuge tube.
- Repeat wash steps above with 750µL wash solution.
- Centrifuge at 13,000rpm for 2 minutes at room temperature.
- Cut the lid from a sterile Eppendorf tube for each sample.
- Transfer the spin column to the sterile Eppendorf tube, taking care not to transfer any of the wash solution.
- Add 50µL Buffer EB to the spin column.
- Incubate at room temperature for 1 minute.
- Centrifuge at 13,000rpm for 1 minute at room temperature.
- Discard the column. Transfer the eluate from above to a sterile Eppendorf tube (with cap). Samples can be stored at -20°C or below.
To ensure that transformation has occurred in the cells, we will perform a quick electrophoresis step. Follow the procedures outlined earlier using 5µL of your sample. Include wells for each of your vectors and each of your Mini-Prep samples.
The ultimate check of whether we have been successful in generating our mutant version of E7 is to sequence the gene. The procedures to do this are based on PCR and are largely automated. Your samples will be given to one of the Diamantina Institute’s scientists who will process them in time for analysis.
To begin with, we will need to prepare the samples through PCR. The method for this is based on the BigDye kit and is listed below :
For each of your samples, prepare :
|Tube Label||BigDye||BigDye Seq.
|Sample||1 µL||3.5 µL||2 µL||2 µL||11.5µL|
The conditions for the PCR are :
|50°C||5 seconds||25 cycles|
Ethanol / EDTA / Sodium Acetate Precipitation
- To each tube, add the following reagents :
- 2 µL 125mM EDTA
- 2 µL 3M Sodium Acetate (pH 5.2)
- 50 µL 100% Ethanol
- Mix by inverting 4 times
- Incubate at room temperature for 15 minutes
- Spin at 13,000rpm at 4°C for 30 minutes
- Invert tube and spin up to 500rpm for 1 minute
- Add 70µL of 70% ethanol
- Spin at 13,000rpm for 30 minutes
- Invert tube and spin up to 500rpm for 1 minute
- Place samples in 96°C thermocycler with the lid open for 10 minutes to dry. Pellets can be stored at -20°C
When the PCR and precipitation processes are complete, your tutors will take your samples for analysis