PCR Amplification

In order to begin the cloning process, individual portions (amilCP, spisPink, PcArsR) of our insert must be amplified to facilitate digestion and ligation of our plasmid. This PCR involves two different primer pairs; one pair matches to the chromoproteins and the other matches to the arsenic regulator.

The basic PCR master mix includes 1x GoTaq Flexi Green Buffer, 0.2mM dNTPs, 2mM MgCl2, 1.25U GoTaq Polymerase, 1μM FWD primer, 1μM REV primer, 1ng template DNA, and molecular water to a final volume of 25μL.

The typical thermocycler protocol heats the reaction at 95C for 5 min. Then it cycles through steps of denaturing (95C for 60s), annealing (60C for 10s), and extension (72C for 30-60s) for 35 cycles. There is a final extension of 72C for 5 min before the reaction is complete and held at 10C.

Agarose Gel Electrophoresis

1. To prepare 1% agarose gel, measure 0.5g of agarose. Add to 50mL of 1X TAE buffer in flask. Swirl to combine.
2. Boil until liquid is homogenous and clear. Let cool. Add 5µL of Sybr Safe dye. Swirl gently until mixed.
3. Pour gel. Add desired comb. Let cool until translucent.
4. Add 1X TAE buffer solution into tank until gel is slightly covered.
5. Add purple loading dye to DNA sample, in necessary.
6. Add 10µL of ladder to your rightmost/leftmost well. The ladder is 1kb Plus Ladder from ThermoScientific (SM1333). Load 10-20µL of DNA per well.
7. Set for 120V for 15-30 minutes.
8. Image gel using blue light.

Gel Extraction

1.After taking a picture of the results of the gel electrophoresis, use blue light along with an orange filter to help find the DNA bands.
2. Place a cut right underneath the band of DNA that is intended for extraction.
3. Cut a piece of filter paper and dialysis tubing to an appropriate size so that it would fit inside the slit.
4. Stick the tubing and filter paper together. Place this into the slit so that the side with the filter paper faces the band of DNA.
5. Place the gel in a gel box. Run the gel at 120V for 10 minutes or until the DNA moves into the filter paper and stays.
6. After running the gel, place the casting tray under blue light once again to ensure that the DNA has successfully moved onto the filter paper.
7. Take a 0.5mL centrifuge tube and pierce a small hole at the bottom of the tube. Place this small tube in a 1.5mL centrifuge tube.
8. Place the filter paper with the DNA into the small centrifuge tube. Centrifuge both tubes. This allows the supernatant to flow out of the small tube into the large one.
9. Take the small centrifuge tube out and discard it. The DNA has been extracted.

Restriction Digest

1. Use SnapGene to determine which enzymes to use for digesting the desired DNA. Use NEB.com to determine which buffer is compatible with these enzymes.
2. Make master mix (1X Buffer, 1U enzyme A, 1U enzyme B, DNA, and molecular water to 25uL) in a centrifuge tube. Remember to keep the enzymes on ice and to add them to the master mix last.
3. Keep DNA on ice. Add DNA to the centrifuge tube with the master mix. Mix using the pipette.
4. Centrifuge the tube with the digestion to make sure any liquid on the sides of the tubes is at the bottom of the tube.
5. Place the digestion reaction in a water bath set at 37ºC for anywhere between an hour to overnight.

Ligation

1. After the insert DNA and the vector backbone have been digested with the same enzymes, they can now be ligated.
2. The reactions contained a volume of digested insert, a volume of digested linear pSB1C3, 1uL T4 DNA ligase, and 1x T4 ligase buffer, and Molecular Water to a final volume of 10μL.
3. Incubate the ligation at 16°C overnight.

Ligation Screening

1. Before transforming the ligation, we tested the ligation reaction using PCR to determine whether we could detect the presence of inserted DNA in the vector. This is a step that we have not seen many labs use and we were excited to implement it.
2. Using 1 uL of ligation reaction, perform PCR as usual using primers that target sites flanking the insertion site.
3. If the agarose gel shows a product of the correct size, proceed with transformation. If not, then re-ligate using a different proportion of Insert:Vector DNA and try again.

Transformation

1. Label one 1.5 mL tube as a positive control (with vector and insert ligation) and one 1.5 mL as a negative control (vector only).
2. Thaw competent cells on ice. Pipette 50 uL of competent cells into both the positive and negative control tubes.
3. Pipette 2 uL of DNA into the competent cells. Gently mix by flicking the tubes.
4. Incubate both tubes on ice for 30 minutes.
5. Heat shock the tubes at 42ºC for 45 seconds. Put the tubes back on ice for 2 minutes.
6. Pipette 900 uL SOC media to the positive and negative control tubes. Be careful not to contaminate the SOC.
7. Place the tubes in a shaking incubator. Incubate at 37ºC for 1 hour at, shaking at 200 rpm. About 15 minutes before incubation finishes, remove plates from the fridge, label appropriately and leave them at room temperature.
8. Take tubes out of incubator. Spin cells at 6000 rpm for 30-60 seconds.
9. Pour off the supernatant, leaving 100-200 uL in each tube.
10. Gently resuspend the pellet in the tubes using the supernatant left.
11. Pipette the resuspended cells onto the selection plate.
12. Add 15-20 glass beads to each plate. Put lid back on each plate. Shake both plates so that the cells can spread around. Once done shaking, pour the beads out and dispose of them properly.
13. Incubate both plates upside down at 37ºC for 14-18 hours.

Colony PCR Screening

The goal of this PCR is to take single colonies from transformation plates and to amplify the DNA in the positive control (ligated vector and insert) and negative control (vector only). After screening, gel electrophoresis will occur to determine whether or not the vector and insert ligated to each other successfully (do this by checking the base pair count).
1. After running a transformation, prepare a master mix (1x GoTaq Flexi Green Buffer, 0.15mM DNTP, 1.5mM MgCl2, 0.63u GoTaq Polymerase, 0.38μM FWD primer, 0.38μM REV primer, and molecular water to a final volume of 10μL). Add an extra 20% uncertainty to each volume of reagent to ensure there is enough mastermix for all PCR tubes.
2. Make sure master mix is mixed using pipette. Then, put 20 uL of the master mix into each PCR tube.
3. Take out the transformation plates from the refrigerator. Also get a new LB-CAM plate (will be used to streak a single colony onto new plate).
4. Select a colony, place a toothpick on it, and dab the same toothpick onto the new petri plate. Then use the same toothpick to inoculate a PCR reaction. Finish streaking the plate using a round-ended toothpick. Properly discard both toothpicks. Repeat until completed for all PCR tubes.
5. Centrifuge the PCR tubes. Then, set the thermocycler so that it has the correct settings (since each PCR reaction is different. Place the PCR tubes in a thermocycler, and start the reaction.
6. Incubate the new petri plate (with the streaks) upside down at 37ºC for 14-18 hours.