Continuing Wildtype Hemoglobin Expression
- Centrifuge cultures grown overnight at 4500rpm at 4°C for 15 minutes.
- Resuspend each pellet in 1mL of LB.
- Add each pellet to 1L of LB with 100μg/mL of ampicillin (four of these).
- Incubate at 37°C at 165rpm until OD600 = 0.6.
- Add 1mL 0.1 M Isopropyl β-D-1-thiogalactopyranoside (IPTG) to each flask.
- Prepared fresh today: 0.12g + 5mL distilled H2O, sterile-filter
- Add 37.5mL of hemin solution to each flask.
- The hemin solution was prepared by Dr. Hartings on 6/20/12. It was prepared by combining 160mg hemin with 1.38mL triethanolamine, and diluting the solution to 40mL. The pH of the solution was then adjusted to 7.8, and then water was added to a final volume of 150mL.
- Let cells continue to grow and express for 4 hours at 30°C at 165rpm.
- Spin the cells at 4500rpm at 4°C for 15 minutes.
- Resuspend the cells in sterile-filtered 25mM Tris, 50mM NaCl, pH 8 (35mL total).
- Freeze the cells in liquid nitrogen and store at -20°C.
None of the cultures started for the minipreps had any growth last night (the "colonies" were very small). This means no minipreps will be done today. Two colonies did however appear on the NovaBlue + M103S transformation plate from two days ago. The plate was just left at room temperature overnight.
DNA ligation with T4 DNA Ligase
This will be done with both the wildtype Asc Hb double-digested insert and the M8S/M33S/M103S Asc Hb double-digested insert. The procedure is based off of the protocol on the NEB website.
- Mix 2μL of 10x T4 DNA Ligase Buffer, 5μL of the double-digested pQE-80-L-Kan vector, 10μL of the wildtype insert, 2μL of nuclease free H2O, and 1μL of T4 DNA Ligase.
- Mix 2μL of 10x T4 DNA Ligase Buffer, 5μL of the double-digested pQE-80-L-Kan vector, 6μL of the triple mutant insert, 6μL of nuclease free H2O, and 1μL of T4 DNA Ligase.
- Place at room temperature for 10 minutes.
- Place on ice.
My calculations yesterday led me to a 5:1 volume ratio, but I thought about it again and here is my reasoning for for today for a 3:1 insert to vector molar ratio:
- Since the vector is about 4700bp and the insert is about 450bp, then the molecular weight of the vector is probably about 10 times bigger than the molecular weight of the insert.
- I then reasoned that the molarity ratio for the wildtype insert to the vector was about 1.5:1 because of the concentration calculated and the relative molecular weights.
- This seems to mean that the ratio of volumes for insert to vector should be 2:1 for the wildtype insert
- I also reasoned that the molarity ratio for the triple mutant insert to the vector was about 2.6:1.
- I could basically stay close to a 1:1 ratio for the volume ratio, but the real volume ratio would be about 1.15:1
- I was also aiming for about 50ng of each, but since the molar ratio seems to be most important, I will only aim for 50ng of the vector. This means there will be more vector than insert in terms of mass, but not molarity.
Running an Analytical DNA Gel
- Make a 1.2% agarose gel (0.3g agarose + 25mL TAE buffer, microwave until boiling, then pour into gel chamber, and place comb for wells).
- When the gel has solidified, add TAE buffer to the chamber until the gel is completely submerged in buffer.
- Load 5μL of DNA ladder into the 1st well.
- Load 5μL of the M8S/M33S/M103S/A71M/L40M PCR product from yesterday with 1μL 6x loading buffer into the 3rd well (mix before pipetting into well).
- Load 5μL of the wild-type Asc Hb and pQE-80-L-Kan ligation from this morning with 1μL 6x loading buffer into the 5th well (mix before pipetting into well).
- Load 5μL of the triple mutant (M8S/M33S/M103S) Asc Hb and pQE-80-L-Kan ligation from this morning with 0.5μL 6x loading buffer into the 7th well (mix before pipetting into well).
- Run the gel at 100V until the "dye line" has moved about 3/4 of the way down the gel
- Place the gel in Ethidium Bromide Stain for about 30 minutes
- Move the gel to TAE buffer to destain for about 20 minutes
- View under a UV light
- Be careful when working with ethidium bromide and UV light.
Again, there are no band on the gel. I still may try to transform in case the DNA concentration is low, but usually bands at least show up bright after a successful PCR.
I need to make sure that there is actually DNA in my mini-preps from Tuesday because my two PCR reactions this week did not seem to work. To do this I will quantify the DNA in the colonies 2 and 3 minipreps of M8S/M33S/M103S/A71M Asc Hb. This procedure was repeated for both minipreps:
- Using a smaller volume quartz cuvette, 95μL of dH2O was placed in the spectrophotometer and the baseline was corrected with this.
- 5μL of DNA was added to the 95μL of water. This was pipetted up and down with a pipette to mix.
- The absorbance was taken at 260 and 280nm.
Note: the absorbances are for 20x dilute DNA samples
- A260 = 0.147
- A280 = 0.081
- Absorbance Ratio = 1.8089
- 20x Dilute DNA concentration = 7.3350 μg/mL
- Non-dilute DNA concentration = 146.7 μg/mL
- A260 = 0.181
- A280 = 0.104
- Absorbance Ratio = 1.7404
- 20x Dilute DNA concentration = 9.0500 μg/mL
- Non-dilute DNA concentration = 181 μg/mL
So it seems there is DNA, which means there are other things that can probably be troubleshooted.