In order to attain our results in an effective manner, we carried out a multitude of experiments, all of which we list here and which can easily be recreated.
Competent Cell Check
The purpose of the competent cell check was to find the optimal concentration of the competent cells. We tested five different concentrations and found that 10pg/ul of the competent cells had the most efficient growth.
- Competent cell kit/ Transformation Efficiency Kit
- 2.0m; microcentrifuge
- Pipette: 1ul, 50ul, 200ul- Incubator- Water bath- SOC media- Float rack- Agar plates
1. Spin down the DNA tubes from the Competent Cell Test Kit/Transformation Efficiency Kit to collect all of the DNA into the bottom of each tube, prior to use. A quick 20-30 second spin at 8,000-10,000 rpm will be sufficient.
Note: There should be 50 µL of DNA in each tube sent in the Kit.
2. Thaw competent cells on ice. Label one 2.0ml microcentrifuge tube for each concentration and then pre-chill by placing the tubes on ice.
3. Pipette 1 µL of DNA into each microcentrifuge tube. For each concentration, use a separate tube.
4. Pipette 50 µL of competent cells into each tube. Flick the tube gently with your finger to mix.
5. Incubate on ice for 30 minutes.
6. Preheat the water-bath to 42°C.
7. Heat-shock the cells by placing them into the water-bath for one minute. Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
8. Immediately transfer the tubes back to ice, and incubate on ice for 5 minutes. This helps the cells recover.
9. Add 200 µL of SOC media per tube, and incubate at 37°C for 2 hours. Prepare the agar plates during this time: label them, and add sterile glass beads if using beads to spread the mixture.
10. Pipette 20 µL from each tube onto the appropriate plate, and spread the mixture evenly across the plate. Do triplicates (3 each) of each tube if possible, so you can calculate an average colony yield.
11. Incubate at 37°C overnight or approximately 16 hours. Position the plates so the agar side is facing up, and the lid is facing down.
12. Count the number of colonies on a light field or a dark background, such as a lab bench. Use the following equation to calculate your competent cell efficiency. If you've done triplicates of each sample, use the average cell colony count in the calculation. Make sure to measure the area of each colony to see how effective our cells are.
The purpose of the Gibson Assembly was to synthesize DNA fragments of the E. coli bacterium with Andersen promoters of three different strengths. In performing this procedure, we were able to determine the optimal promoter strength for the growth of COX-2 and c-Myc. We discovered that the medium strength promoter was more competent with COX-2 and the strong promoter was more competent with c-Myc. The COX-2 promoter and c-Myc promoter were then synthesized by IDT and used for making the BioBrick.
- Master Mix
- Pipette: 1ul, 10ul, 11ul
- COX-2 gene
- C-Myc gene
- Anderson promoter
- Distilled water
Set up the following reaction on ice:
* Optimized cloning efficiency is 50–100 ng of vectors with 2–3 fold of excess inserts. Apply 5 times as many inserts if size falls below 200 bps. Total volume of unpurified PCR fragments in Gibson Assembly reaction should not exceed 20%.
** Control reagents are provided for 5 experiments.
*** If greater numbers of fragments are assembled, additional Gibson Assembly Master Mix may be required.
2. Incubate samples, using a thermocycler at 50°C, for 15 minutes when 2 or 3 fragments are being assembled, or 60 minutes when 4-6 fragments are being assembled. Following incubation, store samples on ice or at –20°C for subsequent transformation.
Note: Extended incubation up to 60 minutes may help to improve assembly efficiency in some cases (for further details see FAQ section).
The chemically competent cell transformation was for the purpose of observing phenotypic results from our previous Gibson Assembly procedure in the E. coli bacterium. A phenotypical reporter system in the presence of black light, part of our Biobrick, was what we searched for after the final incubation. This was done by examining the cell cultures in black light after an overnight process of full recovery.
- NEB 5-alpha Competent E. coli cells
- Pipettes: 50ul, 2ul, 950ul, 200ul
- 1.5ul microcentrifuge tube
- Ice bath
- SOC media
- Inoculating loop
- LB plate
- CAMr plate
1. Thaw New England Biolab (NEB) competent cells on ice.
2. Chill approximately 5 ng (2 μl) of the ligation mixture in a 1.5 ml microcentrifuge tube.
3. Add 50 µl of competent cells to the DNA. Mix gently by pipetting up and down or flicking the tube 4–5 times to mix the cells and DNA. Do not vortex.
4. Place the mixture on ice for 30 minutes. Do not mix.
5. Heat shock at 42°C for 30 seconds*. Do not mix.
6. Add 950 µl of room temperature SOC media* to the tube.
7. Place tube at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.
8. Warm selection plates to 37°C.
9. Spread 50–100 µl of the cells and ligation mixture evenly onto the plates.
10. Incubate overnight at 37°C.
11. Check for phenotypical expression of cell cultures in black light.
* Please note: For the duration and temperature of the heat shock step as well as for the media to be used during the recovery period, please follow the recommendations provided by the competent cells’ manufacturer.
The procedure of electrophoresis was used to verify that all our Biobrick components had been successfully assembled during the Gibson Assembly protocol. Through this process, we were able to confirm success, through determining and comparing the results to each corresponding construct’s predicted plasmid length.
- Electrophoresis chamber + power supply
- Agarose solution
- Pipette: 1ul
- TBE buffer
- Bromophenol blue
Cast 0.8% Agarose Gel
1. Seal ends of gel-casting tray with tape, and insert well-forming comb. Place gel-casting tray out of the way on lab bench so that agarose poured in next step can set undisturbed.
2. Carefully pour enough agarose solution into casting tray to fill to depth of about 5mm. Gel should cover only about one-third the height of comb teeth. Use a pipette top to move large bubbles or solid debris to sides or end of tray, while gel is still liquid.
3. Gel will become cloudy as it solidifies (about 10 minutes). Be careful not to move or jap casting tray while agarose is solidifying. Touch corner of agarose away from comb to test whether gel has solidified.
4. When agarose has set, unseal ends of casting tray. Place tray on platform of gel box, so that comb is at negative (black) electrode.
5. Fill box with TBE buffer, to a level that just covers entire surface of gel.
6. Gently remove comb, taking care not to rip wells.
7. Make certain that sample wells left by comb are completely submerged. If “dimples” are noticed around wells, slowly add buffer until they disappear.
Load Gel and Electrophoresis
1. Add loading dye to each reaction.
a. Add 1ul of loading dye to each reaction tube. Close tube tops, and mix by tapping tube bottom on lab bench, pipetting in and out, or pulsing in a microfuge. Make sure tubes are placed in a balanced configuration in rotor.
2. Use micropipette to load entire contents of each reaction tube into separate well in gel as shown in diagrams. Use fresh tip for each reaction.
a. Steady pipette over well using two hands
b. If there is air in the end of the tip, carefully depress plunger to push the sample to the end of the tip.
c. Dip the pipette tip through the surface of the buffer, left it over the well, and gently depress the pipette plunger to slowly expel sample. Sucrose in the loading dye weighs down the sample, causing it to sink to the bottom of the well.
3. Close top of electrophoresis box, and connect electrical leads to a power supply, anode to anode and cathode to cathode. Make sure both electrodes are connected to same channel of power supply.
4. Turn power supply on, and set to 100-150 volts. The ammeter should register approximately 50-100 milliamperes. If current is not detected, check connections and try again.
5. Run the gel through the electrophoresis machine for 40-60 minutes. Good separation will have occurred when the bromophenol blue band has moved 4-8 cm from wells. If time allows, electrophorese until the bromophenol blue band nears the end of the gel. Stop electrophoresis before bromophenol blue band runs off end of gel.
6. Turn off power supply, disconnect leads from the inputs, and remove top of electrophoresis box.
7. Carefully remove casting tray from electrophoresis box, and slide gel into disposable weigh boat or other shallow tray. Label staining tray with your name.
We used restriction digest to find the restriction enzyme that cut our construct into the least number of parts. The restriction enzyme that we choose will allow us to separate our construct from the plasmid backbone.
- 1.5 ml tubes
Restriction enzymes: PstI, EcoRI, SpeI, Xbal
- Pipette: 4 ul, 5ul, 10ul, 1ul, 6ul
1. Use permanent marker to label four 1.5-ml tubes, in which restriction reactions will be performed
P = PstI
E = EcoRI
S = SpeI
X = Xbal
- = no enzyme
2. Use table below as a checklist by adding reagents to each reaction.
3. Collect reagents, and place in test tube rack on lab bench.
4. Add 4 μl of DNA to each reaction tube. Touch pipet tip to side of reaction tube, as near to the bottom as possible, to create capillary action to pull solution out of tip.
5. Always add buffer to reaction tubes before adding enzymes. Use fresh tip to add 5 μl of restriction buffer to clean spot on each reaction tube.
6. Use fresh tips to add 1 μl of PstI, EcoRI, SpeI, XbaI to appropriate tubes.
7. Use fresh tip to add 1μl of dH20 to tube labeled “-”.
8. Close tube tops. Pool and mix reagents by pulsing in a microfuge or by sharply tapping the tube bottom on lab bench.
9. Place reaction tubes in 37 C water bath, and incubate for a minimum of 20 minutes. Reactions can be incubated for a longer period of time.
Bradford Protein Assay
1. The standard protocol can be performed in three different formats, 5 ml and a 1 ml cuvette assay, and a 250 µl microplate assay. The linear range of these assays for BSA is 125–1,000 µg/ml, whereas with gammaglobulin the linear range is 125–1,500 µg/ml.
2. Remove the 1x dye reagent from 4°C storage and let it warm to ambient temperature. Invert the 1x dye reagent a few times before use.
3. If 2 mg/ml BSA or 2 mg/ml gamma-globulin standard is used, refer to the tables in the 11 4110065A.qxp 9/25/2007 2:39 PM Page 17 appendix as a guide for diluting the protein standard. (The dilutions in the tables are enough for performing triplicate measurements of the standards.) For the diluent, use the same buffer as in the samples (refer to Troubleshooting section for more information). Protein solutions are normally assayed in duplicate or triplicate. For convenience, the BSA or gamma-globulin standard sets can be used, but blank samples (0 µg/ml) should be made using water and dye reagent.
4. Pipette each standard and unknown sample solution into separate clean test tubes or microplate wells (the 1 ml assay may be performed in disposable cuvettes). Add the 12 4110065A.qxp 9/25/2007 2:39 PM Page 18 1x dye reagent to each tube (or cuvette) and vortex (or invert). For microplates, mix the samples using a microplate mixer. Alternatively, use a multichannel pipet to dispense the 1x dye reagent. Depress the plunger repeatedly to mix the sample and reagent in the wells. Replace with clean tips and add reagent to the next set of wells. Assay Volume of Volume of Standard and Sample 1x Dye Reagent 5 ml 100 µl 5 ml 1 ml 20 µl 1 ml Microplate 5 µl 250 µl
5. Incubate at room temperature for at least 5 min. Samples should not be incubated longer than 1 hr at room temperature. 13 4110065A.qxp 9/25/2007 2:39 PM Page 19
6. Set the spectrophotometer to 595 nm. Zero the instrument with the blank sample (not required for microplate readers). Measure the absorbance of the standards and unknown samples. Refer to Section 3 for data analysis. Note: If the spectrophotometer has a reference and sample holder, the instrument can be zeroed with two blank samples. If the effect of buffer on absorbance is required, zero the instrument with a cuvette filled with water and dye reagent in the reference holder.
1. Make the separating gel:
- Set the casting frames (clamp two glass plates in the casting frames) on the casting stands.
- Prepare the gel solution (as described above) in a separate small beaker.
- Swirl the solution gently but thoroughly.
- Pipet appropriate amount of separating gel solution (listed above) into the gap between the glass plates.
- To make the top of the separating gel be horizontal, fill in water (either isopropanol) into the gap until a overflow.
- Wait for 20-30min to let it gelate.
- Make the stacking gel:
- Discard the water and you can see separating gel left.
- Pipet in stacking gel until overflow.
- Insert the well-forming comb without trapping air under the teeth. Wait for 20-30min to let it gelate.
2. Make sure a complete gelation of the stacking gel and take out the comb. Take the glass plates out of the casting frame and set them in the cell buffer dam. Pour the running buffer (electrophoresis buffer) into the inner chamber and keep pouring after overflow until the buffer surface reaches the required level in the outer chamber.
3. Prepare the samples:
- Mix your samples with sample buffer (loading buffer).
- Heat them in boiling water for 5-10 min.
4. Load prepared samples into wells and make sure not to overflow. Don't forget loading protein marker into the first lane. Then cover the top and connect the anodes.
5. Set an appropriate volt and run the electrophoresis when everything's done.
6. As for the total running time, stop SDS-PAGE running when the downmost sign of the protein marker (if no visible sign, inquire the manufacturer) almost reaches the foot line of the glass plate. Generally, about 1 hour for a 120V voltage and a 12% separating gel. For a separating gel possessing higher percentage of acrylamide, the time will be longer.
Note: Various factors affect the properties of the resulting gel.
• Higher concentration of ammonium persulfate and TEMED will lead to a faster gelation, on the other hand, a lower stability and elasticity.
• The optical temperature for gel gelation is 23°C-25°C. Low temperature will lead to turbid, porous and inelastic gels.
• The pH is better to be neutral and the gelation time should be limited in 20-30 min.