Team:Heidelberg LSL/Notebook methods
Transformation is a process in which linear or circular DNA, in other words your designed plasmid, is transferred into competent bacteria without the help of bacteriophages. This is done foremost with one of the following two intentions: firstly, you want to amplify your chosen BioBrick DNA. Transformed bacteria multiplies quickly which doubles the amount of DNA at every proliferation step. It is a very fast and convenient way. Afterwards you have to extract your chosen BioBrick out of your bacteria. Secondly, when you completed the cloning of your construct, you might need to bring it into your final bacteria cells to test if it works.
For amplifying DNA from the registry, we transformed chemocompetent Top10 cells by applying the following protocol:
DNA from Registry Plate: 10 µl water is added to the corresponding well containing the desired BioBrick DNA; DNA is solved by incubation for 15 min at room temperature. 5 µl of DNA solution are added to 50 µl of chemocompetent Top10 cells and incubated on ice for 20 min. Afterwards, a heat shock is performed by incubation of the bacteria on 42°C/50 s. After additional 2 min on ice, the bacteria are plated onto LB-Amp agar plates.
Inoculation of LB agar plates
Bacteria are cultivated on agar to be amplified and to be able to pick single colonies directly from the solid medium.
LB agar is made of LB medium, substituted with 100 ug/ml ampicillin and 1.5 % Agar, which makes the LB media solidify.
When plating out transformed bacteria, you should start by carefully labeling the plate with the construct name, date, bacterial strain, antibiotic resistance and your initials and preheat the plate to 37° C. Using a bunsen burner, you form an inoculation loop out of a small glass pipette. After adding 20-100 µl of bacterial suspension to the plate, you evenly distribute it on the agar plate surface with your glass tool. Now, you can incubate the inoculated LB agar plates at 37 degrees Celsius over night.
Inoculation of liquid LB medium
A bacterial culture in liquid medium is inoculated in order to grow a culture, which is required for a miniprep. Liquid cultures where used during our tests. To inoculate a liquid culture the pellet which has been centrifuged out of another liquid culture is diluted with LB Medium containing the required antibiotics. To inoculate a liquid culture on the base of an agar plate culture you have to pick the colony from the plate with a pipette tip. Then the tip is dropped into the falcon tube with the medium. After the inoculation the falcon or the flask is put in an incubator at 37°C shaking
With a Miniprep you extract DNA out of liquid bacterial suspensions such as overnight cultures. There are various companies that offer kits for this process. We followed the instructions of the Qiagen Miniprep kit or the Promega Miniprep kit according to the manufacturer's protocols.
DNA was eluted from the column using either 50 µl of water or TE buffer.
A restriction digest describes the process in which restriction enzymes, which can be seen as small DNA-scissors, cut DNA in pieces. These enzymes bind selectively at specific sequences and cut double-stranded DNA either blank or sticky (i.e. they produce up to 4 bp non-paired overhangs). You have to think carefully about the choice of the right restriction enzymes when developing your cloning strategy. For iGEM standard cloning in BBa-format as it was applied here, the following protocol can be used for all Enzymes used (EcoRI, SpeI, XbaI, PstI).
We took the NEB restriction enzymes and buffers for setting up the digestions. Those were done as follows:
- 1 µg of DNA
- 3 µl of NEB buffer 2
- 3 µl of BSA
- 1 µl of each restriction enzyme used
- Water was added to reach a final volume of 30 µl
After setting up the digestions, the reaction mixtures were mixed briefly, spun down in a centrifuge (quick run up to 5.000 rpm) and then incubated on a thermomixer heating device for 1 h at 37 °C and stored at -20 °C afterwards. After Digestion, DNA was purified via a qiagen nucleotide removal or gel extraction kit.
Shrimp alkaline phosphatase can be used for dephosphorylation of the vector backbone. This ensures, that the vector backbone can neither religate with itself nor with any dephosphorylated insert (that might occur during digestion of the vector backbone) during the ligation process.
For dephosphorylation of the vector backbone DNA, 1 unit of FastAP enzyme was added to the digestion mix (after digestion was performed for 1 h) and the mix (including SAP) was incubated at 37 °C for 1 h again.
Oligo design and synthesis
Oligos were designed using WinSerial Cloner and VectorNTI and synthesyzed by Sigma (http://www.sigmaaldrich.com/life-science/custom-oligos.html, 0.025 µM synthesys range).
Gel electrophoresis is used to analyze the size of linear DNA fragments and to separate them physically based on this property. In order to do so the DNA is loaded together with a loading dye indicating the progress of a running gel onto a agarose gel. The negatively charged nucleotides move towards the positive pole of the applied electric field. The longer a DNA fragment the larger is the drag caused by the web structure of the agarose gel. To make the DNA bands visible a fluorescent dye like ethidium bromide is added. The EtBr molecules intercalate with the DNA and start to glow after by UV-induced activation. To have a size reference a DNA ladder with fragments of known lengths is also loaded onto the gel. The gel can be photographed with a camera or bands containing required DNA can be cut out easily.
The gel is prepared by adding about 1g/100ml agarose to buffer TAE or TBE, then boiling it in a microwave, and finally by adding ethidium bromide. Afterwards the gel is poured into the gel tray that was sealed before and a comb is mounted. The gel tray is transferred to the gel box which is filled with buffer TAE or TBE. Now it can be charged and it is ready to run.
A gel extraction is performed to extract the DNA fragment out of an excised gel band. The DNA is washed using a membran column. Gel extractions are performed to purify DNA parts and to seperate them from other restriction- or PCR products. This can be done before a ligation assembly is conducted. We did gel extractions using the QIAquick Gel Extraction Kit.
QIAquick nucleotide removal kit
This kit is applied to purify DNA in solution, e.g. after a restriction digest. This kit has got the advantage compared to the QIAquick Gel Extraction Kit, that smaller DNA fragments are conserved too. This kit was used according to the standard manufacturer's protocol.
Ligation digest is the process of connecting compatible ends of DNA or RNA by means of the enzyme ligase. Visually, you can close your open plasmid with a ligation back to circular DNA. Our Ligation reactions were set up using 3-6 fold access of insert (RecA) and 50 ng of each backbone (roughly). Then we add 2 µl (10x) T4 ligase buffer for 20 µl total volume and 1µl of the enzyme T4 ligase. The rest of the volume is water. The reaction mix is incubated for 50 min at room temperature. To inactivate the enzyme T4 ligase you heat up the mix to 70 °C for 5 min. Afterwards, the ligation mix (chilled) can be used for transformation of E.coli chemocompetent cells.
Annealing is performed to gain double-strand DNA out of two synthetic oligo nucleotides. These are obtained as single-stranded DNA. During the annealing process hydrogen bonds are formed and base pairs are established. This process yields double-stranded DNA. By the design of the correct overlaps no further restriction digest is required. Besides, the use of synthetic DNA fragments increases the reliability of the assembly of small parts (<100bp).
To perform the annealing process 5 µl of each oligo (diluted to a concentration of 100 mM) and 5 µl of NEB buffer 2 are added to 35 µl of water. After mixing, the reaction mix is heated up to 95 °C/5 min and then cooled down slowly to room temperature and stored at -20 °C.
With this PCR you test if your construction of your designed plasmid worked as you planned it. With intelligent primer selection you amplify the crucial construction fragments. You can now estimate the length of the expected PCR product. Then you analyse your amplicon with gel electrophoresis and measure the length of the fragments. Only those fragments with the suitable length show that the construction was successful.
After restriction digest, ligation digest, transformation of bacteria and inoculation on agar plates, you pick colonies and put them into some liquid medium. You incubate it and take some of it for your PCR. The reaction mix consists of 0,2 µl of each primer, 9,6 µl water, and 10 µl of 2x PCR mastermix (fermentas). The PCR program was done as follows: 94°C/3min||94°C/30s|60°C/30s|72°C/3min 45s||30x 72°C/10min|4°C/forever
Sequencing was performed by GATC Biotech (http://www.gatc-biotech.com/en/index.html) using the registry standard sequencing primers VF2 (5’-tgccacctgacgtctaagaa-3’) or VR (5’-attaccgcctttgagtgagc-3’).
For characterizing the precA,B and psulA constructs with a lacZ reporter in a close-to application context, we performed X-Gal assays. Primarily we exposed our samples to UV-radiation in the gel electrophoresis chamber. After the samples were incubated for 1 h at @37 °C/80 rpm we added X-Gal substrate (final concentration of 200 ug/ml) to all samples. Coloring of the wells were monitored 15 min after X-gal addition and quantifications of the coloring intensity were done.
In order to determine the response of our SulA and RecA promoters to small UV doses, we used the highly sensitive ONPG assay.
ONPG (Ortho-nitrophenyl-β-D-galactopyranoside) is a synthetic LacZ substrate and produces a yellow color (o-nitrophenol) upon cleavage by LacZ. The formation of the yellow color can be easily determined using a photometer at 420 nm absorbance wavelength.
Concentration measurement with nanodrop
The nanodrop spectrophotometer is used to measure DNA concentrations fast and accurate. The system can measure a drop of 1µl. This renders dilutions and measurements in a standard cuvette photometer unnecessary. Before the measurement is done the photometer is initialized and blanked with 1µl of H2O. Then 1µl of the sample can be pipetted onto the device after the previous drop is wiped away with a lab towel. After the measurements have been completed another sample of water is loaded to check whether the blank measurement was correct.
Use of the photometer
A photometer is used to measure the light absorption of a colored solution. The solution is irradiated by monochromatic light of a given wavelength. The light passes through a cuvette containing the sample. Behind the cuvette the light beam hits a detector, a photocell which measures the intensity of the transmitted light. This value is then compared to the intensity of light when passed through a substance with the same solvent but without the colored substance. With this ratio it is possible to calculate the concentration for the colored substance using Beer‘s law. A more saturated substance would have a higher absorption of light.
We have used the photometer to measure the optical density of the bacteria solution before the exposure to UV-light to confirm bacterial growth using a photometer at 600 nm wavelength.
Use of the plate reader
The plate reader is a automatic photometer to detect the absorbance of the bacteria suspensions. The samples are loaded into a 96-well (8 by 12 matrix) with 50 µL of the bacteria suspension and 50 µL ONPG per well. A monochromatic light illuminates the sample using a wavelength of 420 nm at the maximum absorbance of ONPG. A light detector located on the other side of the well measures how much of the initial light is transmitted through the sample. All samples can be analyzed in one run, and the results are given in a clearly arranged spreadsheet.
To get further information we highly recommend to visit www.openwetware.org or the pages of the kit distributors.