Team:Freiburg/Methods
Protocol
Add Instant blue stain to the gel, incubate 30-60 min until bands are visible.
Protocol
- Add 0.5 g of resin to the column. Ad 1 mL lysis buffer and incubate for 10 min.
- Add lysate to the column and let incubate for 5 min. Allow the column to drain by gravitation.
- Wash twice with 5 mL wash buffer. Allow the column to drain by gravitation.
- Eluted 3 times with 1 mL elution buffer in different microcentrifuge tubes. Allow the column to drain by gravitation.
- Perform a SDS-Page to see in which microcentrifuge tube most of the protein got eluted.
To analyse the components of a protein mixtures, one can seperate them according to their size using a polyacrylamide gel. The detergent SDS is used in this experiment to denature the proteins and to attach negative charges to each fragment. This causes all proteins of the sample mixture to move towards the positive pole of an electric field which is applied during the gel electrophoresis. Due to their different sizes they move at different speeds through the crosslinked polyacrylamide matrix of the gel, which leads to a separation of proteins in bands running at different heights according to their size.
Protocol
- Cast a separating gel first according to the following recipe:
| 10% | 15% | 20% | |
| water | 4.1 mL | 2.45 mL | 0.8 mL |
| Bis-acrylamide 30% (w/v) | 3.3 mL | 4.95 mL | 6.6 mL |
| TRIS 1,5M pH=8.8 | 2.5 mL | 2.5 mL | 2.5 mL |
| SDS (10%) | 100 uL | 100 uL | 100 uL |
| TEMED | 10 uL | 10 uL | 10 uL |
| APS | 100 uL | 100 uL | 100 uL |
depending on the size of the proteins to be separated it can be helpful to add 3.6 g urea to the separating gel (equals 6M) d
for smaller samples a gradient gel offers a better separation of proteins. To cast a gradient gel, decide on the percentage the gel should have at the top and at the bottom part and mix half of the volume of each gel recipes. Next use a pipetboy to draw first the higher percentage and then the lower percentage gel mixtures into the same stripette. Draw an air bubble into the stripette which will raise to the top of the fluid level and mix the two fluids that are stacked onto each other in the process. Then add the mixture into the gel casting system evenly.
- let polymerize for approx. 20 min
- Pour a stacking gel on top of the separating gel, which will condense the loaded protein sample into a sharp band:
| standard | sucrose | |
| water | 6.1 mL | |
| Bis-acrylamide 30% (w/v) | 1.3 mL | |
| TRIS 0,5M pH=6.8 | 2.5 mL | |
| SDS (10%) | 100 uL | |
| TEMED | 10 uL | |
| APS | 100 uL |
- place a comb in the stacking gel
- let polymerize for approx. 20 min
- remove the comb, place the gel in an electrophoresis chamber filled with SDS running buffer (RECIPE)
- mix the samples with the appropriate volume of 5x Laemmli buffer (RECIPE) and load them into the pockets of the gel
- close the electrophoresis chamber, connect the electrodes with an energy source and let the gel run at 2 mA until the dye front reaches the bottom of the gel
Protocol:
- Load 20-30 μL protein sample into the wells of a SDS-PAGE gel
- Run the gel for 1–2 h at 100 V.
- Transfer the protein from the gel to the membrane
- Block the membrane for 1 h at room temperature or overnight at 4°C using blocking buffer.
- Incubate the membrane with appropriate dilutions of primary antibody in blocking buffer at 4°C overnight.
- Wash the membrane in three washes of TBST, 5 min each.
- Incubate the membrane with diluted conjugated secondary antibody in blocking buffer at room temperature for 1 h.
- Wash the membrane in three washes of TBST, 5 min each.
- Remove excess reagent and cover the membrane in transparent plastic wrap.
- Visualize image using darkroom development techniques for chemiluminescence or normal image scanning methods for colorimetric detection
To ensure a sufficient amount of input phages for each new panning the eluted phages of the previous round of panning have to be amplified.
<Protocol
1. Prepare a culture of ER2738 in 20 mL LB and let them grow to OD600 0.5 in a big Erlenmeyer flask, at 37°C, shaking at 200 rpm.
2. Add the phage eluate from the previous panning round to the culture.
3. Incubate the cultures for 30 min at 37°C without movement for infection.
4. Incubate at 37°C shaking at 200 rpm o/n, (14 h). → Continue with isolation of phages.
1. Prepare a culture of E.coli in 10 mL LB and let them grow to OD600 0.1 - 0.5.
2. Transfer 1 mL into a culture tube for each phage clone to be amplified.
3. Put picked plaques with a pipette tip in the culture tubes.
4. Incubate at 37°C shaking at 200 rpm for 4 h 30 min.
5. Transfer the culture to microcentrifuge tubes and add to each 500 μL glycerol. Store at -20°C.
→ Continue with colony PCR and gel extraction and sequencing.
To saturate all remaining unspecific binding sites for proteins on the plate the wells were blocked with either BSA or gelatine. This was done to avoid unspecific binding of phages and their displayed ligands to the plastic surface of the well which would result in false positive, non target-specific binders upon elution of the phages with glycine buffer. We used different blocking reagents in the first and second panning, respectively to eliminate all of the phages that bind to BSA despite our internal negative controls. Those phages are lost in the second panning where no BSA is present.
Protocol
1. Slap off the remaining coating solution.
2. Add 300 µL of blocking solution (either 5 mg/mL BSA in 0.1 M NaHCO3 or 0.1% (m/v) gelatine in 0.1 M NaHCO3)
3. Incubate at 4°C for 1 h.
We work with a two-step "sandwich" interaction of streptavidin binding the biotinylated target which binds to a phage. As a first step Streptavidin is coated onto an ELISA plate and adheres via nonspecific hydrophobic interactions. For surface panning the biotinylated target was coated on the streptavidin coated plate as a second step.
Streptavidin Coating
Add to each well (96-well plate) 10 μg Streptavidin in 150 µL 0.1 M NaHCO3.
Incubate the plate in a humid container with damp paper towels o/n at 4°C, gently shaking.
Target Coating
Dilute the target in 150 µL 0.1 M NaHCO3 to a final concentration of 0.44 µM and add it to each well (96-well plate) after slapping off the streptavidin coating solution.
Incubate the plate in a humid container with damp paper towels o/n at 4°C, gently shaking.
To isolate the phages that express target-binding ligands from our library of 1011 different phages, we performed a four-step protocol. First we added the phages to a well which was only coated with streptavidin and blocked with BSA (or gelatine respectively). All phages that are specific for streptavidin or BSA (or gelatine) should should be captured in this well as a negative selection. The phages remaining in the supernatant after this negative selection were then transferred to a microcentrifuge tube with a solution of the biotinylated target to allow phage - target binding without steric hinderances. Afterwards in the third step the phage-target-solution was added to another streptavidin-coated well to capture the target-phage-complexes, taking advantage of the biotin/streptavidin interaction. Finally in the fourth step the supernatant of this well was transferred to a fresh streptavidin-coated well to capture the remaining target-phage complexes. This is meant to ensure that all target-phage complexes can be bound to the plates, reducing the loss of phage-target complexes. Afterwards nonbinding phages were washed away and the remaining, target-bound phages were eluted. For elution we used a chaotropic glycine buffer that dissolves all protein-protein interactions in an unspecific way.
Protocol
1. Add 10 µL of the the phage library in 100 µL TBS to the internal control well (Streptavidin-coated, blocked) and incubate at RT for 30 min, gently shaking.
2. Transfer the unbound phages by pipetting to a centrifuge tube containing the target, diluted to a final concentration of 50 mg/mL in NaHCO3. Incubate at RT for 30 min, gently shaking.
3. Transfer the precomplexed phage-target solution to a streptavidin-coated well. Incubate at RT for 30 min, gently shaking.
4. Transfer the supernatant to another streptavidin coated well and refill the well with NaHCO3 . Incubate at RT for 30 min, gently shaking. supernatant? or unbound phages as in step 2? clarify!
5. Wash 15 times with TBST 0.1-0.5%.
6. Add 150 µL of elution buffer to all wells but the internal control well and incubate for 10-60 min at RT, gently shaking (max. 20 min if using the glycine elution buffer).
7. Neutralize by adding 25 µL TRIS-HCL pH 9.1 to the wells.
8. Transfer the eluates from both wells to a centrifuge tube and store at 4°C.
Mix for colony PCR:
| Green Dream Taq Mastermix | 25 µL |
| Nuclease free water | 15 µL |
| Primer 10 µM M13KE for (GCTAAGTAACATGGAGCAGG) | 5 µL |
| Primer 10 µM M13KE rev (CCCTCATAGTTAGCGTAACG) | 5 µL |
Pick plaques with a pipette tip, shortly swirl in the PCR tubes, fill with the PCR Mastermix and then place in a culture tube containing 1 mL of an 1:100 dilution of an overnight E. coli culture for amplification.
| step | temperature (°C) | time | repetitions |
| initial denaturation | 95 | 10min | - |
| denaturation | 95 | 30s | 35x |
| annealing | 48 | 40s | 35x |
| elongation | 72 | 60s | 35x |
| final elongation | 72 | 7min | - |
20 μL of PCR product should be loaded on a 1-1.5% agarose gel to have a good band (approx. 700kb).
Following the phage display a phage ELISA is performed to verify the binding of the individual phage clones to the target. This is necessary because it cannot be excluded that due to stochastic imbalance of the library or simply by chance some unspecific binders were enriched, even with an internal negative control in place and extensive washing steps. Those phage clones might express peptides that do not bind to the target but rather to streptavidin, polystyrene or one of the blocking agents that are present in the phage display selection wells. This phenomenon is well-characterized and some amino acid motives of streptavidin, BSA and polystyrene binders are described in the literature. While a comparison of these motivs with the sequences of the potential peptide ligands, that were obtained from the phage display is certainly reasonable, this should be verified with a wetlab experiment.
To identify the phage clones that bind selectively to the target, their binding to a plate coated with target was compared to their binding to plate without target. The coating strategy was analogous to the phage display ( streptavidin +/- biotinylated target, blocking with BSA) The phages werde detected with an anti M13 antibody coupled to a horseradish peroxidase (HRP) .
Protocol
Coating of plates:
Day 1:
1. Coat desired wells with strep o/n, 4°C
Day 2:
2. Slap off strep solution, coat every second well with D-PSMα3 o/n, 4°C
Fill wells without target with NaHCO3, use as negative control
3. Incubate 6 μL of each e.coli clone in 250 μL LB o/n
Day 3:
4. Block plates as before 1 h 30 min with BSA
5 .Wash plates 4 times with 300 μL TBST 0.1%
6. Pellet cells at 12,000 g, 3 min, 4°C
7. Add 100 μL of supernatant (containing phages) to target coated well and incubated for 1 h 30 min at 37 °C / on shaker at RT
8. Wash plate 6 times with 150 μL TBST 0.1%
9. Add 100 μL of HRP-conjugated anti-M13 antibody (1:5000, Sino Biological, Inc., Beijing), incubate for 2 h at 37°C
10. Wash plates 3 times with 150 μL TBST 0.1%
11. Add 100 μL of TMB
12. Stop reaction with 50 μL 2 M sulfaric acid after about 3-4 min (or desired color)
13. Use a platereader to detect absorbance at 450 nm
After amplification of the phages in ER2738 the phages need to be extracted and purified from the bacterial suspension. This is done with two serial PEG precipitations.
Protocol
1. Transfer the culture to a centrifuge tube and spin for 10 min at 12,000 g at 4°C.
2. Transfer the supernatant to a fresh tube and discard the pellet. Re-spin 10 min at the same velocity.
3. First PEG-precipitation:
3.1 Transfer the upper 80% of the supernatant to a fresh tube and add 1/6 volume of 20% PEG/2.5 M NaCl. Allow the phage to precipitate at 4°C for at least 2 h, preferably o/n.
3.2 Spin the PEG precipitation at 12,000 g for 15 min at 4°C.
3.3 Discard the supernatant, even if the pellet is not visible. Respin for 15 min at same velocity.
3.4 Discard the remaining supernatant with a pipette, carefully to not disturb the pellet.
3.5. Suspend the pellet in 1 mL / 300 μL of TBS to solubilize the phages again, let solution shake for 1 h 30 min at 4°C. Vortex for 10 seconds. Transfer eluate to a microcentrifuge tube.
To isolate the phages that express target-binding ligands from our library of 1011 different phages we performed a two-step protocol. First we added the phages to a well which was coated with streptavidin and blocked with BSA (or gelatine) but lacks the target. All phages that are specific for streptavidin or BSA (or gelatine) should should be captured in this well as a negative selection. The phages remaining in the supernatant after this negative selection were then transferred to the target-coated well. Nonbinding phages were washed away and the remaining, target-bound phages were eluted. For elution we used either a chaotropic glycine buffer that dissolves all protein-protein interactions in an unspecific way.
Protocol
1. Add 10 µL of the the phage library in 100 µL TBST, 0.1% to the internal control well (streptavidin coated, blocked) and incubate at RT for 30 min, gently shaking.
2. Transfer the unbound phages by pipetting to the emptied target-coated well, incubate at RT for 1 h, gently shaking.
3. Wash 15 times with TBST 0.1-0.5%.
4. Add 150 µL of elution buffer to the target-coated well and incubate for 10-60 min at RT, gently shaking (max. 20 min if using glycine elution buffer).
5. Neutralize by adding 25 µL TRIS-HCL pH 9.1 to the wells.
6. Transfer the eluate to a centrifuge tube and store at 4°C.
To determine the titer (amount of plaque forming units, = infectious phages, per microliter) of the eluates we make serial dilutions and add the diluted phages to an excess of LacZ-/- E-Coli (MOI<1). This ensures that each bacterium is only transfected with one phage. Streaked out on Agar Plates each phage-infected E-coli will form a "plaque" containing the same genotype of phages, which can be distinguished by a diffuse area of diminished growth (as M13 are non-lytic phages). The phages of the library and thus the infected E.coli contain the LacZ reportergene, which results in a blue colour of the plaques. Unlike them uninfected LacZ-/- E.Coli and also plaques formed by potential contaminating phages from the environment will appear colourless. Counting the number of plaques (equalling the number of phages that originally infected an E.coli and founded a colony) for each dilution helps to determine the titer. Dilutions are necessary to have plates with a countable number of plaques.
Protocol
1. Preparations:
1.1 Inoculate a culture of E.coli by picking a clone from the ER2738 plate and inoculating it in 10 mL LB medium. Let it grow, shaking at 200 rpm at 37°C until the OD600 of 0.5 for later use.
1.2 Prepare LB Agar Plates with X-Gal/IPTG (40 mg/mL X-Gal; 0.133 mM IPTG) by adding the IPTG/X-Gal Solution to the plates and letting it settle. Prewarm plates at 37°C for at least an hour.
1.3 Melt Top Agar and pour 3 mL each into a 15 mL centrifuge tube and store it at 45°C to keep it liquid.
2. Dilute the phage solutions appropriately in LB medium (10-1 to 10-4 is recommended for unamplified eluates, 10-5 -10-10 for amplified eluates)
3. Add 10 μL of phage dilutions to 200 μL of the ER2738 culture for infection and vortex, let the infection take place for at least 2 minutes at most 5 minutes.
4. Add the 210 μL of infected ER2738 to the Top Agar, vortex again and quickly pour it onto the IPTG /X-Gal plates. Let the plates cool down for 10 min.
5. Incubate at 37°C overnight.
Evaluation of titers:
Next day: Count the blue plaques on each plate. If plates are overfilled with blue plaques make an approximation. Use plates with 50-100 plaques to calculate the titer. If there are more than one calculate the arithmetic mean. The amount of plaques equals the number of phages in 10 μL of the diluted phage solution.
Protocol
1. Slap off the remaining phage solution and slap the plates on fresh paper towels to remove the remaining liquid. Be careful to avoid cross contamination between the wells/plates and use a new paper towel for each slap.
2. Add 300 µL of TBST (0.1% - 0.5%) and swirl the plate gently.
Repeat several times.
One of the most accurate methods to analyse the interaction of two molecules with minimal influence of the assay conditions is Bio Layer Inferometry and can be performed using the instrument OctetRED. The principle of Bio Layer Inferometry is based on optical measurements of the refractory index of a sensor which is coated with a target protein. If another molecule attaches to this sensor the optical thickness of the layer atop the sensor tip increases and so does its refractory index. This leads to a shift in wavelength which can be measured by the OctetRED.
Sample preparation:
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-
Calcium is an intracellular second messenger which plays a crucial role in the signalling of almost every celltype and has a role in many important cellular functions. For example muscle contraction or neurotransmitter exretion upon nervous stimulus and immune cell degranulation rely on calcium dependent signalling pathways. Upon certain upstream stimuli, like the activation of a receptor, calcium ions are released from the endoplasmatic reticulum. This Ca2+ flux causes a spike in the usually low cytosolic calcium concentration which can in turn activate calcium dependent proteins that execute functions such as exocytosis or cell migration. Generally speaking the cytosolic Ca2+ levels can be interpreted as a marker for the receptor-mediated activation of a cell by exogenous stimuli.
To visualize the levels of calcium that are present in a cell one can make use of the properties of Indo-1, a fluorophore which changes its emission spectrum from approx. 475 nm to 400 nm in the presence of Ca2+ ions. The ratio between Ca2+ bound an unbound Indo-1 can be visualized by measuring the emission at the different wavelengths via FACS and serves as a readout for the cytosolic Ca2+ concentration.
Protocol
- harvest a number of cells that express the receptor of interest according to the setup and timecourse of the experiment. For one minute of measurement at the standard flow rate of the cytometer 500 000 cells are needed. For experiments like the ones described here approximately 7 minutes of analysis are fitting.
- centrifuge the cells at 300 rpm for 3 minutes and resuspend them in 600 uL RPMI with 10% FCS in a centrifuge tube.
- add 1 uL pluronic acid and 0.8 uL of Indo-1 per 106 cells and incubate for 30 - 60 minutes. Flip the tube every 15 minutes but do not vortex.
- centrifuge the cells at 300 rpm for 3 minutes and resuspend them in 100 uL RPMI with 1% FCS per 106 cells.
- prepare the stimuli to be tested at appropriate concentrations, including the unspecific ionophore ionomycine at a final concentration of XXXXX as a positive control.
- dilute the cells needed for one sample in 1,5 mL RPMI with 1% FCS, prewarmed to 37 °C.
Measurment
- place the sample in the FACS and measure the baseline of the cells for 2 minutes without any stimulus
- at a set timepoint add the stimulus of interest in a volume of at least 40 uL and mix well using a syringe
- measure the cellular response to the stimulus for 3 to 4 minutes
- add ionomycine and measure the positive control response for 1.5 to 2 minutes
To research natural processes it is best to simulate the natural conditions in the closest was that is possible. Mammalian cells can be grown outside of their natural environment. They undergo a process of different adaptations to the culture conditions which is called immortalization but they can still be analyzed as a reliable model for studying cellular processes and the reaction of cells to their environment and different compounds such as for example toxins.
Adherent HEK293T cells and nonadherent Jurkat T-cells were cultured in cell culture treated 10cm polystyrene dishes with 10 mL RPMI media supplemented with 10% FCS and HEPES (1:100). The cells were splitted every 2-3 days according to their confluence and the experiments that were planned. All cells were kept in an 37°C, 5% CO2 Incubator.
Transfection
Transient transfection is a means to induce the expression of a desired gene in mammalian cells. It is possible to study the function of a gene or protein of interest, for example its role in cellular signalling cascades, with this model. A gene of interst is introduced on a plasmid which should be equipped with a mammalian promotor and it should be codon optimized for mammalian cells. Since the plasmid is not integrated in the genome of the cells the expression of the gene of interest is lost after several days.
Protocol:
- On the day before transfection split HEK293T cells with a confluence of 80% 1:6 and let them grow to 60% confluence overnight.
- on the next day, feed the cells with RPMI containing 2% FCS and incubate for 2 hours
- Prepare the transfection mix of 520 uL RPMI without supplements, 30 uL PEI and 5 ug plasmid DNA in a 15 mL reaction tube, vortex twice and let it incubate for 10 min.
- Add the transfection mix dropwise to the dish with HEK cells and take care to distribute it evenly
- gently swirl the cell culture dish
- incubate the cells for 3 - 8 h
- Replace the medium with 10% FCS RPMI medium.
Cells were incubated at minimum 36h after transfection before analysis.
Fluorescence assisted cell sorting is used to determine the surface expression of selected antigens in a cell population. This is achieved by staining cells with fluorescence - coupeled antibodies which bind specifically to the selected antigen. The analysis of multiple surface expression patterns at once is made possible by the use of different fluorphores with non overlapping emission spectra that can be excited by lasers of different wavelength. FACS analysis allows a complex analysis of different cell populations according to their surface markers on single-cell basis.
Protocol
- Harvest approximately 105 to 106 cells, spin them down at 300 g for 3 minutes
- add PBS and spin down again at the same speed (washing)
- repeat the washing step
- Add the appropriate dilution of the antibody (1:100 for anti FPR2-APC) to the cellsuspension in PBS
- let incubate for 20 -30 min on ice
- wash the cells twice with PBS
- resuspend the cells in 500 uL PBS for measuring fluorescence at the FACS
When researching toxins their capability to lyse mammalian cells is of vital interest. To assess celldeath upon exposure to the toxin a sensitive method is needed. Dying cells release the mcontents of their cytoplasm, amongst others the ubiquitous enzyme lactate dehydrogenase, LDH which is roughly proportional to the amount of dead cells. In the LDH Cytotoxicity assay the LDH which is released by dying cells catalyzes the reaction of NAD+ to NADH. This serves then as a substrate for the diaphorase, an enzyme which can then in turn convert INT to the coloured product formazane (diaphorase and INT both provided by the CyQUANT™ LDH Cytotoxicity Assay (Invitrogen™)). The absorbance of formazane at a wavelength of 490 and 680 nM can be measured using a plate reader.
Protocol
- seed 105 Jurkat cells in 500 uL RPMI in a 12-well plate
- add the toxin of interest to the desired concentration and incubate for 1 h at 37°C. As negative control use the solvent of the toxin. Any detergent in high concentration can serve as positive control.
- centrifuge at 300g, 3 minutes.
- transfer 50 uL of the supernatant into a flat bottom 96 well plate
- proceed according to the manufacturers instructions
- measure absorbance at 490 and 680 nM in a plate reader
The literature indicates that peptides made from D-aminoacids, in contrast to their L-spiegelmers, cannot be degraded by proteases because of their mirrored orientation. In order to test the degradability of a protein by a protease a straightforward degradation assay can be performed. Both enantiomers are incubated with the unselective protease proteinase K. The result, undegraded protein or protein fragments can be visualized by SDS Page.
Protocol
- Add XXXXmg/mL Proteinase K to a microcentrifuge tube of XXXXmg/mL of the protein of interest
- Incubate for 1 - 24 h at 37°C
- To inactivate the Proteinase K and prevent further digest beyond the desired timepoint add XXXX Phenylmethylsulfonylfluorid (PMSF), XXXXX mM dissolved in ethanol
- Incubate at RT for 1 minute
- Denature all protein components of the digest for 5 min at 92°C.
After an Page the bands that were separated from a protein sample need to be visualized. The usual method for this is western blotting which allows to visualize only one protein in a selective manner by using an antibody to detect it. However for visualizing the products of a digest or a protein for which there is no detection antibody available the silver staining is an ideal method because it stains all proteins, regardless of their sequence. This relies on a chemical reaction of the silver ions from the silver nitrate buffer which is added to the gel with negatively charged sidechains of proteins. Upon addition of formaldehyde the silver ions are reduced to elementary silver particles which precipitate within of the protein bands and there by make them visible.
Silver staining is a very sensitive method which can detect bands down to 1ng of protein mass. Aditionally it is a very fast process preventing the diffusion of protein bands during the staining. Therefore it is perfectly suited to visualize small protein bands.
Follow the instructions of the manufacturer of the Thermo Scientific™Pierce Silver Stain Kit™ or a comparable product.
Transient transfection is a means to induce the expression of a desired gene in mammalian cells. It is possible to study the function of a gene or protein of interest, for example its role in cellular signalling cascades, with this model. A gene of interst is introduced on a plasmid which should be equipped with a mammalian promotor and it should be codon optimized for mammalian cells. Since the plasmid is not integrated in the genome of the cells the expression of the gene of interest is lost after several days.
Protocol:
- On the day before transfection split HEK293T cells with a confluence of 80% 1:6 and let them grow to 60% confluence overnight.
- on the next day, feed the cells with RPMI containing 2% FCS and incubate for 2 hours
- Prepare the transfection mix of 520 uL RPMI without supplements, 30 uL PEI and 5 ug plasmid DNA in a 15 mL reaction tube, vortex twice and let it incubate for 10 min.
- Add the transfection mix dropwise to the dish with HEK cells and take care to distribute it evenly
- gently swirl the cell culture dish
- incubate the cells for 3 - 8 h
- replace the medium with 10% FCS RPMI medium.
Cells were incubated at minimum 36h after transfection before analysis.
Protocol:
- Pick your colony from the agar plate with a pipette tip and press it into bottom of the PCR tube.
- Prepare 1:1 mix of primers (100 nM) and Q5/Phusion Master Mix.
- Put 6 uL of mixture in each PCR tube (on ice).
- Spin down 60 s, 14000 rpm
- Run PCR with your preferred protocol, depending on the plasmid you want to amplify..
- Analyze with gel electrophoresis.
Protocol:
- Use 50 ng of backbone, calculate mg of DNA needed of the remaining constructs with NEBuilder.
- Fill up with nuclease free water up to 5 µl. Add 5µl of HiFi Ligase master mix (NEB).
- Incubate for 15-50 minutes. If more than 3 parts will be ligated, always incubate 50 minutes.
- Transform the ligated constructs into competent bacteria.
Protocol
- Prepare an agarose gel with amount relative to concentration for the fragment (0.7-2 % (w/v)) in TAE buffer.
- Heat mixture until fully dissolved.
- Add 3-6 μL of DNA stain.
- Cast the solution in a gel chamber, add comb and wait 20-30 mins, until it has completely solidified.
- Add loading buffer to each of your DNA samples.
- Transfer gel into gel box and cover gel with TAE buffer. Load the gel with your sample (10-30 μL).
- Let it run at 120V for 60 minutes.
- Visualize DNA fragments under UV illuminator.
Ligation:
- Add 17 µl extracted PRC product into a PCR reaction tube.
- Add 2 µl 10X T4 DNA Ligation Buffer.
- Add 1 µl T4 DNA Ligase.
- Incubate overnight at 4°C.
- Restriction Digest:
- In a PCR reaction tube, add:
-Template DNA (4-5 μg)
-0,5 μL Restriction Enzyme of choice
-10x Cutsmart Buffer (NEB)
-fill up with dH2O to 40 μL - Incubate for one hour at 37°C.
Protocol
Miniprep was performed with ZymoPURE Plasmid Miniprep Kit from Zymo Research.
- Centrifuge 0.5-5 ml of bacterial culture in a clear 1.5 ml tube at full speed for 15- 20 seconds in a microcentrifuge. Discard supernatant.
- Add 250 μl of ZymoPURE P1 (Red) to the bacterial cell pellet and resuspend completely by vortexing or pipetting.
- Add 250 μl of ZymoPURE P2 (Green) and immediately mix by gently inverting the tube 6-8 times. Do not vortex! Let sit at room temperature for 2-3 minutes. Cells are completely lysed when the solution appears clear, purple, and viscous.
- Add 250 μl of ice cold ZymoPURE P3 (Yellow) and mix thoroughly by inversion. Do not vortex! Invert the tube an additional 3-4 times after the sample turns completely yellow. The sample will turn yellow when the neutralization is complete and a yellowish precipitate will form.
- Incubate the neutralized lysate on ice for 5 minutes.
- Centrifuge the neutralized lysate for 5 minutes at 16,000 x g.
- Transfer 600 μl of supernatant from step 6 into a clean 1.5 ml microcentrifuge tube.Be careful not to disturb the yellow pellet and avoid transferring any cellular debris to the new tube.
- Add 275 μl of ZymoPURE Binding Buffer to the cleared lysate from step 7 and mix thoroughly by inverting the capped tube 8 times.Place a Zymo-Spin II-P Column in a Collection Tube and transfer the entire mixture from step 8 into the Zymo-Spin II-P Column.
- Incubate the Zymo-Spin II-P/Collection Tube assembly at room temperature for 2 minutes and then centrifuge at 5,000 x g for 1 min. Discard the flow through1.
- Add 800 μl of ZymoPURE Wash 1 to the Zymo-Spin II-P Column and centrifuge at 5,000 x g for 1 min. Discard the flow through.
- Add 800 μl of ZymoPURE Wash 2 to the Zymo-Spin II-P Column and centrifuge at 5,000 x g for 1 min. Discard the flow through.
- Add 200 μl of ZymoPURE Wash 2 to the Zymo-Spin II-P Column and centrifuge at 5,000 x g for 1 min. Discard the flow through.
- Centrifuge the Zymo-Spin II-P Column at ≥ 10,000 x g for 1 minute in order to remove any residual wash buffer.
- Transfer the Zymo-Spin II-P Column into a clean 1.5 ml tube and add 25 μl of ZymoPURE Elution Buffer directly to the column matrix. Incubate at room temperature for 2 minutes, and then centrifuge at ≥ 10,000 x g for 1 minute in a microcentrifuge. Store the eluted plasmid DNA at ≤ -20°C.
For polymerase mastermixes of Phusion Flash and Q5:
Use 2.5 µL of each primer, 1 µl DNA (or 2 µl if concentration or purity are low). Fill up to 25 µl with nuclease free water, then add 25 µL of Phusion Flash or Q5 NEB master mix.
Calculate the annealing temperatures of the primers with TM-Calculator from NEB.
Calculate elongation time depending on your Polymerase. Q5 needs about 30 seconds for 1 kb of DNA amplification, while Phusion Flash needs around 15 seconds. Add around 15 seconds extra time on your calculated timespan.
Denaturation temperature should be set between 94°C and 98°C, elongation temperature is 72°C.
Troubleshooting:
If the PCR does not work, or Primer dimers are prominent, add 1-2 µL DMSO or MgCl to the PCR mix. Adjust Water correspondingly.
Also recommendable: Elevate the template DNA concentration to 2-3 µL (max 2-3 ng).
Protocol
-
-Into a PCR reaction tube add:
-a 3:1 relation of Insert to Backbone(150ng)
-1.5 µL T4 DNA Ligase Buffer
-1µL BsaI
-1µL T4 DNA Ligase
-fill up with dH2O to 15 µL
|
Step |
Temperature |
Time |
|
Restriction x25 Ligation |
37°C
16°C |
3 min
4 min |
|
Final Restriction |
50°C |
5 min |
|
Denaturation |
80°C |
5 min |
|
Hold |
4°C |
- |
Protocol
- Thaw Bacteria on ice (Top 10 competent E. Coli (Ag Di Ventura, BIOSS)).
- Add 2µl of the assembly mix to the bacteria and flick (do NOT vortex, since competent bacteria have fragile membranes).
- Incubate on ice for 20-30 minutes (best is 25).
- Heat shock for 45 seconds at 42°C and put back on ice for 2 mins.
- Add 500 µL LB medium without antibiotics (give the bacteria time to express the resistance gene).
- Make sure the LB medium is transparent and has not been contaminated.
- Then incubate on shaker at 37°C for 45-60 minutes, but never longer.
- Start warming the plates at 37°C (if the plates have chloramphenicol, it is recommendable to heat them longer, around 60 mins).
- Plate 50-200 µL (rather 200).
Protocol
For sequencing purify your DNA from a PCR or a agarose gel electrophoresis.
Gel extraction was performed with the QIAquick PCR Purification Kit from Qiagen:
- Excise the DNA fragment from the agarose gel with a clean, sharp scalpel.
- Weigh the gel slice in a colorless tube. Add 3 volumes Buffer QG to 1 volume gel (100 mg gel ~100 μl). The maximum amount of gel per spin column is 400 mg. For >2% agarose gels, add 6 volumes Buffer QG.
- Incubate at 50°C for 10 min (or until the gel slice has completely dissolved). Vortex the tube every 2–3 min to help dissolve gel. After the gel slice has dissolved completely, check that the color of the mixture is yellow (similar to Buffer QG without dissolved agarose). If the color of the mixture is orange or violet, add 10 μl 3 M sodium acetate, pH 5.0, and mix. The mixture turns yellow.
- Add 1 gel volume isopropanol to the sample and mix.
- Place a QIAquick spin column in a provided 2 ml collection tube or into a vacuum manifold. To bind DNA, apply the sample to the QIAquick column and centrifuge for
- 1 min or apply vacuum to the manifold until all the samples have passed through the column. Discard flow-through and place the QIAquick column back into the same tube. For sample volumes >800 μl, load and spin/apply vacuum again.
- If DNA will subsequently be used for sequencing, in vitro transcription, or microinjection, add 500 μl Buffer QG to the QIAquick column and centrifuge for 1 min or apply vacuum. Discard flow-through and place the QIAquick column back into the same tube.
- To wash, add 750 μl Buffer PE to QIAquick column and centrifuge for 1 min or apply vacuum. Discard flow-through and place the QIAquick column back into the same tube.Note: If the DNA will be used for salt-sensitive applications (e.g., sequencing, blunt- ended ligation), let the column stand 2–5 min after addition of Buffer PE.
- Centrifuge the QIAquick column in the provided 2 ml collection tube for 1 min to remove
- residual wash buffer.
- Place QIAquick column into a clean 1.5 ml microcentrifuge tube.
- To elute DNA, add 50 μl Buffer EB (10 mM Tris·Cl, pH 8.5) or water to the center of the QIAquick membrane and centrifuge the column for 1 min. For increased DNA concentration, add 30 μl Buffer EB to the center of the QIAquick membrane, let thecolumn stand for 1 min, and then centrifuge for 1 min. After the addition of Buffer EB to the QIAquick membrane, increasing the incubation time to up to 4 min can increase the yield of purified DNA.
- If purified DNA is to be analyzed on a gel, add 1 volume of Loading Dye to 5 volumes of purified DNA. Mix the solution by pipetting up and down before loading the gel.
- To process a larger sample (>100 μl), either increase proportionally the amount of Binding Buffer (Step 1), or divide the larger sample into several 100 μl aliquots and process each as a separate sample.
- 1 After PCR is finished, adjust total volume for each PCR tube (reaction components + DNA product) to 100 μl: – Add 500 μl Binding Buffer to each 100μl PCR tube.
- Mineral oil or wax does not need to be removed from the PCR solution before adding the Binding Buffer
- Mix sample (Binding Buffer + PCR solution) well.
- Insert one High Pure Filter Tube into one Collection Tube.
- Transfer the sample from step 1 using a pipette to the upper reservoir of the Filter Tube.
- Centrifuge 30 – 60 s at maximum speed in a standard table top centrifuge at +15 to +25°C.
- Disconnect the Filter Tube, and discard the flowthrough solution. – Reconnect the Filter Tube to the same Collection Tube.
- Add 500 μl Wash Buffer to the upper reservoir.
- Centrifuge 1 min at maximum speed (as above).
- Discard the flowthrough solution.
- Recombine the Filter Tube with the same Collection Tube. – Add 200 μl Wash Buffer.
- Centrifuge 1 min at maximum speed (as above).
- This second 200 μl wash step ensures optimal purity and full removal of Wash Buffer from the glass fibers.
- Discard the flowthrough solution and Collection Tube.
- Reconnect the Filter Tube to a clean 1.5 ml microcentrifuge tube
- Add 50 – 100 μl Elution Buffer to the upper reservoir of the Filter Tube.– Centrifuge 1 min at maximum speed.
- Do not use water for elution since alkaline pH is required for optimal yield.
- The microcentrifuge tube now contains the purified DNA.
- When subsequent OD260 determination is planned, centrifuge the eluate for more than 1 min
- at maximum speed to remove residual glass fibers from the eluate, because they may disturb absorbance measurement. Use an aliquot of the supernatant to determine concentration.
- Either use the eluted DNA directly or store the eluted DNA at +2 to +8°C or −15 to −25°C for later analysis.
Protocol was performed with the High Pure PCR ProductPurification Kit from Roche
Expressions were carried out in 25 µl reactions and assembled on ice in RNase-free tubes with 10 µl of solution A, 7.5 µl of Solution B, 1 µl RNase-Inhibitor, 250 ng/µl DNA template and the necessary amount of nuclease-free water. Coupled transcription and translation happened during an incubation of 2 hours at 37°C.
Cells were grown in 500 mL TB medium in a 2 L Erlenmeyer flask and incubated at 30° C and 220 r.p.m. to OD600 of 2.5. Cells were pelleted by centrifugation for 15 min at 5000 × g and 4 °C, washed three times with cold S30 buffer (10 mM tris-acetate pH 8.1, 14 mM magnesium acetate, 60 mM potassium acetate, 2 mM dithiothreitol), and stored at − 80 °C. To make cell extract, the thawed cells were suspended in 0.8 mL of S30 buffer per 1 g of wet cell mass and lysed by sonication using the Diagenor Bioruptor Next Gen Sonicater 3 x 6 cycles, each 30 seconds. To minimize heat damage during sonication, samples were placed in an ice-water bath. The extract was then centrifuged at 12,000 g and 4°C for 10 min. The supernatant was flash-frozen using liquid nitrogen and stored at -80°C until use.
Components and amounts of the components were adjusted in different reactions. A 15 μL CFPS reaction was prepared in a 1.5 mL microcentrifuge tube by mixing the following components: 1.2 mM ATP, 13.3 ng/µl plasmid, 10 mM dithiothreitol, 1 µl RNase Inhibitor and 27% of cell extract.
This expression was carried out with cell lysates derived from MG1655 grown in LB, MG1655 grown in LB and C321.∆A grown in TB medium
CFPS reactions containing additional PURExpress components were prepared in a 1.5 mL microcentrifuge tube by mixing the following components: 1 µl RNase-Inhibitor, 13.3 ng/µl plasmid, 5 µl solution A PURExpress, 3.5 µl solution B PURExpress and 27% of cell extract.
The CFPS reaction with arabinose induction contained additionally 57 mM HEPES and 1% arabinose.
Expression for BLADE promotor was induced by a 20 hours incubation at 30°C under 450 nm blue light and an intensity of 90 and for pBAD only 20 hours incubation at 30°C.
Presence of proteins expressed with PURExpress® was detected by SDS gel electrophoresis with TGX 12% Tris-Glycine gels and 16.5% Tris-Tricine gels, ran at 200 V for 30 minutes. Gels were stained with Instant Blue for 1 hour. Active full-length sfGFP protein yields were quantified by measuring fluorescence in a 6% native gel with an Amersham Typhoon laser scanner.
DNA Amplification
The PCR is an essential technique used in molecular biology to rapidly generate numerous copies
of a DNA sequence. In multiple cycles, the template DNA is first denatured at approx. 95 to
separate it into two single strands. Next, the temperature is lowered so that primers can specifically
anneal to the 3’ end of each strand. A heat-resistant DNA polymerase can now bind to
this complex and initiate elongation using deoxynucleotidetriphosphates (dNTPs) supplied in
the reaction mixture. Once elongation is complete, the reaction can be repeated, now also using
the newly generated DNA sequences. This way, 2 n DNA copies can be generated for n cycles.
In this work, the Q5 ®High-Fidelity DNA Polymerase was used, due to a low error rate and
high processivity [Wang et al., 2004] to amplify gene inserts and plasmids for cloning procedures.
The reaction mix was prepared according to an adapted protocol from NEB 5:
| Volume [μl] | Component | Final Concentration |
|---|---|---|
| 25 | Q5 High-Fidelity 2X Master Mix | 1X |
| 1 | DNA template | |
| 1 | forward primer | 0.5 [μM] |
| 1 | reverse primer | 0.5 [μM] |
| 22 | ddH 2 O |
The Master Mix contains dNTPs, Mg 2+ and a broad-use buffer. Reaction mixture must be
prepared on ice.
The PCR reaction was carried out using a Biometra TOne 96 6 Thermocycler using the following
protocol:
Steps 2-4 were repeated for 30 cycles.
| cycle step | temperature [C] | time [s] |
|---|---|---|
| Initial Denaturation | 98 | 30 |
| Denaturation | 98 | 5 |
| Annealing | according to NEB T m Calculator | 30 |
| Elongation | 72 | 30/kB |
| Final Extension | 72 | 120 |
| Hold | 16 | 1 |
Site-directed Mutagenesis
PCR can not only be used to amplify DNA templates, but also to introduce specific mutations to
the amplicon by altering the sequence of a primer complementary to the desired mutation site.
The resulting DNA will then also possess the altered sequence.
New Sequences can be inserted as well by addition to the 3’ end of the primer during synthesis. SDM is therefore
a useful tool for the creation of overhangs needed for cloning, and also to introduce overlapping
regions to a newly assembled plasmid, which automates re-circularization.
In this work, SDM was mainly utilized to introduce the 15 - 80 bp overhangs necessary for
Gibson Assembly, and also to introduce stem loop structures and Amber stop codon (ACS)
mutations to pBAD33 inserts. All oligonucleotide sequences used for SDM are marked in blue
in the plasmid listing.
DNA Digestion
Prior to plasmid transformation into bacterial cells, it is essential to remove what is remaining of
the template DNA, since otherwise it can be tedious to locate the successfully cloned construct
in a mix of transformed colonies. For this purpose, DPn1 restriction digestion was used, since
the restriction enzyme only targets methylated cleavage sites. DNA methylation occurs inside
the E.coli cell, and only the original template, but not the synthetic amplicons, will therefore
be digested. Digestion was performed by:
1. Either Adding 0.5 μl DpnI (NEB) directly to reaction mix after Gibson Assembly
2. Or Adding 0.5 μDpnI (NEB) and 1.5 NEB CutSmart Buffer to purified PCR product
The reaction mix was prepared on ice and incubated at 37 °C for 1 hour.
Ligation
In cases where no overlaps are added during SDM, the blunt ends of a plasmid must be ligated
to recover the circularized state. In this work, the T4 Ligase was used to join phosphorylated
ends of linearized amplicons by adapting the NEB T4 Ligation protocol:
volume [μl] component
1 T4 Ligase
2 T4 Ligase Buffer (10X)
17 DNA as PCR Reaction Product mix
Reaction mix was prepared on ice and incubated at room temperature for 2 hours prior to transformation.
Gibson Assembly
DNA inserts were cloned into their respective expression vectors using Gibson Assembly. This
isothermal cloning reaction is performed in one step, allowing for the seamless, easy construction
and joining of genes and genetic circuits, and is therefore often used in synthetic biology. The
assembly itself is best described as the consecutive activity of three enzymes.
1. The 5’ T5 Exonuclease removes nucleotides from the 5’ ends of the supplied doublestranded
gene fragments, creating one-stranded 3’ overhangs. The gene fragments must
be so designed that the revealed overhangs should be complementary to those of the gene
fragments, enabling the subsequent annealing of both strands. The Exonuclease is heat
inactivated shortly after.
2. a Phusion DNA Polymerase, which does not compete with digestion in the first step,
fixes any gaps left and proofreads the new double-stranded DNA sequence containing the
inserted gene fragment.
3. a Taq DNA Ligase finishes the process by sealing the nicks left in the sequence.
For all cloning processes in this study, the 2X NEBuilder HiFi DNA Assembly Master Mix ©,
has been applied following an adjusted protocol from NEB:
The Master Mix contains all enzymes described above in a broad-use buffer.
The reaction mix was prepared on ice and incubated at 50 °C for 30 minutes.
| volume [μl] | component | Final Concentration |
|---|---|---|
| X | Plasmid Vector | 50-100 ng/ μl |
| X | DNA insert | 150-300ng μl |
| 10-X | ddH2O | |
| 10 | NEBuilder HiFi DNA Assembly Master Mix | 1X |
PCR Purification
To directly purify DNA after a PCR reaction, the QIAprep Spin Miniprep Kit was used following
the respective Qiagen protocol:
1. Add 250 μl of buffer PB to PCR product.
2. transfer reaction mix to QIAprep 2.0 Spin Miniprep Columns, centrifuge for 60s, discard
flow-through
3. Add 750 μl Buffer PE for Washing, centrifuge for 60s, discard flow-through
4. Transfer column to 1.5 ml Eppendorf Tube, add 25 μl buffer EB or 50 °C water for elution
5. Centrifuge for 60s, discard column
6. Measure DNA concentration
Centrifugation steps carried out at 14,000 PRM.
Transformation of Bacteria
The procedure of introducing recombinant DNA through transient expression vectors, also
known as plasmids, into bacterial cells is known as Transformation.
For each transformation in this work, chemically competent cells were taken from -80 °C storage
and transformed using the following protocol:
1. Keep competent cells and plasmid DNA on ice until fully thawed
2. Add 10ng of plasmid DNA for single / 200-300 ng for plasmid DNA for double retransformation
to competent cells. After cloning, add whole reaction mix to cells.
3. Incubate on ice for 15-20 minutes
4. Heat shock at 42 °Cfor 90 seconds
5. Incubate on ice for 5 minutes
6. Add 300 μl LB media to cells
7. Incubate at 37 °C degrees and 750 RPM for 1 hour
8. Spread cells on pre-warmed LB Agar plate containing relevant antibiotics
9. Incubate plates at 37 °C overnight
Culture growth
Bacterial liquid cultures were grown by transferring bacterial cells from a single colony into a
culture tube containing 2-5 ml of LB media with necessary antibiotics added. Cultures were
incubated at 37 and 220 RPM for 6-10 hours before plasmid purification.
Colony PCR
Recombinant DNA can not only be amplified from purified plasmid constructs, but also directly
from bacterial cultures. The initial heating step is sufficient for lysing the cell, releasing the
plasmid from the cytosol and making it readily available for amplification. Primers binding to
either the plasmid backbone or the DNA insert can be selected to check the presence of the insert,
its length and orientation, and also screen for multiple inserts. To extract this information,
the amplified DNA must later be analyzed via gel electrophoresis in the presence of a fitting
DNA size ladder. The reaction mix was prepared using the following protocol adapted from
NEB:
Volume [μl] Component Final Concentration
25 Q5 High-Fidelity 2X Master Mix 1X
1 forward primer 0.5 [μM]
1 reverse primer 0.5 [μM]
23 ddH 2 O -
| Volume [μl] | Component | Final Concentration |
|---|---|---|
| 25 | Q5 High-Fidelity 2X Master Mix | 1X |
| 1 | forward primer | |
| 1 | reverse primer | |
| 23 | ddH 2 O |
The reaction mix was transferred to tubes that each contained cells from one selected bacterial
colony.
The Master Mix contains dNTPs, Mg 2+ and a broad-use buffer. Reaction mixture was prepared
on ice.
The PCR reaction was performed in a Biometra TOne 96 8 Thermocycler using the following
protocol:
Steps 2-4 were repeated for 25 cycles.
| cycle step | temperature [°C ] | time [s] |
|---|---|---|
| Initial Denaturation | 98 | 30 |
| Denaturation | 98 | 5 |
| Annealing | according to NEB T m Calculator | 30 |
| Elongation | 72 | 30/kB |
| Final Extension | 72 | 120 |
| Hold | 16 | 1 |
Miniprep
To obtain a plasmid from the corresponding bacterial growth culture, the cells must be lysed,
and the pure plasmid DNA extracted from the cell debris. Here, the QIAprep Spin Miniprep
Kit was used according the following adjusted protocol from Qiagen.:
1. Centrifuge 2 ml of bacterial liquid culture for 60 seconds, discard supernatant
2. Resuspend bacterial pellet in 250 μl Buffer P1
3. Add 250 μl Buffer P2 and mix gently for Cell Lysis
4. Add 350 μl Buffer N3 and mix gently for Neutralization
5. Centrifuge for 60s
6. transfer supernatant to QIAprep 2.0 Spin Miniprep Columns, centrifuge for 60s, discard
flow-through
7. Add 750 μl Buffer PE for Washing, centrifuge for 60s, discard flow-through
8. Transfer column to 1.5 ml Eppendorf Tube, add 25 μl buffer EB or 50 °C water for elution
9. Centrifuge for 60s, discard column
10. Measure DNA concentration
Centrifugation steps carried out at 14,000 PRM.
Concentration Measurement
DNA concentrations were measured using a NanoDrop™One Microvolume UV/VIS Spectrophotometer
9, which employs spectrophotometrical measurement to determine concentrations via
light absorption. Nucleotides, and consequently DNA, absorb light at 260 nm proportionally to
their amount inside the sample. The concentration is calculated in comparison to a sample-less
blank. Purity is also determined by scanning for carbohydrates, phenols and protein at 230 nm
and 280 nm. 1.2 μl of the sample are required for measurement.
Sequencing
The nucleotide sequence of a newly constructed plasmid must be verified. Even if gel electrophoresis
delivers the correct size information, only the definitive sequencing can rule out that
gene fragment have been inserted incorrectly, and no part is missing or mutated, which would be
detrimental to the whole experiment. Sequences are typically analyzed using Sanger Sequencing
This method requires a DNA template, a Taq Polymerase, a mixture of
dNTPs and special ddNTP, which are not able to form a phosphodiester bond with the next
3’ dNTP due to their lack of an OH-group, and therefore cause the DNA strand to break off.
ddNTP are usually marked with distinctive fluorescent labels, so that the whole sequence can be
analyzed by running the amplified fragments of different sizes on an agarose gel and measuring
the fluorescent signals. All samples were sequence with the GATC SupremeRun Custom DNA
Sequencing Service10. Sample DNA was dissolved in double distilled water (ddH20) to a final
volume of 20μl and final concentration of min. 50 ng/μl. Accompanying primers were dissolved
to a final concentration of 10ng/μl in 20 μl ddH20.
Agarose Gel Electrophoresis
Agarose Gel Electrophoresis is a key method of molecular biology to separate RNA and DNA
molecules according to their respective size and charge. In this study, it was used to verify results
of PCR amplification, Gibson Assembly, Restriction Enzyme Digestion and Colony PCR. 1 %
agarose gels were prepared by weighting the necessary amount of agarose powder and dissolving
it in a stock of 0.5 % TAE buffer, afterwards heating the mixture until the liquid was clear. This
stock can now be stored at 60 °C. To cast a gel, a plastic mold was filled with max. 100 ml of the
stock, depending on the needed gel size and thickness. 2.5 to 3.5μl of Ethidium Bromide (EtBr)
was added to the liquid, for visualization of DNA, which was then left to solidify for 20 minutes
with a comb inserted to leave space for loading pockets.
After the gel solidified, it was submerged in 0.5 % TAE buffer inside a Mini Gel II 11 gel running
chamber, and pockets were loaded with DNA samples (previously mixed with 6X DNA
Gel Loading Dye) and 1.5 μl GeneRuler 1 kb DNA Ladder 12. An electric current of 130V and
200mA was applied for 30 minutes to the gel, causing the negatively loaded DNA molecules
to move towards the positively charged anode of the chamber, with smaller fragments moving
faster through the gel than larger ones. After running, gels were analyzed and documented using
a UVP UVsolo-touch GelDoc System and a UVP Transilluminator 13. Sample volumes varied
depending on application:
1. Colony PCR, PCR Amplification Analysis prior to PCR Purification: 6 μDNA sample
mix, 1 μLoading Dye
2. Whole PCR Product or Restriction Enzyme Digestion Analysis: 50 μl DNA sample mix,
7 μLoading Dye.
Gel Extraction
To purify PCR products after agarose gel electrophoresis, the QIAprep Spin Miniprep Kit was
applied following the accompanying Qiagen protocol:
1. Cut bands carefully from gel using a UV Transilluminator and transfer into Eppendorf
tubes
2. Add 3 vol. buffer QG relative to weight of gel
3. Incubate at 50 °C for 10 min, vortex
4. Add 1 vol. of isopropanol relative to gel weight, mix
5. transfer samples to QIAprep 2.0 Spin Miniprep Columns, centrifuge for 60s, discard flowthrough
6. Add 750 μl Buffer PE for Washing, centrifuge for 60s, discard flow-through
7. Transfer column to 1.5 ml Eppendorf Tube, add 25 μl buffer EB or 50 °C water for elution
8. Centrifuge for 60s, discard column
9. Measure DNA concentration
Centrifugation steps carried out at 14,000 PRM.
Glycerol Stocks
Bacterial cultures transformed with valuable plasmids can be preserved for a long time as Glycerol
Stocks.
To prepare stocks, 0.5 ml of overnight bacterial culture and 0.5 ml of 50 % Glycerol (diluted in
ddH2O) were gently mixed in a screw top tube and stored at -80 °C.
Plasmid Induction Protocol
Genes expressed by the pULTRA plasmid vector are controlled by the LacI promoter, while
genes expressed from the pBAD plasmid vector are under the control of an AraC promoter.
The LacI promoter is typically induced with Isopropyl-beta-D-thiogalactopyranosid (IPTG), a
stable lactose analogue, the ara-Promoter with L-Arabinose. Both inductions follow a similar
mechanism: The induction substance binds to a repressor protein, which then changes its conformation
and detaches itself from an operator sequence, so transcription of the downstream
genes can commence.
Following induction protocol was devised for all experiments:
8 ml bacterial cultures containing the respective plasmids and antibiotics were grown overnight
at 30° C and 220 revolutions per minute (RPM) and diluted to an optical density (OD) 600 of
0.1 in the morning. OD600 was measured spectrophotometrically using light absorption at 600
nm. Cultures were again grown in same conditions for 2 to 4 hours until they reached the
exponential growth phase at 0.4 - 0.6 OD. After reaching this value, each individual culture was
split evenly, with one of the resulting samples being induced with 0.1 % L-Arabinose solution
(40 μl in 4 ml from a 10 percent stock solution) and 10 nM IPTG (8 μl in 4 ml from a 500nM
stock solution). Cells were again incubated at the abovementioned conditions. Incubation times
of 2, 6 and 10 hours were tested. After each inoculation, dilution and induction step, respective
samples were supplemented with 100nM D-Phenylalanine. Samples were finally prepared for
either microscopy, FACS or spectrophotometrical analysis.
Cell Washing Procedure
To eliminate autofluorescence and background signal, cells were washed prior to FACS analysis.
After the induction protocol was completed, bacterial liquid cultures were centrifuged at 14.7 x
10 3 RPM for 60 seconds, the supernatant was discarded, and the pelleted bacterial cells were
resuspended in 1ml of 1 % phosphate-buffered saline (PBS) solution to prevent osmotic lysis of
cells. After resuspension, cells were centrifuged again at the same conditions, the supernatant
was discarded, and cells resuspended in a final volume of 2 ml 1 % PBS solution.
Preparation of Microscopy Slides
To fixate the moving bacterial cells in one place and at the same time protect them from damage
and dryness, Agarose slides needed to be prepared for microscopic analysis. First, an 18 x 18
mm glass cover slip 14 was attached to one side of the opening of a custom metal microscopy
slide using industrial grease. The round opening was then filled with 10
percent Agarose dissolved in 10 percent PBS solution and left do try under another coverslips.
After 5 minutes, the coverslip was removed, and the dried Agarose gel covered with 10 μl of
bacterial liquid culture and again left to dry for approx. 15 minutes. Finally, the upper side of
the slide was again covered with a glass cover slip.
Microscopy
A Zeiss AxioObserver Wide-Field Microscope was used for microscopy analysis. All samples
were microscope with a 100x objective using Carl Zeiss Immersol™518 F Immersion Oil. In
a wide-field microscope, the entire specimen which is mounted on the stage is illuminated by
the light source, which allows for fast imaging. In return, concentrating on single regions of
interest and obtaining optical sections of a specimen, as it is done in a confocal microscope, is
not possible. Available light sources are either regular white light or fluorescence light sources
that excite fluorescent molecules at a specific wavelength. In this setup, images were taken using
1. Differential interference contrast (DIC) Microscopy, where polarized light from the light
source is split into two orthogonally polarized light rays, which pass through the sample at
slightly adjacent areas. Later, the two resulting images are recombined at a polarization
where the previously orthogonal rays can interfere again, and an image with brightened
and darkened spots resembling a three-dimensional image is generated due to the difference
in the optical path of both light rays.
2. Fluorescence microscopy, where a filtered light beam of a specific wavelength is used to
excite the fluorophores contained in the sample, which then absorb it and emit light of
a longer wavelength. The which was used to illuminate the sample must be filtered out
before it reaches the ocular, since it would otherwise overshadow the weaker light emitted
by the fluorophore.
Microscopy Analysis
The resulting images were analyzed using the ImageJ ver. 1.52p Image Analysis Software and resulting
data quantified with Microsoft Excel. Fluorescence intensity was obtained by calculating the mean intensity
of 50 measurements per image, with background intensities previously subtracted. Error bars calculated as standard errors.
Fluorescence Activated Cell Sorting
Fluorescence-activated cell sorting (FACS) can be employed to reliably characterize the properties
of a large quantity of cells and is also less susceptible to biased measurement than microscopy
analysis. In a Flow Cytometry device, a liquid suspension of cells is hydrodynamically focused
using a sheath fluid, so that only one cell can pass the laser light source placed horizontally
towards the screen each time. Fluorescence intensity can then be detected, as well as forward
and side scatter of the light, which correlates to cell size and granularity, respectively. This
way, different cell populations can later be discerned. For this study, a BD LSRFortessa ™cell
analyzer was used, set to an excitation wavelength of 488nm, an emission wavelength of 530nm
and a 30nm optical filter was used. 10000 cells were quantified per sample.
Data was analysed using FlowJo v.10.6 CL software.
