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Contents 1 Agarose Gel Electrophoresis 1.1 Pouring the Gel 1.2 Loading the Gel 1.3 Running the Gel 1.4 Staining the Gel 2 Cloning Protocols 2.1 Cloning Protocols: Restriction (double digest) 2.2 Cloning Protocols: Ligation 2.3 Cloning Protocols: Chemical Transformation 3 Competent Cells 4 Transmission Electron Microscopy, negative staining with 2 % PTA 4.1 Strains 4.2 Media and buffers 4.3 Execution 5 Library Selection 5.1 Attractants 5.2 Media 5.3 Execution 5.3.1 First round of selection 5.3.2 Second round of selection 5.3.3 Third round of selection 5.3.4 Plating of the selected clones 5.3.5 Minipreparation and sequencing of plasmid DNA 5.3.6 Retransformation of the plasmid DNA 5.3.7 Determination of the swimming behaviour of the freshly transformed BL21 cells 6 Overlap PCR for removal of restriction sites 7 Separation Assay 7.1 Strains 7.2 Media 7.3 Execution 7.3.1 Applying the mixed culture to the 0.3 % tryptone swimming agar plates 7.3.2 Separation of the different strains 8 Sequencing 9 Standard PCR 9.1 -> 50 μl / reaction: 9.2 Program: 10 Swimming/Chemotaxis Assay 10.1 Strains 10.2 Media 10.3 Execution 11 QuikChange Protocol 11.1 -> 20 μL / reaction: 11.2 Program: -> 1 min 96 °C 11.3 -> 5 min 72 °C -> Store at 4-8 °C 12 Quantitative Real-Time PCR 12.1 -> Experimental design 12.2 -> Primer for qrtPCR: 12.3 -> 20 μl / reaction: 12.4 Program: 12.5 -> Analysis via the 2–ΔΔCT (Livak) Method

Agarose Gel Electrophoresis

For the analysis of PCR-amplified products, agarose gel electrophoresis is the method of choice. This method takes advantage of the separation of DNA in dependance of the charge-mass ratio. The separation is based on the electric attraction of the negative charged DNA which is guided towards the positive charged anode upon application of a current. The PCR samples are run on agarose gels with different percentages according to the product sizes: small products run faster than bigger products. Later on, these fragments within the gel are made visible by examination under the UV light to ensure the correct DNA fragment length synthesized in the PCR reaction. Prior to UV analysis, a staining method of the DNA, here using ethidium bromide (EtBr), is obligatory. EtBr is an intercalating agent which embeddes itself within the DNA helix. Thus, the absorption spectrum is biased so that it is suitable for DNA detection. The determination of separated molecule sizes is done accodrding to a common DNA size standard.

Pouring the Gel

1 % agarose gels are standard to separate DNA. The percentages and thus the degree of polymerisation of the gel influences the degree of separation, i.e. PCR products of similar size can be distinguished by applying a percentage lower than 1 %. To pour a 1 % agarose gel weight 1 g of Ultra Pure Agarose for every 100 ml of 1x TAE buffer. For preparation of 1x TAE buffer fill 10 mL of 50x TAE buffer stock ad 500 mL ddH₂O. Ensure that the 1 % agarose in 1x TAE buffer is boiled throughly and dissolved completely without streaks. Incomplete boiling will bias the separation results. The liquid is poured in a gel tray with the wanted comb which are assembled in a holder. After solification the gel is placed in a gel chamber and is fully covered with 1x TAE buffer post removal of the comb.

Loading the Gel

Prior to loading, the DNA samples are homogenized with 6x Loading Dye (LD) resulting in a 1x final concentration of the LD (4 μL 6x LD ad 20 μL PCR reaction). Be aware of the fact, that PCR samples should not be mixed with LD if further experiments with the samples are needed to be done. If this is the case, mix the PCR samples with LD on parafilm. Meanwhile, the PCR samples are kept on ice. The 1 kb DNA ladder is stored in the 4 °C fridge and is suitable for larger PCR products ranging from 250 bp to 10000 bp (GeneRuler 1 kb DNA Ladder (Fermentas)). Load the gel with x μL of LD-mixed samples next to the first well with 5 μL marker. The amount loaded for PCR samples depends on the concentration and the size of the products. Note that PCR tubes should be closed to prevent drying up exposed to air.

Running the Gel

When all samples have been loaded, connect the power supply. Ensure that the plus pol is at the opposite site of the loaded wells. Run big gels applying 100 V for a more accurate separation for about 1 h. The time may be adjusted according to the loading front.

Staining the Gel

To make the separated DNA bands visible under the UV light, the samples were stained in a EtBr bath for about 10 min. Then, the gel was rinsed in water for approximately 5 min. The EtBr stained gel was exposed to UV light under a gel documentation station, a gel photo was taken and saved in the iGEM2012 folder. The file should be labelled with the date, group number and short description of what was analyzed. A printed version should be pasted into the group notebook.

Cloning Protocols

Cloning Protocols: Restriction (double digest)

• choose the correct restriction enzymes depending on your backbone and insert (e.g. backbone: SpeI and PstI; insert: XbaI and PstI)
• calculate the appropriate amount of enzyme which is necessary for a 5 fold over-digestion. Therefore fill in all parameters of the following equation:
• Keep in mind that you have to calculate the enzyme amounts separately for backbone AND insert (if the differ in size). The result of th the equation must be divided by 10 because most enzymes have a concentration of 10 U / µL. Finally, you have to multiply the value by factor 5 for 5 fold over-digestion in order to guarantee a complete digestion of the parts.</nowiki>
• choose the buffer by using the following tool: [http://www.fermentas.com/en/tools/doubledigest click]
• prepare reaction mix: 1 µg template, buffer, enzyme 1 and enzyme 2 (dependent on the concentration it is often necessary to add purified water)
• Incubation for 1.5 h at 37 °C

Cloning Protocols: Ligation

• Combination of 50 ng of vector with a 2-fold molar excess of insert.
• Addition of an appropriate volume of T4 Buffer.
• Addition of 0.5-1 μL T4 ligase.
• Adjustment of volume to 20 μL with ddH₂O
• Incubation over night at 16 °C
• Deactivation of ligase for 10 min at 65 °C

Cloning Protocols: Chemical Transformation

• Thawing of competent cells on ice
• Addition of 1 μL DNA
• Incubation on ice for 20 min
• Heat shock for 1 min at 42 °C
• Incubation on ice for 5 min
• Addition of 270 μL LB
• Incubation in thermoblock for 45 min (incubation time dependent on used vector!) at 37 °C and 300 rpm
• Smearing of cells on plates and incubation at 37 °C (centrifuge it, discard supernatant, resuspend cells in rest-medium, 20 μL on plate)

Competent Cells

Preparation of CaCl₂ buffer for competent cells! Before you start make sure that the CaCl₂ buffer is ice-cold when needed and the centrifuge is cooled to 4 °C.

• Inoculate LB-liquid medium with 1 ml of overnight culture.
• Let cells grow to OD₆₀₀ 0.4-0.6.
• Transfer them to 50 mL Falcons and centrifuge for 5 min at 1500 g and 4 °C.
• Resuspend pellet in 5-10 mL CaCl₂ buffer; here 7 mL (prechilled).
• Centrifuge 5 min 1500 g, 4 °C
• Resuspend pellet in 5-10 mL CaCl₂ buffer; here 7 mL (prechilled).
• Incubate cells 30-60 min on ice; here 30 min; Eppis prechilled in 5-10 mL on dry ice (or in liquid nitrogen)
• Storage at -80 °C

Transmission Electron Microscopy, negative staining with 2 % PTA

The TEM and the sample preparation was conducted by Dr. Michael Hoppert.

Strains

• MG1655
• BL21 J61002_rfp
• BL21 pSB1C3_QC_fliC_18C
• BL21 pSB1C3_QC_flhDC_18C

Media and buffers

• LB-broth
• LB-agar
• 2 % PTA (phosphotungstic acid)

Execution

• Inoculate the strains of interest in 5 mL L-broth with necessary antibiotics respectively
• Incubate them at 37 °C with 100 rpm (to minimize the chance of the flagella breaking of) over night
• Inoculate fresh 5 mL LB with necessary antibiotics with the overnight culture
  • For electron microscopy of flagella it is necessary to reach a high cell density without reaching a growth state where the flagella are discarded
    • The first time electron microscopy was conducted the new cultures were inoculated with 200 and 500 µL of the overnight culture, but for the cell density was not high enough the one ml of the cultures was spun down
  • One possibility to reach a high OD is to pour LB-agar (with antibiotics) in the glass test tubes and cover it with LB-broth (with antibiotics) and inoculate it, nutrients can freely diffuse through the agar as well as the broth but the cells are constricted to the liquid phase and thus a higher density can be reached
    • The second time electron microscopy was conducted the method described above was applied
• Let grow until a suitable OD₆₀₀ is reached and check the motility of the cells with the light microscope
• Prepare the samples
  • Place a drop of the culture, a drop of ddH₂O and a drop of 2 % PTA on a slice of parafilm
  • Pick a TEM grid (the microscopic slide in TEM) with forceps and place it on the drop of the culture for a suitable amout of time
  • Pick the grid up and let the cell suspension dry for a suitable amount of time
  • Place the grid on the 2 % PTA for a suitable amount of time
  • Let the grid dry for a suitable amount of time
  • Wash the grid through placing it on the ddH₂O drop for a suitable amount of time and let it dry afterwards
• Place the prepared grid in the electron microscope

Library Selection

The library containing vectors with the mutagenized tar-gene was transformed into the E. coli strain BL21. In order to determine certain receptor derivates that enables chemotaxis to a certain molecule a "Library Selection" protocol was determined. For hints and advice in general for swimming assays view the method "swimming/chemotaxis assay".

Attractants

• Coffein: stock solution x mM
• 2-Ethyl-1-Hexanol
• Geraniol
• Sodium-Cyclamat
• D-aspartic acid
• L-aspartic acid 4-benzylester
• Vanillin

Media

• LB-broth
• 0.3 % tryptone-swimming agar(1 % tryptone, 0.5 % NaCl, 0.3 % agar)
• LB agar

Execution

First round of selection

• Thaw one 1 ml cryostock of the library in BL21 and pour into a 200 mL flask filled with LB-media with chloramphenicol
• Inoculate the BL21 strain with the parent plasmid in 5 mL LB broth with chloramphenicol
• Grow the cultures over night at 37 °C with approx. 180 rpm
• Fill 7 + 1 control (whatmanpaper with H2O) 12 cm petridishes with 0.3 % tryptone-swimming agar with chloramphenicol
• Apply 100 µL of the attractant to a steril 2x2cm whatmanpaper respectivly and position it in the center of a petridish
• Spin down at least 1.5 mL of the culture containing the library and at least 1.5 mL of the culture containing the BL21 strain with the parent plasmid with 1.5 X g for 10 minutes
• Discard the supernatant and resuspend the pellet in the remaining medium
• Drop 3 times 5 µL of the library and once 5 µl of the reference strain (BL21 with the parent plasmid) respectively, on each plate
• Let the drops dry for at least 20 minutes until inverting the plates and placing in the incubator at 33 °C over night

Second round of selection

• Determine the drop with the fastes and most directed swimming behaviour on each plate
• In order to select the fastest cells the cells containing agar is cut out:
  • Cut the yellow eppendorf tips of to the first mark (approx 1 cm)
  • The first cut out is shortly befor the swimming front --> I
  • The second cut out is on the swimming front --> II
  • The third cut out is shortly behind the swimming front --> III
  • Place each cut out either with or without the tip in at least 0.5 ml LB media in an test tube or an E-cup
• Incubate the cultures for at least 1 h at 37 °C with approx. 180 rpm
• Meanwhile fill 7 + 1 control (whatmanpaper with H₂O) 12 cm petridishes with 0.3 % tryptone-swimming agar with chloramphenicol
• Apply 100 µL of the attractant to a steril 2x2 cm whatmanpaper respectivly and position it in the center of a petridish
• Transfer the culturesinto an E-cup and spin them down with 1.5 X g for 10 minutes
• Discard the supernatant and the resuspend the pellet in the remaining medium
• Drop 5 µl of the 3 different library cut outs and 5 µl of the reference strain (BL21 with the parent plasmid) respectively on each plate
• Let the drops dry for at least 20 minutes before inverting the plates and placing them in the incubator at 33 °C over night

Third round of selection

• See second round of selection

Plating of the selected clones

• The plates of the third round of selection are treated as described before, but the cultures are not spun down
• 100 µl of a 10^-2 dilution is plated on LB-plates containing chloramphenicol respectively
• Incubate the plates in an incubator over night at 33 °C

Minipreparation and sequencing of plasmid DNA

• A suitable amount of clones are selected from each plate and used to inoculate 5 mL LB media with chloramphenicol respectively
• Incubate the cultures over night at 37 °C with approx. 180 rpm
• Isolate the plasmid DNA according to the instructions of the the peqlab kit
• Sequence the plasmid DNAas described

Retransformation of the plasmid DNA

• In order to determine wether the observed chemotaxis is dependent on the cells themselfes or on the inserted vector the isolated plasmid DNA is transformed into fresh BL21 cells according to the described protocol

Determination of the swimming behaviour of the freshly transformed BL21 cells

• Colonies of the freshly transformed BL21 cells (Retrafo) as well as of the selected BL21 clones (Trafo) are used to inoculate 5 mL LB-media with chloramphenicol, respectively and grown over night at 37 °C with approx. 180 rpm
• Pour 7 x 2 x 3 0.3 % tryptone-swimming agar plates
  • Each attractant has 2 additional controls: one time the whatmanpaper is soaked with H₂O_dest. and the other time with aspartate
  • The whole approach is conducted for the "Trafos" as well as for the "Retrafos"
• Treat and drop the cultures as described
• Let the drops dry for at least 20 minutes before inverting the plates and placing them in the incubator at 33 °C over night

Overlap PCR for removal of restriction sites

The DH10B fliC gene contained three PstI sites and one SpeI site that had to be removed. In order to achieve this in a relatively short time we applied overlap PCR. If you are not familiar with this method you can find a detailed description here.

Separation Assay

One of our goals was to be able to separate two strains that swim with a different speed from each other when they are in a mixed culture. Only when they are in a mixed culture they are incubated at the exactly same conditions and thus the true effect of an attractant can be determined. In this case we utilized different resistance markers that were inserted into two different strains. This version of the separation assay is the final version determined through many tests with the strains Δtar+pSB1C3 (ampicillin resistance, rfp) and Δtar+pSB1C3_tar_QC_18C (chloramphenicol resistance). Specifications for this strain combination are marked in navy, variations in purple. For hints and advice in general for swimming assays view the method "swimming/chemotaxis assay".

Strains

Two strains containing plasmids with different resistance markers for example the biobrick vectors pSB1C3 (chloramphenicol resistance) and J61002 (ampicillin resistance) with different inserts.

Media

• LB broth
• LB agar with two different resistance markers respectively
• 0.3 % tryptone swimming agar (1 % tryptone, 0.5 % NaCl, 0.3 % agar)

Execution

Applying the mixed culture to the 0.3 % tryptone swimming agar plates

• Inoculate the two strains of interest in 5 mL LB broth with the neccessary antibiotic respectively
• Incubate cultures over night at 37 °C with approximately 180 rpm
• Fill a suitable number of 12 cm petridishes with 0.3 % tryptone-swimming agar without antibiotic and let the agar solidify
  • Alternatively M9 agar can be used
• Apply 100 µL of the to be tested attractant to a steril 2x2 cm whatmanpaper respectivly and position it in the center of the petridish
  • When the strains Δtar+pSB1C3 and Δtar+pSB1C3_tar_QC_18C were tested aspartate was used as a attractant
• Measure the OD₆₀₀ of the over night cultures
• calculate the neccessary amout you have to take from each culture to gain a cell ratio of 1:1
• Mix the cultures
• Spin down the mixed culture and 1 ml of the not mixed ultures repectively with 1.5 X g for 10 minutes
  • No change in the results were observed when this step was not conducted
• Remove the supernatant completly and add 100 µL of fresh LB broth
  • Discard the supernatant and resuspend the pellet in the remaining medium, this can only be applied, when the cultures are dropped on the plated immediatly
• Drop two times 5 µL of the mixed culture and once 5 µL of each of the not mixed strains (references)
• Let the drops dry for at least 20 minutes until inverting the plates and placing them in the incubator at 33 °C over night

Separation of the different strains

• Determine the drop of the mixed culture with the fastes and most directed swimming behaviour on each plate
• In order to determine the faster strain and to separate them the agar is cut out at three different positions:
  • Cut the yellow eppendorf tips of to the first mark (approx 1 cm)
  • The first cut out is at the swimming front --> I
  • The second cut out is between the swimming front and the center od the original drop -->II
  • The third cut out is in the center of the original drop --> III
  • Place each cut out either with or without the tip in at least 0.5 mL LB media in an test tube or an E-cup
• Incubate the cultures for at least 1 h at 37 °C with approx. 180 rpm
• Meanwhile prepare the same amout of LB agar plates (9 cm) with the different selection makers
  • When the strains Δtar+pSB1C3 and Δtar+pSB1C3_tar_QC_18C were tested LB agar plates containing either chloramphenicol or ampicillin were poured
• Prepare a dilution series from 10^-1 to 10^-5 of each of the three cultures
• Plate out 100 µL of the dilutions 10^-3, 10^-4 and 10^-5 on LB agar plates with the two different selection markers, respectively
  • Alternatively: Plate out the 10^-2 and the 10^-4 dilution. Dependent on the incubation time a bacterial lawn will be observed on the 10^-2 plates
• Incubate the plates in an incubator over night at 33 °C
• Count out the colonies

Sequencing

Sequencing is used to identify the nucleotide arrangement of a given DNA or RNA template. For example we used sequencing to check if our new designed plasmid-constructs include the correct insert. But before we could sequence our samples or better before we could give our samples to the sequence laboratory, we had to prepare the samples. First of all, the plasmids have to be isolated from cells. Therefore we used the PeqGOLD MiniPrep Kit I from Peqlab. After isolation of the plasmids the concentration has to be determined via Nanodrop. For sequencing we needed a concentration of 250-300 ng in a maximal volume of 4 µl purified water. Finally, 1 µL of the sequence primer (concentration of 5 pmol) was added. In most cases we used the standard Biobrick primer VF2 and VR.

Standard PCR

The polymerase chain reaction (PCR) is a method for in vitro-amplification of DNA sequences. For the amplification of a DNA fragment the heat resistent enzyme DNA polymerase is responsible. There are several types of DNA polymerases purchaseable, e.g. some of which are very fast or are not error-prone due to proof-reading activity. In order to choose the appropriate DNA polymerase, this link might be of interest: [http://barricklab.org/twiki/bin/view/Lab/ProtocolsTaq http://barricklab.org/twiki/bin/view/Lab/ProtocolsTaq; 06/30/2012.]

To allow binding of the DNA polymerase primer are required. Thus, only the flanking sites for the sequence of interest needs to be known to synthesize specific primer. Each primer has a specific annealing temperature according to it GC-content. Primers used here were ordered by Sigma^®.

A single PCR cycle encompasses three steps: denaturation, annealing and elongation. In the first step the DNA double strands are separated at about 95 °C. Next hybridization of the primers at their annealing temperature happens and finally the DNA seqthesis takes place at the optimal working temperature of the chosen DNA polymerase. This amplification cycle is normally conducted between 25-35 cycles depending on the amount of PCR product one wants to synthesize. In one PCR reaction following components are mixed: DNA template, forward and reverse primer, DNA polymerase with appropriate buffer and deoxynucleotides. Note that an increasing cycle number is prone to incorporate errors in the amplified DNA fragment. This is due to the fact that the dNTPs will be depleted and the saturation phase is reached.

-> 50 μl / reaction:

• 5 μL 10x Buffer
• 5 μL dNTPs (0.2 mM each)
• 2.5 μL Primer (0.5 µM) (2x forward and reverse)
• 0.5 μL Template (ca. 20-100 ng)
• 0.5 μL Pfu-Turbo (1.25 U)
• ad 34 μL ddH₂O

Program:

• 1 min 95 °C (initial denaturation)
• 30 s 95 °C
• 30 s 58 °C
• 2 min/kb 72 °C
• 10 min/kb 72 °C (final elongation)
• 30 Cycles

Swimming/Chemotaxis Assay

Another one of our aims was to invent a protocol whith enables us to observe and quantify swimming and chemotaxis of different E. coli strains. Here we use different agar combination dependent on the desired effect and soaked 2x2 cm Whatmanpapers as attractants for chemotaxis.

Strains

In general any strain can be used but some like for examle BL21 are more suitablte for chemotaxis assays, for they exibit the ability to swim.

Media

• LB broth
• 0.3 % tryptone swimming agar
• alternatively: M9 swimming agar
  • this agar is more suitable to observe chemotaxis for the nutrient content is lower
  • keep in mind that some E. coli strains are not able to grow on M9 media without additional supplements e.g. the strain DH10B requires the aminoacid leucin

Execution

• Inoculate the strains of interest in 5 mL LB broth with the neccessary antibiotic respectively
• Incubate cultures over night at 37 °C with approximately 180 rpm
• Fill a suitable number of 12 cm petridishes with 0.3 % tryptone-swimming agar or M9 agar with antibiotics or additional supplements when necessary
  • in some cases the swimming ability could be enhanced through the addition L-methionine to the M9 agar
  • pour the plates fresh before use but let them dry with a slightly open lid under the hood until the condensation water is evaporated to reduce the risk of smearing of the drops
• Apply 100 µL of the test-attractant to a steril 2x2 cm whatmanpaper and position it in the center of the petridish
• Spin down 1 mL of the cultures with 1.5 X g for 10 minutes
  • No change in the results were observed when this step was not conducted, but for the cultures have a higher viscosity the risk of spillage and smearing is lower
• Remove the supernatant completly and add the amount of fresh LB broth neccessary for the number of desired drops
  • Alternatively: Discard the supernatant and resuspend the pellet in the remaining medium
• Drop 5 µL of the cultures on the petridish, at maximum 4 drops per dish
  • The best results were observed when the drop was placed in a distance of 2-2.5 cm from the whatmanpaper
• Let the drops dry for at least 20 minutes until carefully inverting the plates and placing them in the incubator at 33 °C over night

QuikChange Protocol

To remove disturbing restriction sites within the gene for the successful usage of BioBrick system, the QuikChange reaction is used.

-> 20 μL / reaction:

• 2 μL 10x Buffer
• 0.4 μL dNTPs (10 mM each)
• 0.8 μL Primer (10 µM) (2x or as premix)
• 0.4 μL Template (ca. 20-100 ng)
• 0.4 μL Pfu-Turbo
• ad 20 μL ddH₂O

Program: -> 1 min 96 °C

• 20 s 96 °C
• 20 s 58 °C
• 1.5 min/kb 72 °C
• 15 Cycles

-> 5 min 72 °C -> Store at 4-8 °C

After PCR add 1 μL DpnI directly into PCR tube. Incubate reaction 1-2 h at 37 °C. Transform 5 µL into 50 µL competent cells.

Quantitative Real-Time PCR

Quantitative real-time PCR is a powerful tool for quantitation of nucleic acid. The quantitative rtPCR was performed by using SYBR® green 1 (5’ Prime). All real-time PCR systems detect a fluorescent dye and correlate this fluorescence signal to the amount of PCR product in the reaction. There are several methods that quantify the mRNA amount via fluorescence including the method SYBR® green 1 that is used in our experiments. SYBR® green 1 is a nonsequence-specific DNA binding dye that intercalates into dsDNA (it does not bind to single stranded DNA). It emits a strong fluorescent signal upon binding to dsDNA.

-> Experimental design

• 5 mL of LB-media with 50 µg/mL chloramphenicol were inoculated with E. coli strain BL21DE3 haboring the vector pSB1C3 with following combinations of promoters and Tar receptor constructs:
  
• Constitutive promoter constructs expressed in E. coli:
  • Tar receptor under the control of constitutive promoter J23100
    
  • Tar receptor under the control of constitutive promoter J23104
    
  • Tar receptor under the control of constitutive promoter J23105
    
  • Tar receptor under the control of constitutive promoter J23106
    
  • Tar receptor under the control of constitutive promoter J23109
    
  • Tar receptor under the control of constitutive promoter J23112
    
  • Tar receptor under the control of constitutive promoter J23113
    
  • Tar receptor under the control of constitutive promoter J23114
    
  • (For more details click Parts Submitted)

• Cultures were incubated overnight at 37 °C, shaking.
• For quantitative real-time PCR, RNA was isolated from E. coli BL21DE3 by using the RNA isolation Kit “Nucleus Spin II (Machery and Nagel)”. Cultures are inoculated over night at 37 °C to 10⁹ cells. The isolated RNA is then converted to cDNA using the “Reverse Transcription Kit from Qiagen”
• Each primer (forward or reverse) concentration in the mixture was 5 pmol/μl
• The housekeeping gene rrsD (ribosomal RNA, 16S) was used as reference
• The experiment was set up by following the PCR reaction mix illustrated in the table
• All experimental set ups were prepared in triplets for calculating the average
• In addition: to avoid contaminations, a non-template control for our PCR assays was always carried with the samples.
• The quantitative real-time PCR was performed by using ”LightCycler Software Version 4.05 (Roche)”

-> Primer for qrtPCR:

• Primers for amplification of reference cDNA rrsD
  • rrsD_rev_transcript_fw:
    • cgtcagctcgtgttgtgaaatg
  • rrsD_rev_transcript_rev:
    • cgtgtgtagccctggtcgtaag
• Primers for amplification of promoter constructs cDNAs: Primer Set 1
  • query_L1_fw:
    • tgacgtcaacctgggattta
  • query_R1_rev:
    • ggaggaatccatcatcatcc

-> 20 μl / reaction:

• 9 µL 2.5x real master mix/ 20x SYBR
• 2 µL sense primer (5 pmol)
• 2 µL antisense prime (5 pmol)
• 1 µL cDNA (diluted 1:10 in H2O)
• 6 µL water

Program:

• 3 min 95 °C (initial denaturation)
• 30 s 95 °C (denaturation)
• 30 s 61 °C (annealing for our experiment)
• 2 min/kb 72 °C (elongation)
• 42 Cycles

-> Analysis via the 2^–ΔΔCT (Livak) Method

• First, we normalized the CT of the target gene to that of the reference (ref) gene, for both the test sample and the calibrator sample (calibrator sample was the weakest expressed promotor construct in our case):
  • ΔCT(test) = CT(target, test) – CT(ref, test)
  • ΔCT(calibrator) = CT(target, calibrator) – CT(ref, calibrator)
• Second, we normalized the ΔCT of the test sample to the ΔCT of the calibrator:
  • ΔΔCT = ΔCT(test) – ΔCT(calibrator)
• Finally, we calculated the expression ratio:
  • 2^–ΔΔCT = Normalized expression ratio

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