1) Preparation of starting plasmids
Preparation restriction of components:

• restriction using biobrick assembly enzymes
  
• this preparation step is needed to create sticky ends on the cassettes
  
• this step is only performed once
  
• restriction with EcoRI and PstI (see restriction protocol) for all components

Ligation of cassettes into plasmids:

ligation using T4-ligase (see ligation protocol)

plasmid	cassettes
psB1C3	1	2	3
psB1A3	1	2	3
psB1K3	1	2	3

Plasmid cassettes psB1C3 1 2 3 psB1A3 1 2 3 psB1K3 1 2 3 Final preparation steps - transformation of 9 different combinations into competent cells (see transformation protocol) - selection with corresponding antibiotics Next day - colony pcr and gel run to check for sizes of cassettes - miniprep and check concentration via nanodrop - many aliquots needed (small volume because thawing time) for future reactions! 3) 3A-assembly - the 3-A-assembly will be used to add more and more cassettes in a row (see 3-A-assembly protocol) - it is important to only combine cassettes with the same number (1, 2 and 3 have varying spacer length) - we will add cassettes and test frequently for viability to determine the maximum target-sequence length - we want to combine about five cassettes Assembly scheme - in each assembly cycle, there will be two cassettes (same number/length) added to one linearized plasmid - in the first step, we can either just use the prepared plasmid with cassettes already inserted or use an empty one, because the part in between will be cut out anyway - the be able to select for plasmid with higher cassette content the resistances will cycle - the resistance cycle (for plasmids) is K C A - for the inserts, the resistance signals from which plasmid they will be cut - the plasmids resistance determines the selection antibiotics for that step! Legend - CX – Chloramphenicol (Insert/Plasmid) from step X - AX – Ampicillin (Insert/Plasmid) from step X - KX – Kanamycin (Insert/Plasmid) from step X - E – EcoRI - S – SpeI - X – XbaI - P – PstI

Colony PCR with ALLin™ Red Taq Mastermix, 2X:

aim:
Is the insert DNA in the plasmid present or absent?
Much easier than to isolate, purify the vector

good to know before the start:
- Take typical measures to prevent PCR cross over contamination, keep your bench clean, wear gloves, use sterile tubes and filter pipet tips
- Include a no-template control and positive control in parallel.
- Thaw and keep reagents on ice
- Mix well before use.
- The longer the amplicon, the longer the extension time: Use 15 sec/kb extension.
- Use 90 sec extension for multiplexing
- Run an annealing temperature gradient from 55 °C to 65 °C to choose the best specificity conditions. Do not use fast cycling for multiplexing.
- ALLin™ Red Taq Mastermix, 2X is premixed with red dye and density reagents for direct loading on the gels after the PCR. In a 2% agarose TAE gel the dye migrates with~350 bp DNA, in 1% agarose TAE gel with ~ 600 bp DNA fragments

step by step for E.coli:


- Prepare a PCR master mix (always prepare at least 10 % more, use the excel sheet by Sophia to calculate)
- 20 μl would be good, Fabian suggested 5 μl
Mix gently, avoid bubbles.
- Aliquote the 22.5 μl of PCR master mix into each PCR tube
- Add 2.5 μl of the resuspended colony or overnight culture

Do not forget the negative control!

- Close tube
- Perform the PCR using Thermocycler as follow:

Initial denaturation	1 cycle	95°C	60s
Denaturation	30-40 cycles	95°C	15s
Annealing	30-40 cycles	55-65°C	15s
Extension	30-40 cycles	72°C	15-90s
Final extension	1 cycle	72°C	5 min

- Store probes for short time on ice, for long at -20°C
- Load probes on the agarose gel e.g. 10 μl (so in case you have enough left for another round).

step by step for yeast: - If resuspended colonies are to be used: pipette 50 μl of a 0.02 M NaOH solution into each of a set of appropriately labelled PCR tubes or wells of a PCR plate. Using sterile pipette tips or toothpicks, transfer transformants to individual tubes/wells. The amount of cells resuspended must just be visible. Resuspend cells by pipetting or vortexing and incubate for ≥ 5 min at 37 °C.
If overnight cultures are to be used: pipette 40 μl of a 0.1 M NaOH solution into each of a set of appropriately labelled PCR tubes or wells of a PCR plate. Transfer 10 μl of each overnight culture to be tested to the appropriate tube/well and mix by pipetting up and down. Incubate for ≥ 5 min at 37 °C.
- Prepare a PCR master mix (always prepare at least 10% more, use the excel sheet to calculate) - Aliquot 22.5 μl of PCR master mix into each PCR tube.
- Add 2.5 μl of the resuspended colony or overnight culture mixed with NaOH to the appropriate PCR tube.
- Close the tubes
- Perform the PCR using the following cycling profle:

Initial denaturation	1 cycle	95°C	60s
Denaturation	30-40 cycles	95°C	15s
Annealing	30-40 cycles	55-65°C	15s
Extension	30-40 cycles	72°C	15-90s
Final extension	1 cycle	72°C	5 min

- Load probes on the agarose gel
- Store probes for short time on ice, for long at -20°C

Depending on the PCR product Binding of DNA 1. Insert SV Minicolumn into Collection Tube. 2. Transfer dissolved gel mixture or prepared PCR product to the Minicolumn assembly. Incubate at room temperature for 1 minute. 3. Centrifuge at 16,000 ×g for 1 minute. Discard (verwerfen) flowthrough and reinsert Minicolumn into Collection Tube. Washing 4. Add 700 μl Membrane Wash Solution (ethanol added). Centrifuge at 16,000 × g for 1 minute. Discard flowthrough and reinsert Minicolumn into Collection Tube. 5. Repeat Step 4 with 500 μl Membrane Wash Solution. Centrifuge at 16,000 × g for 5 minutes. 6. Empty the Collection Tube and recentrifuge the column assembly for 1 minute with the microcentrifuge lid open (or off) to allow evaporation of any residual ethanol. Elution 7. Carefully transfer Minicolumn to a clean 1.5 ml microcentrifuge tube. 8. Add 50 μl of Nuclease-Free Water to the Minicolumn. Incubate at room temperature for 1 minute. Centrifuge at 16,000 × g for 1 minute. 9. Discard Minicolumn and store DNA at 4°C or –20°C

What is it ? – standard lab procedure for separating DNA by size (e.g., length in base pairs) for visualization and purification
– uses an electrical field to move the negatively charged DNA through an agarose gel matrix toward a positive electrode
- Shorter DNA fragments migrate through the gel more quickly than longer ones

Why are we doing it ?
– to determine the approximate length of a DNA fragment by running it on an agarose gel alongside a DNA ladder (a collection of DNA fragments of known lengths)

Protocol :
- Pouring a Standard 1% Agarose Gel:


- Microwave for 1-3 min until the agarose is completely dissolved (but do not overboil the solution, as some of the buffer will evaporate and thus alter the final percentage of agarose in the gel. Many people prefer to microwave in pulses, swirling the flask occasionally as the solution heats up.).


Pouring of the gel
- Let agarose solution cool down to about 50°C (about when you can comfortably keep your hand on the flask), about 5 mins.

- Add ethidium bromide (EtBr) to a final concentration of approximately 0.2-0.5 μg/mL (usually about 2-3 μl of lab stock solution per 100 mL gel). EtBr binds to the DNA and allows you to visualize the DNA under ultraviolet (UV) light.

- Pour the agarose into a gel tray with the well comb in place.
- Let the newly poured gel sit at room temperature for 20-30 mins, until it has completely solidified.


Loading Samples and Running an Agarose Gel:

- Add loading buffer to each of your digest samples.
Note: Loading buffer serves two purposes: 1) it provides a visible dye that helps with gel loading and will also allows you to gauge how far the gel has run while you are running your gel; and 2) it contains a high percentage of glycerol, so it increases the density of your DNA sample causing it settle to the bottom of the gel well, instead of diffusing in the buffer. 2. Once solidified, place the agarose gel into the gel box (electrophoresis unit). 3. Fill gel box with 1xTAE (or TBE) until the gel is covered. 4. Carefully load a molecular weight ladder into the first lane of the gel. Note: When loading the sample in the well, maintain positive pressure on the sample to prevent bubbles or buffer from entering the tip. Place the very top of the tip of the pipette into the buffer just above the well. Very slowly and steadily, push the sample out and watch as the sample fills the well. After all of the sample is unloaded, push the pipettor to the second stop and carefully raising the pipette straight out of the buffer. 5. Carefully load your samples into the additional wells of the gel. 6. Run the gel at 80-150 V until the dye line is approximately 75-80% of the way down the gel. Note: Black is negative, red is positive. (The DNA is negatively charged and will run towards the positive electrode.) Always Run to Red. Note: A typical run time is about 1-1.5 hours, depending on the gel concentration and voltage. 7. Turn OFF power, disconnect the electrodes from the power source, and then carefully remove the gel from the gel box. 8. Using any device that has UV light, visualize your DNA fragments. Note: Fabian or Lena will give you a short introduction on how to work with UV light !!! Note: When using UV light, protect your skin by wearing safety goggles or a face shield, gloves and a lab coat. Note: If you will be purifying the DNA for later use, use long-wavelength UV and expose for as little time as possible to minimize damage to the DNA. Note: The fragments of DNA are usually referred to as ‘bands’ due to their appearance on the gel. Analyzing Your Gel: Using the DNA ladder in the first lane as a guide (the manufacturer's instruction will tell you the size of each band), you can interpret the bands that you get in your sample lanes to determine if the resulting DNA bands that you see are as expected or not. For more details on doing diagnostic digests and how to interpret them please see the Diagnostic Digest page. Purifying DNA from Your Gel: If you are conducting certain procedures, such as molecular cloning, you will need to purify the DNA away from the agarose gel. For instructions on how to do this, visit the Gel Purification page

What is the PCR ? Method to make multiple copies of a the specific DNA-sequence Protocol for PCR with Q5 High- Fidelity 2x Master Mix Please note that protocols with Q5 High-Fidelity DNA Polymerase may differ from protocols with other polymerases. Conditions recommended below should be used for optimal performance. Reaction Setup: – assemble all reaction components on ice, work on ice while assembling – preheat the thermocycler to the denaturation temperature( 98 °C) – prior to use all components should be mixed – work quickly when transferring the reactions to a thermocycler 1. on ice Assemble all components for the reaction : Component 25 μl Reaction 50 μl Reaction Final Concentration Q5 High-Fidelity 2X Master Mix 12.5 μl 25 μl 1X 10 μM Forward Primer 1.25 μl 2.5 μl 0.5 μM 10 μM Reverse Primer 1.25 μl 2.5 μl 0.5 μM Template DNA variable variable < 1,000 ng Nuclease-Free Water to 25 μl to 50 μl Notes: Two Primers have to be diluted 1:10 ! Notes: Gently mix the reaction. Collect all liquid to the bottom of the tube by a quick spin if necessary. Overlay the sample with mineral oil if using a PCR machine without a heated lid. 2. Transfer PCR tubes to a PCR machine and begin thermocycling. Steps of PCR: 1.Denaturation : double- stranded template DNA is heated to separate it into two single stands 2. Annealing : temperature is lowered to enable the DNA primers to attach to the template DNA 3. Extending : temperature is raised and the new strand of DNA is made by the polymerases Thermocycling Conditions for a Routine PCR: STEP TEMP TIME Initial Denaturation 98°C 30 seconds 25–35 Cycles 98°C 5–10 seconds *50–72°C 10–30 seconds 72°C 20–30 seconds /kb Final Extension 72°C 2 minutes Hold 4–10°C hold is not necessary 1. Template: Use of high quality, purified DNA templates greatly enhances the success of PCR. Recommended amounts of DNA template for a 50 μl reaction are as follows: DNA AMOUNT DNA Genomic 1 ng–1 μg Plasmid or Viral 1 pg–1 ng 2. Primers: Oligonucleotide primers are generally 20–40 nucleotides in length and ideally have a GC content of 40–60%. Computer programs such as Primer3 can be used to design or analyze primers. The best results are typically seen when using each primer at a final concentration of 0.5 μM in the reaction. 3. Mg ++ and additives: The Q5 High-Fidelity Master Mix contains 2.0 mM Mg ++ when used at a 1X concentration. This is optimal for most PCR products generated with this master mix. 4. Deoxynucleotides: The final concentration of dNTPs is 200 μM of each deoxynucleotide in the 1X Q5 High- Fidelity Master Mix. Q5 High-Fidelity DNA Polymerase cannot incorporate dUTP and is not recommended for use with uracil-containing primers or templates. 5. Q5 High-Fidelity DNA Polymerase concentration: The concentration of Q5 High-Fidelity DNA Polymerase in the Q5 High-Fidelity 2X Master Mix has been optimized for best results under a wide range of conditions. 6. Denaturation: An initial denaturation of 30 seconds at 98°C is sufficient for most amplicons from pure DNA templates. Longer denaturation times can be used (up to 3 minutes) for templates that require it. During thermocycling, the denaturation step should be kept to a minimum. Typically, a 5–10 second denaturation at 98°C is recommended for most templates. 7. Annealing: Optimal annealing temperatures for Q5 High-Fidelity DNA Polymerase tend to be higher than for other PCR polymerases. The NEB T m Calculator should be used to determine the annealing temperature when using this enzyme. Typically use a 10–30 second annealing step at 3°C above the T m of the lower T m primer. A temperature gradient can also be used to optimize the annealing temperature for each primer pair. For high T m primer pairs, two-step cycling without a separate annealing step can be used (see note 10). 8. Extension: The recommended extension temperature is 72°C. Extension times are generally 20–30 seconds per kb for complex, genomic samples, but can be reduced to 10 seconds per kb for simple templates (plasmid, E. coli , etc.) or complex templates < 1 kb. Extension time can be increased to 40 seconds per kb for cDNA or long, complex templates, if necessary. A final extension of 2 minutes at 72°C is recommended. 9. Cycle number: Generally, 25–35 cycles yield sufficient product. For genomic amplicons, 30-35 cycles are recommended. 10. 2-step PCR: When primers with annealing temperatures ≥ 72°C are used, a 2-step thermocycling protocol (combining annealing and extension into one step) is possible. 11. Amplification of long products: When amplifying products > 6 kb, it is often helpful to increase the extension time to 40–50 seconds/kb. 12. PCR product: The PCR products generated using Q5 High-Fidelity 2X Master Mix have blunt ends. If cloning is the next step, then blunt-end cloning is recommended. If T/A-cloning is preferred, the DNA should be purified prior to A-addition, as Q5 High-Fidelity DNA Polymerase will degrade any overhangs generated. Addition of an untemplated -dA can be done with Taq DNA Polymerase ( NEB #M0267 ) or Klenow exo – ( NEB #M0212 )

Protocol is following!

Salkowski Assay for colony screening Large qualitative screening of IAA-producing colonies at the same time to see if our constructs are still functional in our E.coli/yeasts Helps us pic kthe right colonies for colony-PCR and GC-MS measurements Reagent: 2% 0.5M FeCl3 in 35% perchloric acid Perchloric acid is highly corrosive and dangerous!! Always uses protective gear and work under a fume hood! Reagent will always be mixed together on the spot, FeCl3 stock solution is finished, acid will be taken from the chemicals sheld from the AG plant physiology (has been negotiated) What happens? Reagent reacts to IAA (and other indolic compounds) to make several colored products IAA will be seen as bright red (other compounds brown or yellowish) Assay conditions Plates were inoculated in a grid pattern and overlaid with an 82mm-diameter disk of Nitrocellulose membranes Plates are overlaid with Nitrocellulose immediately after inoculation with toothpicks After normal incubation (i.e. overnight) time, the membrane was removed and soaked in reagent (or reagent-saturated [2.5 mL] filter paper, here “Whatman grade 2” had best results)) After 30 - 60 minutes, coloring reaction is finished and fading began Best results with colony sizes between 0.5 to 2 mm Addition of Tryptophan greatly enhances color reaction but does not interfere with distinguishing IAA positive and negative colonies (yellow background and strong red to pink positives) Other indolic compounds (i.e. indolepyruvic acid) are distinguishable by a more yellow-brownish color

Tranformation Protocol What is it? Transmission of genetic information into competent cells (or plants, algas, mushrooms) (the target organismn) Übertragung von genetischer Information durch Aufnahme freier DNA in kompetente Zellen oder auch in Pilzen/Algen/Pflanzen DNA in Zielorganismen einzuschleusen Why we are doing it? Transmission of genetic information into competent cells What you need: *1.5 mL tube, pipette (50µL), pipette(1000µL), plate *Ice, SOC Medium, competent cells How we are doing it? 1. Prechil 1.5 ml tube on ice 2. Thaw a tube competent E. coli cells on ice for 10 minutes. 2.1. mix gently 2.2. Pipette 50 µl of the cells into the 1.5ml tube (Temperature over 0°C decrease the efficiency of the transformation!) 3. Add 1-5 µl (containing 1 pg-100 ng of plasmid) DNA to the cell mixture. (as soon as as the last bit of ice in the tube is disappeared!) 4. Flick the tube 4-5 times to mix cells and DNA.  (No vortexing!) 5. Place the mixture on ice for 30 minutes (without mixing!) (2-fold loss in transformation efficiency for every 10 minutes this step is shortened!)  6. Heat shock at exactly 42°C for exactly 30 seconds. (without mixing!) (temperature and timing specific to transformation volume and vessel) 7. Place on ice for 5 minutes. (without mixing!) 8. Pipette 950 µl of room temperature SOC into the mixture. 9. Place at 37°C for 60 minutes and shake vigorously (800 rpm in thermo mix block) (2-fold loss in transformation efficiency for every 15 minutes this step is shortened) (SOC gives 2-fold higher transformation efficiency than LB medium) (incubation with shaking or rotating the tube gives 2-fold higher transformation efficiency than without) 10. Warm selection plates to 37°C (plates can be used warm or cold, wet or dry…efficiency is nearly the same… warm plates easier to spread and allow most rapid colony formation) 11. Mix the cells thoroughly by flicking the tube and inverting 12. Spread 50-200 µl onto a selection plate and incubate overnight at 37°C. (Alternative: incubate at 30°C for 24-36 hours or 25°C for 48 hours.) (For low efficiency cloning reactions: spin down the whole transformation mixture, remove nearly the complete supernatant (approx. 900 ul), resuspend cells in remaining liquid and plate completely)

Zymo Research: Preparation of Competent Cells Grow yeast cells at 30 ° C in 10 ml YPD broth until mid-log phase (~5 x 10 6 - 2 x 10 7 cells/ml or OD 600 of 0.8-1.0). The following steps are accomplished at room temperatur e. 1. Pellet the cells at 500 x g for 4 minutes and di scard the supernatant. 2. Add 10 ml EZ 1 solution to wash the pellet. Repellet the cells and discard the supernatant. 3. Add 1 ml EZ 2 solution to resuspend the pellet. At this point, the competent cells can be used for transformations directly or stored frozen at or bel ow -70 ° C for future use. It is important to freeze the cells slowly. To accomplish this, either wrap the aliquotted cell s in 2-6 layers of paper towels or place in a Styrofoam box before placing i n the freezer. DO NOT use liquid nitrogen to snap-freeze the cells. Transformation This part of the procedure is the same for both fro zen stored (thawed at room temperature) and freshly prepared competent yeast cells. 1. Mix 50 μ l of competent cells with 0.2-1 μ g DNA (in less than 5 μ l volume); add 500 μ l EZ 3 solution and mix thoroughly. 2. Incubate at 30 ° C for 45 minutes. Mix vigorously by flicking with f inger or vortexing (if appropriate for your DNA) 2-3 times during this incubation. 3. Spread 50-150 μ l of the above transformation mixture on an appropr iate plate. It is unnecessary to pellet and wash the cells before spreading. Incubate the plates at 30 ° C for 2-4 days to allow for growth of transformants .

Ligation Protocol with T4 DNA Ligase (M0202) 1. Set up the following reaction in a microcentrifuge tube on ice. T4 DNA Ligase Bu ff er (10 x) 2 ul T4 DNA Ligase 1 ul Vector DNA Insert DNA Nuclease-free water to 20 ul 1. Calculation of the DNA 2. Example calculation 1:3 vector to insert mass Vector DNA: 100 ng Vector DNA: 10 kb Insert DNA: 3 kb 2. T4 DNA Ligase should be added last. 3. Use nebiocalculator.neb.com/#!/ to calculate molar ratios. 4. The T4 DNA Ligase Bu ff er should be thawed and resuspended at room temperature. 2. Gently mix the reaction by pipetting up and down and microfuge briefly. 3. Incubation 1. cohesive (sticky) ends 1. 16°C overnight or room temperature for 10 minutes. 2. blunt ends or single base overhangs 1. 16°C overnight or room temperature for 2 hours (alternatively, high concentration T4 DNA Ligase can be used in a 10 minute ligation) . 4. Heat inactivate at 65°C for 10 minutes. 5. Chill on ice and transform 1-5 μ l of the reaction into 50 μ l competent cells

Promega “Wizard Plus SV Miniprep Purification System“ mehr Infos zum Kit sind in der Anleitung zu finden (ausgedruckt hinter dem “Miniprep-Protokoll“ 1. Production of cleared lysate 1. Isolation of the bacteria 1. harvest 1–5ml (high-copy-number plasmid) or 10ml (low-copy-number plasmid) of bacterial culture 2. centrifugation for 5 minutes at 10,000 x g in a tabletop centrifuge 3. pour o ff the supernatant 4. reinsert again bacterial culture to the pellet and repeat step 2 and 3 5. blot the inverted tube on a paper towel to remove excess media 2. Resuspension of the cells 1. add 250 μ l of Cell Resuspension Solution 2. completely resuspend the cell pellet by vortexing or pipetting 3. it is essential to thoroughly resuspend the cells 3. Lysing 1. add 250 μ l of Cell Lysis Solution 2. mix by inverting the tube 4 times - do not vortex 3. incubate until the cell suspension clears (clear ≠ colorlessly) ( approximately 1– 5 minutes ) 4. Splitting proteins 1. add 10 μ l of Alkaline Protease Solution 2. mix by inverting the tube 4 times - do not vortex 3. incubate for 5 minutes at room temperature 5. Neutralization 1. add 350 μ l of Neutralization Solution 2. immediately mix by inverting the tube 4 times - do not vortex 6. Isolation of the plasmids 1. centrifuge the bacterial lysate at maximum speed (around 14,000 × g ) in a microcentrifuge for 10 minutes at room temperature