• Place gel slice in a 1.5 mL microcentrifuge tube.
• Pre-warm an aliquot of sterile milli-Q at ~40°C.
• Add 10 µL Membrane Binding Solution per 10 mg of gel slice. Vortex and incubate at 50-65 °C until gel slice is completely dissolved.
• Insert SV Minicolumn into Collection Tube and label both of them according to the labelling of your samples.
• Transfer the dissolved gel mixture to the Minicolumn assembly. Incubate at room temperature for 1 minute. NOTE: When pipetting into the column, aim the pipette to the wall not the membrane to avoid damaging it.
• Centrifuge the SV Minicolumn assembly at maximum speed for 1 minute.
• Discard the flowthrough and reinsert the SV Minicolumn into the Collection Tube.
• Add 700µL of Membrane Wash Solution (if it is the first use, dilute it with 95% ethanol following the bottle's instructions).
• Centrifuge the SV Minicolumn assembly at maximum speed for 1 minute.
• Discard the flowthrough and reinsert the SV Minicolumn into the Collection Tube.
• Repeat steps 9-11 with 500µL of Membrane Wash Solution and centrifuging for 5 minutes.
• Once the Collection Tube is empty, centrifuge the Minicolumn assembly at maximum speed for 1 minute with the microcentrifuge lid open to allow ethanol full evaporation.
• Transfer the SV Minicolumn to an empty, labelled 1.5 mL tube.
• Add 50 µL of the pre-warmed water (30 µL for higher concentrations or when small amounts of DNA are suspected) directly to the centre of the SV Minicolumn, without touching the membrane with the pipette tip.
• Incubate at room temperature for 5 minutes.
• Centrifuge at maximum speed for 1 minute.
• Discard the SV Minicolumn, cap the tube containing the eluted DNA and keep at 4°C (for immediate use) or -20°C (for storage).

• Add x µL of sterile milli-Q to dissolve the DNA material. This creates a 100 µM stock solution.
• Heat the primer stock solution to 65 °C for 20 minutes.
• Centrifuge the primer stock solution at maximum speed (~17,000x g) for 2 minutes.
• Prepare a 10x diluted work solution (10 µm) by dilution with sterile milli-Q.

Please note that protocols with Q5 HighFidelity DNA Polymerase may differ from protocols with other polymerases. Conditions recommended below should be used for optimal performance.

• Assemble all reaction components on ice.
• Transfer quickly the reactions to a thermocycler preheated to the denaturation temperature (98°C). All components should be mixed prior to use.
• (Optional) Dilute Q5 HighFidelity DNA Polymerase in 1X Q5 Reaction Buffer just prior to use in order to reduce pipetting errors. 
• Transfer PCR tubes to a PCR machine and begin thermocycling.

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.

• Ethanol treat all working areas for sterility.
• Inoculate 250 ml of SOB medium with 1 ml vial of seed stock and grow at 20°C to an OD600nm of 0.3. Use the "cell culture" function on the Nanodrop™ Spectrophotometer to determine OD value*. 

OD value = 600nm Abs reading x 10 

*This takes approximately 16 hours.

*Controlling the temperature makes this a more reproducible process, but is not essential.  Room temperature will work. You can adjust this temperature somewhat to fit your schedule.  Aim for lower, not higher OD if you can't hit this mark

• Fill an ice bucket halfway with ice. Use the ice to pre-chill as many flat bottom centrifuge bottles as needed.
• Transfer the culture to the flat bottom centrifuge tubes. Weigh and balance the tubes using a scale*.

*Try to get the weights as close as possible, within 1 gram

• Centrifuge at 3000g at 4°C for 10 minutes in a flat bottom centrifuge bottle. Flat bottom centrifuge tubes make the fragile cells much easier to resuspend 
• Decant supernatant into waste receptacle, bleach before pouring down the drain.
• Gently resuspend in 80 ml of ice cold CCMB80 buffer. Pro tip: add 40ml first to resuspend the cells. When cells are in suspension, add another 40ml CCMB80 buffer for a total of 80ml 

-Pipet buffer against the wall of the centrifuge bottle to resuspend cells. Do not pipet directly into cell pellet!

-After pipetting, there will still be some residual cells stuck to the bottom. Swirl the bottles gently to resuspend these remaining cells

• Incubate on ice for 20 minutes
• Centrifuge again at 3000G at 4°C. Decant supernatant into waste receptacle, and bleach before pouring down the drain. 
• Resuspend cell pellet in 10 ml of ice cold CCMB80 buffer. 

  -If using multiple flat bottom centrifuge bottles, combine the cells post-resuspension

• Use Nanodrop to measure OD of a mixture of 200 μl SOC and 50 μl of the resuspended cells

  -Use a mixture of 200 μl SOC and 50 μl CCMB80 buffer as the blank

• Add chilled CCMB80 to yield a final OD of 1.0-1.5 in this test. 
• Incubate on ice for 20 minutes. Prepare for aliquoting

  -Make labels for aliquots. Use these to label storage microcentrifuge tubes/microtiter plates

  -Prepare dry ice in a separate ice bucket. Pre-chill tubes/plates on dry ice

• Aliquot into chilled 2ml microcentrifuge tubes or 50 μl into chilled microtiter plates.
• Store at -80°C indefinitely.

  -Flash freezing does not appear to be necessary

• Perform test transformations to calculate your competent cell efficiency.

  -Thawing and refreezing partially used cell aliquots dramatically reduces transformation efficiency by about 3x the first time, and about 6x total after several freeze/thaw cycles

  -Good cells should yield around 100 - 400 colonies

  -Transformation efficiency is (dilution factor=15) x colony count x 10⁵/µgDNA

  We expect that the transformation efficiency should be between 1.5x10⁸and 6x10⁸ cfu /µgDNA 

• Place a small glass bottle with sterile SOC (10 ml) in the 37°C incubator to pre-warm. Alsopre-warm at this point the plates for spreading out the transformation.  
• Thaw on ice the Eppendorf tube (200 µl) containing self-made competent Top10 cells per ligation to be transformed (Keep the competent cells as much as possible on ice!) 
• Add the entire ligation (20 µl) to the cells (or 0.1 to 1 µl for plasmids to be re-transformed) and mix by tapping gently. Do not mix cells by pipetting, their membrane is fragile and shear force can kill cells at this point!
• Incubate on ice for 30 minutes (if possible, mix once or twice by gently tapping again).
• Switch on the water bath immediately after adding the DNA to the cells and set to 42°C (check temperature with a thermometer, sometimes the setting needs to be adjusted to achieve measured 42°C. In our lab, set to 39.5°C to achieve 42°C!).
• Heat shock the cells by incubating the tube for 60 seconds in the 42°C water bath. Do not mix or shake!
• Remove the tubefrom the 42°C bath and quickly place on ice, incubate for 2 min. 
• Add 500 μLof pre-warmed SOC medium (SOC is a rich medium; use proper sterile technique to avoid contamination) .
• Secure the tubein a small microcentrifuge rack with tape and place in the 37°C shaking incubator for 60- 90 min (the rack should stick down with its short side so that the tube lie parallel to the platform for maximum aeration). Warning: if you label only the lids, the tape will remove the labelling when you strip it off after incubation. Label the side of the Eppendorf and secure the label with masking tape! 
• Plate 50- 100 µl of the cells on one selective pre-warmed plate. Spin down the remainder of the cells at 3800 rpm for 3 min at RT and pour off the SN until about 50 µl of liquid remain in the tube. Resuspend the pellet in this remaining liquid by pipetting up and down with a 200 µl pipette and plate the entire resuspension onto a second plate. 
• Wrap the plates loosely in cling film and incubate upside down at 37°C Warning: if you use the thin blue/black permanent marker to label the bottom of the plate, the cling film can remove it while unwrapping. Use the thick black markers to label the plates! 

• Calculate the vector to insert molar ratios.

As a starting point, usually people try a 1:3 vector to insert ratio first. This ratio is about the number of fragments and as fragments of vector and insert are most likely of different lengths, it is not enough to just use the same volumes of an identical DNA concentration for both, i.e. 5 µl of a 10 ng/ µl digested vector backbone of 5000 bps will have less ‘DNA pieces’ than 5 µl of a 10 ng/ µl digested insert of 1000 bps. The overall DNA amount would be the same, but there would be 5 times more insert fragments than vector fragments in this example.  

• To calculate the vector, insert molar ration:

ng insert = (ng vector x bps insert) /bps vector x (insert/vector) molar ratio 

• Alternatively, or to double check you can use this NEB online tool:

NEBioCalculator (http://nebiocalculator.neb.com/#!/ligation)  

• The total the DNA amount in a ligation reaction should not exceed 100 ng (or at least not by much). So for the formula above you can start with 50 ng vector.
• In case your insert is bigger than your vector I would treat the insert as vector for calculating the molar ratio.
• Ratios of 1:5 and 1:10 (vector: insert) are also commonly used, if you have enough DNA of both you can try different ratios at the same time. 
• Also important, plan a negative control with only linearized vector, i.e. use the same amount of vector as in your ligation with the insert, but replace the insert with dH₂ This control should ideally give you no colonies on a selective plate. If you get colonies it means usually that your vector was not fully digested or illegal ligation has been taking place between restriction enzyme sites that should not be compatible.
• Set up the following ligation reaction in a PCR tube on ice,exactly in the order shown in the table below.
• T4 DNA Ligase should be added last.

Make sure that you leave the T4 Ligase in the freezer until you need it. Only then do you bring it to your place in the yellow enzyme bucket and as soon as you have pipetted the ligase to your reaction you bring the yellow bucket with the ligase back to the freezer!! Same as for Q5 polymerase!

• Thaw the T4 DNA Ligase Buffer in your hand, give it a brief vortex and short spin. Once thawed it should be stored on ice. The T4 DNA Ligase Buffer (10X)* contains ATP that degrades quickly over time if not stored on ice and during repeated freeze and thaw cycles.
• Gently mix the reaction by pipetting up and down and microfuge briefly.
• For cohesive (sticky) ends, incubate at 23 °C for 15 minutes or 16°C overnight 
• For blunt ends or single base overhangs, incubate at 16°C overnight or 23 °C for 2 hours.
• Heat inactivate at 65°C for 10 minutes.
• Chill on ice and transform the entire reaction into 200 μl self-made DH5 alpha competent cells.

• Design your plasmid and order primers
• Generate DNA segments by PCR
• Run PCR product on an agarose gel to check for size and yield. If there are significant amounts of undesired product, gel purify DNA segments. Otherwise PCR purification or even the raw PCR mix can work fine in an assembly if you want to save time.
• Combine segments in Gibson Assembly Reaction. Note:Yields will be best when the DNA fragments are present in equimolar concentrations.

 The Gibson Cloning Master Mix consists of three different enzymes within a single buffer. Each enzyme has a specific and unique function for the reaction:

• T5 Exonuclease- creates single-strand DNA 3’ overhangs by chewing back from the DNA 5’ end. Complementary DNA fragments can subsequently anneal to each other.
• Phusion DNA Polymerase- incorporates nucleotides to “fill in” the gaps in the annealed DNA fragments.
• Taq DNA Ligase- covalently joins the annealed complementary DNA fragments, removing any nicks and creating a contiguous DNA fragment.Incubate the mix for 1 hour at 50°C or follow manufacturer's instructions.
• Transform the DNA into bacteria and screen for the correct plasmid product by Restriction Digest.
• Sequence the important regions of your final plasmid, particularly the seams between the assembled parts.

• Prepare liquid cultures according to theliquid (starter) culture (10 mL) 
• Under sterile conditions, take a 200 µL aliquot as a back-up and store at -80 °C.

Prepare Lysate

• Centrifuge the Falcon tubes for 10 minutes at maximum speed.
• Pour off the supernatant and resuspend in 600µL of TE buffer. If they are not already in a microcentrifuge tube, transfer the resuspended cells to a sterile 1.5 ml microcentrifuge tube(s).
• Add 100 µL of Cell Lysis Buffer (blue), and mix by inverting the tube 6 times.
• Add 350 µl of cold (4–8 °C) Neutralization Solution, and mix thoroughly by inverting.
• Centrifuge at maximum speed in a microcentrifuge for 3 minutes.
• Transfer the supernatant (~900 µl) to a PureYield™ Minicolumn without disturbing the cell debris pellet.
• Place the minicolumn into a Collection Tube, and centrifuge at maximum speed in a microcentrifuge for 15 seconds.
• Discard the flowthrough, and place the minicolumn into the same Collection Tube.

Wash

• Add 200 µl of Endotoxin Removal Wash (ERB) to the minicolumn. Centrifuge at maximum speed in a microcentrifuge for 15 seconds.
• Add 400 µl of Column Wash Solution (CWC) to the minicolumn. Centrifuge at maximum speed in a microcentrifuge for 30 seconds.

Elute

• Transfer the minicolumn to a clean 1.5 ml microcentrifuge tube, then add 30 µl of sterile milli-Q directly to the minicolumn matrix. Let stand for 1 minute at room temperature.
• Centrifuge for 15 seconds to elute the plasmid DNA.
• Following the NanoDrop protocol, determine the concentration of the miniprepped samples. Store at -20 °C.

• Add 230 µL of LB into each well 
  Use at least 2 wells for blank, add 250 µL LB in these.
• Inoculate 18 wells per culture with 10 µL of the starter culture
• An hour after inoculation, the cells are induced in triplo with the following final IPTG-concentrations: 0 mM; 0.1 mM; 0.3 mM; 0.5 mM; 0.7 mM; 1 mM 
• After induction, the growth and fluorescence was monitored with a plate-reader.

• Turn on the NanoDrop UV-VIS Spectrophotometer.
• Press the button for dsDNA to measure the concentration of double-stranded DNA in your samples.
• Clean the measurement surface with a piece of tissue and ethanol.
• Use 1-1.5 µL of sterile milli-Q as a blank.
• Clean the measurement surface with a piece of tissue and water.
• Use 1-1.5 µL of sample to measure its concentration.
  NOTE: It is best to measure the same sample in triplo and use the average value.
• If you have multiple samples, clean the measurement surface in between measurements.