Here you will find various protocols we have used during the development of our project. We have separated them into 4 main sections:

#General Protocols
#Cyanobacteria Protocols
#Bactomithril Protocols
#Characterization Protocols

LB media

Final concentrations:

• 1% (w/v) tryptone
• 0.5% (w/v) yeast extract
• 86 mM NaCl

Per liter:

• 10g Tryptone
• 5g Yeast extract
• 5g NaCl




• 15g Agar -Optional-
• Adjust pH to 7.5 with NaOH 1M

SOB media

Final concentrations:

• 0.5% (w/v) yeast extract
• 2% (w/v) tryptone
• 10 mM NaCl
• 2.5 mM KCl
• 20 mM MgSO4

Per liter:

• 5 g yeast extract
• 20 g tryptone
• 0.584 g NaCl
• 0.186 g KCl
• 2.4 g MgSO4




• 15g Agar -Optional-
• Adjust pH to 7.5 with NaOH 1M

BG-11

For 1L:

• 1.5gr NaNO3
• 1L distilled H2O
• 10mL Concentrated salts solution 100X (4°C)
• 200µL H2PO4 1M (Room T°)
• 10gr BactoAgar -optional-

BG-110

For 1L:

• 1L distilled H2O
• 10mL Concentrated salts solution 100X (4°C)
• 200µL H2PO4 1M (Room T°)
• 10gr BactoAgar -optional-

BG-11C

For 1L:

• 1.5gr NaNO3
• 1L distilled H2O
• 10mL Concentrated salts solution 100X (4°C)
• 200µL H2PO4 1M (Room T°)
• 0.84gr NaHCO3
• 10gr BactoAgar -opcional-

Concentrated salts solution 100X

For 1L:

• 7.5gr SO4Mg * 7H2O
• 3.6gr CaCl2 * 2H2O
• 0.6gr Citric Acid
• 0.6gr Ammonium Ferric Citrate
• 0.093gr EDTA-Na2
• 2.0gr CO3 Na2
• 0.286gr BO3H3
• 0.181gr MnCl2 * 4H2O
• 0.022gr ZnSO4 * 7H2O
• 0.039gr NaMoO4 * 2H2O
• 0.0079gr CuSO4 * 5H2O
• 0.0049gr CoCl2 * 6H2O
• 1L distilled H2O

Autoclave and store at 4°C.

5X Gibson Assembly Isothermal Buffer

Final Concentrations:

• 25% PEG MW 8000
• 500mM Tris HCl pH 7.5
• 50mM CaCl
• 50mM DTT
• 1mM dATP
• 1mM dTTP
• 1mM dGTP
• 1mM dCTP
• 5mM NAD+
• H20 nuclease free

For a 2 mL stock (equivalent to 20 100uL 5X isothermal buffers)

• 0.5g PEG MW 8000
• 1.5mL Tris-HCl pH 7.5 1M

Swivel for 30 minutes in a orbital oscillator for PEG to dissolve fully

• 50uL CaCl 2M
• 100uL DTT 1M
• 20uL dATP 100mM
• 20uL dTTP 100mM
• 20uL dGTP 100mM
• 20uL dCTP 100mM
• 200uL NAD+ 200mM
• H20 nuclease free up to 2mL

Alicuot into 100uL stocks

Store at -80°C

1.33X Gibson Assembly Master Mix

For a 375ul 1.33X Gibson Assembly Master Mix

• 100uL 5X Gibson Assembly Isothermal Buffer
• 6.25uL Phusion Polymerase 2 U/uL (cat N° F-350S) from Thermo Scientific
• 2uL T5 Exonuclease 1U/uL (cat N° T5E4111K) from Epicentre

It is important to dilute the T5 Exonuclease stock from 10U/uL to 1U/ul to measure the volume correctly

• 50uL Taq DNA Ligase 2000 U/uL (cat N° M208S) from NEB
• 216.75uL of nuclease free H20

Alicuot 9uL in 0.2mL PCR tubes. This will yield about 42 reactions

Standard Assembly

The standard assembly reaction relies on the standarization of Biobricks to join different DNA parts. Plasmids from the registry of standard parts allow joining of DNA parts by using a combination of specific enzymes to cut a Upstream Biobrick with the restriction enzymes EcoRI and SpeI, and a Downstream Biobrick with the restriction enzymes XbaI and PstI into a Destination plasmid which has been cut with the restriction enzymes EcoRI and PstI. The reaction yields the Upstream and Downstream part joined by a mixed restriction site and allows further elongation of the construct by the same strategy. It is important to notice that such a reaction requires purification of the digested parts if any of the plasmids (Upstream, Downstream or Destination) share a resistance marker.

• Upstream part enzymes: EcoRI & SpeI
• Downstream part enzymes: XbaI & PstI
• Destination backbone enzymes: EcoRI & PstI

Digestion reaction

• X volume of DNA to 500ng of plasmid
• (42.5 - X)uL of nuclease free water
• 5uL of NEB buffer 2
• 0.5uL of BSA 100X
• 1uL of Enzyme 1
• 1uL of Enzyme 2
• Incubate at 37°C for 2 hours
• Heat inactivate enzymes at 80°C for 20 minutes

Ligation reaction

• 2uL of digested Upstream part
• 2uL of digested Downstream part
• 2uL of digested Destination plasmid
• 2uL of T4 DNA ligase buffer
• 11uL of nuclease free water
• 1uL of T4 DNA ligase
• Incubate at room temperature for 10 minutes
• Heat inactivate at 80°C for 20 minutes
• Transform

Gibson Assembly

Gibson Assembly is a DNA assembly method created by Daniel Gibson during the development of the first Synthetic Genome (Synthia) 2. Its adaptation to a cloning method allows fast and accurate production of increasingly complex constructions. The strategy behind the method relies on PCR to obtain different parts which share a 40bp homology region, and a 3 enzyme reaction which produces cohesive ends, fills the gaps between the parts and ligates the resulting construct into a scarless assembly of various (>2) parts.

Primer design

The easiest way to design primers to obtain amplicons with the required overlaps (40bp final overlaps) is to make an in sillico design of the final construct

• Design forward primer of right amplicon of joint in 5'->3' direction
• Calculate length of annealing part of primer as to reach a Tm of approximately 63°C
• Add 20 bp of overlap
• Design reverse primer of left amplicon of joint in 5'->3' direction
• To do this, select full joint and "reverse complement" it (be sure to be able to discriminate between right and left parts of joint)
• Calculate length of annealing part of primer as to reach a Tm of approximately 63°C
• Add 20 bp of overlap
• Apply same principle in all joints

Obtaining parts

• PCR parts using Phusion Polymerase datasheet indications. It is important to use a low ammount of template plasmid (10pg) as to reduce possibility of carry-over during band purification
• Run agarose gel electrophoresis on a adecuate gel (50bp to 200bp parts should be run on a 3% w/v agarose gel, larger parts should be run on a 1% w/v to 1.5% w/v agarose gel) until clear distinction of bands is achieved. It is recommended that gels should be exposed as little as possible to UV transilluminators as UV light damages DNA.
• Cut band and proceed with band purification. Elute in smallest volume as possible according to your kit specification
• Quantify purified DNA

Assembly reaction

The assembly reaction is composed of 9uL of 1.33X Gibson assembly master mix + 3uL of purified parts DNA

• The calculation of the ratio at which your DNA parts are should be in correspondence to the level of competence of your cells. We have found that with our cells (5*10^⁸ colonies/ug of pUC19 DNA) the following ratios work well
• Calculate the amount of pmoles of each purified DNA part using the following equation: pmoles of DNA = weigth in ng * 1000 (conversion factor from nano to pico) / (650 Daltons * base pair length of part)
• We have seen that using 0.01 picomoles of template (part with the selection resistance) and the rest of parts at 0.03 picomoles and above, yield a high ratio of true positive transformants and a decent number of colonies to check
• Thoroughly mix the 3uL of purified parts DNA with the 9uL of 1.33X Gibson assembly master mix in ice
• Directly incubate the reaction at 50°C for 1 hour
• Transform competent cells with 8uL of assembled DNA (that leaves 4uL to check in an agarose gel if the reaction fails)

Checking assembly

• Check transformant colonies by colony PCR using primers for whole insert
• Grow positive colonies in media with corresponding antibiotic
• Miniprep colonies and digest plasmid with EcoRI & PstI restriction enzymes
• If checking protocols validate until now, proceed by sequencing to corroborate

• Keep 50uL of chemically competent cells in ice (for no more than 10 minutes until transformation)
• Mix 0.01 to 1 total ng of plasmid with competent cells (depending on size of plasmid and competence of your cells)

Flick very softly to mix (competent cells are very fragile)

• Leave on ice for 30 minutes
• Heat shock at 42°C for 60 seconds
• Add 250uL of LB media
• Incubate at 37°C in a rotating shaker for 1 hour (Ampicillin) or 2 hours (Chloramphenicol and Kanamycin)
• Plate 50uL of transformed cells to a LB agar petri dish with corresponding antibiotic (30ug/mL Kanamycin or 30ug/mL Chloramphenicol or 100ug/mL Ampicillin)

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