Protocol for ligating a synthetic double stranded DNA cassette that lacks 5’ phosphates
into a plasmid cut with BsaI so that it has sticky ends (single stranded overhangs) with 5’
phosphates that match with the cassette. Generally the cassette is double stranded with
sticky ends to match the cut plasmid.
Cartoon outline:
Protocol outline. (a) The BsaI
enzyme binds to DNA (shown as
red arrows) adjacent to where it
cuts DNA (either edge of the
orange rectangle). Plasmids
engineered with inverted BsaI sites
as illustrated will digest to leave
sticky ends that can be completely
engineered so that they result in a
seemless ligation that does no
change the parental sequence
except for as desired. (b) Digested
plasmid will contain 5’ phosphates
that can efficiently ligate to a
cassette lacking 5’ phosphates.
Note, cutting with an second
enzyme between the BsaI sites
(SphI as outlined in the figure, but
can be other enzymes) reduces
background self ligation reactions.
I. Prepare plasmid and ensure that ends have phosphates (DO NOT use CIP).
1. Digest plasmid with BsaI in a 50 L reaction setup in a PCR tube.
a. ~36 L deionized water.
b. 5 L NEB3 (even though NEB recommends NEB CS, NEB3 works better)
c. 0.5 L 100X Bovine Serum Albumin (passed through a spin filter for
sterilization)
d. 800 ng of plasmid DNA (usually about 3 L).
e. 5 L BsaI enzyme
Pipette up and down gently in order to mix
Incubate in a PCR machine at 50 C for 1 hour
Remove from PCR machine and allow to cool for 10 minutes to room temperature
2. Add second endonuclease (often SphI or EcoRV).
a. Add 1 L of the second enzyme and pipette gently in order to mix.
b. Incubate at 37 C for 1 hour
3. Column purify the DNA away from enzymes.
a. Add 300 L of binding buffer (e.g., ADB or QG). This buffer has a high
concentration of GdmCl that promotes DNA binding to silica columns.
b. Pipette gently to mix.
c. Add solution to a silica column (recommend small column)
d. Spin in a microcentrifuge at 13000xg for 1 minute
e. Discard flow thru
f. Add 500 L of wash buffer with ethanol to the column
g. Spin in a microcentrifuge at 13000xg for 1 minute
h. Discard flow thru
g. Spin in a microcentrifuge at 13000xg for 1 minute to thoroughly dry the
column
i. Transfer the column to a clean microfuge tube
j. Add 40 L of elution buffer (10 mM Tris, pH 8).
k. Spin in a microcentrifuge at 13000xg for 1 minute
l. Remove the column, save and label the microfuge tube containing purified
vector DNA.
m. Note that the concentration of this vector DNA should be roughly 5 nM. 20
ng/L for a 6kb plasmid corresponds to 5 nM.
II. Prepare cassettes. Synthesized DNA generally comes without phosphates at the
termini. To anneal both oligos of the cassette together, mix them in equal amounts
then heat to 95 C for 5 minutes to get rid of any intramolecular interactions, then
allow to cool slowly to room temperature to anneal the two oligos together.
1. Dissolve dry oligos in 10 mM Tris pH 8, 1 mM magnesium chloride (the
magnesium chloride is important to facilitates stable annealing). Dissolve oligos
at a concentration of 50 M.
2. In a separate PCR tube add 50 L of both oligonucleotides that form a single
annealed double stranded cassette (e.g. to form a cassette containing a single
randomized codon) and mix by inverting tubes, or pipetting. Note the double
stranded cassette concentration is 25 M.
3. Label each cassette tube and put them into a rack that can float in water.
4. Boil 1 L of water in a glass beaker by microwaving for about 8 minutes.
5. Using heat protective gloves, pour the boiling water into a Styrofoam container.
6. Put the floating rack holding the tubes of cassettes into the boiling water.
7. Put a lid on and leave closed for five minutes.
8. Open lid about 2 inches and leave for 3 hours to slowly cool to room temperature.
9. Dilute each cassette to 0.5 M.
a. In a microfuge tube, add 4 L of the 25 M annealed cassette to 200 L of 10
mM Tris, 1 mM MgCl2.
b. Mix by vortexing.
III. Ligate cassettes into prepared vectors. This method relies on a gross molar excess
of cassette to plasmid (for a 50 bp cassette and a 5 kb plasmid, the molar ratio of this
recipe is about 100 molecules insert to 1 molecule plasmid). You can use more
cassette, but too much cassette can inhibit ligation by binding to the Mg in the
reaction. The protocol below is for ligating 10 separate cassettes into one vector (e.g.,
to generate randomized codons at 10 consecutive positions)
1. Prepare a master mix with
a. 33 L of vector (~5 nM)
b. 7.2 L of 10X ligase buffer
2. Pipette 3.6 mL of this master mix into 11 separate microfuge tubes.
3. Add 3 L of cassette at 0.5 mM (aka 500 nM) to the first 10 tubes.
4. Add 3 mL of water to the 11th tube (this is a control reaction to assess self-ligation
of the plasmid with no cassette insert)
5. To each of the 11 tubes:
d. Add 0.3 L of T4 DNA Ligase
e. Pipette gently to mix
f. Incubate at room temperature for 30 minutes.
2. Transform ligated DNA into competent bacteria. With each of the 11 tubes:
a. Put ligation reactions onto ice for 5 minutes to cool.
b. Thaw a frozen tube of chemically competent bacteria taking care to keep the
bacteria cold (e.g. thaw just to point of melting and then put onto ice).
c. Add 85 L of competent cells to each tube of ligated DNA.
d. Invert or flick tubes to mix.
e. Incubate on ice for 15 minutes.
f. Prepare water bath at 42 C (either using hot water from the sink in a
Styrofoam container or by adding water to a heatblock at 42 and leaving
enough time for it to temperature equilibrate). Check temperature of water
with a thermometer.
g. Add tubes with competent bacteria to 42C water bath for 45 seconds, then put
back onto ice.
h. Incubate on ice for at least 1 minute.
i. Add 1 mL of LB to each tube of cells.
j. Incubate at 37 C for 1 hour to allow cells to take up plasmid and express
resistance marker.
k. Plate 10 L of cells onto selectable plate (e.g., LB + Kanamycin). Put plate
into 37 C incubator overnight.
l. Put the remainder of the cells (~990 L) into a 50 mL flask with 15 mL of
liquid media (e.g. LB + Kanamycin). Grow this flask overnight at 37 C in a
shaking incubator. Note, the control reaction without insert does not need to
be put into liquid culture.
IV. Analyze plates
1. For a cassette with a single randomized codon there are 64 possible mutations. It
is good to oversample about 10-fold. This means that you want to have >640
colony forming units in your liquid cultures. If you have >6 colonies on the plates
this will be the case.
2. You also want to have a low background meaning fewer colonies on the no-insert
control than in your samples with cassettes. Generally a ratio of 5:1 or greater is a
very good indication that things are working well. Of note, lower ratios can yield
good libraries, so proceed with the protocol even if this analysis is not as good as
you would like.
V. Prepare DNA from liquid cultures.
1. You will have 10 cultures corresponding to randomized codons at adjacent
position in the gene (e.g. randomization of amino acids 11-20). Prepare a DNA
sample that contains an approximately equal mix of each of these ten cultures.
a. Add 1 mL of each of the ten cultures to a 14 mL falcon tube. Invert to mix.
b. Transfer 3 mL of the mixed bacteria to a round bottom tube.
2. You should also prepare DNA from an individual culture as a control.
a. Transfer 3 mL of one of your individual cultures (e.g. containing mutations at
position 11) to a round bottom tube.
3. Prepare DNA from both of theses 3 mL samples (10 amino acid library and single
position control) in round bottom tubes.
a. Spin samples at 3900 RPM in the Sorvall centrifuge for 5 minutes.
b. Pour supernatant into waste container (bleach this before disposing in sink).
c. Add 250 uL of P1 buffer to your cell pellets. Pipette up and down 20 times to
resuspend. Transfer sample to a microfuge tube.
d. Vortex the microfuge tube.
e. Add 250 uL of P2 buffer to your sample. Invert 10 times to mix. The sample
should look viscous, but not have large cloudy clumps.
f. Add 350 uL of N3 buffer to your sample. Invert 20 times to mix. The sample
should be uniform in color with lots of fluffy precipitate.
g. Spin at maximum speed in a microfuge for 10 minutes.
h. Transfer the supernatant to a silica column (recommend large format that
holds 25 ug of DNA).
i. Spin at 13000xg for 1 minute.
j. Discard flow thru (save for chemical waste).
k. Add 500 uL of Wash buffer with Ethanol to the column.
l. Spin at 13000xg for 1 minute.
m. Discard flow thru (save for chemical waste).
n. Spin at 13000xg for 1 minute to thoroughly dry the column.
o. Transfer the column to a clean microfuge tube.
p. Add 60 uL of elution buffer (10 mM Tris, pH 8)
q. Spin at 13000xg for 1 minute.
r. Save and label the microfuge tube with your purified plasmid DNA.
4. Perform quality control checks of your purified DNA.
a. Measure the absorbance spectrum and estimate the DNA concentration from
the A260 value.
b. Perfrom a restriction analyses. Digest a small portion of your DNA samples
with enzymes that bracket the open reading frame as well as an enzyme that
cuts the parental vector, but not after cassette ligation (e.g., the enzyme used
in section I, part 2 – often SphI or EcoRV). Run the digested DNA on an
agarose gel stained to visualize the DNA.