The overall goal of this protocol is to show how to construct and screen mutant libraries in Saccharomyces Cerevisiae for directed evolution experiments. This method can help answer key questions in laboratory creation use for focused directed evolution experiments such as how to assign overlapping areas for in the assembly and cloning. The main advantage of this technique is its simplicity and robustness since in just one transformation step it's something levels with good quality can be assigned.
This protocol for creating and screening mutant libraries will be demonstrated for a fungal aryl-alcohol oxidase or AAO. The regions for focused directed evolution are first chosen with the help of computational algorithms based on the available crystal structure or homology models of the enzyme. Two regions of AAO will be targeted for random mutagenesis and recombination:the M1 region and the M2 region.
Mutagenic PCR will be performed to amplify the targeted areas. To promote in vivo splicing in yeast, overlapping areas between segments of approximately 50 base pairs each will be created by superimposing PCR reactions of the defined regions. Prepare 50 microliter volume mutagenic PCRs, each containing 46 nanograms of DNA template 90 nanomolar each of sense and antisense primers 0.3 milimolar of DNTPs, 3%of DMSO, 1.5 milimolar of magnesium chloride, 05 milimolar of manganese chloride, and 05 units per microliter of tack DNA polymerase.
Use the following PCR program. 95 degrees Celsius for two minutes, 95 degrees Celsius for 45 seconds, 50 degrees Celsius for 45 seconds, and 74 degrees Celsius for 45 seconds for 28 cycles, and 74 degrees Celsius for 10 minutes. The remainder of the AAO gene, the HF region, will be amplified by high fidelity PCR with the corresponding areas overlapping the mutagenic segments, and or linearized vector overhangs included.
Prepare the high-fidelity PCR in a final volume of 50 microliters containing 10 nanograms of DNA template, 250 nanomolar each of sense and antisense primers, 0.8 milimolar of DNTPs, 3%of DMSO, and 02 units per microliter of iProof ultra high-fidelity DNA polymerase. Use the following PCR program. 98 degrees Celsius for 30 seconds, 98 degrees Celsius for 10 seconds, 55 degrees Celsius for 25 seconds, and 72 degrees Celsius for 45 seconds for 28 cycles, and 72 degrees Celsius for 10 minutes.
All the PCR fragments are then purified with a commercial gel-extraction kit according to the manufacturer's protocol. The next step is to linearize the vector to create flanking regions of approximately 50 base pairs that are homologous to the five prime and three prime ends of the target gene. Prepare a 20 micoliter linearization reaction mixture containing 2 micrograms of DNA, 7.5 units of BamH-one, 7.5 units of XHO-one, 20 micrograms of BSA, and two microliters of 10x buffer BamH-one.
Incubate the reaction mixture at 37 degrees Celsius for two hours and 40 minutes. After that, inactivate the reaction at 80 degrees Celsius for 20 minutes. To purify the linearized vector to avoid contamination with the residual circular plasmid, load the digestion reaction mix into the mega-well of a semi-preparative low-melting point Agarose gel.
Load five microliters of the reaction mix in the adjacent well as a reporter. Run DNA electrophoresis at four degrees Celsius and five volts per centimeter between electrodes. Then separate the Agarose gel corresponding to the mega-well and store it at four degrees Celsius in one XTAE.
Stain the lane with the molecular weight ladder and the reporter. Visualize the bands under UV light, and nick the position where the linearized vector places. In the absence of UV light and using the guidance of the nicks in the stained reporter lane, identify the linearized vector in the mega-well fragment and excise it.
Extract the linearized vector from the Agarose and purify it with a commercial gel extraction kit according to the manufacturer's protocol. Subsequently, the purified linearized vector is mixed with the PCR fragments and this DNA mixture is used to transform competent yeast cells using a commercial kit according to the manufacturer's instructions. Plate the transformed cells on SC dropout plates and incubate them at 30 degrees Celsius for three days.
To prepare for this assay, pick individual colonies from the SC dropout plates and transfer them to 96 well-plates containing 50 microliters of minimal medium per well. In each plate inoculate column number six with the parental type as an internal standard. As a negative control, inoculate well H-one filled with SC mediums supplemented with uracil with URA3 negative ESI visceral cells with no plasmid.
Cover the plates with their lids, wrap them in para film, and incubate at 30 degrees Celsius in a humid shaker for 48 hours. After 48 hours, add 160 microliters of expression medium to each well and reseal the plates. Incubate for a further 24 hours.
After centrifuging the plates use a liquid-handling robotic multi-station to transfer 20 microliters of the supernatant from the wells in each plate to a replica plate. Add 20 microliters of two milimolar P-methoxybenzyl alcohol and 100 milimolars sodium phosphate buffer ph 6.0 to each well. Stir the plates briefly with a 96-well plate mixer and incubate them for 30 minutes at room temperature.
Next, add 160 microliters of the FOX reagant to each replica plate and stir briefly with the mixer. Read the plates at 560 nanometers on a plate reader. Incubate the plates at room temperature until the color develops.
Measure the absorption again. The relative activity is calculated from the difference between the absorption value after incubation and that of the initial measurement normalized to the parental type for each plate. This representative gel image shows the linearized vector in lane two, the M1 PCR segment in lane three, the M2 PCR segment in lane four, the HF PCR segment in lane five, and the in vivo reassembled vector linearized with NHE one in lane six.
This next gel shows the plasmid mini-prep of the reassembled vector in lane two, the plasmid linearized with NHE one in lane three, the plasmid linearized with BamH one and XHO one in lane four, and the linearized plasmid obtained by gel extraction and cleanup in lane five. Mutational loads were adjusted by sampling mutant libraries with different landscapes calculating the number of clones with less than 10%of the parental enzyme activity and further verification by sequencing a random sample of active and non-active variants. An evaluation of the detection limit of this assay showed that the assay was linear in the presence of sorbitol represented by black circles.
In the absence of sorbitol represented by the white squares, linearity was more persistent but the response was weaker. A linear correlation was observed between AAO concentration and absorbance. A mutant library of 2, 000 clones was constructed and screened with this assay.
The shadowed square indicates the AAO mutants with notably improved secretion and activity against P-methoxybenzyl alcohol. Once mastered, mutant libraries can be made in approximately four hours if it is performed properly. When attempting this procedure it is important to remember to assign proper overlapping areas for each PCR family and linearized vector for in vivo splicing and cloning in a single transformational step.
Following a similar approach, other methods like in vivo DNA sampling, saturation mutagenesis library or DNA assembly can be performed in order to construct mutant libraries and, or metabolic pathways. After its development, this technique paved the way for researchers in the field of bioengineering to explore activities, stabilities, or even to invent infinite kinds of interaction. After watching this video, you should have a good understanding of how to harness this Saccharomyces Cerevisiae device for library construction and screening
