Selex

SELEX: Systematic Evolution of Ligands via Exponential Selection is the process of identifying specific aptamers.

To select a binding aptamer, you don't look for epitopes. It simplifies the process as you don't need to design a determined structure, but to reduce little by little an already binding population.

The SELEX screening process starts with a random library of nucleotide oligomers of a fixed size with known sequences in each end (for further amplification by PCR). Then the library is incubated with the target molecule. A number of this random sequences will bind to the target and the unbound sequences will be discarded.

The bound sequences are now separated from the target molecule (elution step) and amplified to create a new library.

With every round, more and more oligonucleotides with low binding-affinity for our protein of interest will be discarded leaving only strong binding aptamers at the end:

This process used to imply a high amount of time, money and expertise to be performed correctly. This is why it has been so difficult to earlier iGEM teams to work with aptamers, or why they skip the aptamer developing process and work only with already discovered aptamers.

For the first time in iGEM we present a successful, affordable, and fast SELEX process. You could see the details on how to perform it in our protocol page:

Selex Protocol

We hope that with this approach, in recent years, we will see the aptamers registry page getting larger and larger.

Our experiment:

To separate the bound sequences from the unbound ones, we decided to use a nitrocellulose membrane.

Nitrocellulose membranes can bind protein through hydrophobic interactions, so when applying the incubated library pool with the target protein to the membrane and forced the liquid to pass through it, the protein will stay in the membrane as well as the bound sequences.

The pore size was chosen after folding the aptamers and testing both 0,22 um and 0.45 um pore size. After analyzing the results we conclude that the 0.22 um pore was too small and the aptamers were being trapped because of their size and not by unspecific interactions.

The other improvement and the most relevant one was the column system we designed and 3D printed to conduct our experiment. Its design was calculated to fit in a 1.5 ml Eppendorf tube, allowing us not only to lose as little as possible flow-through in each experiment ( take into account that the volumes we use are in a microlitre scale) but also to save material.

Another change we introduced with our columns, was to use a centrifuge instead than a vacuum bomb, to force the liquid to pass through the membrane. A centrifuge is almost an essential item in a laboratory and extremely easy to use, so in the end, we simplified the protocol.

You can download the columns stl files from our github repository and see further instructions on how to print it, pre-treat it and mount it at our protocol page:

Selex Protocol

Initially, we tested two different kinds of material: PLA ( the most commonly used when 3D printing) and PTEG. Our first choice was the PLA, however, when using the columns made with PLA, we observed a bit of contamination in the nanodrop curve around 260 nm. With the PTEG material, the curve showed no contamination whatsoever.

We tried in parallel two different proteins:

1. BSA

2. Ole-e1

BSA

We tested the detection limit of the ponceau dye to visualize the protein on the nitrocellulose membrane, which was 25 ug. This was decided to allow us to be able to detect, during the process, if the protein was being correctly binding with the membrane:

Figure 1. It is shown the OD600 measurement by the first calibration experiment. The corrected Abs600 is made subtracting H2O measurements from LUDOX CL-X ones. The OD600/Abs600 result (3,652) is the multiplication factor, which you have to use after a cell density measurement with the plate reader to have a correct analysis.

From that point on, the ratio between the aptamer/protein was 1:1. The recommended one is 1: 10, protein/aptamer, so there is competence for the binding sites and only the sequences with the best affinity are selected.

Using our ratio means that further rounds of selection will be needed in order to achieve the same range of affinity than other SELEX selection using less protein.

We start the Selex by dismissing the sequences that bind unspecifically to the system itself. We fold the aptamers and forced them through the membrane and keep the flowthrough.

With each round, we will repeat this process before incubating with the target protein and see that the amplified libraries did not bind to the membrane alone, only after incubation with the protein:

Table 1. Nmol of aptamers trapped in the membrane and the % of sequences retain by unspecifically interactions for each round of Selex with BSA protein.

After the incubation with the target, we forced the mix through the membrane again but this time keeping the membrane. Because of the amount of protein, we use in each round and as in can be seen in figure 1, not all the protein was being trapped by the membrane and in all the measures done by the nanodrop the curves from the protein concentration and DNA concentration overlap, making impossible to measure the flowthrough.

During the consecutive rounds, the number of cycles to amplified by PCR were reduced from 30 to 10:

- The second round of selection..

Figure 2: A:Agarose gel after 10 cycles of amplification. B: Agarose gel after 30 cycles of amplification.

- Fourth round of selection:

Figure 3: Agarose gel after 10 cycles of amplification.

OLE-E1

We start the SELEX protocol as we did with BSA, with the exception that we followed the 1:10 ratio of aptamer/protein proportion, so for this experiment, we used 2 ugs of protein for each round.

We had problems with the amplification of the rounds of selection so instead of checking the proper number of amplification cycles, we fixed 50 cycles. We eventually could manage to see enough DNA in the agarose gel, but the number of cycles boosts the creation of concatamers. Concatamers are long continuous DNA molecules that contain multiples copies of the same DNA sequences linked in series.

Not only were clearly seen in the agarose gels. The amount of DNA that was being trapped in the membrane before the incubation with the target protein increased too.

Figure 4. A: Agarose gel after the second round. B: Agarose gel after the sixth round.

Table 2. Nmol of aptamers trapped in the membrane and the % of sequences retain by unspecifically interactions for each round of Selex with Ole-E1 protein.

At the end we finish for each protein:

- BSA: six rounds.

- Ole-1: seven rounds.

qPCR

The real-time PCR can show the evolution and enrichment of your selection process. When the amount of different sequences is very high, like in the initial population, the fluorescence star to grow and reaches a peak before decreasing.

This happens because as the SELEX id performed, the number of different sequences are drastically reduced, therefore the amplification can be done as usual and the characteristic sigmoid curve finally appears.

This happens because as the SELEX id performed, the number of different sequences are drastically reduced, therefore the amplification can be done as usual and the characteristic sigmoid curve finally appears. With each round of selection, we are able to reduce the number of sequences until the ideal curve it's achieved.

Figure 5: Graph with the cycles of the rounds of qPCR against Relative Fluorescence Units (RFU) for the different round of Selex.

Using the qPCR to check the selection and the numbers of sequences we are left with improves the work of the UBonn iGEM team characterization because they use next-generation sequencing to see the number of sequences and their representation. With our technique, we still don’t the sequence, but we can know the number of sequences there are, and until confirming their affinity with the target, the sequence is less important. The final advantage is that using qPCR saves time: a day instead of 1 month and can be used frequently to check rounds.