(Cell-)SELEX
To generate the aptamers for our test, we are using the Systematic Evolution of Ligands by EXponential Enrichment (SELEX) technique.
Aptamers are short single-stranded oligonucleotide molecules that bind specifically to a variety of target molecules1. These target molecules can be anything, from small organic molecules over proteins up to whole viruses and cells1. The high specificity and affinity of the aptamers to the target molecule are caused by the folding of the aptamers into a unique 3D structure that interacts specifically with the target1. We are using aptamers for our test instead of traditional antibodies, because aptamer sequences can be evolved quicker than antibodies and for a wider range of targets2. They are also cheaper and more versatile in production as they can be easily produced in vitro. Additionally, in contrast to antibodies, they do not require animals for production2.
The Systematic Evolution of Ligands by EXponential Enrichment (SELEX) is a technique in molecular biology to evolve aptamers2. The procedure is based on the random specific binding and washing of non-specific aptamers to and from the target molecule and their subsequent amplification1.
There is a variety of SELEX methods available for various targets3. We used the Cell-SELEX technique, because we wanted to evolve aptamers that bind specifically to the cell surface of the plant pathogen Pseudomonas syringae which we used as a proof of concept for the detection.
Since Cell-SELEX is a difficult method, we invested a lot of time and effort to establish it in our lab. There were a lot of things we had to consider before we could even start with the different parts of our individual Cell-SELEX protocol. Needless to say, we had a lot of troubleshooting to do, once we actually started. To help future iGEM-teams that will use the Cell-SELEX method, we provide a detailed Cell-SELEX description. This will hopefully lead to a more efficient workflow.
Figure 1: Schematic representation of aptamer evolution through Cell-SELEX.
Target cells
In the Cell-SELEX method, the aptamers bind to molecules on the extracellular surface1. Cell-SELEX uses live cells and reducing the amount of dead cells in the solution significantly enhances the enrichment of the potential target aptamers1. Our targets were the cells of the plant pathogen Pseudomonas syringae. We calculated the amount of cells needed for each SELEX-cycle using the OD of an overnight culture. We then made a cryo culture and aliquoted it, so the culture wouldn’t have to go through multiple freeze-thaw-cycles, which can be harmful for the cells. Before using a cell-aliquot in the cycle, we centrifuged and washed the cell suspension two times to receive a clean cell pellet. The supernatant was removed and the pellet was resuspended in a binding buffer. The binding buffer consists of DPBS, BSA and Tween 201, 2.
Library
The library contains about 1014 different oligonucleotides, which share a randomized region of 30 - 50 nucleotides2. The randomized region is flanked by two conserved regions2. These conserved regions function as primer binding sequences in the amplification step. The regions we used were designed by Prof. Dr. Günter Mayer and supposedly reduce the amount of miss amplification4. Because of the sheer number of sequences it is likely that a few of them are able to bind to the target with high specificity. Before use, the library was mixed with the binding buffer, heated to 95 °C for 10 minutes to break the nucleotides down into their primary structure. They were then snap-cooled on ice to keep this structure. This is essential for the aptamers to exhibit their maximal binding potential.
Positive selection
After mixing the library with the target cells, some of the oligonucleotides will bind to the target with varying binding affinities. To increase the specificity of the aptamers, you could also combine the positive selection with a subsequent negative selection. This method is useful to eliminate nontarget cell-specific oligonucleotides5. This is possible due to the binding of the aptamers to the specific molecules on the extracellular surface of the cells1. So the negative selection filters out sequences that may bind to molecules existing on the surface of both the target and the different cell lines1. After some negative selections most of the nonspecific bound aptamers can be eliminated and they will not be amplified in the selected pool1.
After we mixed the target cells and the library together, we incubated the mix on a shaker at 500rpm for 1h at 4 °C.
Extracted bound sequences
Now sequences that are bound to our target cells and the unbound sequences need to be seperated. To remove the aptamers that did not bind to our cells, we used Amicon Filters (0.5 ml 100 kDa cut off). The Amicon Filters let the unbound aptamers flow through and the aptamers bound to the target cells remain in the residue, as the cells cannot pass through the filter. The target cells with the bound sequences then get heat shocked at 95 °C, to denature the proteins on the cell surface and to revert the aptamers binding1. After the heat shock, the mix has to be centrifuged again, to separate the aptamers from the cells.
After a couple of centrifugation and washing steps, we heat shocked the cells to destroy them. Through centrifugation, we created a pellet again, to remove all the cell debris. The aptamers are located in the supernatant and can now be collected.
PCR amplification
To amplify the extracted sequences, we used PCR amplification. The determination of the optimum cycles is very important, because too many cycles can lead to nonspecific amplicons1.
The primers we used for our Cell-SELEX were also designed by Prof. Dr. Günter Mayer4. We used his primers to reduce misamplification during the PCR. The primers were also 5’-phosphorylated, to create single-stranded DNA in the next step. First we did a preparative PCR with 8 cycles to increase the amount of sequences in general. After that we used 5 different samples to determine the right amount of cycles. We stopped the reactions after 4, 6, 8, 10 and 12 cycles. We used a gel electrophoresis to determine the right amount of cycles, but as there were no visible bands, we could not determine the right amount. Because of the lack of time we were not able to establish a finished PCR procedure for our cycle.
Figure 2: Electrophoresis to determine the right amount of PCR cycles
Gel:
3 % TAE Agarose
5 µl Gel red in 50 ml
Lane (from left to right):
- 100 bp ladder
- ssDNA pool
- After 8 rounds of PCR
- After 8 rounds of PCR + 4 rounds of PCR
- After 8 rounds of PCR + 6 rounds of PCR
- After 8 rounds of PCR + 8 rounds of PCR
- After 8 rounds of PCR + 10 rounds of PCR
- After 8 rounds of PCR + 12 rounds of PCR
Digestion
After the PCR reaction the oligonucleotides are now double-stranded. But the aptamers need to be single-stranded DNA. For the next cycle, the 5’-phosphorylated strands are digested by a lambda exonuclease. In our case we used the lambda exonuclease from New England Biolabs (catalog #M0262S). The Lambda Exonuclease is a DNA-specific exonuclease. It is a highly processive 5’-3’ exonuclease that selectively degrades 5’-phosphorylated strands6. The exonuclease prefers 5’-phosphorylated strands, due to the formation of inert enzyme-substrate complexes7. Since we had to prove that the lambda exonuclease works as intended for our cycle, we worked with different methods. First we tried to prove it by measuring the concentration of single-stranded DNA with the Nanodrop by ThermoScientific. But a better and more visual verification was done with a 2 % TBE Agarose gel. The digested samples (single-stranded DNA) traveled a longer observable distance through the gel compared to the undigested samples(double-stranded DNA).This indicates the successful digestion of our sample.
Figure 3: Electrophoresis for digestion verification.
Gel:
2 % TBE Agarose
2,5µl Gel red
75V for 90 minutes
Lane (from left to right):
- single-stranded DNA (Clean up after digestion)
- double-stranded DNA reference
- single-stranded DNA (no clean up after digestion)
- 50 bp ladder
Synthesised aptamers
The whole process can now be repeated. After a variable amount of positive SELEX-cycles (typically 6-20 cycles), aptamers with a high binding affinity and specificity can be evolved for the cells of our target, Pseudomonas syringae.
Ampelpflanze
Figure 4: Infiltration of Nicotiana benthamiana for transient expression of the 'RUBY35S'. N. benthamiana and construct were kindly provided by Vanessa Reichel Deland.
As a contribution, we designed a reporter system to detect drought and pathogenic stress in plants. The German word "Ampelpflanze" translates to "traffic light plant". In our project we aimed to generate a plant which gives a signal when the plant is suffering from stress.
To that end we planned to transform the model organism Arabidopsis thaliana with plasmids that contain different promoter-reporter constructs, which upon induction by their respective stressor-induced promoter lead to the change of the plant's color. That color change is visible to the naked eye and is meant to be an indicator to visualize potential abiotic and biotic stresses within the plant.
We chose Arabidopsis thaliana as it’s genome was fully discovered with many established cloning methods available.
We wanted to generate two reporter systems. The first one consists of the CAP160 promoter which is induced upon drought stress, and the yellow Chromoprotein sequence (amilGFP) that serves as a reporter. The other construct is made of the pathogenesis-related gene 1 (PR1) promoter8 and the RUBY-reporter that generates a red colored pigment9.
The PR1-expression is induced when the plant is infected by Pseudomonas syringae. We want to use its promoter to express the RUBY which is a reporter that encodes three enzymes, P450 CYP76AD1, L-DOPA 4,5-dioxygenase and a glucosyltransferase which are needed to catalyze the betalain synthesis, a red tyrosine derived pigment.
The CAP160 promoter is induced by drought stress and in our case will express the amilGFP that has a yellow color. We obtained the chromoprotein from the iGEM registry and it serves as the reporter gene. This yellow protein originates from the coral Acropora millepora and was codon optimised for Arabidopsis thaliana via jcat.
These constructs were first built with a P35S promoter as control. We used the P35S, as it is a constitutive promoter that has been tested before. This way we could make sure that our reporter systems worked. To be able to introduce our reporters into the plant we used the B415p9o Vector. It is a binary vector which means it contains selection markers for both plants and bacteria.
While many other reporters like GFP or GUS need more expensive equipment to monitor and often require plant sacrifice, RUBY is easy to monitor through many cell cycles as the generated pigment is visible to the naked eye.
You can learn more about RUBY and how it works here!
CAP160 (RD29B) This drought induced promoter was chosen because its specific expression stated in literature 10.
Methods
The following parts were used to generate each construct:
Vector - B415p9o Binary plasmid Promoter - CAP160, PR1 or P35S Reporter - RUBY, Chromoprotein/AmilGFP Terminator - Arabidopsis HSP18.2
The following constructs were designed using Snapgene:
Drought induced yellow construct B415p9o_CAP160 promoter_Chromoprotein_Terminator
Constitutive yellow construct as control B415p9o_P35S_Chromoprotein_Terminator
Pathogen induced red construct B415p9o_PR1 promoter_RUBY reporter_Terminator
Constitutive red construct as control B415p9o_P35S promoter_RUBY reporter_Terminator
gDNA was extracted from Arabidopsis thaliana wild type Col-0 and used as a template for PCR to obtain the PR1 and CAP160 promoters that occur in its genome. We first tried to obtain the promoter sequence from UDP-glucosyltransferase 74F2 (UDP), instead of the PR1 promoter. As we weren’t able to amplify it, we swapped to the PR1 promoter as our pathogen induced promoter.
The chromoprotein was first codon optimised for Arabidopsis thaliana via jcat.de and then ordered as a synthesized sequence. To obtain the RUBY Cassette we ordered the RUBY35S plasmid on Addgene. This Plasmid was also used to amplify the P35S promoter and the Arabidopsis HSP18.2 Terminator.
All primers were designed with snapgene and ordered from IDT.
The restriction sites we used were SfiI, SbfI, SpeI, PmeI, AscI and KpnI. All parts were ligated into the B415p9o vector using the T4 ligase. The vector contains a spectinomycin resistance cassette for bacteria and a kanamycin resistance cassette for selection in plants. To make sure the digestion was successful the digested parts were run through a 1% agarose gel (w/v).
After ligation, the plasmid was used to transform Escherichia coli. After growing for 1 day at 37°C, a colony PCR was performed to confirm the ligation of the desired insert. The plasmid was then isolated and sequenced.
The generated constructs were used to transform Agrobacterium tumefaciens using electroporation. Positive colonies were screened for via colony PCR.
Figure 5: Two plates with positive colonies.
Rifampicin (Rif), Gentamicin (Gent) and Spectinomycin (Spec) were added to the YEP medium. Rif and Gent, to select the Agrobacterium strain - GV3101 pPMP90 and Spec as the selection marker of our plasmid. Out of these colonies, we successfully isolated and sequenced the plasmids.
Both control plasmids for the RUBY and the chromoprotein were confirmed by sequencing. These plasmids were then used to transform Arabidopsis thaliana Col-0 plants.
For this, we inoculated 300ml YEP media with Rif, Gent and Spec with transformed Agrobacterium. The Agrobacterium was then transferred into small containers and 5% sucrose was added. The silques of the Arabidopsis plants were removed carefully. Then we dipped the inflorescence into the solution and let them soak for 30 sec to 1 min. The plants were left in the dark overnight at room temperature and put in a long day greenhouse until new silques developed.
Figure 6: Two pots of transformed Arabidopsis thaliana.
Figure 7: Dried Arabidopsis thaliana two transformed and one wildtype.
We planted the seeds we harvested on selective 0.5x Murashige-Skoogagar with kanamycin to prevent further contamination and to assure that only plants can grow that contain our vector with the corresponding resistance gene.
Results & Discussion
The first sequencing results showed that our RUBY and chromoprotein control plasmids with the P35S promoter were successfully introduced into Agrobacterium.
The plant dipping also resulted in a transient expression in the transformed plant, which was an indicator that our RUBY was working!
Figure 8: Arabidopsis plant one week after the transformation showing transient expression.
Unfortunately the plants with our chromoprotein construct died one week after transformation. Here we could observe a color change to yellow, as expected. Whether this happened naturally due to drought or insufficient light or as a consequence of our transformation is debatable. Agrobacterium tumerferensis usually does not seriously harm plants. Although the plant died, we could harvest approximately 15 seeds.
Figure 9: Transformed Arabidopsis thaliana dying one week after dipping.
The seeds of the transformed plants were able to grow on the 0.5x MS medium with kanamycin, indicating that they had the kanamycin resistance from our plasmid. The seedlings themselves did not become red during their growth. This lack of red coloration indicates an unsuccessful transformation, however since the seeds were unconfusably red, this option seems unlikely. Another potential cause for the loss of pigmentation could be low expression levels of tyrosine in seedlings. More transformation events are required for the assessment of the functionality of these reporter systems in the model organism Arabidopsis thaliana. In addition, the sequencing of the T1 generation is of vital importance for the confirmation of correct transformation of the plants.
Figure 10: Seedlings grown from seeds from *Arabidopsis thaliana* plants transformed with the P35S_RUBY-construct.
iGEM wikisync tool
Thanks to last year's iGEM Team BITS Goa and their very useful tool called igem-wikisync we could head start our wiki and have a well thought out workflow. Unfortunately for us (and we are sure for some other teams too) it had some minor flaws regarding UTF-8 encoding and special characters.