Design of our NeuroDrop device

Team Grenoble-Alpes has developed a Clickable Outer Membrane design to generate a sensitive and flexible system. The goal was to have a common biological basis that can be easily adaptable for all types of biomarkers. This biological part has to be able to produce a detectable signal when any type of biomarker is recognized.
In this attempt, it is crucial that the system must be easily modular in function of the selected biomarker in addition to have a very strong reporter system.



Etymologically aptus means to fit, and mer is to design the smallest unit of repeating structure. Aptamers are single stranded oligonucleotides with a various affinity and specificity according to the target. The targets can be as various as cells (bacteria or eukaryotic), proteins, small molecules and ions.

Most of the time they are produced by the SELEX method (Systematic Evolution of Ligands by Exponential Enrichment). However, non-SELEX methods exist[1][2]. These biochemical tools can have biotechnological applications in diagnosis or therapeutic according to the ligand. A comparison can be made with the antibodie’s properties.

Brief History

Aptamers have first been isolated in the 90’s by Larry Gold and Craig Tuerk. They observed some RNA strands, which can recognise DNA polymerase of T4 bacteriophage. On the other hand, Andy Ellington and Jack Szostak used RNA ligands against organic dyes. They called it in vitro selection. This method was then automatized in 2001, in order to reduce selection cycles from six weeks to only three days. The SELEX method was born.

This technique was patented by Gilead Sciences, a pharmaceutical firm, in order to select DNA oligonucleotides against thrombin. In 2002, natural aptamers are discovered in living organisms. Researchers have found a genetic controller based on nucleic acids. They called it riboswitch. They act by interacting with the DNA to control the expression.
Since, a lot of research groups all over the world work on aptamers.

The SELEX Method

The Systematic Evolution of Ligands by Exponential enrichment is the technique commonly used for the selection and the isolation of aptamers. This method combines both chemistry and molecular biology fields.

All of this begins with the synthesis of a library constituted of random DNA or RNA oligonucleotides. These sequences of oligonucleotide are incubated with the target, and thanks to their 3D folding the oligonucleotides can link the target or not. A rinsing permits the elimination of not linked fragments. Those which are linked are eluted and amplified by PCR for the next round of selection. Throughout the cycles, the conditions are more and more stringent in order to identify sequences with the better chemical property for the target.

Figure 1: SELEX Method.

Recognition Systems

Several mechanisms are used in apta-analysis. They are based on the folding structure of the aptamer. The modification of the structure changes the fluorescent signal detected and allows the detection or not.

First of all, the allosteric switch works with one single fluorescent modified aptamer (Figure 2). When the aptamer recognises the target, its 3D folding is structured. Some examples of these 3D structures are hairpins or G-quadruplex.

Figure 2: Allosteric Switch.
In purple the aptamer, the blue star represents the fluorescence.

A second technique is based on the use the aptamer and its fluorescent complementary strand (Figure 3). When the aptamer detects its target, the complementary fluorescent strand dissociates.

Figure 3: Motion of the complementary strand.
In green the aptamer, in purple its complementary strand, the blue star represents the fluorescence.

There also exists “sandwich” techniques, which are complex to implement, and require lots of optimisation.
The kissing complex (Figure 4) is a good example. It consists on coating hairpin DNA aptamer in a well. The potential target is recognised by the coated aptamer and then, a specific hairpin RNA aptamer can link the DNA aptamer by a loop-loop interaction. The affinity of this kind of system is very strong.

Figure 4: Kissing complex.
In purple the DNA aptamer, in red the RNA aptamer, the blue star represents the fluorescence.

There also exists one split aptamer system (Figure 5). However, the sequence is therefore shorter and the specificity may decrease. To avoid this loss of specificity a “sandwich” with two aptamers can be done (Figure 6). The condition is to have a sufficiently large target such as a macromolecule.

Figure 5: Split aptamer.
In purple the aptamer, the blue star represents the fluorescence.

Figure 6: Two aptamers "sandwich".
In purple and red two different aptamers, the blue star represents the fluorescence.

In our system, we use a “sandwich” technique with two aptamers, because our target is large enough to have two recognition sites. These two aptamers are clicked to OmpX proteins mutated from the E Coli membrane by a cyclo-addition mechanism called click chemistry (see Click chemistry below). The mutated OmpX proteins are each linked to a subunit of Adenylate cyclase either T18 or T25. By recognizing their target, the two aptamers of the sandwich system will induce a convergence. This will allow the reconstitution of adenylate cyclase and permit the production of cAMP, which will activate the transcription of the Nanoluciferase gene.


Aptamers have some interesting properties in the field of health. They can be used as therapeutics, for diagnosis or for analytical aim, environment and food safety.

As a matter of fact, aptamers can link small molecules, which can include contaminants, pollutants, drugs… Their high specificity makes them particularly efficient for analysis. Several approaches of their use were described, such as extraction cartridges with coated aptamers coated on it, combination with chromatography and diverse processes of apta-analysis.

In therapeutic, their low immunogenicity offers them a considerable potential. It is also very simple to add groups with various properties to them, and therefore use them for different functions, such as agonist or antagonist of a biological target. Moreover, their short half-life in the bloodstream because of their degradation by the large number of nucleases and their low molecular weight allow an easy kidney clearance, that decreases its toxicity. As consequence it can safely be used in the treatment of temporary phase like blood clotting, or on targeted organs like eyes.

Indeed, one of the best-known medicines with aptamers, and also the first one FDA approved is the Macugen (Pegaptanib), which treats the Age-related Macular Degeneration or AMD. In that case the oligonucleotides are modified with polyethylene glycol (PEG). They bind the Vascular endothelial Growth Factor with a high affinity and inhibit its activity.

One of the properties of aptamers is to specifically recognize a particular target leading them to diagnostic application. Each aptamer would diagnose a individual biomarker with high specificity. We have taken advantage of this feature, which makes our project very powerful because by selecting the appropriate aptamer, it is possible to diagnose any target biomarker. This makes our system universal.


In our project, we want to create an engineered E.coli bacterial strain acting as a “Biosensor”. This bacterium detects the presence of a biomarker of interest present in the tears, and in response will produce a detectable bioluminescent signal. For this purpose, we first selected a specific aptamer of the biomarker we want to detect, which is present in the tears.
This specific aptamer has been functionalized by a cyclooctyne group allowing its click chemistry fixing to the azide-OmpX. We had to modify the transmembrane protein, OmpX, expressed at the surface of the bacteria (within the outer membrane) by adding an azide functional group in one amino acid of OmpX, that will allow the click reaction with the specific aptamer. The signaling cascade, which is present inside the bacteria and conducts to the production of the bioluminescence, was based on a Bacterial Adenylate Cyclase Two Hybrid Assay (See further).

The aptamer part of the biosensor

The use of aptamers in our project gives a great flexibility and modality allowing to detect any biomarker whose aptamer sequences are identified. These aptamers will be the recognition elements of our biosensor.
Aptamers have been appointed as the solution because they have the advantage of disposing of a small size and a high stability while offering affinities similar to those of antibodies. Once functionalized by a cyclooctyne group [7], these aptamers can theoretically be fixed, by a click chemical reaction, on the transmembrane protein serving as a scaffold.

Figure 7: The amino acid functionalized with an azide allows any molecule functionalized by DBCO/DIBO to be attached to the cell membrane by the SPAAC click chemistry.

What is Click-Chemistry and why we use it in our project?

The click chemistry reaction used in our project is a chemical reaction class used to attach to a given substrate a specific biomolecule (Bioconjugation Reaction). This type of reaction is increasingly used in biology because it is a part of the so-called "bio-orthogonal" reactions that are able to be performed in biological conditions without interfering with other internal biochemical processes.

In other words, with this click-chemistry reaction, we could easily change the biomolecule clicked to a protein at the surface of the bacteria. But how does it work?

One of the most used chemical groupments in click chemistry is azide (Figure 2), which has several advantages : it is small, metabolically stable and lacks of reactivity[8].
Figure 8: The Azide Group.

Azide allows 3 bio-orthogonal click reactions :

  • Staudinger's ligature
  • Copper catalyzed azide alkyne cycloaddition (CuAAC)
  • Strain promoted Azide-Alkyne Cycloaddition (SPAAC)
One main advantage of azide, is that it could be incorporated in unnatural or synthetic amino acids during translation and protein expression, thus conferring a "clickability" on the proteins they integrate[9].

Which Clik-Chemistry have we chosen in NeuroDrop?

It has been shown that the binding of Staudinger is not sufficiently bio-orthogonal for some living organisms such as mice and that the kinetics of its reaction is slow which restricts its use[10].
It has also been described that CuAAC requires the intervention of copper, the latter representing a major inconvenient due to its high cytotoxicity for organisms[11].

Thus the choice was made for the SPAAC, which does not need the presence of copper, the latter being replaced by cyclooctyne group. This group can react selectively with azide thus ensuring a click chemistry reaction effective, specific and above all without toxicity (Figure 9)[12].

Figure 9: Principle of the SPAAC click chemistry used here: the molecule in green (A) is functionalized by an azide group; the orange molecule (B) is functionalized by a cyclooctyne group.

The transmembrane protein OmpX and its modification with the azide group

In order to create a signaling pathway on the outer membrane, our system requires intracellular signaling domains, that's why we chose the OmpX protein. This 171 amino acids protein is naturally expressed in the outer membrane of E. coli. In addition, its three-dimensional and crystallographic structure is well described[13].

Figure 10: A) The structure of the OmpX protein was elucidated by NMR and X-ray crystallography. B) Square residues are important for the secondary structure of OmpX. To keep the structure intact, we introduce an amber stop codon into loop 3 (L3). Adapted from:

In order to be able to click the OmpX protein, it must be modified so that it has an azide group outwardly to allow aptamers to bind by SPAAC reaction. This protein has also a second role because on its periplasmic C-terminus will be fixed one of the subunits of adenylate cyclase (T18 / T25) to ensure its function of signaling inside bacteria (More informations).
To make this modification, a mutation in its genetic sequence is necessary, this mutation will incorporate an unnatural amino acid functionalized by an azide group that allows the reaction of click chemistry.

Figure 11: A) For our signaling proteins, we chose OmpX, which were functionalized by introducing the amber stop codon into its loops (red). The signaling domains are fused to OmpX C-terminally.


In order to allow the expression of genetically modified OmpX with azide group in unnatural amino acid, we need to change our expression system. The expression system must integrate the unnatural amino acid. Our bacteria has to ensure the expression of the 2 key proteins for our system. This is why we must prepare them and equip them with the necessary tools to incorporate the unnatural amino acid into their proteins.

To realize our system, the bacteria we have chosen will be co-transformed with 2 plasmids: the first, pGRE-Alps, will contain our 2 transmembrane proteins fused with the subunits of adenylate cyclase. The second plasmid, pEVOL-pAzF, will provide the molecular tools necessary for the incorporation of the unnatural amino acid into the protein of the outer membrane (Figure 12).

Figure 12: As a proof of concept, we will construct and express our device within E. coli Cya-. To obtain this device, the E.coli cells will be co-transformed with two plasmids. The first plasmid, pGre-Alpes, carries the genes for the outer membrane proteins. The second plasmid, pEVOL-pAzF, is necessary for the incorporation of the unnatural amino acid within the outer membrane proteins.

Incorporation of the unnatural amino acid

In order to perform the click chemistry reaction, our membrane proteins must be functionalized with an azide group. We therefore need to incorporate the unnatural amino acid p-Azido-phenylalanine (pAzF) to OmpX. This incorporation is only possible by the suppression of the Amber stop codon, the expression of the aminoacyl tRNA synthetase of pAzF and finally the presence of pAzF in the medium.

DNA is known as the carrier of hereditary information, this information is composed of 4 basics pairs: adenine, guanine, cytosine and thymine. To express it 2 mechanisms are involved: the first is to transcribe the information in the form of messenger RNA (ribonucleic acid). Once obtained, it will be used for the second step which is the translation. The base pair sequences are translated into a succession of amino acids in the form of a codon (triplet). These triplets are recognized by the transfer RNAs (tRNAs) which carrie the right amino acid to incorporate (Figure 13).

Figure 13: There are many different types of tRNAs. Each type reads one or a few codons and brings the right amino acid matching those codon. In elongation, the mRNA is read one codon at a time, and the amino acid matching each codon is added to a growing protein chain. Adapted from Khan Academy Website.

In addition to coding for amino acids, some triplet known as stop codon do not correspond to any amino acid. They have the role of stopping the elongation of the peptide chain during translation. On a total of 64 codons, there are 3 stop codons which, instead of fixing the tRNA, will allow the recruitment of ribosome dissociation factors, thus liberating the amino acid chain formed in the cytoplasm. It will undergo various changes in its structure to ensure its function.

In nature there is no tRNA capable of recognizing stop codons and all the natural proteins are constituted and formed from a score of natural amino acid.
In 2002, a team of researchers in California was able to develop a molecular system capable of inserting the unnatural amino acid into E.coli proteins[14] by incorporating unnatural tRNAs that recognize a stop codon. This tRNA is used to compete with the ribosome dissociation factors to incorporate the unnatural amino acid.

In the cytosol, these tRNAs will carry the unnatural amino acid, and as they have a high affinity for the chosen stop codon, a competition will take place between them and the dissociation factors. If the tRNA binds to the ribosome, an unnatural amino acid will be incorporated into the peptide chain. This results in a protein that incorporates an unnatural amino acid.
Among the 3 existing stop codons, it is the Amber codon (TAG) which was chosen, because it is the least abundant in E.coli. In our project we will insert this codon so that the amino acid once incorporated and will be exposed to the outside at the 3rd loop, ensuring the feasibility of the click chemistry reaction.

In E. coli, out of a total of 4290, there are only 326 reading frames that ends with the Amber codon[15]. That is why it was chosen because it should cause as little damage as possible to E. coli cells. It was also shown that the misclassification of E. coli genetic code of this way resulted in phenotypic changes and this was also observed in our results, because bacteria whose genetic system was diverted, were smaller than the control.

Clickable amino acid: p-Azido-Phenylalanine (pAzF)

As explained in the "click chemistry" section, SPAAC is a bio-orthogonal, rapid and non-toxic reaction that requires the presence of two groups: azide and cyclooctyne.

In our project the azide function will be present on pAzF. In 2002 a team of researchers succeeded in mutating an existing tRNA synthetase so that it could incorporate pAzF[14] This will make it possible to use cells for synthesizing proteins to which any DIBO / DBCO-modified compound may be attached and which will allow, in our case, to have clickable membrane proteins which will serve as scaffolds for the aptamers.

This strategy has already been used by 2 iGEM teams from Eindhoven in 2015 and Eindhoven in 2014.

OmpX and the Clickable Amino Acid (pAzF)

The residue to be substituted with the unnatural amino acid has been chosen so that the aptamer will have no difficulty attaching to. Moreover, it was chosen so that the structure of OmpX remains as stable as possible. And that's why it seemed obvious to us to choose the Tyrosine residue located at the 3rd loop, because the tyrosine points outward of OmpX (Figure 14), the tyrosine strongly resembles the unnatural amino acid functionalized with the azide function, which ensures stability to the OmpX structure.
The azide function of p-AZIDO-L-phenylalanine (pAzF) will enable the realization of the click reaction and thus fix the aptamers spontaneously and efficiently.

Figure 14: The unnatural azide-functionalized amino acid closely ressembles Tyrosine.


To functionalize our cells, we will co-transform two plasmids. The first plasmid is pGre-Alps: which is a pUT18 plasmid to which the sequences of the two proteins have been added. The second vector is the pEVOL-pAzF plasmid which contains the tRNA sequence and the aminoacyl-tRNA synthetase which are necessary for the incorporation of the unnatural amino acid.


It is a vector designed and optimized for the incorporation of the unnatural amino acid (pAzF) into proteins in E. coli. The coding sequence that it contains codes for Aminoacyl tRNA synthetase, which makes it possible to translate the sequence of the Amber stop codon by the incorporation of the non-natural amino acid. This vector has been optimized and improved to make incorporation of the amino acid 250% more effective[15]. Note that one of the first unnatural amino acids to be incorporated into endogenous E. coli proteins using the pEVOL vectors was pAzF. We will use exactly the same vector to build our mutant proteins in vivo.

  • The p15A origin of replication ensures a wide range of compatibility with other plasmids. The compatibility of the origins of replication is very important, because if origins of replication of two plasmids are similar, the bacteria can not distinguish between them and may possibly lose one or the other plasmid because the amount of two plasmid is limited to a single copy number. This is why plasmid compatibility is defined as the failure of the stable inheritance of two co-resident plasmids in the absence of external selection[16].
  • It also features a chloramphenicol resistance gene.
  • Aminoacyl-tRNA synthetase is under control of Arabinose inducible promoter (AraBAD).

Figure 15: Simplified vector map of pEVOL-pAzF.


It is a vector that we have created. It is derived from pUT18, itself derived from the pUC19 high copy vector, expressing an ampicillin resistance marker, and encoding for the T18 fragment (amino acids 225 to 399 of CyaA) which is expressed under the transcriptional control of a lactose promoter. The T18 open reading frame is located downstream of a MCS with 9 unique restriction sites. This plasmid is designed to express chimeric proteins in which a heterologous polypeptide is fused to the N-terminus of T18 (OmpX). In our case, the sequence of OmpX binds to the N-terminal end of T25 will be added to it, and the inducible promoters will be replaced by constitutive promoters, thus forming our new plasmid pGre-Alps which will be able to express the 2 key proteins of our System.

Figure 16: Simplified vector map of pGre-Alps.


Many experiments were planned to verify the feasibility of our system and more precisely the feasibility of the click reaction. Click validation allows as well to validate the expression system.


Because of the efficiency and selectivity of SPAAC and the fact that no additional reagent has to be added to the reaction mix, SPAAC was chosen to further explore functionalizing bacterial membranes.

Currently, the fastest known cyclooctine is BARAC. However, this cyclooctine has been found to be sensitive to the addition of Michael by thiols, which is not favorable when the system is applied to a protein, since proteins contain many thiols (cysteins). Dibenzoannulated cyclooctine (DIBO) and Dibenzocycloocytne (DBCO) have a lower rate constant, but show no sensitivity to Michael addition[17]. In addition, there are already several commercially available DBCO or DIBO functionalized polymers and fluorophores.

To verify the location of the amino acid functionalized by an azide in the OmpX loops, we used fluorophores functionalized by DIBO (Click-iT Alexa Fluor 488 sDIBO) on proteins of the outer membrane. After a few washing and spinning steps, we expected the cells to remain fluorescent.

Figure 17: To check the reaction of the click, the cells will be incubated with Click-iT Alexa Fluor 488 sDIBO. If the COMP protein is functionalized with the pAzF which represents the azide group, the dye will bind covalently proteins. In this case cells will remain fluorescent after some washing steps.

To analyze the fluorescence at the cell level, fluorescence activated cell sorting (FACS) and epifluorescence microscopy were used.

FACS: Flow Cytometry Cell Sorting

FACS is a specialized flow cytometry that provides information on the size, granulometry and fluorescence of cells. The relative granulometry of the cells is measured using the size dispersion (SSC). The relative cell size is measured using direct scattering (FSC). Fluorescence is induced by a laser and is captured by a wide range of filters. And all these features can be combined to sort the cells.

Figure 18: Simplified function of the FACS. Retrieved from:

Fluorescence Microscopy

Fluorescence (or epifluorescence) microscopy is a technique using an optical microscope taking advantage of the fluorescence phenomenon. Fluorescence is the property of some bodies to emit light after absorbing photons of higher energy. Fluorescence microscopy is based on the formation of an image by detection of this emitted light. The Stokes shift describes the difference between the wavelength absorbed by the object (emitted by the light source of the microscope) and emitted by the object. The greater the difference between the two wavelengths, the easier it is to observe the fluorescence.

Figure 19: Simplified function of the fluorescent microscopy. Retrieved from ThorLabs Website.

Bacterial Adenylate Cyclase Two-Hybrid (BACTH)

Bacterial Two-Hybrid - Overview

In 1989, Fields and Song demonstrated a new genetic system allowing the detection of protein-protein interaction [18]. At first, it was performed in Saccharomyces cerevisiae yeast and was named the yeast two-hybrid assay (Y2H). In 1998, Ladant and al. described the system in bacteria[19]. Nowadays, this biological technique is mostly used to show and characterize the physical interaction between two cytosolic proteins or internal membrane proteins in vivo[20].

Bacterial Adenylate Cyclase Two-Hybrid (BACTH)

The BACTH principle lies on the interaction-mediated reconstitution of a signaling cascade in Escherichia coli. The messenger molecule involved in this cascade is the cyclic adenosine monophosphate (cAMP) produced by the adenylate cyclase. Adenylate cyclase is an enzyme catalysing the cAMP production from ATP (Figure 20). It physiologically participates to the cellular transmission.

Figure 20: Production of cAMP from ATP, catalyzed by the adenylate cyclase enzyme[21].

This system involves the adenylate cyclase (AC) of Bordetella pertussis, which is the responsible agent for the pertussis disease. The catalytic domain of the adenylate cyclase has the possibility to be split in two distinct parts: T18 and T25 fragments, unable to have any enzymatic activity unless they are physically close. To perform a Bacterial Adenylate Cyclase Two Hybrid (BACTH), each subparts of the AC is fused to proteins of interest - either the bait or the prey protein chosen beforehand by the examinator -, thus constituting two hybrids[22]. These two hybrids are co-expressed in strains of Escherichia coli deficient in adenylate cyclase (cya-) like BTH101 strains.

The physical closeness of these hybrids induces their heterodimerization and results in a functional complementation between T18 and T25 fragments. Therefore, the AC enzyme is operational again and can produce cAMP (around 17,000 mmol of cAMP formed per mg of adenylate cyclase per minute[23]), which molecule responsible of the signal transduction in the bacterium (Gif 1).

Gif 1: Animation highlighting the two possible pathways in a BACTH assay. (NO interaction) is when the two proteins of interest analyzed don’t interact, there is no cAMP production. (Interaction) is when an heterodimerization is performed by the two proteins of interest, involving the reconstitution of a functional adenylate cyclase, able to produce cAMP.

The cAMP produced by the reconstitution of the AC will act as a messenger by interacting with catabolite activator proteins (CAP) in a ratio 1 to 1. Then two cAMP/CAP complexes are needed to activate the expression of the gene regulated by the lactose promoter.
The high enzymatic activity [23] of Bordetella pertussis Adenylate Cyclase (AC) generates a high production of cAMP in presence of ATP in the bacterium (Figure 1) thus activating the signaling cascade with the CAP-cAMP dependant promoter. Because of the high quantity of cAMP diffusing in the cytoplasm of the bacterium [24], the reporter gene is continuously activated as long as cAMP is produced (Gif 2). Hence this system is promising because it might have a great sensitivity and may drive a great signal amplification for a low amount of target molecules to be detected.

Gif 2: Animation highlighting the signaling cascade inside the bacteria. cAMP molecule, after its diffusion in the cytoplasm of the bacteria, is coupled to CAP in order to activate the promoter lactose plac.

In this system, a cAMP inducible CAP-dependent lactose promoter is needed and has to be inserted upstream a given reporter gene. Two major factors affect this promoter :

  • IPTG, known to have a positive effect on the transcription of the gene by removing the lac repressor from the DNA.
  • CAP, known to have a positive effect on the transcription when it binds cAMP by helping the fixation of RNA-polymerase on the DNA (Figure 21).
To be able to bind the CAP sites on the promoter, the CAP protein has first to interact with a cAMP molecule. As soon as two cAMP-CAP complexes are bound to the CAP sites, the RNA Polymerase initiates the transcription.

Figure 21: Structure of the complete lac operon and effect of cAMP on CAP-dependent promoter.

Outer-Membrane Bacterial Two-Hybrid System (mBACTH)

In NeuroDrop, the two adenylate cyclase fragments are fused to an engineered transmembrane protein named COMP – for Clickable Outer Membrane Protein – expressed in the external membrane of E.coli. This protein has been initially engineered with the OmpX protein.
OmpX is an outer membrane protein with the C- and N-termini in the intracellular domain. To be able to use OmpX as a scaffold, a unnatural amino acid needs to be introduced. This can be done by implementing the amber stop codon TAG in one of the loops of OmpX via a mutation. With a specific tRNA an azide-functionalized amino acid can be built in, which can be used for the SPAAC click chemistry reaction with DBCO functionalized groups, this modified protein is called COMP. The complex aptamer fixed to a COMP is then called a COMB for Clickable Outer Membrane Biosensor[25].
In this case, the two AC subparts - either T18 or T25 - are fused to the C-terminal ends of COMPs with a Gly-Gly-Ser Linker (GGS) of 54 amino acids, in order to ensure a sufficient flexibility (Gif 3).

Gif 3: Animation showing the genetic constructions performed in order to obtain the mBACTH in NeuroDrop project. Each COMP protein is fused to a subpart of the AC with a flexible linker. The reporter gene needs to be under a plac promoter (or another cAMP dependent catabolite promoter).

When COMBs catch the extracellular target, they get closer, thus allowing the reconstitution of a functional adenylate cyclase due to the physical proximity of the two sub-parts (Gif 4). The enzyme is operational again and produces a high quantity of cAMP, the molecule responsible for the signal transduction in bacteria.
ATP is not naturally present in large amount in the periplasm of the bacteria, thereby it has to be added in the bacteria medium to enhance its periplasm diffusion and to be available for the adenylate cyclase catalytic reaction.

Gif 4: Animation showing the recognition of a specific biomarker with a specific aptamer fixed to COMPs proteins. T18 and T25 are the two subparts of the adenylate cyclase that enable the BACTH to work.

Advantages of the BACTH system

The strain used is E.Coli cya- : BTH101, which enables an easy screening and characterization of protein-protein interactions with standard molecular biology techniques. The growth of these bacteria is fast and they can be easily transformed.

The BACTH system relies on a signaling cascade which utilizes the diffusible regulatory molecule, cAMP. As a consequence, the physical association of the two interacting chimeric proteins can be spatially separated from the transcription activation readout. Hence, it is possible to analyze protein-protein interactions that occur either in the cytosol, at the inner membrane level, on the DNA, or even in the periplasm[22].

This system allows a reduction of potential false positives, because the heterodimerization of the two hybrids is reversible. This means that if the closeness of the hybrids is not stabilized by the target and is reached randomly due to the free movements of the proteins, the signaling cascade will not last in time.

The NanoLuciferase reporter gene

What is this reporter gene?

For the NeuroDrop project, the main issue considering the reporter gene was to choose between a fluorescent protein (ex: GFP) and a bioluminescent one, the Nanoluciferase. In order to answer this question, we firstly had to understand: what is the Nanoluciferase ?

Brief History

The concept of luminescence is really old. Greeks and Romans described it in around 350 before JC as “cold light” in many marine species. Since this period, many travellers talked about “burning sea” phenomenon like Christopher Columbus. Several years later, luminescence appeared in literature as fireflies and glow-worm (Hamlet, Shakespeare, 1603)[26][27][28].
During the 19th century, Raphael Dubois extracted two components of bioluminescence and created light with it. He named it luciferin and luciferase.

The major improvement was during the 20th century, thanks to E.Newton Harvey who published a book where he referenced almost all luminescent species. In 1956, luciferin was isolated for the first time by Green and McElroy[29]. Nowadays, several luciferin types and many luciferases are commercialized. Nanoluciferase is a part of them.

The NanoLuciferase

Nanoluciferase is a small and monomeric enzyme (19.1kDa) and is about 150 times brighter than other luciferases. Its new substrate called furimazine produces a high luminescence intensity. Moreover, in contrast with classic bioluminescence proteins like Luciferase, the Nanoluciferase do not consume ATP as a substrate which is a very good point for our BACTH system that need ATP as much as possible for the cAMP production. After looking at the bibliography, Nanoluciferase gene was chosen because it allowed us to have a stronger signal than other classic reporters. Indeed, this protein isn’t secreted but unfused. Furthermore, its intracellular half-life is superior to 6 hours and the Nanoluciferase has a low autoluminescence. Thus giving us a maximum sensitivity.

The NanoLuciferase is used in biomedical research for several applications, including the study of protein-protein interactions, the monitoring of protein stability, BRET-based sensors, molecular imaging or even genetic regulation and cell signaling investigations, such as in our NeuroDrop Project[30][31]. The emission pic of NanoLuciferase is located at 460nm.

Figure 22: NanoLuciferase reaction with its substrate called furimazine. The enzyme creates an oxidative process. Retrieved from Promega’s website.

How did we use this gene in our project?

Genetic Constructions

We used Nanoluciferase following the Nano Glo Luciferase Assay System from Promega, which was offered to us within the scope of their sponsorship. The gene was received in a pNL1.1 vector (Figure 23). In this vector, the nanoluciferase gene size is 513bp.

Figure 23: pNL1.1 vector from Promega.

This vector doesn’t contain a promoter. That is why a promoter had to be included in a plasmid to form a promoter-RBS-Nanoluciferase-terminator biobrick. For this, an inducible promoter was used in order to regulate the gene promoter with a signaling pathway. As seen in the BACTH part, an induction was generated thanks to the cAMP molecule (Figure 24).

This is why a plac promoter was used : the lactose operon promoter. Indeed, it is regulated by cAMP and IPTG as it has been demonstrated with the BBa_J04450[30] part characterization. This construct was made thanks to restriction site additions by PCR. This work was performed in endogeneous adenylate cyclase deficient bacteria, which are called BTH101.

Figure 24: Structure of the complete Lac operon and the effect of cAMP on CAP promoter. To have more explanations, please refer you to the characterization document.

Use in the project

The part containing the PLac-RBS-Nanoluciferase-Terminator is included in the final plasmid. This plasmid is composed of OMPX-T18 and OMPX-T25 which are mutated in their third loop for an amino acid in OMPx allowing click-chemistry. Moreover, this plasmid contains the Nanoluc parts and a resistance to ampicillin. (Figure 25).

Figure 25: Our final plasmid called pGre-Alps composed by the COMP-T18 biobrick, the COMP-T25 biobrick, and the Nanoluciferase gene (BBa_K3128001). It contains an ampicillin resistance cassette which is not visualized in this figure.


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