Please set your mobile browser to desktop mode, or display this website on a computer for a better experience


Respiration is a vital step in all living systems, which is simply defined as shuttling an energy from the fuels (consumed food) to the biological systems. While biological systems can be anything from energy collection in coenzymes or ATP to protein folding and replication, shuttling happens of always the same particle - an electron. But just like in computer circuitry, living creatures require “a ground negative” for depositing sequestered electrons from the fuels, or the continuity of the energy will halt. Oxygen, currently all-available, is a perfect example of how life has found its “ground negative,” as it serves as an electron acceptor, but before it was "invented" 3 billion years ago [1], creatures of earth were limited to anaerobic respiration. What’s more, due to high likelihood of reactive oxygen species production by oxygenation, life had avoided this currently indispensable molecule.

In the world without oxygen, living systems adapted to depositing sequestered electrons to other elements of high electronegativity, or simply elements with a “tendency to accept electrons”. This evolutionary feat has currently turned into a rosetta stone of modern energy-innovation in living species. At this crossroads, one of the most studied, understood and engineered is Shewanella oneidensis MR-1 bacteria, which has an entire concert of proteins dedicated for solving the evolutionary crisis of anaerobic respiration. It is Shewanella and its proteins that have inspired us when developing our project.

S. oneidensis is a gram-negative, facultative anaerobic bacteria. In anaerobic environment, it swiftly switches to anaerobic respiration, governed by metal reducing proteins (Mtr). Mtr respiration pathway spans both inner and outer membranes, while inner membrane is where Cytochrome A is situated [2], and outer membrane bears MtrB, MtrA and MtrC proteins. Electron transfer begins at CymA, which after oxidizing quinol to quinone passes electrons to the cytochromes inside the periplasmic pool. Compared to Shewanella’s well-known periplasmic shuttles-like STC and FccA, flavocytochromes are also readily available in E. coli [3], which may serve as a burden-relieving factor when minimizing the system. Following the periplasmic shuttling, electrons are delivered to outer membrane proteins. Here, MtrB acts as a porin [4], functionalizing respiratory-active protein - MtrA, which shuttles electrons along its heme molecules along the outer membrane. Lastly, MtrC is anchored to the MtrAB complex, and acts as a final harbor of electrons, before they are passed to the free-moving flavins or the target element - in our case electrodes.

Electron shuttling is one of three general pathways employed for extracellular electron transfer (EET). Shuttle-mediated EET (in the media) is an alternative to biofilm formation on the anode (direct contact between bacteria and anode), since it guarantees delivery of electrons at a distance, via shuttles [5]. Shuttles can either be low-molecular weight organic molecules, freely present in the bacterial environment, or endogenously produced melanin, phenazine, quinones flavin and riboflavin [6–10].

Canthaxanthin production

Interestingly, Socco et al. [10] have described a promising EET shuttle: canthaxanthin which belongs to the carotenoid class of compounds [11] which unifies antioxidants usually known for their vivid colors. For health, color and stability canthaxanthin has become renowned in a list of human industries from cosmetics to health products. It is likely due to versatility of canthaxanthin applications, that its market value in 2017 has reached $75 million [12]. Among other intriguing factors, as a carotenoid, it can be expressed via heterologous engineering of endogenous MEP pathway. The latter, combined with its appealing red-orange color makes canthaxanthin a perfect target for engineering and streamlining biosynthetic pathways.

Since E. coli is not a natural host for Mtr proteins, compared to S. oneidensis, its EET efficiency is limited. To amend for the latter, we have introduced canthaxanthin in our engineered E. coli. By introducing an electron shuttle, we have guaranteed EET in the whole volume of the bacterial chamber, given that canthaxanthin binds to the MtrC. For production, purification and engineering successes, detailed design-build-test-learn (DBTL) cycle can be read below.


The canthaxanthin production pathway (Figure 1) can be divided in un upper (E. coli endogenous) module consisting in production of the intermediate farnesyl diphosphate (FPP), the middle module for the production of ß-carotene and the bottom module for the conversion of ß-carotene to canthaxanthin.

From ß-carotene to canthaxanthin

Canthaxanthin is derived from ß-carotene through the action of the enzyme ß-carotene-ketolase (EC which oxidizes the two rings of ß-carotene sequentially and converts it first to echinenone then to canthaxanthin [13]. Two structurally different ß-carotene-ketolases, the CrtW-type and the CrtO-type, were identified in various organisms where, depending on their substrate specificity, are able to catalyze only the first reaction (ß-carotene to echinenone), only the second one (echinenone to canthaxanthin) or both. Moreover, some CrtW and CrtO enzymes are promiscuous and able to oxidize other carotenoids like for example zeaxanthin or adonixanthin.

To achieve the canthaxanthin production, we choose to use the CrtW148 gene from Nostoc punctiforme PCC 73102 which was shown to efficiently catalyze both steps of the canthaxanthin synthesis from ß-carotene [14].

In addition, we also choose to use the gene responsible for the canthaxanthin production in Dietzia sp. RNV-4 strain [10]. To the best of our knowledge, no biochemical analysis was described in the literature for any CrtW or CrtO enzymes of any Dietzia sp. Moreover, the genome of the Dietzia sp. RNV-4 strain is not available, nor the sequence of a CrtW or a CrtO enzyme. However, based on the 16S rRNA sequence analysis, the Dietzia psychralcaliphila ILA-1 species, whose genome is available, has 99% sequence similarity with the Dietzia sp. RNV-4 strain. Using different CrtW and CrtO protein sequences for the EC enzymes found in the Kegg database, we performed protein homology searches on the D. psychralcaliphila ILA-1 genome (GenBank Acc. N° CP015453) and thus uncovered a potential CrtO-like enzyme encoded by the A6048_06315 locus (annotated as a FAD-dependent oxidoreductase by automated computational analysis using protein homology gene prediction method). No CrtW-like enzyme was spotted.

From FPP to ß-carotene

One prerequisite for canthaxanthin production is the ß-carotene synthesis. This can be ‘easily’ achieved in E. coli by using parts constructed by previous iGEM teams, like for instance the BBa_K274220 in which four genes CrtE, CrtB, CrtI and CrtY from Pantoea ananatis were assembled in an operon under the control of pBad promoter inducible by L-arabinose by the iGEM 2009 Cambridge team.

To FPP in E. coli

E. coli is not able to naturally produce carotenoids, but is able to synthesize the farnesyl diphosphate (FPP) which is a key intermediate in the isoprenoids biosynthesis. In E. coli, FPP production is achieved via the MEP (2-C-methyl-D-erythritol 4-phosphate) pathway which is a ten steps pathway starting from pyruvate and glyceraldehyde 3-phosphate. An alternative, the mevalonate pathway using as precursor acetyl-CoA is present in eukaryotic organisms (including examples of fungi and algae) and archaea.

Microbial biosynthesis of carotenoids is a well-studied and optimized metabolic engineering case thanks to the introduction of a biosynthetic pathway to classical E. coli chassis. The first metabolic engineering approaches for the carotenoid biosynthesis rapidly included optimizing the flux through the FPP to increase production yields. Overexpressing endogenous rate limiting enzymes of the MEP pathway [15,16] or expressing heterologous mevalonate pathway [17] have both been used. The latter gave better results so far, but MEP pathway is nonetheless promising because its theoretical yield is higher [18].

To maximize FPP production through the MEP pathway, understanding its regulation is critical to bypass cellular control, since it is an endogenous pathway [19]. The new tools provided by the systems biology field established that the steps catalyzed by the 1-deoxyxylulose-5-phosphate synthase (dxs) and isopentenyl diphosphate isomerase (idi) enzymes are limiting steps of the MEP pathway [20]. Indeed, overexpression of dxs and idi enzymes increased carotenoids production yields in E. coli [21].

Thus, for improving the canthaxanthin yield in E. coli, we decided to overexpress the E. coli dxs and idi genes and thus increase the amount of FPP precursor available.

Figure 1. Biochemical pathway of canthaxanthin synthesis composed of the endogenous E. coli MEP pathway, the heterologous ß-carotene producing pathway (the CrtEBIY genes from Pantoea ananatis) and the canthaxanthin producing step catalyzed either by the CrtW enzyme from Nostoc punctiforme or the CrtO enzyme from Dietzia psychralcaliphila.


To implement the canthaxanthin production pathway in E. coli and enhance its yield we used biobricks assembled by previous iGEM teams, as well as new ones designed by us.

Upper module: enhancing the FPP in E. coli

To overexpress the E. coli idi and dxs genes we assembled, through the Golden Gate technique, an expression vector in both the pSB1A3 and pSB3T5 backbones (BBa_K4432122) in which the idi and dxs genes were placed under the control of a hybrid T5 promoter regulated by LacI. For this, the idi and dxs gene sequences were PCR-amplified from the genome of E. coli and equipped by custom-made RBSes that we specifically designed using the online tools provided by Salis’s De Novo DNA company.

Middle module: from FPP to ß-carotene

We used BBa_K274220 comprising the CrtEBIY operon (containing four genes from P. ananatis allowing the conversion of FPP to ß-carotene) placed by the iGEM 2009 Cambridge team under the control of pBad promoter inducible by L-arabinose. As a control, we also used BBa_K274120 assembled by the same team, in which the CrtY is not present and the carotenoids pathway thus stops at lycopene. Both these biobricks were recovered from the iGEM distribution kits where they were made available in the pSB2K3 backbone.

Bottom module: from ß-carotene to canthaxanthin

The two selected genes, CrtW from N. punctiforme and CrtO from D. psychralcaliphila were placed under the control of either a hybrid T5 promoter regulated by LacI (BBa_K4432120 and BBa_K4432121), or the classical pLac promoter (BBa_K4432220 and BBa_K4432221) and assembled through the Golden Gate technique in the pSB1A3 backbone. For a proper protein expression, custom-made RBSes were specifically designed using the online tools provided by Salis’s De Novo DNA company.

Upper and Bottom modules

Moreover, to lower the burden of plasmid maintenance, we decided to combine the Upper and Bottom modules in a single backbone, the pSB1A3. Thus, both CrtW and CrtO were expressed under the control of the hybrid T5 promoter regulated by LacI as operons together with the E. coli idi and dxs genes (BBa_K4432320 and BBa_K4432321).


To demonstrate the whole-cell bioproduction of canthaxanthin, E. coli MG1655 cells were co-transformed with 2 or 3 plasmids encoding the genes for the bottom, middle and upper parts of the pathway described above. Bacterial cultures were fully grown in LB medium containing the appropriate antibiotics (100 µg/mL ampicillin, 25 µg/mL kanamycin, 5 µg/mL tetracycline) and induced with 100 µM IPTG and 1.5 mM L-arabinose. After 24 hours, 15 mL were centrifuged (15 minutes at 4000 g, 4°C) in order to separate the biomass from the culturing medium, and carotenoids were extracted from the pellets with acetone twice followed by methanol. For this, the cell pellet was resuspended in 2 mL acetone in the presence of 0.3 g of glass beads (diameter 1 mm) and homogenized by vortexing for 5 minutes. After centrifuging the sample for 5 minutes at 4000 g, 4°C, the acetone-soluble fraction was recovered and the pellet was subject to a second round of acetone extraction. Finally, as for canthaxanthin it was shown that methanol improves the extraction yield [22], we performed a third extraction using 2 mL of methanol and the same protocol as for acetone.

Acetone and methanol extracts were analyzed by high pressure liquid chromatography (HPLC) using a Shimadzu Prominence LC20/SIL-20AC equipped with a Kinetex XB-C18 reversed phase column (250 mm × 4.5 mm, 5 μm) and an UV–Vis detector. Mobile phases A (methanol) and B (acetonitrile) were set at a flow rate of 1 mL/min and the separation was performed isocratically with an A:B ratio of 90:10. The sample injection volume was 10 μL, the column was thermostated at 40°C, and the metabolites were monitored at 452 nm and 472 nm. The product quantification was done by interpolation of the integrated peak areas using calibration curves prepared with standard samples for lycopene, ß-carotene and canthaxanthin as detailed on the Measurement page on this wiki. Data was normalized to grams of dry cell weight (gDW) estimated by converting first the cell density (OD600nm) to number of cells (based on the approximate conversion of OD600nm of 1.0 = 8 x 108 cells/mL) and finally to gDW knowing that the dry weight of one E. coli cell is 3 x 10-13 g according to the E. coli Metabolome Database (ECMDB) [23].


Carotenoids production was readily visible by naked eye. Indeed, bacterial pellets were colored in different shades from pale yellow to dark red, while the negative control performed with E. coli cells not expressing any Crt gene had the characteristic beige color (Figure 2).
When only the CrtEBI genes were present, the cells were slightly but visibly red, indicative of lycopene expression, as expected. The color intensity is enhanced in the presence of the idi and dxs expression device. This same trend was observed when the ß-carotene production operon composed of the 4 genes CrtEBIY was expressed in the E. coli cells: strains exhibited higher orange coloration with the induced metabolic boost plasmid (idi and dxs) being it in the high copy plasmid pSB1A3 or the low copy one pSB3T5.

The presence of either CrtW or CrtO genes led to the appearance of a coral color, strongly suggesting the production of canthaxanthin.

Figure 2. E. coli MG1655 cell pellets expressing the lycopene (CrtEBI), ß-carotene (CrtEBIY) or canthaxanthin (CrtEBIYW/O) devices along or not with with the idi and dxs synthetic operon.

To assess each carotenoid production in each strain, both qualitatively and quantitatively, the extracts were subject to reverse phase chromatography and spectrophotometric analysis. Comparaisons with standard commercial lycopene, ß-carotene and canthaxanthin confirmed their production in E. coli cells expressing the lycopene (CrtEBI), ß-carotene (CrtEBIY) and canthaxanthin (CrtEBIYW/O) devices respectively.

Indeed, the reverse phase chromatography allowed us to clearly and effectively separate the 3 carotenoids and thus unambiguously identify them (Figure 3). For instance, on the HPLC chromatogram of the acetone extraction from E. coli cells expressing the lycopene (CrtEBI) operon a single peak with the same retention time as commercial lycopene is observed, while on the HPLC chromatogram of the acetone extraction from E. coli cells expressing the ß-carotene (CrtEBIY) operon, one can observe a single peak with the same retention time as commercial ß-carotene. As expected, no peak is visible on the HPLC chromatogram of the acetone extraction from E. coli cells not expressing any Crt gene.

Moreover, when either CrtW or CrtO genes are present, the characteristic peak of canthaxanthin appears indicating that both enzymes are capable of synthesizing canthaxanthin. It should be noted that residual amounts of ß-carotene are visible on the HPLC chromatogram in the case of CrtO suggesting a less efficient conversion compared to CrtW.

Figure 3. HPLC chromatograms of commercial lycopene, ß-carotene and canthaxanthin and of acetone extractions from E. coli MG1655 cells expressing the lycopene (CrtEBI), ß-carotene (CrtEBIY) and canthaxanthin (CrtEBIYW/O) devices along with the idi and dxs synthetic operon. The negative control was performed using E. coli cells not expressing any Crt gene. Images were produced using Shimadzu’s LabSolutions Postrun Analysis software.

Spectroscopic analysis further supports the chromatography results (Figure 4). Indeed, the acetone extractions from E. coli MG1655 cells expressing the lycopene (CrtEBI), ß-carotene (CrtEBIY) and canthaxanthin (CrtEBIYW) devices have the same spectral properties as comercial lycopene, ß-carotene and canthaxanthin respectively, while, in the case of CrtO, a mixed ß-carotene + canthaxanthin spectrum is observed.

Figure 4. Spectroscopic analysis of commercial lycopene, ß-carotene and canthaxanthin (A) and of acetone extractions from E. coli MG1655 cells expressing the lycopene (CrtEBI), ß-carotene (CrtEBIY) and canthaxanthin (CrtEBIYW/O) devices along with the idi and dxs synthetic operon (B). The negative control was performed using E. coli cells not expressing any Crt gene.

Quantitative analysis of the HPLC data (Figure 5) confirms the conclusions derived from the visual observation of the color of the cells pellets. Indeed, the production of all three carotenoids is strongly enhanced when the MEP pathway is deregulated by the overexpression of the idi and dxs genes. Our best canthaxanthin yield is obtained upon co-expression of the CrtEBIY (BBa_K274220) and CrtW+idi+dxs (BBa_K4432320) operons and it reaches 17.9 µmol (10.1 mg) per gram of E. coli dried weight (gDW) which is in the same range of previous studies that reported 16.1 mg/g of 90% pure canthaxanthin [24].

Figure 5. Canthaxanthin, ß-carotene or lycopene production yields by E. coli MG1655 cells expressing the lycopene (CrtEBI), ß-carotene (CrtEBIY) or canthaxanthin (CrtEBIYW/O) devices along or not with with the idi and dxs synthetic operon.

Phenazine-1-carboxylate (PCA) production

Another interesting EET shuttle molecule is the phenazine-1-carboxylate (PCA). Phenazines are polyaromatic secondary metabolites known as compounds with high redox activity [25]. These secondary metabolites are produced by a variety of bacteria, especially Pseudomonas species.

To improve the electricity generation in our system, previous studies have shown that the use of Pseudomonas species producing phenazine-based metabolites in the anode chambers of MFCs can improve anodic electron transfer [26]. In an MFC, phenazines are found to enable a high electrical conductivity of the multilayered biofilm on the anode, leading to enhanced electricity generation from bacterial metabolism. Overexpression of the phz operon increases the electricity production of the Pseudomonas species-inoculated MFCs [27–30].

Since our E. coli chassis has a limiter EET efficiency (compared to S. oneidensis), we decided to introduce the PCA production pathway, the PhzA1-G1 operon. To obtain the DNA encoding it, we first turned to Parts Registry where we found BBa_I723028 annotated as the “PCA biosynthesis operon”. However, we quickly noticed that this part does not contain the entire PhzA1-G1 operon of P. aeruginosa, but a truncated version in which only the 5’ part of the PhzG1 gene is present. We concluded that is part is not functional and performed experiments to improve it, as detailed on the Improvement page of this wiki.

Thus, we constructed expression cassettes of both full P. aeruginosa PhzA1-G1 and PhzA2-G2 operons (BBa_K4432040 and BBa_K4432041) and succeeded in producing PCA in E. coli (Figure 6).

Figure 6. The efficiency of PCA production in two different E. coli strains carrying the pSB1A3 plasmid harboring the PhzA1-G1 (full and truncated) and PhzA2-G2 operons or the negative control (an empty pSB1A3 backbone).

Shewanella’s MtrCAB nano-conduct complex and CymA cytochrome expression in E. coli

Expressing the Shewanella’s MtrCAB nano-conduct complex in an engineered in E. coli together with the CymA cytochrome is another strategy to increase the EET efficiency of our chassis. However, based on our human practices work, as we just want to get an ON/OFF switch with an electric output, we would not need the Mtr complex to get a proper signal, but only to incorporate the toehold switch controlling the expression of CymA in an engineered E. coli. The toehold unfolding itself, by the only expression of the Cytochrome A (CymA) nano-conduction may provide a sufficient electric output to distinguish both the OFF and ON states of the switch. This is probably the most convenient and rapid way to get proper experimental results, however the electric output might be too low to be detected in the presence of the lncRNA PANTR1. So, false negative output might be a problem to make the difference between samples from sick and healthy people.


We placed CymA upstream of our PANTR1 specific toehold switches (n°5, n°6 and n°8) in the composite parts BBa_K4432135, BBa_K4432136 and BBa_K4432138 respectively (in pSB3T5 backbone). As a control, we built an expression cassette of CymA (BBa_K4432130) under the control of the T7 promoter with the synthetic RBS stem-loop designed for standard toehold switches.
To allow co-expression of both MtrCAB genes and the PANTR1 toeholds cognated triggers, we recovered the BBa_K1316012 from the 2018 iGEM distribution kit and build by classical RFC[10] assembly the composite parts BBa_K4432035, BBa_K4432036, BBa_K4432038, respectively, in the pSB1C3 backbone.


For a fast and high-throughput analysis of the electron transfer capacities of our engineered E. coli cells we set-up a colorimetric assay based on the peroxidase-catalyzed oxidation of the chromogenic commpound 3,3',5,5'-tetramethylbenzidine (TMB). This test was previously described as efficient for characterizing S. oneidensis EET efficiency [31]. Following the described protocol, E. coli cells were grown over-night in LB media containing the appropriate antibiotics and inducers (as indicated in the corresponding sub-chapters of this wiki). After washing them 3 times, cells were resuspended in pure water at an OD600nm of 1 and the colorimetric assay was performed immediately in a transparent 96-well polystyrene microplate (Sarstedt). For this, 40 µL of cell suspension was mixed with either 40 µL of water or 40 µL of a cell suspension of S. oneidensis MR1 (pre-treated as E. coli cells were) and the reaction was started by adding 80 µL of TMB substrate solution (Sigma T4444). After 20 minutes of incubation of room temperature, the reaction was stopped by adding 40 µL of H2SO4 2M and the absorbance at 450 nm was measured using a CLARIOstar (BMGLabtech) plate reader with the pathlength correction option turned on and adjusted to 200 µL. Absorbance values were converted to total number of transferred electrons using a molar extinction coefficient of 5.9 x 104 M-1cm-1 [32] and taking into consideration that each TMB molecule transfers 2 electrons. Data were normalized to number of cells (based on the approximate conversion of OD600nm of 1.0 = 8 x 108 cells/mL for E. coli and 5 x 108 cells/mL for S. oneidensis [31])


We first evaluated the electron transfer capacities of S. oneidensis MR-1 and E. coli alone. The results presented in Figure 7 show a linear relationship between the transfeted electrons and the number of cells for both strains. They also confirm the limited EET efficiency of E. coli compated to S. oneidensis.
Figure 7. Electron transfer capacities of S. oneidensis MR-1 and E. coli cells.

When using our engineered E. coli strains carrying either the phenazine pathway (PhzA1-G1 and PhzA2-G2 operons) or the canthaxanthin producing devices (CrtEBIY + CrtW/O-idi-dxs operons), the results are promissing (Figure 8).

E. coli cells producing PCA (Figure 8A) have the capacity to increase by about 10% the EET capacity of S. oneidensis MR-1. PCA seems to have the same effect on E. coli cells alone. In addition, both our engineered E. coli expressing the full PhzA1-G1 and PhzA2-G2 operons are perfoming better than the cells containing the truncated version of the PhzA1-G1 operon, in line with the PCA qualtification results (Figure 6).

In contrast, E. coli cells producing canthaxanthin (Figure 8B) do not have a significant increased EET efficiency compared to E. coli cells not producing it. Moreover, no effect on the EET of S. oneidensis MR-1 is observed.

Figure 8. Electron transfer capacities of E. coli strains carrying either the PCA pathway (PhzA1-G1 and PhzA2-G2 operons) or the canthaxanthin producing devices (CrtEBIY + CrtW/O-idi-dxs operons). The data and error bars are the mean and standard deviation of at least three measurements on independent biological replicates.

The expression of CymA E. coli does not seem to have an effect on EET capacity of either E. coli or S. oneidensis MR-1 cells (Figure 9). Indeed, no diffrence is observed in TMB tests performed with E. coli cells expressing CymA compated to the negative control performed with an empty backbone. In this context, it's not unexpected that the PANTR1 toehold switches gave the same output in the presence or absence of the cognate triggers.
Figure 9. Electron transfer capacities of E. coli BL21 Star™(DE3) cells carrying the CymA expression casettes under the control of the PANTR1 toehold switches n°5, 6 or 8 in the presence or not of the cognate PANTR1 triggers. The negative controls have been performed with an empty pSB3T5 and the positive control with BBa_K4432130. The data and error bars are the mean and standard deviation of at least three measurements on independent biological replicates.

External electron transfer measurements in our microbial fuel cell hardware

Using our MCF device, that we designed and built as detailed on the Hardware page on this wiki, we have performed in vivo tests using first S. oneidensis MR-1 cells for rapid prototyping. An increasing electrical output was readily observed, thus demonstrating our MCF's functionality (Figure 10). The negative control, performed with LB media, remained constant at a negijable voltage (data not shown).

Moreover, when our engineered E. coli strains were added in the MCF along with S. oneidensis MR-1 cells, a strong correlation is observed between the increase of the electric output and the fact that Shewanella is or not in the presence E. coli expressing the PCA pathway (PhzA1-G1 and PhzA2-G2 operons). This electrical output is due to the PCA electron shuttle mediators transferring more easily electrons from the media to the anode of the microbial fuel cell.

Figure 10. External electron transfer measurements in our microbial fuel cell hardware.


We have successfully produced two extracellular electron transfer shuttle mediators, canthaxanthin and phenazine-1-carboxylate and evaluated their production titer in E. coli through high performance liquid chromatography and spectrophotometric analysis. Moreover we set-up a colorimetric assay in microtiter plates to characterize their electron transfer capacities and showed that PCA expressing E. coli cells are able to lead to an electron flow outside the cell, which can be detected in real-time via our microbial fuel cell device connected to an Arduino chip (see the Hardware page on this wiki).

We also designed and built parts to express the Shewanella’s Cytochrome A (CymA) gene under the control of our best PANTR1 toehold switches (see the Engineering page of this wiki) to regulate the external electron transfer depending on the presence / absence of the PANTR1 lncRNA. However, the expression of CymA alone was not sufficient to detected an effect on the EET. The presence Shewanella’s MtrCAB operon seems to be required, but its expression is chalanging (we encountered difficulties to grow in liquid cultures the E. coli cells carrying the Shewanella’s MtrCAB operon, even without induction).


[1] Jabłońska J, Tawfik DS. The evolution of oxygen-utilizing enzymes suggests early biosphere oxygenation. Nature Ecology & Evolution (2021) 5: 442–448.
[2] Edwards MJ, Richardson DJ, Paquete CM, Clarke TA. Role of multiheme cytochromes involved in extracellular anaerobic respiration in bacteria. Protein Science (2020) 29: 830–842.
[3] Caux C, Guigliarelli B, Vivès C, Biaso F, Horeau M, Hassoune H, Petit-Hartlein I, Juillan-Binard C, Torelli S, Fieschi F, Nivière V. Membrane-bound flavocytochrome MsrQ is a substrate of the flavin reductase Fre in Escherichia coli. ACS Chemical Biology (2021) 16: 2547–2559.
[4] Edwards MJ, White GF, Butt JN, Richardson DJ, Clarke TA. The crystal structure of a biological insulated transmembrane molecular wire. Cell (2020) 181: 665-673.e10.
[5] Sriram S, Wong JWC, Pradhan N. Recent advances in electro-fermentation technology: a novel approach towards balanced fermentation. Bioresource Technology (2022) 360: 127637.
[6] von Canstein H, Ogawa J, Shimizu S, Lloyd JR. Secretion of flavins by Shewanella species and their role in extracellular electron transfer. Applied and Environmental Microbiology (2008) 74: 615–623.
[7] Marsili E, Baron DB, Shikhare ID, Coursolle D, Gralnick JA, Bond DR. Shewanella secretes flavins that mediate extracellular electron transfer. Proceedings of the National Academy of Sciences (2008) 105: 3968–3973.
[8] Brutinel ED, Gralnick JA. Shuttling happens: soluble flavin mediators of extracellular electron transfer in Shewanella. Applied Microbiology and Biotechnology (2012) 93: 41–48.
[9] Kotloski NJ, Gralnick JA. Flavin electron shuttles dominate extracellular electron transfer by Shewanella oneidensis. mBio (2013) 4: e00553-12.
[10] Sacco NJ, Bonetto MC, Cortón E. Isolation and characterization of a novel electrogenic bacterium, Dietzia sp. RNV-4. PloS One (2017) 12: e0169955.
[11] Rebelo BA, Farrona S, Ventura MR, Abranches R. Canthaxanthin, a red-hot carotenoid: applications, synthesis, and biosynthetic evolution. Plants (2020) 9: 1039.
[12] Ahuja K, Rawat A. Global canthaxanthin market to exceed $85 mn by 2024. Global Market Insights Inc. (2018).
[13] Sandmann G. Diversity and origin of carotenoid biosynthesis: its history of coevolution towards plant photosynthesis. The New Phytologist (2021) 232: 479–493.
[14] Steiger S, Sandmann G. Cloning of two carotenoid ketolase genes from Nostoc punctiforme for the heterologous production of canthaxanthin and astaxanthin. Biotechnology Letters (2004) 26: 813–817.
[15] Kim SW, Keasling JD. Metabolic engineering of the nonmevalonate isopentenyl diphosphate synthesis pathway in E. coli enhances lycopene production. Biotechnology and Bioengineering (2001) 72: 408–415.
[16] Alper H, Fischer C, Nevoigt E, Stephanopoulos G. Tuning genetic control through promoter engineering. Proceedings of the National Academy of Sciences of the United States of America (2005) 102: 12678–12683.
[17] Yoon S-H, Lee S-H, Das A, Ryu H-K, Jang H-J, Kim J-Y, Oh D-K, Keasling JD, Kim S-W. Combinatorial expression of bacterial whole mevalonate pathway for the production of beta-carotene in E. coli. Journal of Biotechnology (2009) 140: 218–226.
[18] Ajikumar PK, Xiao W-H, Tyo KEJ, Wang Y, Simeon F, Leonard E, Mucha O, Phon TH, Pfeifer B, Stephanopoulos G. Isoprenoid pathway optimization for Taxol precursor overproduction in E. coli. Science (New York, N.Y.) (2010) 330: 70–74.
[19] Banerjee A, Sharkey TD. Methylerythritol 4-phosphate (MEP) pathway metabolic regulation. Natural Product Reports (2014) 31: 1043–1055.
[20] Volke DC, Rohwer J, Fischer R, Jennewein S. Investigation of the methylerythritol 4-phosphate pathway for microbial terpenoid production through metabolic control analysis. Microbial Cell Factories (2019) 18: 192.
[21] Albrecht M, Misawa N, Sandmann G. Metabolic engineering of the terpenoid biosynthetic pathway of E. coli for production of the carotenoids β-carotene and zeaxanthin. Biotechnology Letters (1999) 21: 791–795.
[22] Scaife MA, Ma CA, Armenta RE. Efficient extraction of canthaxanthin from E. coli by a 2-step process with organic solvents. Bioresource Technology (2012) 111: 276–281.
[23] Sajed T, Marcu A, Ramirez M, Pon A, Guo AC, Knox C, Wilson M, Grant JR, Djoumbou Y, Wishart DS. ECMDB 2.0: A richer resource for understanding the biochemistry of E. coli. Nucleic Acids Research (2016) 44: D495-501.
[24] Scaife MA, Prince CA, Norman A, Armenta RE. Progress toward an E. coli canthaxanthin bioprocess. Process Biochemistry (2012) 47: 2500–2509.
[25] Chen J-J, Chen W, He H, Li D-B, Li W-W, Xiong L, Yu H-Q. Manipulation of microbial extracellular electron transfer by changing molecular structure of phenazine-type redox mediators. Environmental Science & Technology (2013) 47: 1033–1039.
[26] Rabaey K, Boon N, Höfte M, Verstraete W. Microbial phenazine production enhances electron transfer in biofuel cells. Environmental Science & Technology (2005) 39: 3401–3408.
[27] Feng J, Qian Y, Wang Z, Wang X, Xu S, Chen K, Ouyang P. Enhancing the performance of E. coli-inoculated microbial fuel cells by introduction of the phenazine-1-carboxylic acid pathway. Journal of Biotechnology (2018) 275: 1–6.
[28] Bosire EM, Rosenbaum MA. Electrochemical potential influences phenazine production, electron transfer and consequently electric current generation by Pseudomonas aeruginosa. Frontiers in Microbiology (2017) 8: 892.
[29] Pham TH, Boon N, De Maeyer K, Höfte M, Rabaey K, Verstraete W. Use of Pseudomonas species producing phenazine-based metabolites in the anodes of microbial fuel cells to improve electricity generation. Applied Microbiology and Biotechnology (2008) 80: 985–993.
[30] Jayapriya J, Ramamurthy V. Use of non-native phenazines to improve the performance of Pseudomonas aeruginosa MTCC 2474 catalysed fuel cells. Bioresource Technology (2012) 124: 23–28.
[31] Zhou S, Wen J, Chen J, Lu Q. Rapid measurement of microbial extracellular respiration ability using a high-throughput colorimetric assay. Environmental Science & Technology Letters (2015) 2: 26–30.
[32] Josephy PD, Eling T, Mason RP. The horseradish peroxidase-catalyzed oxidation of 3,5,3’,5’-tetramethylbenzidine. Free radical and charge-transfer complex intermediates. The Journal of Biological Chemistry (1982) 257: 3669–3675.