PLANT SCIENCE

An algal photoenzyme converts fattyacids to hydrocarbonsDamien Sorigué,1 Bertrand Légeret,1 Stéphan Cuiné,1 Stéphanie Blangy,1

Solène Moulin,1 Emmanuelle Billon,1 Pierre Richaud,1 Sabine Brugière,2 Yohann Couté,2

Didier Nurizzo,3 Pavel Müller,4 Klaus Brettel,4 David Pignol,5 Pascal Arnoux,5

Yonghua Li-Beisson,1 Gilles Peltier,1 Fred Beisson1*

Although many organisms capture or respond to sunlight, few enzymes are known to bedriven by light. Among these are DNA photolyases and the photosynthetic reaction centers.Here, we show that the microalga Chlorella variabilis NC64A harbors a photoenzymethat acts in lipid metabolism. This enzyme belongs to an algae-specific clade of theglucose-methanol-choline oxidoreductase family and catalyzes the decarboxylation offree fatty acids to n-alkanes or -alkenes in response to blue light. Crystal structure ofthe protein reveals a fatty acid–binding site in a hydrophobic tunnel leading to thelight-capturing flavin adenine dinucleotide (FAD) cofactor. The decarboxylation is initiatedthrough electron abstraction from the fatty acid by the photoexcited FAD with a quantumyield >80%.This photoenzyme, which we name fatty acid photodecarboxylase, may be usefulin light-driven, bio-based production of hydrocarbons.

Inboth photosynthetic andnonphotosyntheticorganisms, light is involved in biological pro-cesses such as growth, development, metab-olism, and circadian entrainment (1–4). Inmost cases, the effects of light on cell phys-

iology are mediated by photoactive proteins,which include light-sensitive ion channels andpumps, photoreceptors, photosynthetic antennaproteins, and light-dependent enzymes (5). Amongthe latter, two different types can be distinguished.Light-activated enzymes require only a flash oflight to become active, whereas photoenzymesrequire a continuous flux of photons to remaincatalytically active—their activity is driven by light.Photoenzymes are a rare type of catalyst. In

nature, two families of proteins bearing a light-driven activity are DNA photolyases, which areinvolved in repairing ultraviolet (UV) damage inDNA in many organisms, from bacteria to plantsand animals (6–8), and light-dependent proto-chlorophyllide reductases, which catalyze chlo-rophyll synthesis (9). Chlorophyll f synthasemightbe a third type of photoenzyme, but the light-driven nature of the reaction has not been es-tablished (10). The multimeric complexes thatform the basis of oxygenic photosynthesis—plastocyanin:ferredoxin oxidoreductase (oftenreferred to as photosystem I) (11) and water:plastoquinone oxidoreductase (photosytem II)

(12), as well as related proteins in nonoxygenicphotosynthetic bacteria—may also be classifiedas photoenzymes (5).Photoenzymes are useful tools for research and

biotechnology. For example, light-controlled catal-ysis allows the study of rapid enzymatic processesin real time (13). And, the use of a photoexcitedcofactor to design an enzyme with a different cat-alytic function (14, 15) as well as the developmentof optogenetics (16, 17) expands the scope of pos-sible applications of enzymes driven by light.Some microalgae, including Chlorella variabilis

NC64A and Chlamydomonas reinhardtii 137C,

have the capacity to convert long-chain fattyacids to alkanes or alkenes in a process depen-dent on light (18). Because no homologs to knownhydrocarbon-forming enzymes have been foundin the genomes of C. variabilis or C. reinhardtii,we hypothesized that microalgae harbor a dif-ferent type of alkane synthase. Conversion ofmicroalgal fatty acids to hydrocarbons has beenmostly studied in Botryococcus braunii (19). Here,we identify an alkane synthase in the greenmicro-algae C. variabilis and show that it is a photo-enzyme belonging to an algae-specific subfamilyof proteins that has been conserved throughoutalgal evolution.

ResultsIdentification of a microalgalalkane synthase

In order to identify the enzyme responsible forhydrocarbon synthesis in microalgae, we under-took an approach of enzyme purification inC. variabilis. Following the observation thatC. variabilis cells can convert exogenous fattyacids into alkanes or alkenes (18), an activityassay for a hydrocarbon-forming enzyme was setup (fig. S1A). This discontinuous assay was sen-sitive enough to allow a partial purification of theprotein responsible for the alkane-forming activ-ity (fig. S1B). Proteomic comparison of three par-tially purified fractions resulted in a list of 10common proteins, nine of which were homol-ogous to well-characterized proteins (Fig. 1A).The most represented protein, and also the onlyone whose function was uncertain, was a mem-ber of the glucose-methanol-choline (GMC) oxido-reductases, a family of flavoproteins comprisinga wide range of oxidases and dehydrogenases(20). A full-length cDNAcoding for theC. variabilisGMC oxidoreductase was cloned and found to

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1Biosciences and Biotechnologies Institute of Aix-Marseille(BIAM), Commissariat à l'Energie Atomique et aux EnergiesAlternatives (CEA), CNRS and Aix-Marseille University, UMR7265 LB3M, CEA Cadarache, F-13108, Saint-Paul-lez-Durance,France. 2University Grenoble Alpes, CEA and INSERM, BIG-BGE,F-38000, Grenoble, France. 3European Synchrotron RadiationFacility, 71 Avenue des Martyrs, CS 40220, F-38043 Grenoble,France. 4Institute for Integrative Biology of the Cell (I2BC), CEA,CNRS, University Paris-Sud, University Paris-Saclay, F-91198,Gif-sur-Yvette cedex, France. 5BIAM, CEA, CNRS and Aix-Marseille University, UMR 7265 LBC, CEA Cadarache, F-13108,Saint-Paul-lez-Durance, France.*Corresponding author. Email: frederic.beisson@cea.fr

Fig. 1. Identificationof an alkane synthasein C. variabilis.(A) Number ofproteins found afterthe final step in threeindependent purifica-tions and list of the10 proteins in commonfound by means of pro-teomic analysis. Pro-teins are ranked bydecreasing spectralcounts. (B) Predicteddomains of theC. variabilis GMCoxidoreductase.(C) Chromatogramof the hydrocarbonsproduced (left) andimmunoblot of the totalproteins (right) in anE. coli strain expressingthe C. variabilis GMCoxidoreductase(alkane synthase).

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encode a polypeptide of 654 amino acids (~69 kDa)(Fig. 1B). Expression in Escherichia coli of thecorresponding codon-optimized synthetic gene(fig. S2) resulted in the production of C13 to C17alkanes or alkenes (Fig. 1C), which demonstratedthat this proteinwas indeed a hydrocarbon-formingenzyme.

Characterization of the fatty aciddecarboxylase activity

To further characterize the activity of the C.variabilis GMC oxidoreductase, we expressedthe recombinant protein in E. coli and purified it.Mass spectrometry analysis revealed the pres-ence of a flavin adenine dinucleotide (FAD)cofactor (fig. S3), as observed in other charac-terized GMC oxidoreductases (20). Characteri-zation of the hydrocarbon-forming activity ofthe recombinant protein showed that this ac-tivity was optimum around pH 8.5 and decreasedsharply when temperature rose above 35°C (fig.S3). No activity could be detected by using eithera fatty acid methyl ester or a fatty aldehyde assubstrate. In order to determine whether thereaction catalyzed by the alkane synthase wasan actual decarboxylation of fatty acid, palmiticacid labeled with 13C on the carboxylic groupwasused as substrate. Upon incubation of the sub-strate with the C. variabilis alkane synthase,production of 13CO2 was detected at more than

20 times its natural abundance (Fig. 2A), thusindicating that the enzyme was a fatty aciddecarboxylase.The purified enzyme was then tested on a

wide range of free fatty acid substrates (C12 toC22) (Fig. 2B). All of them were converted intoCn-1 alkanes or alkenes, with a higher efficiencyfor C16-C17 chains. Saturated fatty acids wereconverted to alkanes, not alkenes, which showedthat the C. variabilis fatty acid decarboxylase didnot introduce a terminal double bond in the car-bon chain, unlike the two fatty acid decarboxylasesidentified in bacteria (21, 22). We also identifieda GMC oxidoreductase from C. reinhardtii withfeatures and substrate specificities similar to itsC. variabilis homolog (fig. S4).

Light-dependence of the C. variabilisalkane synthase

In algal cells, alkane formation requires light (18).The fact that the C. variabilis alkane synthasecontains a chromophore, together with the ob-servation that the purified enzyme does notrequire any additional cofactor to be active,prompted us to check the light dependency ofthe enzyme activity. We first observed that whenthe E. coli strain expressing the C. variabilis al-kane synthase was cultivated under completedark and never exposed to light, no hydrocarbonwas produced (fig. S3G). Measurement of the

activity of the purified C. variabilis enzyme atdifferent wavelengths of visible light showedfurther that the alkane synthase was active inthe presence of blue light (400 to 520 nm) (Fig.3A). The action spectrum correlated with FADabsorbance, thus suggesting that FAD is thelight-capturing part of the protein. However,these observations did not discriminate betweena light-activated enzyme and a photoenzyme.The activity of the purified enzyme on 1-13C–labeled palmitic acid was thus measured underdifferent light conditions by monitoring the re-lease of 13CO2. We observed that 13CO2 produc-tion started immediately upon illumination withblue light and stopped immediately after a shiftto dark or red light (Fig. 3B). This demonstratedthat the C. variabilis alkane synthase is a photo-enzyme because it requires a continuous flux of(blue) photons for its activity. The same featurewas also noted on the purified C. reinhardtiiGMC oxidoreductase (fig. S4D). Alkane forma-tion increased with light intensity (Fig. 3C). Usingpalmitic acid as substrate, we estimated theturnover number of the C. variabilis GMC oxido-reductase at 0.86 ± 0.13 s−1 under a fluence rateof 2000 mmol photons m−2 s−1 white light. Thephotoexcitation rate (supplementary materials,materials and methods) in this experiment was~0.9 s−1, indicating efficient photocatalysis (quan-tum yield >80%). On the basis of these results, wepropose the name fatty acid photodecarboxylase(FAP) for the C. variabilis GMC oxidoreductaseand its C. reinhardtii homolog.The blue light dependency of the FAP prompted

us to investigate the effect of the quality oflight on the formation of hydrocarbons in algalcultures. When C. reinhardtii cultures weregrown in photobioreactors under successivelyblue (<500 nm) and red (>600 nm) lights, al-kane or alkene formationwas correlatedwith thepresence of blue light (Fig. 3D). This experimentthus showed that the light dependency of theFAP observed in vitro was relevant to living cellsand likely to be related to its function in the cell.

Three-dimensional structure of the FAP

In order to gain insights into the structural basisof the light-dependent fatty acid decarboxylase

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Fig. 2. Fatty acid decarboxylase activity of the C. variabilis alkane synthase. (A) Relativequantification of 13CO2 released upon incubation of 1-13C-palmitate with the purified recombinantenzyme. S, substrate; E, enzyme. (B) Relative activity of the purified recombinant enzyme on variousfree fatty acid substrates. Mean ± SD (n = 3 repeats).

Fig. 3. Light dependency of the C. variabilis alkane synthase.(A) Absorption spectrum and action spectrum of the purified enzyme.Mean ± SD (n = 3 repeats). The position of the flavin absorption maximumin the enzyme (467 nm) deviates from typical values (445 to 450 nm formost flavoproteins and free flavins). (B) Activity of the purified enzymeunder successive light conditions. Activity on 1-13C-palmitate was

monitored via release of 13CO2 by using membrane inlet mass spectrom-etry. (C) Dependence of the activity of the purified enzyme on whitelight intensity. Mean ± SD (n = 3 repeats). (D) Variation of totalhydrocarbons in C. reinhardtii cells grown in a photobioreactor underblue and then red light. Mean ± SD (n = 3 repeats). FAMEs, fatty acidmethyl esters.

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activity of the C. variabilis FAP, we determinedits three-dimensional (3D) structure in complexwith a palmitate substrate (table S1). A crystalform diffracting to 3.15-Å resolution was ob-tained. Although the crystal was twinned, wewere able to solve its structure via molecularreplacement by using domains from two pro-teins of the GMC oxidoreductase family withknown3D structures (PDB codes 2JBVand4YNT).The overall architecture of the FAP was similarto other GMC oxidoreductases, with a typicaltwo-domain fold (Fig. 4A). The first domain,mostly shaped by N-terminal residues, stabilizesthe FAD cofactor, whereas the second domainparticipates in substrate/product trafficking,withloops from both domains shielding the activesite from the solvent and holding together bothdomains.The 3D structure of the FAP revealed a narrow

hydrophobic tunnel, open to the solvent on oneend and leading to the tricyclic ring of FAD onthe other end (Fig. 4B). Electron density in thistunnel indicates the stabilization of a palmitatesubstrate that wraps around the side chain ofY466, with the carboxylate moiety interactingwith the side chains of residues R451, C432,and Q486 (Fig. 4C) and pointing toward the

FAD (shortest distance 3.7 Å) (Fig. 4D). Thelower enzyme activity for medium-chain fattyacids (Fig. 2B) may be caused by a less efficientstabilization of short hydrophobic chains in thetunnel (especially around Y466). A multiple se-quence alignment of GMC oxidoreductases ofthe FAP subgroup showed thatmost of the aminoacids of the hydrophobic tunnel present in thevicinity of the FAD or lining the entry site ofpalmitate (vestibule) were conserved (fig. S5).

Role of FAD in the FAP photocycle

To elucidate the role of the FAD cofactor in thedecarboxylation of fatty acids by FAP, we per-formed time-resolved optical spectroscopy mea-surements on the purified recombinant enzyme.The fluorescence kinetics (Fig. 5A) show thatthe substrate quenches the singlet excited state1FAD*, shortening its lifetime from t ~ 5 ns(typical time constant for unquenched flavins(23) to t ~ 300 ps. FAD-based reaction inter-mediates and kinetics were resolved by meansof transient absorption spectroscopy on nano-second to millisecond time scales. We presumedthat the 300-ps kinetics might reflect electrontransfer (ET) from the substrate to 1FAD*. In-deed, at 376 nm (where FAD•− absorbs more

strongly than FAD) (Fig. 5B) an instantaneous(within the instrumental time resolution of 10 ns)absorbance increase was observed (Fig. 5C, blacktrace), whereas at 488 nm (where FAD•− absorbsless than FAD) (Fig. 5B), an initial bleachingoccurred (Fig. 5C, blue trace). Measurements atother wavelengths confirmed the rapid forma-tion of FAD•− (fig. S6, A and B). At most wave-lengths studied, the initial absorbance changesdecayed with t ~ 100 ns (Fig. 5C and fig. S6A), in-dicating reoxidation of FAD•− to FAD. At 515 nm,however, the absorbance rose in ~100 ns (Fig.5C, green trace) and decayed with t ~ 4 ms (Fig.5C, inset). Because the slope of the FAD absorp-tion spectrum in FAP is steep at 515 nm (Fig. 5B),we considered the possibility that after reoxida-tion of FAD•−, the FAD spectrum was red-shiftedbecause of a change in its environment resultingfrom the decarboxylation of the substrate. Spec-tral analysis of the 4-ms phase (fig. S6C) con-firmed such a red-shift.In the absence of substrate, flash-induced

absorbance changes show the formation of theFAD triplet state 3FAD* (fig. S6, D and E), whichis consistent with an intersystem crossing yieldin the order of 70% for unquenched singlet ex-cited flavins (23) and decay of 3FAD* to ground

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Fig. 5. Characterization of C. variabilis FAP by means of time-resolvedspectroscopy. (A) Time-resolved fluorescence of FAP in the presence orabsence of palmitic acid. (B) Absorption spectra of FAD•− [in glucose oxidase(31); red line] and oxidized FAD in FAP (solid black line), scaled assuming amolar absorption coefficient of 11 300 M−1cm−1 for the maximum of the

absorption band in the blue (32). A red-shifted blue band of FAD in FAP (dottedblack line) is shown to demonstrate the effect on the absorption at 515 nm.(C) Examples of transient absorption changes of FAP in the presence of excesssubstrate. Complementary data are shown in fig. S6. (D) Suggestedmodel of theFAP photocycle. Directly observed intermediates and kinetics are shown in red.

Fig. 4. Structural features of the C. variabilis FAP. (A) Overallarchitecture of the enzyme in complex with FAD and palmitate (PLM).The structure is represented as an illustration colored from blue to redfrom the N to the C terminus, with the FAD and palmitic acid representedin stick. The long N-terminal helix is projected toward a noncrystallographicsymmetry–related molecule that is not represented here for clarity.(B) Slice through the surface representation of the fatty acid photo-decarboxylase. For clarity, small cavities in the interior of the enzyme are

not shown. (C) Details of the palmitic acid–binding site with the sidechains of residues within 4 Å of the substrate shown in stick. The omitmap electron density associated to palmitic acid is also shown andcontoured at 0.5s (2Fo-Fc; blue) and 2s (Fo-Fc; green). Single-letterabbreviations for the amino acid residues are as follows: A, Ala; C, Cys;D, Asp; E, Glu; F, Phe; G, Gly; H, His; I, Ile; K, Lys; L, Leu; M, Met; N, Asn;P, Pro; Q, Gln; R, Arg; S, Ser; T, Thr; V, Val; W, Trp; and Y, Tyr. (D) Distances(angstroms) between palmitic acid and FAD.

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state FAD in ~80 ms. In the presence of substrate,formation of 3FAD* should be largely outcom-peted by ET from the substrate to 1FAD* in~300 ps (fig. S6F), which is consistent with ahigh quantum yield of photocatalysis.

FAP clade in phylogenetic tree of GMCoxidoreductases

All members of the GMC oxidoreductase familycharacterized so far are oxidases or dehydro-genases (mostly acting on sugars), except someplant lyases that catalyze breaking of a C–C bond(24) and the FAP, which is also a lyase. Weanalyzed the phylogenetic relationships of 50prokaryotic and eukaryotic members of the GMCoxidoreductase family, including 20 proteins withdemonstrated biochemical activities. C. variabilisand C. reinhardtii FAP did not group with theplant clade but instead belonged to a separateclade that contained only sequences from algae(Fig. 6, table S2, and fig. S7). This algal-specificclade included micro- and macroalgae and algaeof primary or secondary endosymbiotic origin.Among the 22 algal species with sequenced ge-nomes, 18 had a GMC oxidoreductase in theFAP clade. The four other algae (Ostreococcusand Micromonas species, from the same basalMamiellophyceae class of green algae) lacked apredicted GMC oxidoreductase. Algal specieswith several GMC oxidoreductases were all ofsecondary endosymbiotic origin (such asNanno-

chloropsis, Phaeodactylum, or Emiliana). Thus,the algal clade of GMC oxidoreductases is likelyto be a group with a conserved FAP function.

Discussion

Here, we identified a microalgal alkane syn-thase as a member of the GMC oxidoreductasefamily and a photoenzyme. Known enzymeswith hydrocarbon-forming activity include oxido-reductases and polyketide synthases in cyano-bacteria (25, 26), fatty acid desaturase-like proteinsCER1 and CER3 in plants (27), cytochrome P450sin the bacterium Jeotgalicoccus sp. ATTC8456and in insects (21, 28), and nonheme diironoxidoreductases in the bacterium Pseudomonasaeruginosa (22). Algal GMC oxidoreductasesnow join that group but have the added featureof being light-driven. Photoenzymes are an an-cestral type of enzyme and may be rare becauseevolutionary selection has not favored light-drivencatalysis (29). Given the diversity of proteins ableto perform synthesis of alkanes, light activationrequired by the chemistry seems unlikely. Wetherefore speculate that the light-driven natureof the microalgal alkane synthases is related totheir function within the cell. The FAP functionlikely appeared after primary endosymbiosis, waskept during secondary endosymbiosis, but waslost in plants. Conservation of the FAP functionin algae throughout evolution suggests a biolog-ical function specific to this group of organisms.

Because C. reinhardtii and C. variabilis FAPare active on various fatty acids in vitro, the for-mation in C. reinhardtii and C. variabilis cellsof a predominant species of hydrocarbon (7-heptadecene) (18) may be due to the channelingof the precursor (cis-vaccenic acid) to the FAP.This result suggests that the C. reinhardtii andC. variabilis alkane synthases interact with ayet-to-be-identified lipolytic enzyme that releasescis-vaccenic acid from a lipid or cofactor.We propose a model of the FAP photocycle

(Fig. 5D) that combines observed FAD-basedintermediates and kinetics (in red) with in-ferred (not directly observed) correspondingchanges in the substrate. The FAD in FAP isfirst photoexcited (step 1). The singlet excitedstate 1FAD* abstracts an electron from thefatty acid in ~300 ps (step 2). The fatty acidradical decarboxylates yielding an alkyl radi-cal [step 3; not directly observed but analo-gous to the photo-Kolbe reaction (30)]. FAD•−

transfers the electron back to the resulting alkylradical in ~100 ns (step 4). Because of the ex-tremely strong proton affinity of alkyl anions,we suppose that the ET is coupled to protontransfer from a proton donor XH (one of theneighboring amino acid residues or a watermolecule). Alternatively, XH may transfer a hy-drogen atom to the primary alkyl radical, fol-lowed by ET from FAD•− to X• in ~100 ns. Wespeculate that the resulting negative charge ofX− may cause a red shift of the FAD absorptionband in the visible. The FAD band shifts backto the blue in ∼4 ms (step 5), possibly because ofreprotonation of X−. The alkane is released (step 6),and a new substrate is bound (step 7). Judgingfrom the crystal structure, we considered thebest candidates for XH would be cysteine C432and tyrosine Y466 (Fig. 4C). In order to test ourhypotheses about the involvement of a protondonor XH in the catalytic reaction, we havemutated these two residues to alanine, whichshould not be able to function as a proton orhydrogen atom donor. Although 50 to 60% ofpurified wild-type enzyme contained FAD, noneof the mutant proteins bound the FAD cofactor.Thus, the hypothesis could not be tested.Our identification of a photoenzyme acting

on lipids shows that light-driven catalysis is notrestricted to the processes of light capture anduse or to the repair of UV damages in DNA. Thisdiscovery should thus provide an incentive to re-investigate the activity of some enzymes involvedin metabolic or signaling pathways regulatedby light.

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Fig. 6. Unrooted phylogenetic tree of the GMC oxidoreductase family. The neighbor joiningmethod was used. Species are abbreviated to three letters (one for the genus and two for the speciesepithet) (table S2). Proteins with demonstrated biochemical activity are indicated by three additionalcapital letters: AAO, aryl alcohol oxidase; AOX, alcohol oxidase; FAP, fatty acid photodecarboxylase;CBQ, cellobiose dehydrogenase; CHD, choline dehydrogenase; COX, cholesterol oxidase; CKO,compound K oxidase; FDH, fructose dehydrogenase; FOX, fructose oxidase; GDH, glucosedehydrogenase; GOX, glucose oxidase; HFO, hydroxy fatty acid oxidase (genetic evidence only); HNL,hydroxynitrile lyase; and POX, pyranose oxidase.

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ACKNOWLEDGMENTS

We thank J. Ohlrogge for critical reading of the manuscript. Thanksare also due to V. Cardettini, A. Beyly, I. Té, P. Auroy, A. Beynel, J. Alric,C. Sahut, D. Lemaire, and F. Carrière for technical help or discussions.This work was supported by grants from L’Agence Nationale de laRecherche (ANR) (MUSCA ANR-13-JSV5-0005 to Y.L-B. and A*MIDEXANR-11-IDEX-0001-02 to G.P.), the European Union (EU) (Sun2Chemproject of ERASynBio), the French Infrastructure for IntegratedStructural Biology (ANR-10-INSB-05-01), the Proteomics FrenchInfrastructure (ANR-10-INBS-08-01), and the HelioBiotec platformfunded by the EU, the région Provence-Alpes-Côte-d’Azur (PACA), the

French Ministry of Research, and the CEA. PhD studentships wereprovided by the région PACA and the CEA (D.S.) and the École NormaleSupérieure Paris (S.M.). F.B. conceived the project. P.A., D.N., andD.P. acquired x-ray data and resolved the protein structure. S.B. andY.C. performed the proteomic analysis. P.M. and K.B. performedmeasurements and analyzed data of time-resolved spectrometry.D.S. executed other experiments and analyzed data with the help of allother authors. F.B., D.S., K.B., P.M., and P.A wrote the paper withcontributions from G.P., D.P., Y.C., B.L., P.R., and Y.B.-L. The codingsequence of the C. variabilis FAP has been deposited in GenBankunder accession no. KY511411. Atomic coordinates and structurefactors have been deposited in the Protein Data Bank (PDB) underaccession no. 5NCC. All other data needed to evaluate the conclusionsin the paper are present in the paper and/or the supplementarymaterials. D.S., F.B., B.L., S.C., S.B., and G.P. are inventors on patentapplication EPA 16305583 submitted by CEA and CNRS that covers“New fatty acid decarboxylase and its uses.”

SUPPLEMENTARY MATERIALS

www.sciencemag.org/content/357/6354/903/suppl/DC1Materials and MethodsFigs. S1 to S7Tables S1 to S3References (33–46)

11 May 2017; accepted 20 July 201710.1126/science.aan6349

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An algal photoenzyme converts fatty acids to hydrocarbons

Li-Beisson, Gilles Peltier and Fred BeissonSabine Brugière, Yohann Couté, Didier Nurizzo, Pavel Müller, Klaus Brettel, David Pignol, Pascal Arnoux, Yonghua Damien Sorigué, Bertrand Légeret, Stéphan Cuiné, Stéphanie Blangy, Solène Moulin, Emmanuelle Billon, Pierre Richaud,

DOI: 10.1126/science.aan6349 (6354), 903-907.357Science 

, this issue p. 903; see also p. 872Scienceproximity to the flavin adenine dinucleotide cofactor.photons to carry out its catalytic reaction. A long hydrophobic tunnel in the enzyme stabilizes the fatty acid substrates in

showed that hydrocarbon production requires visible light. In fact, the enzyme requires a constant input of bluecoliEscherichiaglucose-methanolcholine oxidoreductase (see the Perspective by Scrutton). Expression of the enzyme in

found aet al.Microalgae make hydrocarbons. In searching for the enzyme responsible, Sorigué Algal enzyme driven by blue light

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