Plant Physiology Preview. Published on June 10, 2016, as DOI:10.1104/pp.16.00462
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Short title: Microalgal hydrocarbon synthesis
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Corresponding author: Fred Beisson
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CEA Cadarache, France; Email: [email protected]
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Tel:
+33442252897
Fax:
+3344225626
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Microalgae synthesize hydrocarbons from long-chain fatty acids via a light-dependent pathway
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Damien Sorigué1, Bertrand Légeret1, Stéphan Cuiné1, Pablo Morales1, Boris Mirabella1, Geneviève
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Guédeney1, Yonghua Li-Beisson1, Reinhard Jetter2, Gilles Peltier1 and Fred Beisson1
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CEA and CNRS and Aix-Marseille Université, Biosciences and Biotechnologies Institute
(UMR 7265), Cadarache 13108, France
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Department of Botany and Department of Chemistry, University of British Columbia,
Vancouver V6T 1Z4, Canada
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Summary : A pathway converting C16 and C18 fatty acids to alkanes or alkenes, involving
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the light-dependent transformation of cis-vaccenic acid into 7-heptadecene, is present in
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various microalgae.
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F.B., R.J. and Y.L.-B. conceived the original research project; F.B., D.S., B.L., S.C., G.G. and G.P.
designed the experiments and analyzed the data; D.S., B.L., S.C., P.M., B.M. and G.G. performed the
experiments; F.B. and D.S. wrote the article with contributions from R.J., Y.B.-L., B.L. and G.P.
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Funding was provided by the Agence Nationale de la Rercherche (grant MUSCA n° ANR-13-
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JSV5-0005 to Y.L-B. and grant A*MIDEX n° ANR-11-IDEX-0001-02 to G.P.), the
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Commissariat à l’Energie Atomique et aux Energies Alternatives (CEA) and the région
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Provence-Alpes-Côte-d’Azur (PACA) (PhD studentship to D.S.). Support was also provided
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by the HélioBiotec platform funded by the European Regional Development Fund, the région
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PACA, the French Ministry of Research, and the CEA.
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Address correspondence to: [email protected]
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ABSTRACT (247 words, 250 words max)
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Microalgae are considered a promising platform for the production of lipid-based biofuels.
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While oil accumulation pathways are intensively researched, the possible existence of a
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microalgal pathways converting fatty acids into alka(e)nes has received little attention. Here,
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we provide evidence that such a pathway occurs in several microalgal species from the green
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and the red lineages. In Chlamydomonas reinhardti (Chlorophyceae), a C17 alkene, n-
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heptadecene, was detected in the cell pellet and the headspace of liquid cultures. The
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Chlamydomonas alkene was identified as 7-heptadecene, an isomer likely formed by
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decarboxylation of cis-vaccenic acid. Accordingly, incubation of intact Chlamydomonas cells
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with per-deuterated D31-16:0 (palmitic) acid yielded D31-18:0 (stearic) acid, D29-18:1 (oleic
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and cis-vaccenic) acids and D29-heptadecene. These findings showed that loss of the carboxyl
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group of a C18 monounsaturated fatty acid lead to heptadecene formation. Amount of 7-
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heptadecene varied with growth phase, temperature and was strictly dependent on light, but
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was not affected by an inhibitor of photosystem II. Cell fractionation showed that ca. 80% of
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the alkene is localized in the chloroplast. Heptadecane, pentadecane as well as 7- and 8-
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heptadecene were detected in Chlorella variabilis NC64A (Trebouxiophyceae) and several
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Nannochloropsis
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(Mamiellophyceae) and the diatom Phaeodactylum tricornutum produced C21 hexaene,
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without detectable C15-C19 hydrocarbons. Interestingly, no homologs of known hydrocarbon
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biosynthesis genes were found in the Nannochloropsis, Chlorella or Chlamydomonas
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genomes. This work thus demonstrates that microalgae have the ability to convert C16 and C18
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fatty acids into alka(e)nes by a new, light-dependent pathway.
species
(Eustigmatophyceae).
In
contrast,
Ostreococcus
tauri
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INTRODUCTION
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Hydrocarbons derived from fatty acids (i.e. alkanes and alkenes) are ubiquitous in plant and
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insect outermost tissues where they often represent a major part of cuticular waxes and play
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an essential role in preventing water loss from the organisms to the dry terrestrial environment
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(Hadley, 1989; Kunst et al., 2005). In several insect species, select cuticular alkenes also act
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as sex pheromones (Wicker-Thomas and Chertemps, 2010). Occurrence of alkanes or alkenes
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has also been reported in various microorganisms (Ladygina et al., 2006; Wang and Lu,
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2013). For example, synthesis of hydrocarbons is widespread in cyanobacteria (Coates et al.,
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2014) and it is thought that cyanobacterial alka(e)nes contribute significantly to the
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hydrocarbon cycle of the upper ocean (Lea-Smith et al., 2015).
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Alka(e)nes of various chain lengths are important targets for biotechnology because
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they are major components of gasoline (mainly C4-C9 hydrocarbons), jet fuels (C5-C16) and
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diesel fuels (C12-C20). The alkane biosynthetic pathway of plants has been partly elucidated
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(Lee and Suh, 2013; Bernard and Joubès, 2013) but the use of plant hydrocarbons as a
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renewable source of liquid fuels is hampered by predominance of constituents with high
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carbon numbers (>C25), which entails solid state at ambient temperature (Jetter and Kunst,
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2008). Therefore, there is great interest in the microbial pathways of hydrocarbon synthesis
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producing shorter chain compounds (C15-C19). In cyanobacteria, hydrocarbons are produced
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by two distinct pathways. The first one comprises the sequential action of an acyl-ACP
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reductase transforming a C15-C19 fatty acyl-ACP into a fatty aldehyde, and an aldehyde-
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deformylating oxygenase catalyzing the oxidative cleavage of the fatty aldehyde into
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alka(e)ne and formic acid (Schirmer et al., 2010; Li et al., 2012). The second pathway
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involves a type I polyketide synthase that elongates and decarboxylates fatty acids to form
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alkenes with a terminal double bond (Mendez-Perez et al., 2011). Additional pathways of
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alkene synthesis have been described in bacteria. In Micrococcus luteus, a three-gene cluster
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has been shown to control the head-to-head condensation of fatty acids to form very-long-
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chain alkenes with internal double bonds (Beller et al., 2010). Direct decarboxylation of a
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long-chain fatty acid into a terminal alkene has also been reported and is catalysed by a
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cytochrome P450 in Jeotgalicoccus spp. (Rude et al., 2011), and by a non-heme iron oxidase
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in Pseudomonas (Rui et al., 2014).
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Among microbes, microalgae would be ideally suited to harness the synthesis of
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hydrocarbons from fatty acid precursors because they are photosynthetic organisms
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combining a high biomass productivity (León-Bañares et al., 2004; Beer et al., 2009; Malcata,
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2011; Wijffels et al., 2013) and a strong capacity to synthesize and accumulate fatty acids (Hu
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et al., 2008; Harwood and Gushina, 2009; Liu and Benning 2013). Studies on microalgal
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alka(e)nes synthesis are scarce however. In some diatoms and other algal species, a very long
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chain alkene, a C21 hexaene, has been found (Lee et al., 1970; Lee and Loeblich, 1971).
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Other very long chain alkenes have been described in the slow-growing colonial
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Chlorophycea Botryococcus brauni, which excretes a variety of hydrophobic compounds
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including C27 n-alkadienes (Metzger and Largeau, 2005; Jin et al., 2015). A decarbonylase
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activity converting a fatty n-aldehyde substrate to a n-alkane has been shown in Botryococcus
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(Dennis and Kolattukudy, 1992), however the corresponding protein has not been identified
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so far. Presence of shorter chain alka(e)nes in some microalga species has been reported in the
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context of geochemical studies (Han et al., 1968; Gelpi et al., 1970; Tornabene et al., 1980;
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Afi et al., 1996) but the biology of these compounds has not been investigated further.
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Here, we show that alka(e)nes with C15 to C17 chains can be detected in several
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model microalgae and originate from fatty acid metabolism. In Chlamydomonas reinhardtii
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and Chlorella variabilis NC64A, 7-heptadecene is identified as the major hydrocarbon
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produced, and we demonstrate that its synthesis depends strictly on light and uses cis-
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vaccenic acid as a precursor. We also show the presence of C15 to C17 alka(e)nes in
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Nannochloropsis sp., a model microalga from the red lineage. Absence of homologs to known
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hydrocarbon synthesis genes in the genomes of Chlamydomonas, Chlorella and
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Nannochloropsis indicates that a hitherto-unknown type of alka(e)ne-producing pathway
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operates in these microalgae. The wide occurrence of microalgae in marine environments
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suggests that microalgal alka(e)nes contribute significantly to the oceans’ hydrocarbon cycle.
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RESULTS
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Chlamydomonas and Chlorella synthesize long-chain alka(e)nes
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To investigate the capacity of microalgae to produce alkanes or alkenes, we first analyzed the
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unsaponifiable fraction of vegetative cells in two prominent green microalga models,
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Chlamydomonas reinhardtii (CC124 strain) and Chlorella variabilis NC64A. The solvent
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extracts of the saponification mixtures were analyzed by gas chromatography coupled to mass
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spectrometry (GC-MS). The chromatograms of Chlorella and Chlamydomonas extracts
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showed a peak at 14.3 min in the region of long-chain hydrocarbons (Fig. 1A). This peak
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displayed a mass spectrum identical to that of a heptadecene standard (Fig. 1B). In Chlorella,
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two close peaks corresponding to heptadecene were present (Fig. 1A). In addition, n-
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heptadecane (peak at 14.6 min) and traces of n-pentadecane (peak at 12.3 min) were detected
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(Supplemental Fig. S1). The finding that all these hydrocarbons were absent from control
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samples generated by saponification of fresh culture medium ruled out that any hydrocarbons
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had been introduced as contamination from solvents, reagents or culture medium (Fig. 1A).
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In order to further exclude the possibility that the harsh conditions of the
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saponification may have caused an artefactual decarboxylation of fatty acids to alka(e)nes,
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volatile compounds present in the headspace (gas phase) of Chlamydomonas and Chlorella
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cultures were analyzed by solid phase microextraction (SPME), which does not involve any
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treatment of cells with chemical reagents or heat. The SPME fiber was put in the headspace of
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a sealed vial containing a Chlamydomonas liquid culture or the fresh culture medium.
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Analysis by GC-MS of the compounds absorbed onto the SPME fiber showed that
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heptadecene was indeed present in the gas phase of a Chlamydomonas culture, but was absent
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when no cells were present in the culture medium (Supplemental Fig. S2). The Chlorella
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hydrocarbons found by saponification were also detected by SPME but no other hydrocarbon
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peak could be found. These results thus establish that the long-chain alka(e)nes detected in C.
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reinhardtii and C. variabilis cells are genuine products of microalgal metabolism.
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Total alka(e)ne amounts per cell volume were similar in Chlorella and
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Chlamydomonas as determined by total transmethylation of cells and GC-MS/FID analysis of
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hydrocarbons and fatty acid methyl esters (FAMEs) (Fig. 1C). This represented ca. 0.62%
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and 1.15% of total fatty acids in Chlamydomonas and Chlorella, respectively (or 0.036% and
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0.08% of biomass dry weight). Heptadecenes formed ca. 80% of hydrocarbons in Chlorella.
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A control performed with pure oleic acid showed that the heptadecene quantified after
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transmethylation was not originating from spontaneous fatty acid decarboxylation
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(Supplemental Fig. S3).
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To reveal the heptadecene double bond positions, extracts of saponified
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Chlamydomonas or Chlorella cells were derivatized with dimethyl disulfide (DMDS) and the
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reaction products were analyzed by GC-MS. In both species, a peak with a mass spectrum
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characteristic of the DMDS derivative of 7-heptadecene (diagnostic fragments at m/z 145 and
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187) was observed (Fig. 2). An additional minor peak with fragments at m/z 159 and 173,
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indicative of 8-heptadecene could be detected in Chlorella, but was absent in
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Chlamydomonas. Therefore, we conclude that the heptadecene produced in Chlamydomonas
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(and the major in Chlorella) is 7-heptadecene. The minor isomer of Chlorella is 8-
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heptadecene (around 20% of the major isomer).
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Long-chain alka(e)nes derive from a formal decarboxylation of fatty acids
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One of the simplest biosynthetic routes to synthesize heptadecenes and other long-chain
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hydrocarbons is the formal decarboxylation of corresponding fatty acids. To determine
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whether Chlamydomonas and Chlorella alka(e)nes originate from fatty acid precursors, cells
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were cultured for three days in Tris Acetate Phosphate (TAP) medium augmented with per-
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deuterated palmitic acid (D31), and total fatty acid content was analyzed by GC-MS/FID
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following transmethylation. In the Chlamydomonas product mixture, D31-16:0 (palmitic), D31-
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18:0 (stearic) and D29-18:1 acids were observed, indicating that the algal cells had taken up
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exogenous D31 palmitic acid and metabolized it (Fig. 3). Interestingly, D29-heptadecene was
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identified based on MS fragmentation of the GC peak eluting at 9.2 min. Incubation of
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Chlorella variabilis NC64A with per-deuterated palmitic acid (D31) also resulted in the
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formation of D31-16:0, D31-18:0 and D29-18:1 acids (Supplemental Fig. S4). Formation of
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D29-heptadecene and additionally of D31-heptadecane was observed for Chlorella (Fig. 3). It
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should be noted that the retention behaviour of per-deuterated fatty acids and hydrocarbons,
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with elution long before corresponding unlabeled compounds, was coherent with previous
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observations on per-deuterated alkanes (Matucha et al., 1991). Overall, labelling experiments
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therefore showed that Chlamydomonas and Chlorella had enzyme(s) that catalyzed the
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conversion of long-chain fatty acids into alka(e)nes by a formal decarboxylation.
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Heptadecene content varies with growth stage and culture conditions
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We next sought to determine the effect of physiological conditions, growth stage or life cycle
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on the content of hydrocarbons in a microalga. We used Chlamydomonas reinhardtii for these
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experiments, because it is the most-studied model for algal physiology, including lipid
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metabolism (Merchant et al., 2012; Li-Beisson et al., 2015). Initial experiments showed no
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significant differences in heptadecene content between the vegetative cells of the strains
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CC124 (mating type: minus) and CC125 (mating type: plus) (Supplemental Fig. S5).
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Similarly, induction of gametogenesis in the vegetative cells of the two mating types did not
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result in any significant decrease or increase.
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Hydrocarbons were then quantified in vegetative cells of CC124 under various
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culture conditions. We first determined the heptadecene content as a function of algal growth
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under standard laboratory conditions, i.e. in flasks under constant illumination, either in
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photoautotrophy (minimal medium, i.e. with light as only source of energy) or in mixotrophy
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(TAP medium, i.e. acetate and light as energy sources). Heptadecene levels increased steadily,
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up to 422 ng/mm3 cell volume at 168 h in TAP media, corresponding to 368 ng/mg biomass
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dry weight. The alkene amounts were at all times substantially higher under mixotrophic
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conditions than under photoautotrophic conditions (Fig. 4A). The increase in heptadecene
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concentration in the cells appeared to correlate with the increase in cell concentration in the
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culture (Fig. 4B). Under TAP medium, transfer to a nitrogen-deprived medium, which causes
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slow-down and eventually arrest of cell division (Merchant et al., 2012), resulted in an initial
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strong reduction of heptadecene content and an arrest of the synthesis of heptadecene (Fig.
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4C). By contrast, under minimal medium (MM) conditions, nitrogen deprivation caused an
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initial reduction of heptadecene, but after 24h the synthesis resumed and at 96h reached
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similar values as in MM (Fig. 4D). When cells were cultivated at various temperatures,
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heptadecene content tended to increase with temperature, with a significant difference
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between 35°C and 4°C (Supplemental Fig. S6).
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Synthesis of alka(e)nes is strictly dependent on light but not on photosynthesis
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In order to gain more insights into the environmental parameters that might affect synthesis of
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heptadecene by Chlamydomonas cells, the effect of light was investigated. When
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Chlamydomonas cells were grown in flasks under a night and day cycle (12 h light/12 h dark),
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it was observed that heptadecene content per cell increased by about 50% during the day,
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decreased steeply during the first three hours of the night and stayed constant during the rest
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of the night (Fig. 5A). This suggested that heptadecene synthesis was dependent on light and
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prompted us to investigate the synthesis of heptadecene in complete absence of light. Cells
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were grown heterotrophically for five days in a flask in the dark and heptadecene was
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quantified. Strikingly, no alkene could be detected in dark-grown cells, in stark contrast to
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cells grown in the same medium under constant light (Fig. 5B). This showed that formation of
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Chlamydomonas heptadecene was strictly dependent on light. A strong light dependence of
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alka(e)ne synthesis was also observed in Chlorella variabilis (Supplemental Fig. S7).
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Interestingly, when Chlamydomonas cells were broken and chloroplasts isolated, 80% of the
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cellular heptadecene was found to be present in the chloroplast fraction (Fig. 5C).
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To further investigate the dynamics of light-dependent heptadecene formation and its
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potential link to photosynthesis, cells were grown for five days in the dark and then moved to
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light. The levels of newly formed heptadecene reached 10 ng per mm3 cell volume already 10
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min after switching the light on, and this amount increased only slightly in the next few hours
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to reach ca. 15 ng per mm3 cell volume at 6 h (Fig. 5D). Similar results were obtained in the
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presence or absence of 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), an inhibitor of
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photosystem II, showing that heptadecene formation was not dependent on photosynthesis.
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Finally, we sought to test the effect of light intensity on alkene synthesis. For this
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purpose, Chlamydomonas cells were grown in a photobioreactor operated as a turbidostat (i.e.
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at constant cell density) to ensure a constant illumination of the culture. Heptadecene content
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per cell volume unit at each light intensity increased in the range 125 to 300 µE (Fig. 5E).
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Heptadecene productivity (calculated from steady state heptadecene cellular contents and
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dilution rates) followed the same trend as biomass productivity, increasing steeply with light
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intensity from 15 to 125 µmol photons m-2 s-1, but kept increasing at high light intensities
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(Supplemental Fig. S8) due to the increase in the heptadecene amount per cell (Fig. 5).
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Microalgae producing long-chain alka(e)nes have no homologs to known alka(e)ne-
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forming enzymes
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The presence of long-chain hydrocarbons in Chlamydomonas reinhardtii and Chlorella
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variabilis NC64A raised the questions whether similar compounds also occurred in other
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microalgae of the green lineage or in species of the red lineage, and whether these compounds
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could be produced by enzymes homologous to the ones found in plants, cyanobacteria or
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other organisms. We thus investigated the presence of hydrocarbons in some other model
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microalgae with sequenced genomes: the Mamiellophycea Ostreococcus tauri (green
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lineage), and the diatom Phaeodactylum tricornutum and the Eustigmatophycea
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Nannochloropsis sp. (red lineage). Heptadecane and heptadecene were observed in all strains
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of Nannochloropsis, and in all species except Nannochloropsis limnetica pentadecane was
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also detected (Fig. 6). The total amounts of hydrocarbons varied up to 10-fold between the
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various Nannochloropsis species, with a maximum of 597 ng/mm3 cell volume in N.
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gaditana. Although we could not detect any C15-C17 hydrocarbons in P. tricornutum and O.
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tauri under the culture conditions used, we identified a C21:6 hexaene, all-cis-3,6,9,12,15,18-
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heneicosahexaene, based on matching mass spectrum and retention time with the same
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compound previously described for several diatoms and various other algae (Lee and
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Loeblich, 1971), namely the one derived from all-cis-4,7,10,13,16,19-docosahexaenoic acid
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(DHA) (Supplemental Fig. S9). Therefore, all the model microalgal species investigated
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produced alka(e)nes, either in the form of C15-C17 alka(e)nes or the C21:6 hexaene.
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Finally, to determine if the microalgal alka(e)nes detected here could be produced
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along known hydrocarbon biosynthesis pathways, the algal genomes were searched for
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potential orthologs to genes encoding enzymes known to be involved in hydrocarbon
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formation. BlastP searches were performed in the genomes of Chlamydomonas (Merchant et
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al.; 2007), Chorella (Blanc et al., 2010), Nannochloropsis (Radakovits et al., 2012; Vieler et
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al., 2012), Phaeodactylum (Bowler et al., 2008) and Ostreococcus (Derelle et al., 2006) using
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the sequences of the cyanobacterial acyl ACP-reductase and Aldehyde Deformylating
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Oxygenase (Schirmer et al., 2010), the plant ECERIFERUM 1 and 3 (Aarts et al., 1995;
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Rowland et al., 2007; Bernard et al., 2012), the insect cytochrome P450 CYP4G1 (Qiu et al.,
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2012), the bacterial cytochrome P450 CYP152/OleT (Rude et al., 2011) and the bacterial non-
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heme oxidase UndA (Rui et al., 2014). The genomes of Ostreococcus tauri and
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Phaeodactylum tricornutum both contained one potential ortholog to the plant ECERIFERUM
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1 and ECERIFERUM 3 (CER1/3) (Table 1). However, no other homologs to known
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hydrocarbon synthesis genes were found in the genomes of Chlorella, Chlamydomonas and
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Nannochloropsis.
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DISCUSSION
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C15-C17 alka(e)nes are produced by various microalgae and are as abundant as in
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cyanobacteria
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The C21 hexaene derived from DHA that is formed by diatoms has been well characterized
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(Lee et al., 1970; Lee and Loeblich, 1971). By contrast, the endogenous or exogenous origin
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of the various shorter chain alkanes reported in geochemical studies for some microalgal
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species has not been investigated further. Possible exogenous origin of hydrocarbons detected
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in minute to trace amounts in biological material is a major concern because medium- and
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long-chain hydrocarbons are common contaminants of organic solvents and culture media.
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Here, using SPME analysis of intact cells and culture medium as well as labeling with
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deuterated fatty acid precursors (Fig. 1, Fig. 2, Supplemental Fig. S1,2 and 3), we show
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unambiguously that C15-C17 alka(e)nes are synthesized by several model microalgae (Table
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1). These include the Chlorophycea Chlamydomonas reinhardtii and the Trebouxiophycea
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Chlorella variabilis NC64A, which belong to the Archaeplastida (green lineage), and the
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Eustigmatophyca Nannochloropsis sp. which is a microalga of the red lineage closely related
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to the Phaeophyceae (brown algae).
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marine species but can thrive in a variety of environments and a freshwater species is known
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(N. limnetica). Interestingly, all 6 reported species of Nannochloropsis including N. limnetica
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produced C15-C17 hydrocarbons. Although Ostreococcus (green lineage) and the diatom
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Phaeodactylum (red lineage) have C16-C18 fatty acid, they did not produce detectable
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amounts of C15-C17 hydrocarbons under the cultivation conditions used. However, they
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formed a C21 hexaene (Supplemental Fig. S9). Taking together, these data suggest that
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hydrocarbons synthesized from fatty acids are likely to be present in a wide array of
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microalgal groups and that in microalgal species devoid of very-long-chain fatty acids
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(>C20), C15-C17 alka(e)nes may be synthesized, preferentially from cis-vaccenic acid.
Nannochloropsis species are mostly considered as
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In Chlamydomonas and Chlorella, C15-C17 alka(e)nes represent 0.04 to 0.1% dry
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weight. This is in the same range as strains for two abundant marine cyanobacteria,
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Prochlorococcus and Synechococcus, which produce between 0.02 and 0.4% dry weight of
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such compounds (Lea-Smith et al., 2015). In the same work on cyanobacterial hydrocarbons,
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it has been suggested that cyanobacterial hydrocarbons may play a significant role in the
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hydrocarbon cycle of upper oceans. Given the relatively high abundance of eukaryotic
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phytoplankton compared to total prokaryotes in sunlit oceans (de Vargas et al., 2015), our
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data suggest that microalgal hydrocarbons may also contribute significantly to this cycle.
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Microalgal alka(e)ne synthesis requires light
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Our data show that light is required for hydrocarbon synthesis in Chlamydomonas and
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Chlorella (Fig. 5B; supplemental Fig. S7). Accordingly, when Chlamydomonas cells are
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cultivated under day and night cycles (Fig. 5A, Supplemental Fig. S10), alkenes are
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produced during the day (quantity of alkenes per cell increases while cell number is constant)
326
and at the beginning of the night the alkene amount per cell decreases in the same proportion
327
as the cell number increases. Decrease of the cellular alkene content during the dark phase
328
seems thus due to cell division in absence of alkene synthesis. Upon illumination, synthesis of
329
heptadecene occurs within a few minutes (Fig. 5D). Taken together these data show that the
330
microalgal synthesis of heptadecene is strongly dependent on light and occurs only during the
331
day. Presence in the chloroplast of 80% of Chlamydomonas heptadecene suggests a
332
choroplastic location of the synthesis (Fig. 5C). Interestingly, the fact that alkene production
333
is not affected by the photosystem II inhibitor DCMU, shows that the alkene biosynthetic
334
pathway is not coupled to photosynthetic electron transfer reactions either in a direct or
335
indirect manner. It seems therefore clear that microalgal synthesis of C15-C17 hydrocarbons
336
is linked to the presence of light but is not related to the photosynthetic activity. To our
337
knowledge, a strict light dependency of alka(e)ne synthesis in photosynthetic cells has never
338
been reported. In cabbage leaves, light has been shown to increase the synthesis of some
339
cuticular wax compounds, including alkanes (Macey et al., 1970). In Euphorbia suspension
340
cells, light has been shown to affect the chain length distribution of the alkanes produced but
341
alkane synthesis can occur in the dark (Carrière et al., 1990). In Arabidopsis, the total wax
342
loads from stems and leaves were ∼20% lower in plants grown in dark conditions for 6 days
343
compared with those grown under long-day conditions (Go et al., 2014). An AP2/ERF-type
344
transcription factor that is preferentially expressed in the epidermis and induced by darkness
345
has been show to downregulate the expression of alkane-forming enzymes. In
346
Chlamydomonas, the complete absence of heptadecene in the dark and the rapid induction of
347
heptadecene synthesis by light (Fig. 5B and D) seems to point to a direct positive regulation
348
by light of alkane-forming enzymes or genes rather than a negative regulation induced by
349
dark. It should be noted that light may not be the only environmental factor affecting
350
heptadecene. Transfer of cells to a nitrogen-deprived medium (Fig. 4) clearly caused an arrest
351
of its synthesis at least for 24h, and the initial drop in heptadecene content after transfer
352
suggests the alkene is degraded or released. Also, heptadecene increases with cultivation
353
temperature (Supplemental Fig. S6), which might indicate a role in membrane fluidity.
10
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354
The major long-chain alka(e)ne produced derives from cis-vaccenic acid
355
Labeling experiments with per-deuterated palmitic acid indicate that heptadecenes result from
356
the loss of the carboxyl group in a C18 monounsaturated fatty acid (Fig. 3). The two major
357
possible origins of heptadecenes are oleic acid (double bond in Δ9 position) and its positional
358
isomer cis-vaccenic acid (Δ11), a bacterial-type fatty acid fairly abundant in some microalgae,
359
including Chlamydomonas reinhardtii (Giroud et al., 1988). Loss of the carboxyl group in
360
oleic and cis-vaccenic acids would yield 8-heptadecene and 7-heptadecene, respectively
361
(Supplemental Fig. S11). Analysis of Chlamydomonas or Chlorella hydrocarbons with
362
dimethyl disulfide (DMDS) revealed that the heptadecene produced in Chlamydomonas (and
363
the major heptadecene in Chlorella and Nannochloropsis) was 7-heptadecene (Fig. 2). The 8-
364
heptadecene is a minor isomer in Chlorella. The major hydrocarbon of Chlorella,
365
Chlamydomonas and Nannochloropsis is therefore derived from cis-vaccenic acid and not
366
from oleic acid. This indicates a high substrate specificity of at least one enzyme in the
367
biosynthetic pathway. A simple explanation to account for this specificity could be the
368
involvement of a phospholipase A1 (Fig. 7). Indeed, in Chlamydomonas the lipid class most
369
enriched in cis-vaccenic acid is phosphatidylinositol and this fatty acid is found almost only in
370
the sn-1 position (Giroud et al., 1988). The free fatty acid released by might be converted to
371
an alkene via a fatty aldehyde intermediate (cyanobacterial- or plant-type pathway involving
372
an acyl-ACP or acyl-CoA reductase). Alternatively, the alkene might be directly produced
373
from the fatty acid by a bacterium-type pathway (Rude et al., 2011; Rui et al., 2014) involving
374
an actual or formal decarboxylation.
375
376
Microalgae harbor new alka(e)ne-forming enzymes
377
Regardless of whether or not the microalgal synthesis of long-chain alka(e)nes involves an
378
aldehyde intermediate, the presence of C15-C17 alka(e)nes in Chlamydomonas, Chlorella,
379
Nannochloropsis and the absence in these species of proteins homologous to alkane-forming
380
enzymes from plants, cyanobacteria, bacteria or insects, clearly indicate that some hitherto-
381
unidentified alka(e)ne-forming enzyme(s) are present in these three microalgae (Table 1).
382
This conclusion is consistent with the unique strong lightdependency of heptadecene synthesis
383
observed in Chlamydomonas and Chlorella. The C21 hexaene found in the Mamiellophycea
384
Ostreococcus tauri and in the diatom Phaeodactylum might be synthesized by the plant
385
CER1/CER3 homolog found in these two species only. Microalgal homologs of CER1/CER3
386
might be thus more specific of very-long-chain fatty acids, but another biochemical role
11
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
387
cannot be excluded. Clearly, further genetic or biochemical work will be needed to identify
388
the enzyme(s) responsible for long-chain alka(e)ne synthesis in microalgae. This will be an
389
important step toward unraveling the function of alka(e)nes within the microalgal cell as well
390
as elucidating its regulation in connection with light and other environmental factors. Possible
391
roles of hydrocarbons in chloroplasts of microalgae include cell signaling or the regulation of
392
the fluidity of photosynthetic membranes in response to changes in light intensity or
393
temperature. Given the high hydrophobicity of hydrocarbons, a role in regulating membrane
394
properties seems more likely., From a biotechnological perspective, the discovery of the
395
microalgal alkane synthase(s) would also expand the repertoire of enzymes available to
396
harness hydrocarbon synthesis in microbes. (Choi et al., 2013; Liu et al. 2014; Chen et al.,
397
2015). Conversion of exogenously added palmitic acid to alkane by Chlamydomonas and
398
Chlorella cells shows that microalgal hydrocarbon-forming enzymes have in fact a broad
399
substrate specificity. They could therefore be used to generate alkanes and alkenes from
400
abundant cellular fatty acids in a bacterial or microalgal host. Combination of this alkane-
401
forming system with a medium-chain specific thioesterase would allow to produce volatile
402
alka(e)nes and circumvent the need for intracellular alkane storage (Choi et al., 2013).
403
Identification of the enzymes forming C15-C17 hydrocarbons in green microalgae is therefore
404
an exciting task from the point of view of microalgal physiology but also from a
405
biotechnological perspective.
406
407
408
MATERIAL AND METHODS
409
Strains and culture conditions
410
Chlamydomonas reinhardtii wild-type strains CC124 (nit1 nit2; mt-) and CC125 (nit1 nit2;
411
mt+) was used. Nannochloropsis species were from the CCMP culture collection:
412
Nannochloropsis oceanica CCMP 531 and 1779; Nannochloropsis gadinata CCMP 527;
413
Nannochloropsis
414
Nannochloropsis salina CCMP 537 and 538 and Nannochloropsis oculata CCMP 525.
415
Chlorella variabilis NC64A was from the laboratory of JL Van Etten (University of
416
Nebraska). Other strains were from the Roscoff Culture Collection: Ostreococcus tauri
417
RCC745 (strain OTTH0595-genome); Phaeodactylum tricornutum RCC 2967 (strain
418
Pt1_8.6). All strains except Ostreococcus were grown routinely in conical flasks in incubation
419
shakers at 25°C (Infors HT) under air enriched with 2% CO2, with agitation at 140 rpm and
420
light intensity at 120 µmol photons m-2 s-1 (µE) (70 µE for Chlorella). Ostreococcus was
granulata
CCMP
529;
Nannochloropsis
limnetica
CCMP
505;
12
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
421
grown in a culture chamber at 25°C in flasks without agitation and using a light intensity of
422
80 µE. Regarding culture media, Chlamydomonas and Chlorella were cultivated in Tris-
423
Acetate-Phosphate (TAP) medium and minimal medium (Harris, 1989) containing 20mM
424
MOPS. Other microalgae were grown in artificial sea water added with various nutrients:
425
Conway’s, Guillard's F/2 (Lananan et al., 2013) and Keller's (Keller et al., 1987) media were
426
used for Nannochloropsis, Phaeodactylum and Ostreococcus respectively. Cells were
427
routinely counted using a Multisizer™ 3 (Coulter). To produce gamete, culture medium of
428
vegetative cells in exponential growth phase (10 million cells ml-1) was changed from TAP
429
medium to TAP nitrogen depleted medium (TAP-N). To be sure to remove all nitrogen, cells
430
were washed three times with TAP-N medium. Cells were kept in TAP-N for 16 hours before
431
analysis.
432
433
Cultures in photobioreactors
434
Chlamydomonas reinhardtii CC124 (mt- nit1 nit2) cells were cultured in minimal medium
435
(Harris, 1989) in one liter photobioreactors (BIOSTAT Aplus, Sartorius Stedim Biotech)
436
operated as turbidostats. A880 was measured continuously using a biomass probe (Excell
437
probe, Exner) and cultures were maintained at constant A880 by injection of fresh medium.
438
The pH was maintained at a constant value of 7 by injection of KOH (0.2N) or HCL (0.2N).
439
The cultures were stirred using a metal propeller (250 rpm). The gas flow rate was adjusted to
440
0.5 liters min-1. Air enriched with 2% CO2 was generated using mass flow meters (EL flow,
441
Bronkhorst). Light was supplied by eight fluorescent tubes (Osram Dulux L 18 W) placed
442
radially around the photobioreactor. Cells were cultivated for three days with the same light
443
intensity before collecting cells. Light intensity was increased gradually from 15 to 300 µmol
444
photons m-2 s-1.
445
446
Cell labeling with deuterated palmitic acid
447
One hundred million cells of Chlamydomonas reinhardtii 137c and 200 million cells of
448
Nannochloropsis gaditana CCMP 527 and Chlorella variabilis NC64A were incubated for 24,
449
48 or 72 hours in sealed tubes with 5 mL of TAP medium containing 100µM of perdeuterated
450
(D31) palmitic acid (from a 10 mM stock solution in dimethyl sulfoxide). Total lipids and
451
alka(e)nes were transmethylated and analyzed by GC-MS/FID as described below.
452
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453
Saponification
454
Cell pellets (100 million cells for Chlamydomonas, 200 million cells for other microalgae)
455
were added with 4 ml of an aqueous solution of KOH 5% (w/v) and heated for 90 min at 85°C
456
in sealed vials. After cooling down, 4 mL of NaCl 0.9% (w/v) was added and unsaponifiable
457
compounds were extracted 3 times with 4 mL methyl-tert-butyl-ether (MTBE). Samples were
458
vortexed for 10 min and the organic phases were recovered by centrifugation at 3000g for 2
459
min and pooled. MTBE was evaporated under a gentle stream of nitrogen gas and
460
unsaponifiables were re-suspended in 500 µL hexane and 1 µl was injected in the GC-MS.
461
462
Transmethylation
463
To quantify hydrocarbons together with fatty acids, transmethylation of whole cells was used.
464
Briefly, cell pellets (one hudred million cells for Chlamydomonas, two hundred million cells
465
for the other microalgae) were added with 2 mL of a solution containing methanol with 5%
466
(v/v) sulfuric acid and 25 µg of triheptadecanoate (from a stock solution 2.5 mg.mL-1 in
467
chloroform) and 5 µg of 16:0-alkane (stock solution 1 mg.mL-1 in chloroform) were included
468
as internal standards. Samples were incubated at 85°C for 90 minutes in sealed glass tubes.
469
After cooling down, FAMEs and hydrocarbons were extracted by adding 500 µL hexane and
470
500 µL NaCl 0.9% (w/v). Samples were mixed for 10 min and the organic phase was
471
separated from the aqueous phase by centrifugation at 3000g for 2 min. The hexane phase was
472
recovered and 1 µl was injected in the GC-MS/FID.
473
474
Solid Phase MicroExtraction
475
One µg of 16:0-alkane internal standard (0.1 mg ml-1 in chloroform) was added to a 250 mL
476
flask and the flask was left at room temperature under a hood for 2 min to evaporate the
477
chloroform. Microalgal cell culture (200 mL at 15 millions cells per ml) or 200 mL of culture
478
medium was then added to the flask. A solid phase microextraction (SPME) fiber (DVB-
479
PDMS fused silica, 65 µm double-polar, Supelco) was mounted on a holder, and inserted and
480
incubated for 16 hours at room temperature in the headspace of the sealed flask. After
481
incubation, the fiber was immediately inserted into the injector of the GC-MS and analytes
482
were desorbed at 250°C (first two minutes of the GC program). GC-MS analysis was then
483
carried out as described below.
484
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485
Determination of double bond position in alkenes
486
After saponification of cell pellets, MTBE extracts containing alkenes were evaporated in a
487
vial under a gentle stream of nitrogen gas. One hundred µL of dimethyl disulfide (DMDS)
488
containing 1.3 mg of iodine was then added, the vial was sealed and the mixture was
489
incubated for 30 min at 35°C. After cooling down the mixture, 0.9 mL of a mixture
490
diethylether:hexane (1v/9v) was added. The whole sample was then loaded onto a mini-
491
column containing silica (Shibahara et al., 2008). The column was washed 5 times with 1 mL
492
of the diethylether:hexane mixture. The entire eluent (5 mL) was evaporated under a gentle
493
stream of nitrogen gas. The residue was re-suspended in 500 µL of hexane and 1 µl was
494
injected in the GC-MS.
495
GC-MS analyses
496
Analyses by gas chromatography coupled to mass spectrometry (GC-MS), which were
497
perfomed after experiments of DMDS adducts, saponification and solid phase microextraction
498
(SPME), were carried out using the following set up. A Thermo-Fischer gas chromatographer
499
Focus series coupled to a Thermo-Fischer DSQII mass spectrometer (simple quadrupole) was
500
used with a DB-5HT (Agilent) apolar capillary column (length 30 m, internal diameter 0.25
501
mm, film thickness 0.1µm). Helium carrier gas was at 1 mL min-1. Oven temperature was
502
programmed with an initial 2 minute hold time at 50°C, then a ramp from 50°C to 300°C at
503
10°C min-1, and a final 3 minute hold time at 300°C. Samples were injected in splitless mode
504
(2 min) at 250°C. The MS was run in full scan over 40-500 amu (electron impact ionization,
505
70eV) and peaks were quantified based on total ion current using the internal standards.
506
507
GC-MS/FID analyses
508
Analyses by gas chromatography coupled to mass spectrometry and flame ionization
509
detection (GC-MS/FID) were performed only after transmethylation reactions in order to
510
quantify fatty acids and hydrocarbons. Analyses were carried out on an Agilent 7890A gas
511
chromatographer coupled to an Agilent 5975C mass spectrometer (simple quadrupole). A
512
Zebron 7HG-G007-11 (Phenomenex) polar capillary column (length 30 m, internal diameter
513
0.25 mm, film thickness 0.25 µm) was used. Hydrogen carrier gas was at 1 mL min-1. Oven
514
temperature was programmed with an initial 2 minute hold time at 60°C, a first ramp from
515
60°C to 151°C at 20°C.min-1 with a 5 minute hold time at 151°C, then a second ramp from
516
151°C to 240°C at 6°C.min-1 and a final 3 minute hold time at 240°C. Samples were injected
517
in splitless mode (1 min) at 250°C. The MS was run in full scan over 40-350 amu (electron
15
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
518
impact ionization at 70eV) and peaks were quantified based on the FID signal using the
519
internal standards.
520
521
Chloroplast isolation
522
The Chlamydomonas reinhardtii cell-wall-deficient strain CW15 was used to isolate
523
chloroplasts. Culture conditions and procedure for chloroplast purification were identical to
524
those previously published (Mason et al., 2006). Chlorophyll of whole cells and purified
525
chloroplasts was extracted with 80% (v/v) acetone and quantified by spectrophotometry.
526
Alkenes were quantified by transmethylation of whole cells or chloroplasts.
527
528
ACKNOWLEDGEMENTS
529
We thank Patrick Carrier for help with GC maintenance and Drs. Xenie Johnson, Jean Alric,
530
Claire Sahut and Frédéric Carrière for helpful discussions. Technical help of Jérémy
531
Monlyade and gift of the Chlorella variabilis NC64A strain by the JL Van Etten laboratory
532
are also acknowledged.
533
534
535
TABLES
Chlamydomonas
reinhardtii
Alka(e)nes
detected
7-heptadecene
Nannochloropsis
sp.
pentadecane,
heptadecane,
7-heptadecene
Chlorella
variabilis
NC64A
pentadecane,
heptadecane,
7-heptadecene
8-heptadecene
Ostreococcus
tauri
Phaeodactylum
tricornutum
21:6-alkene
21:6-alkene
one hit
one hit
(to AtCER1/3,
(to AtCER1/3,
A0A090MA0,
B7G477,
none
none
none
score 478,
score 102,
e value 3e-163,
e value 2e-22,
40% identity)
27% identity)
Table 1. Summary of hydrocarbons detected in the species investigated and potential orthologs to proteins known to
be involved in hydrocarbon synthesis. Potential orthologs were identified by BLASTP reciprocical best hits in
complete genomes. The five microalgal target genomes were first searched with BLASTP 2.3.1 (NCBI website,
default parameters) using as queries the following proteins (Uniprot accession number between brackets): AtCER1
(F4HVY0), AtCER3 (Q8H1Z0), CYP152/OleT (E9NSU2), CYP4G1 (Q9V3S0), UndA (Q4K8M0), AAR(Q54765)
and ADO (Q54764). Possible orthologs were only found for AtCER1 and AtCER3 (in Ostreococcus and
Phaeodactylum). These single hits were then used as a query to search with BLASTP the genome of Arabidopsis
thaliana.
Potential
orthologs to
hydrocarbon
synthesis
genes
536
537
538
539
540
541
542
543
16
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
544
545
546
547
548
Supplemental Data
549
550
SupplementalFig.S1.HeptadecaneandpentadecanearedetectedinChlorellacultures.
551
SupplementalFig.S2.Detectionofhydrocarbonsintheheadspaceofliquidcultures.
552
Supplemental Fig. S3. Heptadecenedoes not result from artefactual decarboxylation of oleic acid
553
during transmethylationof cells.
554
SupplementalFig.S4. Deuterium-labeled(D31)palmiticacidismetabolizedbyChlorellacells.
555
SupplementalFig.S5. Heptadecenecontentinoppositematingtypesofvegetativecellsandgametes.
556
SupplementalFig.S6. Chlamydomonasheptadecenecontentvarieswithgrowthtemperature.
557
SupplementalFig.S7. Alka(e)nesynthesisisstronglydependentonlightinChlorellavariabilisNC64A.
558
SupplementalFig.S8.Biomassandheptadeceneproductivitiesincreasewithlightintensity.
559
SupplementalFig.S9.MassspectrumoftheC21:6hexaenefoundinPhaeodactylum.
560
SupplementalFig.S10.VariationofChlamydomonascellconcentrationdurinday/nightcycle.
561
SupplementalFig.S11.Heptadeceneisomersthatwouldresultfromformaldecarboxylationofoleican
562
dcis-vaccenicacids.
563
564
FIGURE LEGENDS
565
Figure 1. Long chain alka(e)nes are synthesized by cells of Chlamydomonas reinhardtii and
566
Chlorella variabilis NC64A. A, Chromatogram of the unsaponifiable cell material analyzed
567
by GC-MS. The long-chain hydrocarbon region is magnified (inset). Legend: 17:1-alk, n-
568
heptadecenes; 17:0-alk, n-heptadecane; 15:0-alk, n-pentadecane. B, Mass spectrum of the
569
Chlamydomonas peak at 14.3 min in A. This spectrum is identical to that of a commercial
570
standard of 1-heptadecene. C, Quantification of alka(e)nes detected in Chlamydomonas and
17
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
571
Chlorella after transmethylation of cells and GC-MS/FID analysis. Values are mean of three
572
biological replicates and error bars represent standard deviations; nd, not detected.
573
574
Figure 2. The heptadecene isomers are 7-heptadecene and 8-heptadecene. Cells were
575
saponified and a solvent extract was reacted with DMDS before analysis by GC-MS. The
576
major diagnostic ions expected for DMDS derivatives of 7-heptadecene and 8-heptadecene
577
and the mass spectra obtained for the two Chlorella n-heptadecene isomers modified by
578
dimethyldisulfide (DMDS) are shown.
579
580
Figure 3. Chlamydomonas and Chlorella hydrocarbons derive from fatty acids. Cells were
581
cultivated for five days in presence of deuterium-labeled (D31) palmitic acid, collected and
582
directly transmethylated. The transmethylation extract was analyzed by GC-MS/FID. A, Part
583
of the GC-MS chromatogram showing endogenous and labeled Chlamydomonas fatty acids as
584
methyl esters (FAMEs). B,D, Part of the chromatograms showing Chlamydomonas (B) and
585
Chlorella (D) hydrocarbons. C,E, Mass spectra of the Chlamydomonas peak at RT=9.2
586
minutes (C) and the Chlorella peak at RT=8.9 minutes (E). An interpretation of the fragments
587
observed is shown in red and is consistent with the compound being D29 n-heptadecene and
588
D31 n-heptadecane respectively.
589
590
Figure 4. Heptadecene content varies with growth and culture conditions in Chlamydomonas.
591
After cell transmethylation products were analyzed by GC-MS/FID. A and B, Heptadecene
592
content (A) and growth curve (B) of cells grown for nine days in flasks under mixotrophic
593
(TAP medium) or photoautotrophic (minimal medium) conditions. C and D, cells were grown
594
for 2 days in TAP medium (C) or minimal medium (D) and changed to the same culture
595
medium devoid of nitrogen (TAP-N, MM-N). Values are means of four biological replicates
596
and error bars represent standard deviations. Stars denote significant differences at p<0.05 (*),
597
p<0.01 (**) and p<0.001 (***) in a 2-sided t-test.
598
599
Figure 5. Chlamydomonas heptadecene synthesis is strictly dependent on light and increases
600
with light intensity. A, Variation of heptadecene during day-night cycles. Cells were grown in
18
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
601
flasks under a day-night cycle (12 h light/12 h dark) for three days, then diluted at the
602
beginning of the day and samples were collected during the next 24 hours. Values are means
603
of three biological replicates and error bars represent standard deviations. B, Heptadecene
604
synthesis in dark- and light-grown cultures. Cells were grown in flasks in TAP medium at 120
605
µmol photons.m .s and then diluted in the same medium to be grown for an extra five days
606
either in the same condition or in the dark (i.e. under heterotrophic conditions). Values are
607
means of three biological replicates and error bars represent standard deviations; nd, not
608
detected. C, Quantification of heptadecene in isolated chloroplasts. Whole cells or isolated
609
intact chloroplasts were transmethylated and heptadecene was quantified using GC-MS/FID.
610
The proportion of cell heptadecene present in the chloroplast was calculated using the
611
chlorophyll content of isolated chloroplasts and assuming all chlorophyll was present in
612
chloroplasts. Values are means of three biological replicates and error bars represent standard
613
deviations. D, Kinetics of heptadecene synthesis upon light exposure. Cells were grown for 5
614
days in the dark in TAP medium and heptadecene was quantified at various times after
615
exposure to light (120 µE) in presence or absence of the photosynthesis inhibitor 3-(3,4-
616
dichlorophenyl)-1,1-dimethylurea (DCMU). Values are means of three biological replicates
617
and error bars represent standard deviations; nd, not detected. Using 2-sided t-tests, no
618
significant differences were found at p<0.01 with and without DCMU. E, Influence of light
619
intensity on heptadecene content per cell volume. Cells were grown in minimal medium at
620
different light intensities in a photobioreactor operated at constant cell density. Cells cultures
621
were stabilized for three days at a constant light intensity (starting at 125 µmol photons.m .s
622
(µE)) before switching to the next higher intensity (up to 300 µE). Cells aliquots were
623
harvested from stabilized cultures at each intensity to measure heptadecene content. Values
624
are means of three biological replicates and error bars represent standard deviations.
-2 -1
-2 -1
625
626
Figure 6. Long-chain hydrocarbons are synthesized by Nannochloropsis sp. Strains were
627
cultivated for five days and hydrocarbons were analyzed by transmethylation of whole cells
628
and GC-MS/FID. Values are means of three biological replicates and error bars represent
629
standard deviations.
630
631
Figure 7. Possible pathways for 7-heptadecene synthesis in Chlamydomonas. Two major
632
possibilities are presented based on findings of this work and reactions known in other
19
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
633
organisms. The specificity for cis-vaccenic acid would be brought by a phospholipase A1 or
634
another type of lipase depending on the lipid used as substrate. Left side: cyanobacterial- or
635
plant-type pathway with acyl-ACP reductase and formal decarbonylation of aldehyde
636
intermediate. Chloroplast localization would involve acyl-acyl carrier protein (acyl-ACP)
637
substrate while other subcellular localizations would involve acyl-Coenzyme A (acyl-CoA)
638
substrate. Right side: bacterium-type pathway with decarboxylation of a free fatty acid (but
639
without formation of terminal double bond). Decarbonylase and decarboxylase are indicated
640
with quotes because decarbonylation and decarboxylation might be only formal and not actual
641
reactions. Regulation by light may occur at the step of free fatty acid generation and/or at its
642
conversion to heptadecene.
643
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
Fig. 1
1E+06
A
17:1 alk
Signal intensity (TIC)
6E+07
15:0 alk
5E+05
17:0 alk
4E+07
0E+00
12
13
14
15
2E+07
0E+00
7
17
27
Chlorella extract
Chlamydomonas extract
Fresh culture medium
Retention time (min)
B
C
55
100%
700
Relative abundance
80%
RT= 14.3 min
83
97
60%
40%
111
238
20%
Hydrocarbons
(ng mm-3 cell volume)
600
69
Pentadecane
Heptadecane
Heptadecene
500
400
300
200
100
125
nd
0
0%
40
80
120 160 200 240 280 320
Chlorella variabilis
NC64A
nd
Chlamydomonas
reinhardtii
m/z
Figure 1. Long chain alka(e)nes are synthesized by cells of Chlamydomonas reinhardtii and
Chlorella variabilis NC64A. A, Chromatogram of the unsaponifiable cell material analyzed
by GC-MS. The long-chain hydrocarbon region is magnified (inset). Legend: 17:1-alk, nheptadecenes; 17:0-alk, n-heptadecane; 15:0-alk, n-pentadecane. B, Mass spectrum of the
Chlamydomonas peak at 14.3 min in A. This spectrum is identical to that of a commercial
standard of 1-heptadecene. C, Quantification of alka(e)nes detected in Chlamydomonas and
Chlorella after transmethylation of cells and GC-MS/FID analysis. Values are mean of three
biological replicates and error bars represent standard deviations; nd, not detected.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
Fig. 2
7-heptadecene DMDS derivative
7
187
8
145
187
Relative abundance
100%
major isomer
145
80%
60%
40%
.
332 (M+ )
20%
0%
40
80
120 160 200 240 280 320 360
m/z
8-heptadecene DMDS derivative
159
9
8
173
173
100%
Relative abundance
minor isomer
159
80%
60%
40%
.
332 (M+ )
20%
0%
40
80
120 160 200 240 280 320 360
m/z
Figure 2. The heptadecene isomers are 7-heptadecene and 8-heptadecene. Cells were saponified
and a solvent extract was reacted with DMDS before analysis by GC-MS. The major diagnostic ions
expected for DMDS derivatives of 7-heptadecene and 8-heptadecene and the mass spectra obtained
for the two Chlorella n-heptadecene isomers modified by dimethyldisulfide (DMDS) are shown.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2016 American Society of Plant Biologists. All rights reserved.
A
Fig. 3
Signal intensity (TIC)
6E+06
16:0 FAME
5E+06
16:4 FAME
D31-16:0 FAME
4E+06
3E+06
D29-18:1 FAME
2E+06
18:0 FAME
D31-18:0 FAME
18:1 FAME
1E+06
0E+00
20
21
22
23
24
25
26
Retention time (min)
B
C
C3D5
Relative abundance
Signal Intensity (TIC)
3E+05
17:1 alk
2E+05
1E+05
D29-17:1 alk
0E+00
8.5
9
9.5
100%
46
80%
62
C4D7
RT=9.2 min
C5D9
78 94
60%
C6D11
C7D13
110
40%
C17D29H5 (M+.)
20%
267
0%
40
10
80
120 160 200 240 280 320
m/z
Retention time (min)
D
C3D7
C4D9
3E+05
66
100%
Relative abundance
Signal intensity (TIC)
E
2E+05
D29-17:1 alk
17:1-alk
1E+05
D31-17:0 alk
17:0-alk
0E+00
8.5
9
9.5
Retention time (min)
10
50
80%
C5D11
82
60%
C6D13
RT= 8.9 min
98
40%
C17D31H5 (M+.)
20%
271
0%
40
80
120 160 200 240 280 320
m/z
Figure 3. Chlamydomonas and Chlorella hydrocarbons derive from fatty acids. Cells were cultivated for
five days in presence of deuterium-labeled (D31) palmitic acid, collected and directly transmethylated. The
transmethylation extract was analyzed by GC-MS/FID. A, Part of the GC-MS chromatogram showing
endogenous and labeled Chlamydomonas fatty acids as methyl esters (FAMEs). B,D, Part of the
chromatograms showing Chlamydomonas (B) and Chlorella (D) hydrocarbons. C,E, Mass spectra of the
Chlamydomonas peak at RT=9.2 minutes (C) and the Chlorella peak at RT=8.9 minutes (E). An
interpretation of the fragments observed is shown in red and is consistent with the compound being D29 nDownloaded from on June 17, 2017 - Published by www.plantphysiol.org
heptadecene and D31 n-heptadecane
respectively.
Copyright
© 2016 American Society of Plant Biologists. All rights reserved.
Fig. 4
A
Heptadecene
(ng mm-3 cell volume)
Cell concentration
(million cells mL-1)
300
200
100
Mixotrophy (TAP)
50
Photoautotrophy (MM)
40
30
20
10
0
0
24
48
72 96
Time (h)
500
TAP
400
TAP-N
D 500
*
***
300
0
120 144 168 192
***
200
100
Heptadecene
(ng mm-3 cell volume)
Heptadecene
(ng mm-3 cell volume)
400
0
C
B
Mixotrophy (TAP)
Photoautotrophy (MM)
500
0
400
24
48
72 96 120 144 168 192
Time (h)
MM
MM-N
300
200
100
**
**
0
24
48
Time (h)
96
24
48
Time (h)
96
Figure 4. Heptadecene content varies with growth and culture conditions in Chlamydomonas.
After cell transmethylation products were analyzed by GC-MS/FID. A and B, Heptadecene
content (A) and growth curve (B) of cells grown for nine days in flasks under mixotrophic
(TAP medium) or photoautotrophic (Minimal Medium) conditions. C and D, cells were grown
for 2 days in TAP medium (C) or Minimal Medium (D) and changed to the same culture
medium devoid of nitrogen (TAP-N, MM-N). Values are means of four biological replicates
and error bars represent standard deviations. Stars denote significant differences at p<0.05 (*),
p<0.01 (**) and p<0.001 (***) in a 2-sided t-test.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
Fig. 5
A
B
25
Day
120
Heptadecene
(ng mm-3 cell volume)
Heptadecene
(ng million cell-1)
20
140
Night
15
10
5
100
80
60
40
20
nd
0
0
0
3
6
9
12
15
18
21
0
24
Light intensity (µmol photons m-2 s-1)
Time (h)
C
D
18
25
-DCMU
16
Heptadecene
(ng mm-3 cell volume)
Heptadecene
(ng million cell-1)
120
14
12
10
8
6
4
20
15
10
5
2
0
0
Whole cell
+DCMU
nd
0 min 10
min
Chloroplast
E
30
min
1h
Time after dark
250
Heptadecene
(ng mm-3 cell volume)
3h
200
150
100
50
0
0
50
Light
100
150
200
250
300
350
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
-2 Plant
Copyright(µmol
© 2016 American
Society
intensity
photons
mof
s-1) Biologists. All rights reserved.
6h
Figure 5. Chlamydomonas heptadecene synthesis is strictly dependent on
light and increases with light intensity. A, Variation of heptadecene during
day-night cycles. Cells were grown in flasks under a day-night cycle (12 h
light/12 h dark) for three days, then diluted at the beginning of the day and
samples were collected during the next 24 hours. Values are means of three
biological replicates and error bars represent standard deviations. B,
Heptadecene synthesis in dark- and light-grown cultures. Cells were grown
in flasks in TAP medium at 120 µmol photons.m-2.s-1 and then diluted in the
same medium to be grown for an extra five days either in the same
condition or in the dark (i.e. under heterotrophic conditions). Values are
means of three biological replicates and error bars represent standard
deviations; nd, not detected. C, Quantification of heptadecene in isolated
chloroplasts.
Whole
cells
or
isolated
intact
chloroplasts
were
transmethylated and heptadecene was quantified using GC-MS/FID. The
proportion of cell heptadecene present in the chloroplast was calculated
using the chlorophyll content of isolated chloroplasts and assuming all
chlorophyll was present in chloroplasts. Values are means of three
biological replicates and error bars represent standard deviations. D,
Kinetics of heptadecene synthesis upon light exposure. Cells were grown
for 5 days in the dark in TAP medium and heptadecene was quantified at
various times after exposure to light (120 µE) in presence or absence of the
photosynthesis inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU).
Values are means of three biological replicates and error bars represent
standard deviations; nd, not detected. Using 2-sided t-tests, no significant
differences were found at p<0.01 with and without DCMU. E, Influence of
light intensity on heptadecene content per cell volume. Cells were grown in
minimal medium at different light intensities in a photobioreactor operated
at constant cell density. Cells cultures were stabilized for three days at a
constant light intensity (starting at 125 µmol photons.m-2.s-1 (µE)) before
switching to the next higher intensity (up to 300 µE). Cells aliquots were
harvested from stabilized cultures at each intensity to measure heptadecene
content. Values are means of three biological replicates and error bars
represent standard deviations.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
Fig. 6
300
Pentadecane
Heptadecene
Heptadecane
Hydrocarbons
(ng mm-3 cell volume)
250
200
150
100
50
0
N. limnetica
N. oculata
N. gaditana
N. granulata
N. oceanica
N. salina
Nannochloropsis
Figure 6. Long-chain hydrocarbons are synthesized by Nannochloropsis sp. Strains were
cultivated for five days and hydrocarbons were analyzed by transmethylation of whole cells and
GC-MS/FID. Values are means of three biological replicates and error bars represent standard
deviations.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
Fig. 7
cis-vaccenic acid-containing lipid
Light
(Phosphatidylinositol or other lipid)
?
+
Phospholipase A1
or other lipase
11
?
+
12
cis-vaccenic acid (C18:1cisΔ11)
Acyl-activating
enzyme
Fatty acid
“decarboxylase”
cis-vaccenoyl-ACP (CoA)
Acyl-ACP (CoA)
reductase
11-heptadecenal
Aldehyde
“decarbonylase”
8
7
7-heptadecene
Figure 7. Possible pathways for 7-heptadecene synthesis in Chlamydomonas. Two major
possibilities are presented based on findings of this work and reactions known in other
organisms. The specificity for cis-vaccenic acid would be brought by a phospholipase A1 or
another type of lipase depending on the lipid used as substrate. Left side: cyanobacterial- or
plant-type pathway with acyl-ACP reductase and formal decarbonylation of aldehyde
intermediate. Chloroplast localization would involve acyl-acyl carrier protein (acyl-ACP)
substrate while other subcellular localizations would involve acyl-Coenzyme A (acyl-CoA)
substrate. Right side: bacterium-type pathway with decarboxylation of a free fatty acid (but
without formation of terminal double bond). Decarbonylase and decarboxylase are indicated
with quotes because decarbonylation and decarboxylation might be only formal and not actual
reactions. Regulation by light may occur at the step of free fatty acid generation and/or at its
conversion to heptadecene. Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2016 American Society of Plant Biologists. All rights reserved.
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