Abstract
Diatoms are one of the most ecologically successful classes of photosynthetic marine eukaryotes in the contemporary oceans. Over the past 30 million years, they have helped to moderate Earth’s climate by absorbing carbon dioxide from the atmosphere, sequestering it via the biological carbon pump and ultimately burying organic carbon in the lithosphere1. The proportion of planetary primary production by diatoms in the modern oceans is roughly equivalent to that of terrestrial rainforests2. In photosynthesis, the efficient conversion of carbon dioxide into organic matter requires a tight control of the ATP/NADPH ratio which, in other photosynthetic organisms, relies principally on a range of plastid-localized ATP generating processes3,4,5,6. Here we show that diatoms regulate ATP/NADPH through extensive energetic exchanges between plastids and mitochondria. This interaction comprises the re-routing of reducing power generated in the plastid towards mitochondria and the import of mitochondrial ATP into the plastid, and is mandatory for optimized carbon fixation and growth. We propose that the process may have contributed to the ecological success of diatoms in the ocean.
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Change history
19 August 2015
Affiliation number 4 was corrected.
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Acknowledgements
This work was supported by grants from Agence Nationale de la Recherche (ANR-12-BIME-0005, DiaDomOil to C.B., D.P. and G.F.; ANR-8NT09567009, Phytadapt to B.B., G.F. and C.B.; ANR-11-LABX- 0011-01, Dynamo to F.R. and P.J.; ANR-11-IDEX-0001-02, PSL Research University and ANR-10-LABX-54, MEMOLIFE to C.B.), the Région Rhône-Alpes (Cible project) to G.F., the Marie Curie Initial Training Network Accliphot (FP7-PEPOPLE-2012-ITN; 316427) to G.F., D.P., S.F. and V.V., an ERC Advanced Award (Diatomite) and the EU MicroB3 project to C.B., the CNRS Défi (ENRS 2013) to G.F. and L.T., and the CEA Bioénergies program to G.F and D.P. P.C., N.B. and B.B acknowledge financial support from the Belgian Fonds de la Recherche Scientifique F.R.S.-F.N.R.S. (F.R.F.C. 2.4597.11, CDR J.0032.15 and Incentive Grant for Scientific Research F.4520). B.B. also acknowledges a post-doctoral fellowship from Rutgers University and J.P. was funded from the COSI ITN project to C.B. Thanks are due to J.-L. Putaux and C. Lancelon-Pin for help with electron microscopy, to L. Moyet for technical support for the in vivo NMR analysis, to A. E. Allen for the AOX antibody, and to A. Falciatore and F. Barneche for critical reading the manuscript.
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B.B., L.T., C.B. and G.F. designed the study. B.B., N.B., O.M., D.P., J.P., A.T., V.V., R.B., S.F., D.F., A.K-L, F.R., P.J., L.T., P.C. and G.F. performed experiments. B.B., N.B., O.M., D.P., R.B., A.K.-L., S.S., F.R., P.J., L.T., P.F., P.C., C.B. and G.F. analysed the data. B.B., C.B. and G.F. wrote the manuscript, and all authors revised and approved it.
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Extended data figures and tables
Extended Data Figure 1 Deconvolution of the quadratic and linear ECS in P. tricornutum.
a, Absorption difference (ΔI/I) kinetics followed at different wavelengths in P. tricornutum, after a series of six saturating laser flashes, in anaerobic conditions. Solid lines correspond to the global fit of the experimental data with a sum of two exponential decays, with time constants τ and 2τ, respectively, as expected for linear and quadratic dependencies (see Methods). b, ΔI/I spectra are shown at different times during ECS relaxation. All spectra were normalized to 1 at 520 nm for better comparison. The observation that the blue and green parts of the spectrum are homothetic during relaxation, while changes are seen in the red most part of it, reflects the presence of the two ECS components, having different relaxation kinetics.
Extended Data Figure 2 Separation of c-type cytochrome signals from linear and quadratic ECS signals in P. tricornutum.
a, Kinetics of ΔI/I changes at 520, 554 and 566 nm during an ∼10 ms pulse of saturating red light (4,500 µmol quanta m−2 s−1) and the subsequent dark relaxation (top: control conditions; bottom: AA + SHAM). b, Kinetics of ecslin, ecsquad changes and c-type cytochrome redox state, from kinetics in a, after deconvolution as explained in Methods. c, d, Relationship between the quadratic and the linear ECS, before (ecslin, ecsquad, c) and after (ECSlin, ECSquad, d) correction for the dark electric field (see Methods). Dark yellow and magenta symbols correspond to control and AA + SHAM conditions, respectively. The green arrow indicates the value of ΔΨd in control conditions. Data are representative of five independent biological samples. The black boxes in a and b indicate periods of darkness.
Extended Data Figure 3 Cyclic electron flow and water-to-water cycles in P. tricornutum.
a, Representative traces of changes in ECSlin (normalized as explained in Methods, namely expressed in charge separation per PS) to evaluate linear and cyclic electron flow. Cells were illuminated with 1,870 µmol quanta m−2 s−1 of red light, in absence (filled circles) and presence (open circles) of DCMU and then transferred to the dark; r.u., relative units. b, Representative traces of the 16O2 and 18O2 concentrations at the offset of a 280 µmol quanta m−2 s−1 blue light; a.u., arbitrary units. In a and b, light and dark periods are represented by white and black boxes, respectively. c, Photochemical rate corresponding to total electron flow (TEF, dark symbols, data from four independent biological samples) and CEF (red symbols, n = 8 ± s.d.) at different irradiances. TEF and CEF were estimated from the initial slope of the ECS decay, as (SD − SL control) and (SD − SL DCMU), respectively (see Methods). d, Light-dependencies of oxygen uptake (U0, half-filled symbols) and gross photosynthesis (E0, open symbols) in control conditions (dark) and in the presence of DCMU (red). Data from two independent biological samples (squares and circles).
Extended Data Figure 4 ΔΨd and photosynthesis under respiratory inhibition in P. tricornutum.
a–c, Dependency of the ETRPS II (a), ΔΨd (b) and dark respiration (c) upon inhibition of the cyanide-sensitive respiratory pathway with different concentrations of antimycin A, in the presence of saturating SHAM (1 mM). Data from two independent biological samples. Experimental data were fitted with a mono-exponential decay function. d, e, ETRPS II (d), ΔΨd and dark respiration (e), expressed as percentage of control, in the presence of saturating AA (5 µM), SHAM (1 mM) or AA + SHAM (four independent experiments ± s.d.). f, Relationship between ΔΨd and mitochondrial respiration in samples treated with increasing concentrations of AA in the presence of 1 mM SHAM (mean value ± s.e.m. from b and c).
Extended Data Figure 5 Dark respiration, PMF and growth in AOX knockdown lines of P. tricornutum.
a, Respiratory activity of wild-type and AOX knockdown lines. Total respiration rate (red bars) and the contribution of the AOX capacity (white bars, see Methods) were normalized to wild-type values (n = 5 ± s.d.). b, ECS-based measurements of ΔΨd in wild-type (n = 3 ± s.d.) and AOX knockdown lines (n = 2 ± s.e.m.), in control conditions (green), in the presence of AA (dark green) and in the presence of AA + SHAM (white). c, Growth curves of wild-type and AOX knockdown lines in the presence/absence of AA (2 µM). Three independent growth curves are shown for each strain/condition. AA was added every day and cells were grown in continuous light to prevent them from dying in the dark because of lack of respiration.
Extended Data Figure 6 Subcellular localization of AOX in P. tricornutum and plastid–mitochondria interaction in P. tricornutum wild-type cells.
a, Subcellular localization of AOX. Cells were treated with an anti-AOX antibody and then with a secondary Alexa Fluor 488 antibody (see Methods). Positions of plastid and nuclei are indicated by chlorophyll a autofluorescence (red) and DAPI staining (blue), respectively. The pattern of AOX localization is highly similar to that observed with the mitochondria-specific mito-tracker probe (data not shown). Images are representative of 60 cells from three independent biological samples. b, Electron micrographs of the plastid–mitochondria juxtaposition in P. tricornutum. Arrows indicate possible physical contacts between the plastid and mitochondrial membranes. Image is representative of 51 images from seven independent biological samples.
Extended Data Figure 7 Cytochrome b6f turnover in P. tricornutum wild-type and AOX knockdown lines.
a, Schematic representation of the electron-flow reaction steps in the cytochrome b6f complex, which can be evaluated by spectroscopic measurements. b, Slow phase of ECSlin indicating cytochrome b6 activity (top) and time-resolved redox changes of cytochromes c6/f (bottom) in wild-type and AOX knockdown lines (kd-c5 and kd-c9). P. tricornutum cells were exposed to saturating single-turnover laser flashes given 10 s apart. Data were normalized to the amplitude of the fast phase of the ECSlin signal. Cytochrome c and ECSlin were deconvoluted as explained in Methods. Three independent biological samples are shown in red, blue and green colours. Cell concentration was 2 × 107 cells per millilitre. Note that both the slow phase of the ECSlin and the reduction of cytochromes c6/f were completely abolished by the plastoquinone competitive inhibitor DBMIB (10 µM; black arrow).
Extended Data Figure 8 In vivo changes in the NADPH redox state and ATP in wild-type and AOX knockdown lines.
a, Changes in NADPH at different light intensities. Light and dark periods are represented by white and black boxes, respectively. Light intensities were 50, 100, 200 and 400 µmol quanta m−2 s−1 (green, blue, red, and black traces, respectively). Chlorophyll concentration was ∼5 µg ml−1. b, Spectra from cells of wild-type (left) and AOX knockdown lines c5 (middle) and c9 (right) in the dark (red), light (green) and light + AA (blue) conditions are shown, with normalization to the internal standard (methylenediphosphonate; pH 8.9). The positions of the α-, β- and γ-phosphates of NTPs are shown. a, b, Representative of three independent biological samples.
Extended Data Figure 9 Cyclic electron flow in representative diatoms.
a, Total electron flow (filled symbols) was measured at different light irradiances, as illustrated in Extended Data Fig. 3, in T. weissflogii (black, n = 3 independent biological samples), T. pseudonana (blue, n = 2) and F. pinnata (red, n = 4). Cyclic electron flow capacity was also measured for every species (open circles, five independent biological samples ± s.d.). b, CEF capacity was plotted against TEF. CEF and TEF are presented as mean values ± s.d. from a. The green line corresponds to CEF = 5% of the maximal total electron flow.
Extended Data Figure 10 ΔΨd and photosynthesis under conditions of respiratory inhibition in representative diatoms.
Dark respiration (a–d), ΔΨd (e–h) and ETRPS II (i–l) in untreated cells and after treatment with inhibitors of respiration antimycin A, and/or SHAM at saturating concentrations. a, e, i, T. weissflogii (black). b, f, j, T. pseudonana (blue). c, g, k, F. pinnata (red). d, h, l, D. brightwellii (green). The data represent the mean value ± s.d. of three (l), four (d), five (f, h, k), six (a, c, e, j), seven (b, i) or eight (g) independent experiments. All data were normalized to the control value.
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Bailleul, B., Berne, N., Murik, O. et al. Energetic coupling between plastids and mitochondria drives CO2 assimilation in diatoms. Nature 524, 366–369 (2015). https://doi.org/10.1038/nature14599
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DOI: https://doi.org/10.1038/nature14599
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