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Origin and taxonomic diversity of phytoplankton
Who are the phytoplankton organisms? Are they part of the same phylogenetic family as terrestrial plants?
Figure 1-4 presents a phylogenetic tree of eukaryotes based on the analysis of the gene encoding the RNA 18s cells. Photosynthetic organisms are indicated on the colored branches. On this tree, we see that terrestrial plants belong to a single monophyletic family belonging to Chlorophyta. Green algae are close relatives of terrestrial plants and also belong to the Chlorophyta clade. On the other hand, marine photosynthetic eukaryotes are distributed in many branches of the phylogenetic tree of eukaryotes such as Glaucophyta and Rhodophyta, which group together with Chlorophyta in the Archaeplastida (Figure 1-4). We also find marine phototrophs within Alveolata, among which we find dinoflagellates (Dinophyceae), Stramenopila which comprises diatoms, Haptophyta which includes coccolithophores, but also Rhizaria, Discoba or Cryptista (Obornik, 2019).
The ability to perform oxygen photosynthesis is also found in some species of bacteria (not shown on this tree), called cyanobacteria. A question that arises with regard to this tree is: how to explain the presence of photosynthetic organisms within such diverse phylogenetic groups? Has the ability to achieve photosynthesis appeared several times during evolution by evolutionary convergence?
Endosymbiotic events are shown in the hypothetical tree : P-primary endosymbiosis ; C-complex endosymbiosis; S-secondary endosymbiosis. Losses of photosynthetic ability (white rectangles) or losses of the entire plastid (black rectangles) are indicated. SAR: Stramenopila + Alveolata + Rhizaria. From Oborník, 2019 Several arguments led to propose a chimeric origin of photosynthetic eukaryotes. In 1970, based on morphological arguments (the resemblance between chloroplasts and cyanobacteria), Margulis proposed that the chloroplasts of eukaryotic cells come from the incorporation of a cyanobacterium by a heterotrophic unicellular (Margulis, 1970): this is the endosymbiotic theory. Since then, many arguments have accumulated to support this theory, one of the most robust of which is phylogenetic: chloroplast DNA analyses show the close kinship between all chloroplast and cyanobacteria (Rodríguez-Ezpeleta, 2005). Nowaday, it is well admitted that the photosynthetic organelle of algae and plants (the plastid) traces its origin to a primary endosymbiotic event in which a previously non-photosynthetic protist engulfed and enslaved a cyanobacterium (Figure 1-5). This eukaryote then gave rise to the red, green and glaucophyte algae (Bhattacharya, Yoon and Hackett, 2004). However, many algal lineages, such as the chlorophyll c-containing stramenopiles, have a more complicated evolutionary history involving a secondary endosymbiotic event, in which a protist engulfed an existing eukaryotic alga, in this case, a red alga; the additional engulfment of a green alga was suggested (Bhattacharya, Yoon and Hackett, 2004; Prihoda et al. 2012, Moustafa et al., 2009). Another algal group, the dinoflagellates, has undergone tertiary (engulfment of a secondary plastid) and even quaternary endosymbiosis (Figure 1-5).
Gene transfer from the endosymbiont to the host nucleus is shown with the arrows and the colored chromosomes. The mitochondrion has been omitted from these figures. The remnant algal nucleus (nucleomorph) in secondary and tertiary endosymbioses is shown. This genome has been lost in all algae but the cryptophytes (member of the chromalveolates) and chlorarachniophytes (marked with asterisks). From Bhattacharya, Yoon and Hackett, 2004.
Photosynthetic eukaryotes are chimeras. The study of their nuclear genome shows a great phylogenetic diversity whereas the study of the plastid genome shows a close relation of kinship. A question that arises is what about functional diversity? Do photosynthetic eukaryotes have a significant diversity linked to the origin of the host cell that integrated the organelles? Or do they have common mechanisms for using light energy inherited from the common cyanobacterium ancestor?
Different types of photosynthesis
Among the three fundamental domains of living organisms, bacteria, archaea and eukaryotes (Woese et al., 1990), we find photosynthetic organisms within bacteria and eukaryotes only. It is to note that another mechanism of conversion of light energy into chemical energy exists which is fundamentally different from the chlorophyll-based photosynthesis. Halophilic archaea possess bacteriorhodopsins, retinal-binding integral membrane proteins, which can use light to pump protons, generate an electrochemical proton gradient and produce adenosine triphosphate (ATP) from adenosine diphosphate (ADP) through this process (Hayashi, Tajkhorshid and Schulten, 2003). Later, metagenomics approaches revealed widespread retinal-binding rhodopsin proteins, called proteorhodopsins in marine bacteria (Beja et al, 2000). It also exists in eukaryotes (Bieszke et al., 1999). These discoveries have challenged the notion that only chlorophyll-based photosynthesis allows the conversion of solar energy into the marine ecosystem (Beja et al, 2000). The importance of proteorhodopsins in the global conversion of light into chemical energy and biomass is still a matter of debate (Fuhrman et al. 2008). Here, we will not consider the light-driven ATP synthesis catalyzed by bacteriorhodopsins and proteorhodopsins, and the term “photosynthesis” will be used in its historical meaning, i.e. for the conversion of light into chemical energy based on chlorophylls or bacteriochlorophylls.
The use of chlorophyll or bacteriochlorophyll also distinguishes two different types of photosynthesis, whose general principles are the same but which depend on different sources of electrons. Oxygenic photosynthesis uses water as a source of electrons, a reaction that results in the release of molecular oxygen, and those electrons can be transferred through two photosystems to reduce carbon dioxide. But some photosynthetic bacteria can use light to extract electrons from other molecules than water. They use instead inorganic compounds, like H2 or H2S, or inorganic compounds like malate. Because they do not evolve O2, their photosynthesis is named anoxygenic photosynthesis. Another difference with oxygenic phototrophs is that they possess only one type of reaction center: a pheophytin-quinone (type II) reaction center, or an iron-sulfur (type I) one. Among anoxygenic phototrophs using type-II reaction centers are purple bacteria and green filamentous bacteria. On the contrary, green sulfur bacteria and heliobacteria use type-I reaction center (Blankenship, 1992).
In this work, we will only consider oxygenic photosynthesis, which exists in cyanobacteria and in photosynthetic eukaryotes. A major difference between these two groups of organisms is structural: in photosynthetic eukaryotes, photosynthesis occurs in specialized organelles, the plastids, while cyanobacteria do not possess intracellular organelles. In the latter, photosynthesis occurs in membrane invaginations. The following paragraph therefore only concerns eukaryotes.
The chloroplast: an organelle hosting photosynthesis
The site of photosynthesis in eukaryotes is the plastid, a membranous subcellular organelle ( Figure 1-6). Chloroplast numbers per cell vary considerably in plants, from 1 to 1000 depending on species, life stage and environment. Microalgae generally possess one or a few chloroplasts delimited by 2, 3 or 4 membranes (Staehelin, 1986; Falkowski and Raven, 2013). The inner membrane encloses the stroma, a concentrated solution of enzymes that also contains the DNA, RNA and ribosomes involved in the synthesis of several chloroplast proteins. Inside the chloroplast, membrane folds form bags called thylakoids which delimit an internal space, the lumen. In higher plants, thylakoids consist of stacks of disk-like sacs named grana which are interconnected by unstacked stroma lamellae whereas in green algae, the grana organization typical of plants is replaced by a mix of stacked and unstacked thylakoid membranes that still allow a lateral heterogeneity in protein content of these membranes (Staehelin, 1986, Figure 1-6). In secondary phototrophs (photosynthetic species deriving from the second endosymbiosis), the organelle where photosynthesis takes place is not properly speaking a chloroplast, since this term was historically used for chlorophyll a and b containing plastids.
Secondary endosymbionts contain chlorophyll a and c, and we use the generic term plastid (Gould, Waller and McFadden, 2008). Electron micrographs show no thylakoid subdomains in secondary plastids, only loose stacks of two, three or four thylakoids (most of the time 3, Pyszniak and Gibbs, 1992) which still lead to a marked difference in protein content between the outer membrane of a stack and its inner membranes.
The main role of chloroplasts/plastids is the photosynthetic metabolism, but it also hosts nitrate and sulfate reduction, as well as synthesis of chlorophylls, carotenoids, haems, amino acids and fatty acids (Jensen and Leister, 2014).
Oxygenic photosynthesis, a two-step reaction
In the first decade of the 20th century, it was generally assumed that light, absorbed by photosynthetic pigments, directly reduced CO2 which in turn combined with water to form carbohydrates. And under this hypothesis, CO2 was considered as the source of O2 generated by photosynthesis. But in 1931, based on work on purple sulfur bacteria, Cornelis van Niel proposed that photosynthesis is a two-stage process in which light energy is used to oxidize H2O into O2 (van Niel and Muller, 1931). The validity of van Niel’s hypothesis was established unequivocally by two experiments. In 1937, Robert Hill discovered that when isolated chloroplasts that lack CO2 are illuminated in the presence of an artificial electron acceptor, O2 evolved with concomitant reduction of the acceptor. This Hill reaction demonstrates that CO2 does not participate directly in the O2 producing reaction. In 1941, when the oxygen isotope 18O became available, Samuel Ruben and Martin Kamen directly demonstrated that the source of O2 in photosynthesis is H2O.
It is now well understood that photosynthesis is enabled by two steps: the production of reducing power in the form of reduced nicotinamide adenine dinucleotide phosphate (NADPH) and chemical energy in the form of ATP that occurs in the membrane photosynthetic chain thanks to light energy. Carbon fixation then uses products of the photosynthetic chain in the stromal compartment. The result of the « light » phase of photosynthesis is the conversion of light energy into chemical energy, stored in molecules with high energy potential, ATP and NADPH. In the « dark » phase of photosynthesis, ATP and NADPH are used together to catalyze all of the carbon assimilation reactions used to make carbohydrates, called the Benson-Calvin cycle. When the final carbon molecule is glucose, the entire process described so far leads to the following reaction: In the following two sections, I will describe first the Calvin Benson Bassham cycle where reduction of carbon dioxyde occurs (the biosynthetic, or “dark”, phase of photosynthesis), and then the photochemical phase which occurs in the thylakoid membranes (also called “light” phase of photosynthesis).
Discovery and chemical equation of the CBB cycle
The metabolic pathway by which most of the autotrophs incorporate CO2 into carbohydrates was elucidated between 1946 and 1953 by Melvin Calvin, James Bassham and Andrew Benson (Bassham, Benson and Calvin, 1953). The basic experimental strategy they used was to expose growing cultures of the green alga Chlorella to radioactive labelled 14CO2 for varying times and under different illumination conditions and then to drop the cells into boiling alcohol so as to disrupt them while preserving their labeling pattern. The radioactive products were then separated and identified by a two-dimensional paper chromatography coupled to autoradiography. The overall pathway is presented in Figure 1-7 and shows that the reduction of CO2 requires an energy supply in the form of adenosine triphosphate (ATP) and an input of reducing power provided by a coenzyme, the reduced form of nicotinamide adenine dinucleotide phosphate (NADPH). These two substrates are used with a ratio of 3 ATP for 2 NADPH. Although other carbon fixation pathways have been found in autotroph bacteria and archae, CBB cycle is the only carbon fixation pathway operating in eukaryotes (Hügler and Sievert, 2011).
Carbon concentration mechanisms (CCM) enable high rate of carbon fixation
The first step along the reductive pathway of CO2 is catalyzed by the Ribulose-1,5-bisphophaste Carboxylase Oxygenase, RuBisCO, a big protein complex. RuBisCO has a low affinity for CO2 and catalyzes also a reaction of ribulose-1,5-bisphosphate and molecular oxygen (O2) that is competing with carbon fixation. Carbon fixation rate is thus depending on the ratio of CO2 concentration over O2 concentration in the surrounding environment of RuBisCO (Delgado et al., 1995).
The way to deal with CO2 is obviously very different for terrestrial organisms (plants) and for aquatic organisms. CO2 can dissolve into water and its concentration in surface Ocean ranges from 10 to 30 µM (Reinfelder, 2011). Dissolved CO2 is not the only source of inorganic carbon in aquatic environments. CO2(d) can undergo nucleophilic displacement from a water molecule, leading to the formation of H2CO3 that is in chemical equilibrium with bicarbonate and carbonate ions: CO2(g) ⇌ CO2(d) + H2O ⇌ H2CO3 ⇌ HCO3– + H+ ⇌ CO32- + 2H+ The ionic forms do not contribute to the vapor pressure of the gaseous form, thus the concentration of the sum of all dissolved inorganic carbon can greatly exceed the atmosphere/water equilibrium concentration of gaseous CO2. In the surface ocean, total dissolved inorganic carbon is approximately 200 times higher than dissolved CO2 concentration (Falkowski and Raven, 2013). Nevertheless, transient patches of CO2-depleted surface seawater may persist for several days to weeks (Takahashi, 2009) because CO2(d) diffusion rate and bicarbonate dehydration to CO2 are low processes regarding potential consumption by photosynthetic reactions (Falkowski and Raven, 2013).
In all marine phytoplankton, the intrinsic Michaelis constant (Km) of the Rubisco is much higher than the apparent photosynthetic requirement for extracellular dissolved CO2 (K1/2) (Kaplan and Reinhold, 1999). This is attributed to carbon concentration mechanisms, CCM, that increase locally the CO2 concentration. The biophysics and biochemistry of CCMs vary within and among marine phytoplankton clades such as dinoflagellates, haptophytes and diatoms (Reinfelder, 2011). They may involve the activity of external or intracellular carbonic anhydrase and active HCO3 transport. CCM have thus a cost of energy and nutrient: for enzymes production and for active transport of bicarbonate. In general, coccolithophores have low-efficiency CCM, and diatoms and the haptophyte genus Phaeocystis have high-efficiency CCM while dinoflagellates appear to possess moderately efficient CCM (Takahashi, 2009).
The photosynthetic electron transfer chain uses light energy to supply the CBB cycle
Calvin Benson cycle reactions involve consumption of reducing power in the form of NADPH and chemical energy in the form of ATP high energy bond. This is supplied by the photosynthetic electron transfer chain whose mechanistic details have been resolved in the second part of the 20th century. Resolution of the pathway allowing electron transfer from water to NADP+ has first been investigated with chemical tools. Thus, the global pattern of the pathways has been resolved before understanding the molecules and protein complexes that were involved.
In 1953, the groups of Robert Hill working with isolated chloroplast and the one of Cornelius van Niel working with bacteria could show that light energy was used for endergonic electron transfer and not for photolysis of CO2 (van Niel et al., 1953). Arnon and collaborators then highlighted that isolated chloroplasts are capable of ATP synthesis (Arnon, Allen and Whatley, 1954). This is suggesting that light driven electron transfer is followed by a downhill exergonic reaction coupled to ATP synthesis. Cyclic electron transfer has first been proposed as the pathway of photophosphorylation. But in 1958, Arnon’s group could show that ATP synthesis was coupled to a linear electron pathway inducing a donor oxidation and a final acceptor reduction (Arnon, Whatley and Allen, 1957). This raised a thermodynamic problem: it was difficult to imagine a downhill electron transport that could be coupled to the endergonic ATP synthesis.
Emerson, Chalmers and Cederstrand, revealed the existence of two site of photochemistry, based on the spectral dependence of the photosynthetic process (Emerson, Chalmers and Cederstrand, 1957). Those two sites were called photosystem I and photosystem II. Later, two cytochromes were discovered in the chloroplast, named cytochrome b6 and cytochrome f, and Hill and Bendall proposed that those cytochromes allow the electron transfer between the two photosystems and their operation in series (Hill and Bendall, 1960). In agreement with the redox potentials of the two cytochromes and the two reaction centers, the general Z scheme or diagram of the photosynthetic electron transfer, was elucidated (Hill and Bendall, 1960). In addition to the electron transfer chain between PSII.
Describing the Z-scheme of photosynthesis requires the use of oxidoreduction principles. « Oxidation » and « reduction » reactions, refer to the loss or gain of one or more electrons, respectively. The redox potential is a scale (in volts) that has been developed to predict the tendency of a redox reaction. The molecules with the lowest redox potential will tend to be oxidized by molecules of higher redox potential.
When these considerations are applied to the photosynthetic electron transfer chain, it is clear that linear electron transfer from water to NADP+ is unfavorable. Two photochemical steps that occur in photosystem II (PSII) and photosystem I (PSI) are crucial: the light catalyzed electron transfer from the high redox potential special pairs P680 and P700 towards the low redox potential acceptors, QA and iron-sulfur centers (FX, FA and FB), respectively (Nelson and Yocum, 2006). Those are highly endergonic (i.e. thermodynamically unfavorable) reactions. Those two steps are put in series thanks to exergonic (i.e. thermodynamically favorable) electron transfer from PSII to PSI via the membrane plastoquinones (PQ), the cytochrome b6f complex (cyt. b6f) and the luminal plastocyanins (PC) that reduce the PSI primary donor, P700. At the donor side of PSII, Oxygen Evolving Complex (OEC), a metallic center (Mn)4Ca catalyzes transfer of electrons from water to a tyrosine residue, YZ that in turn reduces P680+. At the acceptor side of PSI, FB reduces ferredoxin (Fd) that in turn reduces NADP+.
Table of contents :
Chapter 1: General introduction
1. 1. Marine ecosystems, phytoplankton and photosynthesis
1. 1. 1. Most of the genetic diversity in the Ocean is known
1. 1. 2. Phytoplankton plays a crucial role in fluxes of matter and energy in the ocean
1. 1. 3. Origin and taxonomic diversity of phytoplankton
1. 2. Reactions and mechanisms involved in eukaryotic photosynthesis
1. 2. 1. Different types of photosynthesis
1. 2. 2. The chloroplast: an organelle hosting photosynthesis
1. 2. 3. The Calvin-Benson-Bassham cycle synthesizes organic carbon from CO2
1. 2. 4. The photosynthetic electron transfer chain uses light energy to supply the CBB cycle
1. 2. 5. From light absorption to energy storage
1. 2. 6. Monitoring photosynthesis with chlorophyll fluorescence
1. 3. Regulation of the photosynthetic process and acclimation to changing environments
1. 3. 1. Light stress and photo-inhibition
1. 3. 2. Protecting photosystem I and II from light stress
1. 3. 3. Photosynthetic alternative pathways
1. 3. 4. CEF, a crucial regulative pathway remaining mysterious
1. 4. Thesis outline
1. 5. Bibliography
2 Exploring the diversity of cyclic electron flow around photosystem I in microalgae species
Chapter 2: Probing PSI activity
2.1. How to probe photosystem I with absorption spectroscopy?
2.1.1. Some reminders about photosystem I
2.1.2. Measuring P700 redox state to investigate PSI activity
2.1.3. Electrochromic Shift: an internal voltmeter
2.2. Article: Critical reappraisal of methods to measure photosystem I activity
2.3. Discussion
2.3.1. The P700 pulse method underestimates Y(I) because of reduction of PSI acceptors during the multiple turnover pulse
2.3.2. Technical considerations regarding P700 measurements
2.3.3. Generalization to the case of an active photosystem II
2.3.4. Revisiting the literature based on the P700 pulse method
2.3.5. Partial conclusions and transition
2.4. Bibliography
Chapter 3: Diversity of cyclic electron flow in microalgae
3.1 Introduction: CEF, a still mysterious alternative pathway
3.1.1. Role of CEF in ATP/NADPH adjustment and photo-protection
3.1.2. Regulation of CEF rate
3.1.3. Measuring CEF is a methodological challenge
3.1.4. Chapter outlines
3.2 Material and methods
3.2.1 Strains, growth and sampling
3.2.2 Chemicals
3.2.3 In vivo spectroscopy
3.2.4 ECS spectra and linearity with electric field
3.2.5 Absorption cross section assessment
3.3 Results (I): Exploring CEF diversity
3.3.1 DCMU titration reveals CEF behavior
3.3.2 Which observables?
3.3.3 CEF is not essential to photosynthesis in the dinoflagellate Amphidinium carterae
3.3.4 CEF is independent on LEF photosynthesis in the dinoflagellate Symbiodinium sp.
3.3.5 Chlamydomonas reinhardtii displays a CEF which is dependent on LEF
3.3.6 Partial conclusion on CEF diversity
3.4 Results (II): Evaluating CEF and LEF absolute rates as a function of light irradiance in the green alga Chlamydomonas reinhardtii
3.4.1 ECS-based estimations of the PSI and PSII absorption cross sections
3.4.2 Validation of the method for measurement of absorption cross sections
3.4.3 Evaluating CEF absolute rates from ETR(total) and ETR(II)
3.4.4 Calculations of the ATP/NADPH ratio produced by the photosynthetic chain
3.5 Discussion and future perspectives
3.5.1 A simple and robust protocol to investigate CEF diversity highlights three different behaviors
3.5.2 Limitations of the DCMU titration method
3.5.3 Despite a complex relationship, CEF and LEF remain proportional at all light intensities in Chlamydomonas reinhardtii
3.6 Bibliography
Chapter 4: Photosynthetic physiology of the coccolithophore Emiliania huxleyi Major environmental determinants and signature of viral infection in the field
4.1 Introduction
4.2 Material and methods
4.2.1 Mesocosm setup, treatments and sampling
4.2.2 Flow cytometry/ qPCR
4.2.3 Measurement of environmental (abiotic) parameters
4.2.4 Photophysiology by Fast Induction and Relaxation fluorometry (FIRe)
4.2.5 Statistical analyses
4 Exploring the diversity of cyclic electron flow around photosystem I in microalgae species
4.3 Results
4.3.1 A two phases phytoplankton bloom occured
4.3.2 Photosynthetic physiology
4.3.3 Fluorescence signals were mainly due to Emiliania huxleyi from day 9 to 24th of the experiment
4.3.4 Evolution of environmental parameters during the experiment
4.3.5 Environmental determinants of E. huxleyi photosynthesis
4.3.6 A photosynthetic signature of viral infection?
4.4 Conclusion and discussion
4.5 Bibliography
Chapter 5: General discussion
5. 1. 1. The importance of cross validations of methods
5. 1. 2. Using the flash-induced ECS to estimate the photochemical rate: a good choice?
5. 1. 3. Studying CEF and its abiotic and biotic determinants in the field: an accessible project?
5. 2. Roles of CEF and mechanisms of the regulation of the CEF and LEF
5. 2. 1. ATP:NADPH ratio equilibration
5. 2. 2. A role of CEF when PSII is inhibited?
5. 2. 3. CEF and LEF regulation
5. 3. Three years were too short for…
5. 3. 1. Revisiting literature using P700 method
5. 3. 2. Exploring cyclic electron flow using our methods
5. 3. 3. Harness all data collected in the field
5. 4. Bibliography
