Diatoms are unicellular algae responsible for approximately 20% of global carbon fixation. Their evolution by secondary endocytobiosis resulted in a complex cellular structure and metabolism compared to algae with primary plastids.
The whole genome sequence of the diatom Phaeodactylum tricornutum has recently been completed. We identified and annotated genes for enzymes involved in carbohydrate pathways based on extensive EST support and comparison to the whole genome sequence of a second diatom, Thalassiosira pseudonana. Protein localization to mitochondria was predicted based on identified similarities to mitochondrial localization motifs in other eukaryotes, whereas protein localization to plastids was based on the presence of signal peptide motifs in combination with plastid localization motifs previously shown to be required in diatoms. We identified genes potentially involved in a C4-like photosynthesis in P. tricornutum and, on the basis of sequence-based putative localization of relevant proteins, discuss possible differences in carbon concentrating mechanisms and CO2 fixation between the two diatoms. We also identified genes encoding enzymes involved in photorespiration with one interesting exception: glycerate kinase was not found in either P. tricornutum or T. pseudonana. Various Calvin cycle enzymes were found in up to five different isoforms, distributed between plastids, mitochondria and the cytosol. Diatoms store energy either as lipids or as chrysolaminaran (a β-1,3-glucan) outside of the plastids. We identified various β-glucanases and large membrane-bound glucan synthases. Interestingly most of the glucanases appear to contain C-terminal anchor domains that may attach the enzymes to membranes.
Here we present a detailed synthesis of carbohydrate metabolism in diatoms based on the genome sequences of Thalassiosira pseudonana and Phaeodactylum tricornutum. This model provides novel insights into acquisition of dissolved inorganic carbon and primary metabolic pathways of carbon in two different diatoms, which is of significance for an improved understanding of global carbon cycles.
Citation: Kroth PG, Chiovitti A, Gruber A, Martin-Jezequel V, Mock T, et al. (2008) A Model for Carbohydrate Metabolism in the Diatom Phaeodactylum tricornutum Deduced from Comparative Whole Genome Analysis. PLoS ONE 3(1): e1426. doi:10.1371/journal.pone.0001426
Academic Editor: Juergen Kroymann, Max Planck Institute for Chemical Ecology, Germany
Received: July 13, 2007; Accepted: December 11, 2007; Published: January 9, 2008
Copyright: © 2008 Kroth et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: PGK is supported by the Deutsche Forschungsgemeinschaft (DFG, projects 1661/3-2, 1661/4-1) and the University of Konstanz. AC acknowledges funding from the Australian Research Council and industry partner, Akzo Nobel, Gateshead, UK (Industry Linkage Grant #LP0454982), as well as financial assistance from the Defence Science and Technology Organization of the Australian Department of Defence and the University of Melbourne (MRGS scheme). VMJ was supported by EU Sixth Framework Programme “Diatomics” (LSHG-CT-2004-512035), the University of Nantes and CNRS. TM was supported by a fellowship within the Postdoc program of the German Academic Exchange Service (DAAD). MSP is supported in part by the PNW Center for Human Health and Ocean Studies (NIH/National Institute of Environmental Health: P50 ES012762 and National Science Foundation: OCE-0434087) and by the Gordon and Betty Moore Foundation. MS was supported by the EC Sixth Framework Programme “Diatomics” (LSHG-CT-2004-512035). LC was supported by CNRS. EVA is supported by the Pacific Northwest Center for Human Health and Ocean Sciences (National Institute of Environmental Health: P50 ES012762 and National Science Foundation: OCE-0434087) and a Gordon and Betty Moore Foundation Marine Microbiology Investigator Award. Diatom whole genome sequencing was funded by the US Department of Energy and was performed at the Joint Genome Institute (Walnut Creek, CA, USA).The generation and sequencing of P. tricornutum ESTs was coordinated by CB and was performed at Genoscope (Evry, France) as a part of projects funded by Genoscope, the EU Sixth Framework Programme (Diatomics; LSHG-CT-2004-512035), and the Agence Nationale de la Recherche (ANR; France).
Competing interests: The authors have declared that no competing interests exist.
Diatoms are abundant unicellular algae in aquatic habitats. They can produce enormous amounts of biomass and are thought to be responsible for about 20% of global carbon fixation. As much as 16 gigatons of the organic carbon produced by marine phytoplankton per year, or about one third of total ocean production is thought to sink into the ocean interior preventing re-entrance of this carbon into the atmosphere for centuries . Recent assessments suggest that diatom-mediated export production can influence climate change through uptake and sequestration of atmospheric CO2 , . The role diatoms play in mitigating atmospheric CO2 concentrations is of special interest now with the rising levels of this “greenhouse gas” and consequent global warming. A significant fraction of the organic carbon generated by diatoms remains in the upper ocean and supports production by higher trophic levels and bacteria.
Despite the important role of diatoms in aquatic ecosystems and the global carbon cycle, relatively little is known about carbon fixation and carbohydrate pathways in these algae . For example the exact mode of CO2 fixation is largely unsolved. Ribulose-1,5-bisphosphate carboxylase/oxygenases (Rubisco) from diatoms have half-saturation constants for CO2 of 30–60 µM  despite the fact that typical sea water contains about 10 µM CO2 . To prevent potential CO2 limitation, most diatoms have developed mechanisms to concentrate dissolved inorganic carbon (DIC) via a CO2 concentrating mechanism (CCM) . Although most of the Calvin cycle enzymes in diatoms are very similar to those in land plants, there are indications that they may be differently regulated by light . Furthermore, some metabolic pathways appear to be missing altogether from diatoms . Finally, there is only scarce information available on the localization, synthesis and storage of chrysolaminaran, the principle storage carbohydrate in diatoms. Diatoms may produce and secrete vast amounts of carbohydrates that play important roles in phototrophic biofilms, yet very little is known about synthesis and secretion of these carbohydrates.
Research on diatoms advanced significantly with publication of the whole genome sequences of the centric diatom Thalassiosira pseudonana  and of expressed sequence tags (ESTs) from the pennate diatom Phaeodactylum tricornutum . Recent availability of whole genome sequence and about 100,000 ESTs for Phaeodactylum tricornutum provide additional opportunities to understand unique physiological characteristics of diatoms. Together with new experimental resources such as genetic transformation, now feasible for several diatom species – and various laboratory-based studies of their physiology , diatoms have become model photosynthetic representatives for non-green algae.
Diatoms have an evolutionary history distinct from higher plants. Diatoms are eukaryotic chimeras derived from a non-photosynthetic eukaryote that domesticated a photoautotrophic eukaryotic cell phylogenetically close to a red alga . After incorporation, the endosymbiont was successfully transformed into a plastid that retained a small plastid genome, but lost the nuclear and the mitochondrial genomes as distinct entities. In addition to the genetic consequences that resulted from extensive gene transfer events and genomic reorganization, secondary endocytobiosis also increased the complexity of diatom cell structure, with implications on physiology and biochemistry. A significant difference between diatom plastids and those of higher plants is that diatom plastids are surrounded by four rather than two membranes, the outermost of which is contiguous with the endoplasmic reticulum. This means that import of all nuclear-encoded plastid proteins and the exchange of metabolites like carbohydrates between the plastids and the cytoplasm must take place across four membranes. To accomplish this task, nuclear-encoded proteins imported into diatom plastids possess an N-terminal signal peptide that targets the protein first to the endoplasmic reticulum and a plastid localization peptide that targets the protein to plastid stroma , . Another striking difference between diatoms and green algae/land plants is their different nuclear and mitochondrial backgrounds because they arose from different host cells.
We annotated genes involved in carbon acquisition and metabolism in the genome of the diatom P. tricornutum and compared these gene models to the only other diatom whole genome sequence of Thalassiosira pseudonana. The 5′-most ends of a majority of critical genes were identified based on EST support. This meant that N-terminal leader sequences could be predicted for most proteins and thus their targeting to different compartments within the cell. Here, we present a comprehensive model of the localization of enzymes and pathways involved in carbon assimilation and carbohydrate production and catabolism.
Results and Discussion
Structure of the genome and gene annotation
Following publication of the draft Thalassiosira pseudonana Hasle & Heimdal (CCMP 1335) genome , a majority of sequence gaps were closed at the Stanford Human Genome Center (SHGC; Stanford, CA, USA) and a new version of the genome sequence is now publicly available at http://genome.jgi-psf.org/Thaps3/Thaps3.home.html. A second diatom genome, from Phaeodactylum tricornutum Bohlin (CCAP1055/1), was subsequently sequenced and completed at the U.S. Department of Energy Joint Genome Institute (JGI, http://www.jgi.doe.gov/, Walnut Creek, CA, USA) and SHGC, and is available publicly at http://genome.jgi-psf.org/Phatr2/Phatr2.home.html. In addition, 100,000 ESTs generated from P. tricornutum cells grown in 14 different conditions have been generated by Genoscope (Evry, France) and are available at http://www.biologie.ens.fr/diatomics/EST. Both genomes are approximately 30 Mb and contain between 10,000 and 11,500 genes. Assembly and annotation of the whole genome of P. tricornutum will be published separately (manuscript in preparation). Here we focus solely on those pathways involved in carbohydrate metabolism. In the following sections, we include protein IDs (Prot-ID) from version 2.0 (P. tricornutum) of the JGI sequence database in parentheses. See Table S1 for a list of annotated genes together with the Prot-IDs in T. pseudonana.
Prediction of intracellular targeting
Nuclear encoded proteins are translated in the cytosol and subsequently transported to their respective target locations. In most known cases an N-terminal targeting domain can send the proteins into the ER, mitochondria, plastids, the extracellular space or to other compartments. In land plants relatively similar transit peptides are used to target into plastids or mitochondria, making it sometimes difficult to predict the correct compartment. Mitochondrial import sequences in diatoms are similar to those in other eukaryotes. Diatom plastid presequences, however, differ significantly from those of land plants or green algae . Diatom plastids are surrounded by four membranes, the outermost being studded with ribosomes and continuous with the endoplasmic reticulum (ER) . Nuclear encoded plastid proteins of diatoms contain N-terminal bipartite presequences consisting of a signal peptide followed by a transit peptide-like domain. Such presequences are easily recognized due to an essential targeting motif with a characteristic signature at the signal peptide cleavage site , .
CO2 fixation: a biochemical (C4) or a biophysical CCM-like metabolism?
The apparent photosynthetic affinity of diatoms for inorganic carbon (Ci) is considerably higher than expected based on the affinity of their Rubisco for CO2 . Extensive diatom blooms that occur during large iron fertilization experiments in high nutrient low chlorophyll regions of the oceans ,  suggest that diatoms are not CO2 limited under natural oceanic conditions. Both results imply that diatoms possess efficient CO2 concentrating mechanisms (CCM), although underlying mechanisms (either a biochemical C4 or a biophysical CCM, or both) are still controversial (see , ).
Studies on the biochemistry of photosynthesis in the well-characterized marine diatom Thalassiosira weissflogii suggested that a C4-like pathway could exist whereby a C4 compound such as malate or OAA is decarboxylated, typically within the chloroplast, to deliver CO2 to Rubisco , . The possibility of a C4-like pathway in the related species T. pseudonana was examined based on an in silico analysis of gene content . The T. pseudonana genome appears to encode the enzymes phosphoenolpyruvate carboxylase (PEPC), phospoenolpyruvate carboxykinase (PEPCK) and pyruvate orthophosphate dikinase (PPDK). Each of these enzymes is required for C4-metabolism, although they also play a role in C3-metabolism. Subsequent analysis of transcript abundances for the putative C4-related genes in T. pseudonana indicated that the gene encoding PEPCK was up-regulated about 1.5 fold under reduced CO2 concentrations, whereas expression of genes encoding PEPC and PPDK were unaffected . Despite the presence of typical C4 enzymes in both Thalassiosira species, short 14CO2 labelling experiments showed marked differences between them . In T. weissflogii, about 30% of the 14C label (in 5 sec. experiments) was observed in malate and about 40% in triose phosphates. In contrast, in T. pseudonana production of 14C-labeled C4 products was negligible. Roberts et al.  concluded that a typical C3 metabolism occurs in T. pseudonana, despite the presence of C4 enzymes, whereas an intermediate C3-C4 may function in T. weissflogii.
Genes essential for C4 metabolism were identified in P. tricornutum. A PPDK (21988), which catalyzes the formation of PEP, was identified and includes both a signal peptide and a putative plastid targeting sequence suggesting that PEP is generated in the plastid (Fig. 1). Two genes encoding PEPC have been identified (Fig. 1). The predicted protein sequence for one of them (PEPC1, 56026) has a high degree of identity (ca. 40% amino acid identity) with the PEPCs from green algae and higher plants. It possesses a signal peptide, but a plastidic transit peptide was not detected suggesting that this protein is targeted either to the ER or to the periplastidic space of the plastids . A second PEPC (PEPC2, 20853) has high similarity (ca. 40% amino acid identity) to PEPCs from bacteria and contains a predicted mitochondrial targeting presequence. Decarboxylation of OAA appears to occur via a mitochondrial-localized PEPCK (23074). This enzyme has the greatest similarity to PEPCK from the proteobacterium Campylobacter jejuni (58% amino acid identity). Two additional decarboxylating enzymes belonging to the malic enzyme family were identified (27477, 56501) and apparently both possess a mitochondrial presequence. One of these enzymes (ME1) (56501) is characterized by a dinucleotide binding site that binds NAD rather than NADP. Thus, P. tricornutum and T. pseudonana appear to have two mitochondrial malic enzymes that are either NAD- or NADP-dependent. Genes encoding a mitochondrial-localized malate dehydrogenase (MDH) (51297), a pyruvate-kinase (PK6) (56172) and a pyruvate-carboxylase (PYC1) (30519) were also identified (Fig. 1). The respective substrates for this pathway may be transported into the mitochondria by a putative mitochondrial oxoglutarate/malate transporter (8990).
Figure 1. Model of carbon concentrating mechanisms (CCM) in diatoms based on annotations of the Phaeodactylum tricornutum and Thalassiosira pseudonana genomes.
For discussion of the pathways see text. Enzyme abbreviations: CA: carbonic anhydrase; MDH: malate dehydrogenase; ME1: NAD malic enzyme, mitochondrial; PEPC: phosphoenolpyruvate carboxylase; PEPCK: phosphenolpyruvate carboxykinase; PK: pyruvate kinase; PPDK: pyruvate-phosphate dikinase; PYC: pyruvate carboxylase; RUBISCO: ribulose-1,5-bisphosphate carboxylase.doi:10.1371/journal.pone.0001426.g001
Models were developed to evaluate how such a C4-like carbon fixation pathway could operate in P. tricornutum (see our working scheme, Fig. 1). The first hypothesized step in carbon fixation is delivery of HCO3− into cells either via specific transporters or by diffusion of CO2 and its subsequent conversion to HCO3− through CA activity (see below). The hypothesized localization of PEPC1 (56026) to the ER or to that part of the ER that is connected to the plastid (CER) or to the periplastidic space (PPS) suggests that subsequent fixation of HCO3− into a C4 compound likely occurs within either the ER or the periplastidic space. The localization of the C4 decarboxylation that delivers CO2 to Rubisco for fixation is not clear. Immuno-localization based studies provided early evidence that the decarboxylating enzyme PEPCK is located in the plastids of the centric diatom Skeletonema costatum . Later, Reinfelder et al.  found that PEPCK activity co-localized with Rubisco activity in isolated plastid-enriched fractions from T. weissflogii and concluded that decarboxylation occurred within the plastids. Subsequent in silico analysis of both P. tricornutum and T. pseudonana indicated that the decarboxylating enzymes PEPCK and malic enzyme do not possess plastid targeting sequences. Moreover, there is no evidence that malate and/or oxaloacetate transporters in these organisms are localized to plastid membranes. Finally, addition of oxaloacetate to intact plastids isolated from the diatom Odontella sinensis  does not result in net O2 evolution as would be expected from the malic dehydrogenase reaction due to turnover of NADPH. Combined, these results suggest that subsequent decarboxylation steps required to generate CO2 for Rubisco delivery, at least in P. tricornutum and T. pseudonana, do not occur in the plastid.
Recent evidence with single chlorenchyma cells of the higher plants Bienertia cycloptera and Borszczowia aralocaspica provides support for a compartmentalized separation of CO2 generation via decarboxylation of C4 compounds and subsequent CO2 fixation by Rubisco , . In these plant cells, PPDK is located in chloroplasts where it converts pyruvate to PEP. The PEP is then transported to the cytosol where it is carboxylated (using HCO3−) via PEPC. The C4 acids produced diffuse to the proximal part of the cell where they are decarboxylated in the mitochondria by NAD-malic enzyme. The resulting CO2 may enter the chloroplasts where it is captured by Rubisco.
Both sequenced diatoms possess two malic enzymes that decarboxylate malate to pyruvate. An NADP-malic enzyme has been proposed for diatoms by Granum et al. . A potential NAD-dependent malic enzyme (56501) was also identified that is predicted to be localized to mitochondria and displays sequence similarity to malic enzymes from the C4 plants Amaranthus hypochondriacus , B. cycloptera and B. aralocaspica . The dinucleotide binding site of the malic enzyme from A. hypochondriacus, P. tricornutum and T. pseudonana possesses a similar amino acid composition suggesting that NAD is the preferred co-factor. These data suggest that in diatoms, decarboxylation of malate to generate CO2 may occur within mitochondria, which are often closely associated with plastids. It is important to note however, that any CO2 molecules released from the mitochondria must cross six membranes to enter the plastid stroma. Moreover, it is likely that CO2 would be converted to HCO3− by CA activity during movement between the mitochondria and plastids, thereby reducing at least part of the elevated CO2 concentration. In this case, the C4 pathway would become a futile cycle whereby HCO3− is first fixed and then formed again, thereby dissipating ATP for PEP formation.
Co-occurrence of PEPCK- and malic enzyme-based decarboxylation pathways in the same organism was also observed in the C4-plant Urochloa panicoides , . Apparently, in diatoms both enzymes may contribute to the decarboxylation of the C4-acid. In some of the higher plants which perform C4 metabolism, the pyruvate formed by decarboxylation of malate, using the NAD-malic enzyme, can be used for amino acid synthesis (Fig. 1)  or oxaloacetate formation thereby replenishing mitochondrial pools of C4 acids. Oxaloacetate can also be oxidized in the TCA cycle. Decarboxylation of OAA by PEPCK generates PEP, which can be used for gluconeogenesis or be transformed into pyruvate by a pyruvate kinase (Fig. 1). The presence of a plastid-targeted putative PEP-transporter TPT1 (24610) and the plastid targeted PPDK (21988) suggests that pyruvate is phosphorylated inside the plastid to produce PEP. PEP can be used to produce aromatic amino acids (Shikimate pathway) and lipids or can be exported to provide the acceptor molecule for HCO3− fixation by PEPC (Fig. 1). This pathway is also known from C4 plants that decarboxylate PEP inside the mitochondria (see , –). However, most C4-plants carry out the first carboxylation step in the cytosol . In P. tricornutum and T. pseudonana PEP might be exported from the plastid and carboxylated at the ER/CER/PPC by PEPC1 leading to a micro-compartmentalization of the enzyme to the outer chloroplast membranes. Lee and Kugrens  have speculated that the evolutionary success of heterokonts might be partly attributed to the use of the periplastidic space as an acidic generator of CO2 from bicarbonate.
In summary, the accumulated evidence indicates that a functioning C4 pathway in diatoms (Fig. 1) requires a spatial separation between CO2 production via decarboxylation of OAA and malate in mitochondria, and CO2 utilization by Rubisco in the plastid. Furthermore, localization of the first carboxylation step by PEPC is still unclear. The energetic cost of a futile cycle raises the possibility that the C4 metabolism may help cells dissipate excess light energy, a pathway that would presumably require down-regulation under energy-limiting conditions. This suggests that the flow of metabolites in this pathway would be affected by light intensity. Localization of key enzymes and determination of expression of C4 related genes in cells exposed to low levels of CO2 could shed light on this issue.
Two lines of evidence support the hypothesis that a biophysical CCM operates in diatoms, as in many other aquatic photosynthetic organisms , –. First, we identified in the genome of P. tricornutum three genes that encode different systems for bicarbonate uptake. One protein (45656) shows homology to sodium/bicarbonate transporters in various organisms and appears to be localized to the plastid. A second protein (32359) is homologous to sodium-dependent anion exchangers and also possesses the bipartite presequence for plastid targeting. The third protein (54405) shows similarity to Cl−/HCO3− exchangers abundant in red blood cells. T. pseudonana appears to possess a single sodium/bicarbonate transporter (24021) that is targeted to the plastid. The hypothesis resulting from these observations is that inhibitors shown to prevent HCO3− uptake as shown in Ulva sp. , should inhibit uptake of HCO3− and thereby the rate of photosynthesis in both diatoms.
The second form of support for a biophysical CCM is identification of numerous genes encoding carbonic anhydrases (CA) in the diatom genomes. The overall sequence similarities among CAs are rather low and they are commonly identified by the presence of conserved domains and by their biochemical properties . Seven CAs are predicted for P. tricornutum. Two CAs (51305, 45443) are related to the beta type and show similarity to CAs found in both plants and prokaryotes. Based on the presence of a plastid targeting presequence and physiological experiments , one of these proteins (51305) is located in the plastid as has been demonstrated by GFP fusion proteins . The other (45443) has a signal peptide. The five other identified CAs (35370, 44526, 55029, 54251, 42574) apparently belong to the alpha family and all possess signal peptides. A recent study by Szabo and Colman  provided experimental evidence for the presence of CA in the periplasmic space of P. tricornutum suggesting that at least a subset of the signal peptide-possessing CAs are likely targeted to the periplasmic space. Surprisingly, similarity of the CAs between the two diatoms T. pseudonana and P. tricornutum is rather low. T. pseudonana appears to possess more intracellular CAs without signal and transit peptides than P. tricornutum. One exception is the carbonic anhydrase 22391 from T. pseudonana which also possesses a signal peptide. Whether this CA is secreted or targeted to ER or periplastidic space remains to be investigated, while it seems clear that it is not plastid localized. The difference between the two diatoms may indicate specialization of the enzyme depending on different ecological niches. This is supported by recent findings that expression of beta carbonic anhydrases may be regulated by several factors including CO2 and light .
Despite the extensive in silico analyses described here, the potential mechanism by which CO2 is delivered to Rubisco remains elusive. In the well-studied green alga, Chlamydomonas, a thylakoid-located alpha CA facilitates conversion of HCO3− to CO2, thereby raising its concentration in close proximity to Rubisco . Mutants impaired in this CA demand high CO2 concentrations for growth (see , ). In P. tricornutum, the plastid-localized CA (51305) is a beta type CA rather than an alpha type. However, it too localizes to the thylakoids where it forms particles, most probably close to the girdle lamellae , and its expression is strongly enhanced under low CO2 conditions by a mechanism involving cAMP . The other beta type CA (45443) is probably located in the ER or the periplasmic space and is constitutively expressed even under high CO2 concentrations . The presence of bicarbonate transporters in the chloroplast envelope (Fig. 1) is consistent with the operation of a biophysical CCM but is not essential for a C4-like CCM, where the initial HCO3− fixation occurs outside the plastid. Finally, enhanced uptake of inorganic carbon, both as CO2 or HCO3−, would be consistent with both types of CCM, whereas raising the concentration of Ci within the cells  is more consistent with the biophysical CCM. The high affinity of PEPC for HCO3− would be expected to alleviate the need to accumulate high Ci concentrations internally. Induction of an extracellular CA at low CO2 concentrations was also observed in P. tricornutum. Such a CA would be expected to facilitate the rate of CO2 formation in the unstirred layer surrounding the cells and thereby to supply CO2 for photosynthesis by either CCM type. In summary, clear evidence that supports either CCM mode as the only way to raise the CO2 concentration in close proximity of Rubisco is presently missing due to lack of sufficient biochemical evidence.
Photorespiration and glyoxylate metabolism
Photorespiration is the inevitable consequence of the ability of either CO2 or O2 to cleave the double bond obtained in RuBP after enolization by Rubisco. In higher plants, photorespiration is thought to provide the photosynthetic machinery with some protection against photoinhibition –. Mutants of tobacco (Nicotiana tabacum L.) defective in enzymes of the photorespiratory pathway demonstrated enhanced photoinhibition under high light conditions , . The specificity factor (τ) of Rubisco, a measure of its ability to discriminate CO2 from O2, is considerably higher in diatoms than in cyanobacteria and green algae (reviewed in ) suggesting a lower rate of O2 fixation in diatoms than observed in members of the green lineage. This is supported by studies showing photorespiratory activity in diatoms at a reduced rate than expected from studies with higher plants –.
When O2 out-competes CO2 for RuBP, one molecule of 2-P-glycolate and one molecule of 3-P-glycerate are produced. The latter may enter the Calvin cycle, whereas 2-P-glycolate, a metabolite known to inhibit the Calvin cycle enzyme triosephosphate isomerase  must be degraded via the photorespiratory pathway (see Fig. 2). In higher plants, metabolism of 2-P-glycolate takes place via the C2 cycle . Following cleavage of the phosphate group, glycolate is exported out of the chloroplast and enters the peroxisome where it is oxidized to glyoxylate, followed by transamination to form glycine which enters the mitochondrion. The glycine decarboxylase complex together with serine hydroxymethyltransferase, catalyzes the condensation of two glycines to one serine with the consequent release of ammonium ion and CO2. The serine is further metabolized back to P-glycerate which may then enter the Calvin cycle in the chloroplast. Thus, out of four carbons entering the C2 cycle, three are converted back to PGA and one is released in the form of CO2.
Figure 2. Model for photorespiration and associated pathways in diatoms based on the annotations of the Phaeodactylum tricornutum and Thalassiosira pseudonana genomes.
For simplicity, the number of oragenelle membranes has been reduced in this figure. A gene model for glycerate kinase (GK) could not be found in either genome. The bacterial-type glyoxylate to glycerate metabolism is not shown due to uncertainty in the localization of the enzymes. Enzyme Abbreviations: ACS: acetyl CoA synthetase; CTS: citrate synthase; GDC: glycine decarboxylase; GOX: glycolate oxidase; GK: glycerate kinase; HPR: hydroxypyruvate reductase /glycerate dehydrogenase; ICL: isocitrate lyase; ME1: NAD malic enzyme; MLS: malate synthase; PDH: pyruvate dehydrogenase; PGP: 2-phosphoglycolate phosphatase; RUBISCO: ribulose-1,5-bisphosphate carboxylase; SHMT: serine hydroxymethyltransferase; SPT/AGT: serine-pyruvate/alanine-glyoxylate aminotransferase.doi:10.1371/journal.pone.0001426.g002
In microalgae, the cyclic process of PGA recovery is not well-studied. More often, glycolate metabolism is studied in the context of its excretion as a waste-product to circumvent unfavourable growth conditions –. A large fraction of glycolate produced by fixation of O2 is released from the cell – and may serve as an important source of organic carbon in the water body. Leboulanger et al.  found high concentrations of glycolate in seawater at both oligotrophic and eutrophic sites, suggesting that photorespiration may be ubiquitous in the marine environment. Photorespiration may therefore represent an important loss of fixed carbon, either via released glycolate or CO2. Recent studies on photorespiration in the cyanobacterium Synechocystis sp. PCC 6803 showed considerable rates of glycolate formation even when the cells were exposed to high levels of CO2 such as 5% CO2 in air . Interestingly in diatoms, when T. weissflogii and T. pseudonana were exposed to 14CO2 for 5 sec, a considerable label (15%) was detected in glycolate in T. pseudonana but only 5% in T. weissflogii . This may indicate a higher level of CO2 in the vicinity of Rubisco in the case of T. weissflogii and consequently a reduced oxygenase activity.
Recent studies on the expression of key genes in the C2 cycle in Thalassiosira sp. suggest the photorespiratory pathway is active in diatoms and plays a critical role in carbon and nitrogen metabolism in the cell , . We have identified most of the enzymes likely involved in a C2-type glycolate pathway in the genomes of P. tricornutum and T. pseudonana (Fig. 2). The annotation of enzymes in the C2 pathway confirms several differences between the photorespiratory cycle in diatoms and in higher plants, and corroborates the scheme proposed by . In algae, it has been suggested that two types of glycolate-oxidizing enzymes exist: a glycolate oxidase in Chrysophyceae, Eustigmophyceae, Raphidophyceae, Xanthophyceae and Rhodophyceae, and a glycolate dehydrogenase in Chlorophyceae, Prasinophyceae, Cryptophyceae and Bacillariophyceae . Winkler and Stabenau  further suggest that in diatoms glyoxylate is synthesized via a glycolate dehydrogenase in both peroxisomes and mitochondria. In both P. tricornutum and T. pseudonana, two proteins similar to glycolate oxidases (GOX) or possibly glycolate dehydrogenases (GDH) were identified: one of the two proteins (22568) contains the peroxisomal targeting motif PTS1, which is common in many eukaryotes and characterized by the consensus sequence (S/C/A)(K/R/H)(L/M) located at the extreme carboxy-terminus . The other protein (50804) appears to be targeted to the mitochondria. This suggests that at least one glycolate oxidizing enzyme in each diatom is localized in the peroxisome, although based on the previous biochemical studies of Suzuki et al.  and Winkler and Stabenau , it is unclear whether it catalyzes production of hydrogen peroxide. Moreover, the potential for peroxisomal activity is corroborated by identification of a malate synthase (54478). In both diatoms, the enzymes for serine synthesis and metabolism have been found targeted to the mitochondria: serine-pyruvate/alanine-glyoxylate aminotransferase (SPT/AGT, 49601), glycine decarboxylase and serine hydroxymethyltransferase GDC/SHMT (56477, 22187, 32847, 18665, 17456), hydroxypyruvate reductase (56499) (Fig. 2). Interestingly, we were not able to identify a gene for glycerate kinase in P. tricornutum or in T. pseudonana. This enzyme catalyzes the last reaction of the C2 cycle and appears to be present in cyanobacteria, the green algal lineage, the red algal lineage, but only sporadically in alveolates and heterokonts. The absence of this enzyme poses the question of how, or whether, glycerate can be transformed into 3-P-glycerate to be reintegrated into the Calvin cycle. An alternative to glycerate, and thereby 3-P-glycerate, as the endpoint of photorespiration is the possibility that all the glycine and serine produced from the fixation of oxygen are instead shunted to other pathways. For example, the formation of the antioxidant glutathione from photorespiratory glycine has been previously demonstrated (reviewed in ).
Another pathway for glyoxylate metabolism, the tartronate semialdehyde pathway, is known in cyanobacteria . Synechocystis mutants were used to illustrate that a C2 pathway and glyoxylate/glycerate pathway (via glyoxylate carboligase and tartronic semialdehyde reductase) cooperate in the metabolism of 2-phosphoglycolate . Genes encoding a putative tartronate semialdehyde reductase (45141) and a putative glyoxylate carboligase, also called tartronate semialdehyde synthase (56476), have been found in both diatoms. The closest BLAST matches to the models for tartronate semialdehyde reductase are genes encoding 3-hydroxyisobutyrate dehydrogenases. The two enzymes are part of the same enzyme family, making a definitive assignment difficult. The putative P. tricornumtum tartronate semialdehyde reductase (45141) has a mitochondrial targeting peptide while the targeting for the T. pseudonana model (2669) is unclear. The closest BLAST matches to the predicted glyoxylate carboligase were acetolactate synthase, however, these two enzymes are also closely related and difficult to distinguish. Both predicted carboligases appear to have chloroplast transit peptides, but the evidence is weak and therefore targeting of glyoxylate carboligase remains uncertain in both diatoms. The presence of glyoxylate metabolism is supported by an early study of Paul and Volcani  showing that the activity of glyoxylate carboligase in the diatom Cylindrotheca fusiformis is affected by light intensity. These data suggest that similar to cyanobacteria, diatoms combine C2 and glyoxylate/glycerate pathways to metabolize 2-phosphoglycolate back to the Calvin cycle.
Reductive/oxidative pentose phosphate pathway
Photosynthetic carbon fixation in plants and algae is performed by the Calvin cycle. Some Calvin cycle enzymes in land plants are of cyanobacterial origin, while others have been replaced by protobacterial or eubacterial enzymes . Carbon fixation in land plant plastids is highly regulated, either by substrates and ions like Mg2+ or by light-dependent redox regulation either at the transcriptional ,  or the enzymatic level via the ferredoxin/thioredoxin-system , . In addition to the Calvin cycle (reductive pentose phosphate pathway), plastids from land plants and green algae possess an oxidative pentose phosphate pathway (OPP). This ubiquitous process produces NADPH and pentose-phosphates for biosynthesis of nucleotides, amino acids and fatty acids in the dark by decarboxylation of glucose-6-phosphate.
As both pathways in plastids are interconnected, operating them simultaneously would result in a futile cycle, using up energy in the form of ATP without net CO2 fixation. Thus in plastids of land plants and green algae some of the enzymes of the Calvin cycle like the phosphoribulokinase (PRK), glyceraldehyde-3-phosphate dehydrogenase (GAP-DH), fructose-1,6-bisphosphatase (FBP), and seduheptulose-1,7-bisphosphatase (SBP) are activated in the light via reduction by thioredoxin (and become inactive in the dark), while the key enzyme of the OPP, the glucose-6-phosphate dehydrogenase (G6PDH) is active in the dark, but inhibited after reduction in the light. In contrast to higher plants, there is apparently no complete oxidative pentose phosphate pathway (OPP) in the plastids of several diatoms ,  as well as in P. tricornutum, suggesting diatom plastids in general lack this pathway. Two putative 6-phosphoglucono-lactonases might be targeted to the cytosol (31882) and to the plastid (38631), however, both genes are not yet supported by ESTs. The other two required enzymes glucose-6-phosphate dehydrogenase (G6PDH, 30040, and the G6PDH component of a G6PDH/6PGDH fusion protein, 54663) and 6-phosphogluconate dehydrogenase (6PGDH, 26934, and the 6PGDH component of the G6PDH/6PGDH fusion protein, 54663) were found to be cytosolic enzymes, indicating that the complete OPP is only functional in the cytosol (Fig. 3).
Figure 3. Model of the oxidative and reductive pentose phosphate pathways and related reactions in P. tricornutum.
For simplicity, the number of organelle membranes has been reduced in this figure. The superscript numbers attached to the enzyme names indicate the number of isoenzymes within the respective compartment. Enzyme abbreviations: AL: aldolase; FBA: fructose-1,6-bisphosphate aldolase; FBP: fructose-1,6-bisphosphatase; GAPDH: glyceraldehyde-3-phosphate dehydrogenase; GPI: Glucose-6-phosphate isomerase; GPDH: glucose-6-phosphate dehydrogenase; PGL: phosphor-gluconate lactonase; PRK: Phosphoribulokinase; RUBISCO: ribulose-1,5-bisphosphate carboxylase; PGDH: 6-phospho-gluconolactone dehydrogenase, PGK: phospho-glycerate kinase; RPI: ribose-5-phosphate isomerase; RPE: ribulose-phosphate epimerase; RPI: ribose-5-phosphate isomerase; TKL: transketolase; TAL: transaldolase; TPI: triose-phosphate isomerase; SBP: seduheptulose-1,7-bisphosphatase.doi:10.1371/journal.pone.0001426.g003
The only Calvin cycle enzymes encoded on the plastid genome are the small and the large subunit of the Rubisco (GenBank AY819643) . All other genes encoding primary enzymes of the Calvin cycle have been identified in the nuclear genomes of P. tricornutum and T. pseudonana (see Fig. 3). The only exception is the gene for the sedoheptulose bisphosphatase (SBP). The only SBP gene we identified (56467) encodes a protein that lacks a plastid targeting sequence and thus appears to be localized within the cytosol. This SBP, however, is not contained in the large set of ESTs from P. tricornutum, indicating that it is not actively transcribed under the applied conditions. SBP catalyses the reaction from sedoheptulose-1,7-bisphosphate to sedoheptulose-7-phosphate in the Calvin cycle; it is unclear yet whether the SBP reaction does not occur in diatom plastids or - more likely - whether this reaction is performed by one of the plastidic FBPs as shown for FBP I in cyanobacteria . Interestingly there is a gene encoding a plastidic FBP (FBPC2, 42456) with a bipartite plastid targeting presequence that is located about 500 bases upstream of the SBP gene in the same orientation (similar as in T. pseudonana). There is a theoretical possibility that both genes might be transcribed together and - after excision of a putative intron - might be translated as a fusion protein, thus the SBPase could be imported in a piggy-back manner, although this hypothesis has not yet been supported by transcript analyses (Weber and Kroth, unpublished).
Genes encoding the Calvin cycle enzymes fructose-1,6-bisphosphate aldolase (FBA) and FBP are present in several copies. There are two class II aldolases (22993, bd825) and one class I (24113) aldolase in the plastids, while a class I (42447) and a class II (29014) aldolase are found in the cytosol. Four plastidic FBPases have been identified (FBPC1: 42886; FBPC2: 42456; FBPC3: 31451; FBPC4: 54279) and one cytosolic enzyme (23247). The redundancy of isoenzymes may partially reflect the evolution of diatoms by secondary endocytobiosis , . Some of the isogenes may have either a cyanobacterial or a rhodophytic origin or are related to respective enzymes from oomycetes. Other genes may also have been transferred by lateral gene transfer from bacteria or have been duplicated within the heterokonts .
Redox-regulation of enzymatic activity is critical for plastid functions. Thioredoxin is a small protein that is reversibly reduced in the light by ferredoxin/thioredoxin reductase (FTR) and is able to reduce target enzymes resulting in altered enzymatic activities . Several genes encoding thioredoxins (Trx) were identified in P. tricornutum, including the genes for Trxs f (46280) and m (51357) both possessing typical plastid targeting signals. Three genes encoding Trx h proteins (48539, 56471, 48141/56521) were identified, one of which (48539 plus possibly 48141/56521) contains a presequence for targeting into ER/periplastidic space (respective homologues are also found in T. pseudonana). This is surprising because Trx h is located in the cytosol in all other organisms examined so far. Genes encoding two plastidic Trxs y (33356, 43384), a mitochondrial Trx o (31720) and a ferredoxin-thioredoxin oxidoreductase (50907, needed for Trx reduction) were also identified. These results imply that thioredoxin based light-regulation is functional in diatom plastids, although far fewer plastid enzymes in diatoms than in plants may be actual Trx targets (see ).
Another group of proteins involved in redox-regulation in land plant plastids are glutaredoxins (Glrx), which are involved in fine-tuning of the thioredoxin system . In P. tricornutum we predict two glutaredoxins to be targeted into the plastids (43497, 39133), one to the cytosol (16854), and one to the mitochondria (37615). Similar to the unusual periplastidal/ER associated Trxs h (48539/48141), one glutaredoxin (56497) also contains a presequence for targeting into ER/periplastidic space. Taken together, thioredoxins and glutaredoxins are present in the mitochondria, plastids and cytosol of P. tricornutum and T. pseudonana, although their functionality and specificity is unclear.
The plastidic fructose-bisphosphatase (FBP) is the only enzyme in diatoms for which there is direct evidence of redox-regulation by thioredoxin . The PRK also possesses the conserved cysteines for redox regulation, although due to a shift of the redox midpoint potential of this enzyme, it does not get oxidized in vivo and thus is permanently active . Diatom plastids also possess a different GADPH enzyme compared to green algae and land plants, termed GapC1 (25308), which does not contain the respective cysteines  and which is not affected by oxidation or reduction (Michels, A. Wedel, N., and Kroth, P.G., unpublished). The chloroplast ATPase in land plants is modulated by thioredoxin by lowering the energy threshold of the membrane potential necessary to activate the enzyme. The sequence cassette on the γ subunit containing the necessary cysteines (AtpC, 20657) in land plants is missing in diatoms as well as in red algae.
Other plastidic enzymes which are affected by thioredoxin in land plants are not found in P. tricornutum or T. pseudonana. (i) In land plants and in green algae there are two malate dehydrogenases, one of which is NAD-dependent and one of which is NADP-dependent. The NADP-dependent enzyme is redox-regulated via thioredoxin and serves as a valve for excess NADPH . Based on enzymatic and in silico analyses, the redox-regulated isoenzyme appears to be missing from diatom plastids (Mertens and Kroth, unpublished). (ii) ADP-glucose pyrophosphorylase (AGPase) in land plant plastids produces ADP-glucose, the substrate for starch synthesis . Diatoms do not possess a plastidic AGPase, which is consistent with the fact that they export all carbohydrates immediately from the plastids and store them as chrysolaminaran in cytosolic vacuoles. (iii) The Rubisco activase responsible for activation of Rubisco , is apparently also missing from diatom plastids as no gene for this protein has been found in the genomes of P. tricornutum or T. pseudonana. (iv) Another system regulating the Calvin cycle in land plants is the formation of enzyme complexes of GAPDH and PRK by the small protein CP12 via disulfide bridges . In land plants and in green algae these complexes form in the dark, and in the light they are reduced by thioredoxin in the presence of NADPH, dissociate and release GAPDH and PRK activity . A comparison of native GAPDH and PRK enzymes from stromal extracts of diatoms and land plants by gel filtration revealed that diatoms do not form GAPDH/PRK/CP12 complexes (Michels, Wedel and Kroth, in preparation), accordingly we were not able to identify genes for putative CP12 proteins in diatom genomes.
Interestingly, during our genome analysis we identified a few cases of unusual gene fusions. When transcribed as a single mRNA they may form fusion proteins consisting of two metabolic enzymes that are connected by spacers of 8 to 25 amino acids. We found three metabolic enzyme pairs that apparently are fused to each other because they are transcribed by a single mRNA: the mitochondrial triosephosphate-isomerase/glyceraldehyde-3-phosphatedehydrogenase (TIM-GAPC3, 25308) , a cytosolic UDP-glucose-pyrophosporylase/phosphoglucomutase(UGP/PGM, 50444), and a cytosolic glucose-6-phosphate-dehydrogenase/6-phosphogluconate-dehydrogenase(G6PDH/6PGDH, 54663) fusion protein. The fact that each pair of enzymes catalyzes two subsequent metabolic reactions indicates that fusing these genes may result either in a better regulation or a faster conversion of substrates. However, there is evidence that at least some of these fusion proteins may be cleaved post-translationally  (Majeed and Kroth, unpublished). Interestingly, the TIM-GAPDH (present in T.pseudonana and P. tricornutum) and the UGP/PGM are found in the genomes of the stramenopiles Phytophthora ramorum and Phytophthora sojae, while the G6PDH/6PGDH is not.
Glycolysis is a universal cytosolic pathway for degradation of hexoses and results in pyruvate, which may be targeted to the mitochondria in eukaryotic organisms performing aerobic degradation or may be utilized in various other ways in organisms capable of living in anaerobic conditions. Several enzymes involved in glycolysis occur as a number of isoenzymes in P. tricornutum and T. pseudonana. For instance there are five genes for phosphoglucomutases (PGM) present in the P. tricornutum genome: two of the gene products (48819, 50718) are likely to be targeted to the plastid while the other isoenzymes apparently are located in the cytosol (51225, 50118 and the PGM component of a UDP-Glucose-Pyrophosphorylase/Phosphoglucomutasefusion protein 50444). Similarly there are three phosphoglycerate kinases predicted to be targeted either to the cytosol (51125), the mitochondria (48983) or the plastid (29157). Recent analyses using GAPDH genes from diatoms and other organisms indicate a common origin of all chromalveolates (86). Of the six identified GAPDH enzymes in P. tricornutum, two are targeted to the mitochondria (32747 and the GapC3 component of a TPI/GapC3 fusion protein 25308) and one is targeted to the plastids (22122) . GapC2, assigned to be cytosolic  is present in two copies encoded in the same orientation on chromosome 16, with a distance of approx. 24 kilo base pairs (51128, 51129). A third cytosolic GAPDH enzyme was additionally identified (23598). Three genes for glucose-6-phosphate isomerases (GPI) were found, encoding a plastidic GPI (56512) and two cytosolic enzymes  with genes located next to each other in opposite direction (23924, 53878). We found only genes encoding a plastidic (56468) and two mitochondrial enolases (bd1572, and the apparently unfunctional bd1874) in P. tricornutum. However, in T. pseudonana a cytosolic and a mitochondrial enolase have been found (40771, 40391). This indicates that all reactions of the glycolysis may potentially occur within the plastid (Fig. 4), where some of them simply represent essential enzymes of the Calvin cycle. Also surprising is the fact that there are isoenzymes of the complete second half of the glycolysis possessing mitochondrial presequences (see Fig. 4). In some cases the respective enzymes have been shown to be targeted into the mitochondria by fusing the presequences to GFP (C. Rio Bartulos, personal communication). Similarly the translocation of glycolytic reactions to other organelles has been described in unicellular green algae .
Figure 4. Model of the glycolytic reactions in the cytosol and related pathways within mitochondria and plastids of P. tricornutum.
Enzyme abbreviations: PGM: phosphoglucomutase; GPI: Glucose-6-phosphate isomerase; PFK: Phosphofructokinase; FBA: fructose-1,6-bisphosphate aldolase; GAPDH: glyceraldehyde-phosphate dehydrogenase; PGK phospho-glycerate kinase; PGAM: phosphor-glycerate mutase; PK: pyruvate kinase; PPDK: pyruvate-phosphate dikinase.doi:10.1371/journal.pone.0001426.g004
No gene encoding a hexokinase for phosphorylation of glucose was detectable in either P. tricornutum or T. pseudonana. Instead, genes for glucokinases were detected in both species. This observation conforms to the trend that sugar-specific kinases are typical in prokaryotes and unicellular eukaryotes, whereas hexokinases with broader substrate specificities are typical in multicellular eukaryotes . The P. tricornutum cytosolic glucokinase (48774) might additionally be involved in the chrysolaminaran pathways (see below).
Storage products–synthesis and degradation
Chrysolaminaran is the principal energy storage polysaccharide of diatoms. The relatively high contribution of chrysolaminaran to marine particulate matter underscores this molecule's significant role in the oceanic cycling of carbon –. It generally comprises between 10 and 20% of the total cellular carbon in exponentially growing diatoms but can accumulate to up to 80% of the total cellular carbon in cells whose growth is limited by nitrogen . Chrysolaminaran concentrations undergo a diel rhythm characteristic of an assimilatory and respiratory product, accumulating during the daylight and becoming depleted in the dark , , . The structure of chrysolaminaran is fundamentally based on a β-1,3-linked glucan backbone, which is infrequently branched with mainly β-1,6-linkages –. Vacuolar localization of chrysolaminaran in several diatom species, including P. tricornutum and T. pseudonana, was demonstrated by staining with aniline blue  and by immunolabeling with a monoclonal anti-1,3-β-D-glucan antibody .
The biochemical pathways leading to chrysolaminaran synthesis and degradation have not been elucidated. However, enzyme assays of cell-free extracts from the diatom Cyclotella cryptica demonstrated that the formation rate of UDP-glucose was ≥20-fold greater than for any other nucleoside-diphosphate-glucose and that UDP-glucose served as a substrate for chrysolaminaran synthesis . Furthermore, exo-1,3-β-glucanase activity was detected in several planktonic diatoms and upregulation of this activity coincided with chrysolaminaran degradation in the diatom Skeletonema costatum .
We focused on exo- and endo-1,3-β-glucanases and β-glucosidases as the primary enzymes involved in digesting chrysolaminaran. We found four putative exo-1,3-β-glucanases in P. tricornutum, all belonging to the glycosyl hydrolase family 16 (49294, 56510, 56506, 49610). All orthologues possess an N-terminal signal peptide, except (49610), and all contain a C-terminal transmembrane helix. In addition, one (56510) possesses a putative C-terminal ER-retention signal (REEL). Of the four exo-1,3-β-glucanases, only one (49294) was represented in T. pseudonana (13556). Three putative endo-1,3-β-glucanases were identified in P. tricornutum, two belonging to glycosyl hydrolase family 16 (54681, 54973) and one to family 81 (46976). One of these (54681) consists of 1028 amino acid residues and has both an N-terminal signal peptide and a C-terminal transmembrane domain. The second (54973) has a signal peptide but no transmembrane helix and it is only about half the length. Curiously, the third enzyme (46976) apparently lacks a signal peptide but has an N-terminal transmembrane helix, making it a candidate for a type II transmembrane protein. Among the sequences, 54681 from P. tricornutum and 35711 from T. pseudonana are most similar to each other. In a ClustalW tree of endo-β-glucanases, these sequences clustered with bacterial endoglucanases (Rhodothermos marinus, Bacillus circulans, and Sinorhizobium meliloti), and (54973) only very weakly associated with these. The family-81 endoglucanases from the two diatom species grouped together but were apparently still relatively divergent, the next closest sequences being a pair of family-81 endoglucanases from Arabidopsis thaliana.
Three putative β-glucosidases were identified in P. tricornutum, one belonging to glycosyl hydrolase family 1 (50351) and the other two (45128, 49793) to family 3. In T. pseudonana, only a single β-glucosidase was identified (28413), and this belonged to glycosyl hydrolase family 1. In addition to a signal anchor (50351), only one of the P. tricornutum family-3 β-glucosidases (45128) appears to have a C-terminal transmembrane helix. All three P. tricornutum orthologues were represented by ESTs, but to varying degrees.
Overall, at least 10 enzymes predicted to digest 1,3-β-glucans were identified in P. tricornutum. Presumably, at least one of the exo-1,3-β-glucanases and one of the endo-1,3-β-glucanases act complementarily to digest the principle β-1,3-linkages of chrysolaminaran. The products of efficient digestion by this suite of enzymes would be primarily free glucose, with relatively small amounts of glucosyl oligosaccharides dominated by β-1,6-linkages (e.g. gentiobiose) derived from surviving chrysolaminaran branch points. A β-glucosidase could hydrolyze such oligosaccharides to free glucose. The free glucose generated from complete chrysolaminaran degradation would subsequently be phosphorylated by glucokinase.
The vacuolar localization of chrysolaminaran implies that the degradative enzymes are also localized there. However, the exo-1,3-β-glucanase (56510) possesses a C-terminal ER-retention signal in addition to the signal peptide and C-terminal transmembrane helix. In yeast, transmembrane domains can serve as localization signals for sorting proteins from the ER, with the destination (plasma membrane or vacuole) dependent upon transmembrane helix length and composition rather than on a specified sequence . We identified one gene for a glucokinase in P. tricornutum. As described above we conclude the enzyme to be involved in the cytosolic glycolysis (48774). Although there is no EST support, it is possible that by intron splicing the glucokinase may possess a signal peptide, which might allow targeting to the vacuole (compare to 56514). Interestingly, similar to the glucanases the enzyme possesses a C-terminal transmembrane helix, indicating that it might be integrated into membranes as shown for various hexokinases from plants . In addition to a number of bacterial sequences, the most similar sequence to the diatom glucokinases is the glucokinase of Cyanidioschyzon merolae. This enzyme apparently also lacks a hexokinase and its glucokinase also contains a C-terminal transmembrane helix . The simplest model is that the diatom glucan-digesting enzymes and the glucokinase are anchored at their C-termini to cytosolic membranes like the vacuolar membrane either being oriented towards the cytosol or to the vacuole. The localizations of the β-glucosidases are heterogeneous, however, and for any one of them to serve as a vacuolar gentiobiase would require localization by mechanisms other than those that localize the β-glucanases or the glucokinase.
The hypothesis proposed here for degradation of chrysolaminaran has implications for the generation of glucosyl phosphate intermediates from an energy storage glucan. First, enzymes responsible for chrysolaminaran degradation apparently were recruited during evolution from enzymes normally associated with extracellular polysaccharides. Second, and as a consequence, degradative and phosphorylating steps are decoupled in diatoms. In organisms that metabolize starch or glycogen, the degradative and phosphorylating steps are achieved either concomitantly by an ATP-independent pathway or separately by an ATP-dependent pathway in which phosphorylation is catalyzed by hexokinase (for reviews, see , . The apparent occurrence of only glucokinase in both P. tricornutum and T. pseudonana may, apart from reflecting their evolutionary heritage, be an adaptation to a dedicated ATP-dependent pathway for chrysolaminaran digestion. Bacterial glucokinases, such as those of Escherichia coli, Zymomonas mobilis, Bacillus stearothermophilus, and Streptococcus mutans, have a high specificity and moderately high but relatively narrow KM range for glucose (KM = 0.22–0.61 mM; –) compared with broad-specificity eukaryotic hexokinases (KM = 0.007–2.5 mM; , . In diatoms, such a glucokinase could cope with substantial fluxes in glucose concentrations and ensure that the phosphorylation is efficient as high concentrations of free glucose are liberated during chrysolaminaran degradation. The affinities and kinetics of the diatom glucokinases will need to be characterized, to assess the validity of this hypothesis.
The synthetic pathway of chrysolaminaran is essentially unknown. Based on enzyme activity assays of C. cryptica, UDP-glucose likely serves as the substrate for chrysolaminaran synthesis . The apparent absence of genes encoding ADP-glucose pyrophosphorylase in either diatom species provides further support that UDP-glucose serves as the substrate for chrysolaminaran synthesis. Interestingly, the UDP-glucosyl pyrophosphorylase (UGP) from C. cryptica was not inhibited by 3-P-glycerate or inorganic phosphate, suggesting that the assimilatory glucan is synthesized outside the plastid . Surprisingly, a UDP-sugar pyrophosphorylase of the UGP family was encoded in the genome (23639) and this enzyme is predicted to be targeted to the chloroplast. The plastid localization of a potential UGP in both diatoms suggests that UDP-glucose is used for synthesis of chrysolaminaran within the CER en route to the vacuole. The origin of glucose-6-phosphate as substrate for a plastidal UGP remains unclear but is presumably derived from other sugar phosphates circulating in the plastid. A second candidate for UGP is one derived from a UGP/PGM fusion protein apparently localized in the cytosol (50444). This enzyme could supply UDP-glucose to a membrane-bound glucan synthase (see discussion below).
One or probably more glycosyl transferases likely synthesize the chrysolaminaran polymer, and these could be either orthologous to 1,3-β-glucan synthases in other organisms or perhaps more likely, novel enzymes due to the unique structure and function of chrysolaminaran. A single gene encoding 1,3-β-glucan synthase was identified in P. tricornutum. The deduced protein consists of over 2,100 amino acids and possesses 20 transmembrane domains and a signal peptide. A single 1,3-β-glucan synthase was also identified in T. pseudonana, although no signal peptide was detected for the predicted protein likely because the N-terminus was incompletely defined. The two diatom sequences were most similar to each other and showed similarity to callose synthase sequences from dicots such as A. thaliana and Oryza sativa. This membrane-bound enzyme likely catalyzes the addition of glucosyl residues from cytosolic UDP-glucose on one side of the membrane to the growing, non-reducing terminus of the polysaccharide chain, which protrudes from the enzyme on the opposite side of the membrane. This presumed mode of action is akin to that of plasma membrane-bound polysaccharide synthases such as the Thalassiosira chitin synthases ,  and the cellulose synthases of terrestrial plants and multicellular algae , . Extracellular callose has been reported in diatoms and was suggested to serve as a permeable seal in the girdle regions during cell division , so it is feasible that the identified 1,3-β-glucan synthase is a plasma-membrane-bound callose synthase. The relatively high EST support for this gene under a variety of growth limiting conditions, however, argues for a more active role not limited to cell division. Localization of the 1,3-β-glucan synthase to either the vacuole or the CER would help to determine where UDP-glucose is accessed from–either the cytosol or the CER. If UDP-glucose is accessed from the vacuole, this would support the hypothesis that chrysolaminaran metabolism evolved by relocating to the vacuole enzymes involved in the synthesis and processing of extracellular polysaccharides.
Three additional gene models were identified in P. tricornutum (48300, 56509, 50238) and in T. pseudonana (3105, 4956, 9237) that encode proteins with moderate similarity (up to 28%) to fungal Skn1 and Kre6, enzymes required for synthesis of fungal wall 1,6-β-glucans . The diatom proteins all contain N-terminal signal peptides and single C-terminal transmembrane helices, suggesting they are also associated with the suite of enzymes that process β-glucans. Although the precise function of the fungal enzymes is unclear, they resemble family-16 glycosyl hydrolases and have been interpreted as potential glycosylases and/or transglycosylases . Branching in terrestrial plant starches and mammalian and fungal glycogen is achieved by specific enzymes that hydrolyze internal α-1,4-glycosidic bonds and transfer the released reducing ends to C-6 hydroxyls of the acceptor polysaccharide chain , . If analogous processes occur in diatoms, the putative diatom glucosylase/transglucosylases could act as chrysolaminaran branching/debranching enzymes.
Inositol and Propanoate pathways
Many different cyclitols occur in plants with the most widespread and extensively studied being myo-inositol , . Myo-inositol becomes incorporated into several crucial cellular compounds including those involved in signal transduction (phosphatidylinositol [PI], phosphatidylinositol-4,5-bisphosphate [PIPs]), hormone regulation (indole acetic acid [IAA] conjugates), membrane tethering (glycerophosphoinositide [GPI] anchors), stress tolerance (ononitol, pinitol), oligosaccharide synthesis (galactinol), and phosphorus storage (inositol hexakisphosphate [IP6]). Its primary breakdown product, D-glucuronic acid, is utilized for synthesis of various cell wall pectic non-cellulosic compounds, expanding the list of processes impacted by inositol synthesis and metabolism. In both P. tricornutum and T. pseudonana genes encoding enzymes predicted to be involved in inositol metabolism are well-represented, in particular the methylmalonate-semialdehyde dehydrogenase (acylating) (MMSDH), myo-inositol 2 dehydrogenase (InDH), and triosephosphate isomerase (TIM).
De novo synthesis of inositol has been studied mainly in yeast and proceeds from glucose 6-phosphate through inositol 1-phosphate in two steps catalyzed by inositol phosphate synthase (INPS) and inositol monophosphatase (IMP). Genes encoding these two enzymes are present in the genomes of both P. tricornutum and T. pseudonana. Myo-inositol can be interconverted to scyllo-inosose by the enzyme myo-inositol dehydrogenase (InDH; EC 188.8.131.52). Stein et al.  confirmed the presence of InDH in the red alga Galdieria sulphuraria and a further study by Gross and Meyer  examined the distribution of InDH in algae by confirming its presence through enzyme assays. On the basis of InDH activity they assigned the different algae tested into two distinct groups: one composed of red algae and Glaucocystophyta and the other composed of heterokontophytes and haptophytes. They also proposed an inositol/inosose shuttle across the mitochondrial membrane as an alternative to the mitochondrial NADH dehydrogenase present in higher plants and green algae. The mitochondria of red algae seem to be capable of using myo-inositol for respiration, which was hypothesized to exemplify the divergence of basic metabolism during algal evolution. In plants, neither synthesis nor degradation involves InDH and there was no InDH activity present in the green algae tested by Gross and Meyer . Interestingly, neither synthesis nor catabolism involves scyllo-inosose as a reaction product in the algae studied, whereas in mammals  it seems to be involved in the synthesis of scyllo-inositol.
The sequences predicted to encode InDH from both P. tricornutum and T. pseudonana were compared on the basis of their predicted amino acid sequences to other InDH sequences including those from the red algae C. merolae and G. sulphuraria. The predicted protein sequences from P. tricornutum (51869) and T. pseudonana (8703) formed their own clade separate from the other sequences analysed. This would seem to provide further evidence for the theory of Gross and Meyer  for a divergence in algal metabolism based on InDH, but the other P. tricornutum sequence (34720) was found within the clade formed by the red algal sequences.
MMSDH is an enzyme involved in valine catabolism rather than inositol metabolism. Here it catalyzes the irreversible NAD+- and CoA-dependent oxidative decarboxylation of methylmalonate semialdehyde to propionyl-CoA. It has been suggested that a Bacillus version of the protein is located in an operon and/or involved in myo-inositol catabolism, converting malonic semialdehyde to acetyl CoA and CO2 . Without further investigation its role in inositol metabolism in both P. tricornutum and T. pseudonana is unclear.
There are elementary differences between green algae and diatoms
Due to their evolutionary history diatoms naturally display a completely different host cell/mitochondria/plastid relationship compared to green algae and land plants. There are clear differences between diatoms and the green algae and higher plants in the structure of thylakoids, plastid envelope membranes, the mode of carbohydrate storage and the photosynthetic properties including photoprotection (for a detailed comparison see , ). The genome of the unicellular green alga Chlamydomonas reinhardtii, also representing a single-celled alga but originating from a primary endocytobiosis event, has recently been sequenced  and perhaps not unexpectedly, most Chlamydomonas proteins with a plastidic function display similarity to diatom sequences. However, among the 153 diatom sequences we have analysed (table S1) only 3 of them showed the highest similarity to a Chlamydomonas protein whereas 23 displayed the greatest similarity to a higher plant sequence. Although the general pathways of carbohydrates are similar between diatoms and Chlamydomonas, several peculiar differences were identified that may have resulted from intracellular translocation of enzymes and/or pathways. Two distinctive features stand out: The mode of CO2 concentration in diatoms is still largely unclear, as well as the post-translational regulation of photosynthetic products.
Materials and Methods
We screened sequences from the current JGI (http://www.jgi.doe.gov/) diatom genome sequencing projects for the diatoms Thalassiosira pseudonana v3.0 (http://genome.jgi-psf.org/Thaps3/Thaps3.home.html)  and Phaeodactylum tricornutum v2.0 (http://genome.jgi-psf.org/Phatr2/Phatr2.home.html) using the BLAST algorithm . Comparison with the genome sequences of the red algae Cyanidioschyzon merolae (http://merolae.biol.s.u-tokyo.ac.jp/)  and Galdieria sulphuraria (http://genomics.msu.edu/galdieria) ,  as well as with other publicly available algal sequences helped to delimit gene modeling.
Signal peptides of endoplasmic reticulum (ER) proteins were identified using the program SignalP (http://www.cbs.dtu.dk/services/SignalP/) . In addition ER proteins often possess a C-terminal retention signal. The presence of such a signal (KDEL, DDEL or DEEL) was checked manually.
Plastid proteins of diatoms possess bipartite targeting signals consisting of a signal peptide and a transit peptide-like domain with a conserved “ASAFAP”-motif at the signal peptide cleavage site , . We screened for signal peptides using SignalP. For cleavage site predictions the results of SignalP's Neuronal networks (NN)  or Hidden Markov Models (HMM)  were used. For prediction of chloroplast transit peptide-like domains, the program ChloroP (http://www.cbs.dtu.dk/services/ChloroP/)  was used. The transit peptide-like domains of bipartite plastid targeting sequences often attain poor prediction scores, so we used the NCBI (http://www.ncbi.nlm.nih.gov/) Conserved Domain Search (http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi)  to identify N-terminal extensions from the conserved regions of the respective protein. If a distance of at least 10 amino acids between the predicted cleavage site of the signal peptide and the region of high homology to respective proteins of other organisms was found, also a weakly predicted transit peptide-like domain was accepted. In some cases transit peptide-like domains of plastid proteins are also recognised as mitochondrial transit peptides by the program TargetP (http://www.cbs.dtu.dk/services/TargetP/) . Recent mutational analysis of plastid targeting presequences revealed that only the aromatic amino acids phenylalanine, tryptophan, tyrosine and the bulky amino acid leucine at the +1 position of the predicted signal peptidase cleavage site allow plastid import . Proteins which (i) possess a signal peptide but no ER retention signal (ii) possess a N-terminal extension longer than the signal peptide with some transit peptide-like features (iii) contain F, W, Y or L at the signal peptide cleavage site, were considered to be plastid targeted.
Mitochondrial transit peptides were identified using the program TargetP . Putative enzymes without recognizable targeting sequences were considered cytosolic although the possibility cannot be excluded that they might be targeted to further compartments. For a detailed description of protein localization prediction see also .
Genes involved in carbohydrate pathways in the diatom Phaedactylum tricornutum as assessed from the genome publicly available at http://genome.jgi-psf.org/Phatr2/Phatr2.home.html. For every identified gene the following information is given: enzyme name; common abbreviation; designated pathway; Protein ID; GenBank accession number (if available); number of isogenes identified; genomic coordinates; best BLAST hit: gene, organism, % identity, GenBank accession number; targeting predictions: mTP: mitochondrial targeting peptide score, SP: signal peptide score, other: probability for other localization, Loc: Prediction of localization, based on the scores of TargetP, RC: reliability class, 1 = strong, 5 = poor prediction, TPlen: length of transit peptide, regions of proposed signal peptide cleavage site; assigned localization: overall targeting prediction; respective data for homologous T. pseudonana genes if identified. Targeting predictions were performed by TargetP (http://www.cbs.dtu.dk/services/TargetP/)  and SignalP's (http://www.cbs.dtu.dk/services/SignalP/) Neuronal networks (NN)  or Hidden Markov Models (HMM) . For a detailed description of protein localization prediction see also . The Protein IDs and genomic coordinates are directly linked to the genomic database.
(0.31 MB XLS)
We like to thank the diatom sequencing team of the Joint Genome Institute (Walnut Creek, CA, USA), especially R. Otillar and A. Kuo for valuable help with the annotation of the diatom genomes.
Conceived and designed the experiments: AC PK VM TM MP MS AG. Performed the experiments: AC PK VM TM MP MS AG. Analyzed the data: CB UM AC EA AK PK VM TM MP MS LC AG TW. Wrote the paper: CB AC EA AK PK VM TM MP MS AG.
- 1. Falkowski PG, Barber RT, Smetacek V (1998) Biogeochemical controls and feedbacks on ocean primary production. Science 281: 200–205.
- 2. Brzezinski MA, Pride CJ, Franck VM (2002) A switch from Si(OH)4 to NO3-depletion in the glacial Southern Ocean. Geophys. Res Let 29: 5-1 to 5-5.
- 3. Granum E, Raven JA, Leegood RC (2005) How do marine diatoms fix 10 billion tonnes of anorganic carbon per year. Can J Bot 83: 898–908.
- 4. Roberts K, Granum E, Leegood RC, Raven JA (2007) C3 and C4 Pathways of Photosynthetic Carbon Assimilation in Marine Diatoms Are under Genetic, Not Environmental, Control. Plant Physiol 145: 230–235.
- 5. Badger MR, Andrews TJ, Whitney SM, Ludwig M, Yellowlees DC, et al. (1998) The diversity and coevolution of Rubisco, plastids, pyrenoids, and chloroplast-based CO2-concentrating mechanisms in algae. Can J Bot 76: 1052–1071.
- 6. Riebesell U, Wolf-Gladrow , Smetacek V (1993) Carbon dioxide limitation of marine phytoplankton growth rates. Nature 361: 249–251.
- 7. Giordano M, Beardall J, Raven JA (2005) CO2 concentrating mechanisms in algae: mechanisms, environmental modulation, and evolution. Annu Rev Plant Biol 56: 99–131.
- 8. Wilhelm C, Büchel C, Fisahn J, Goss R, Jakob T, et al. (2006) The regulation of carbon and nutrient assimilation in diatoms is significantly different from green algae. Protist 157: 91–124.
- 9. Michels AK, Wedel N, Kroth PG (2005) Diatom Plastids Possess a Phosphoribulokinase with an Altered Regulation and No Oxidative Pentose Phosphate Pathway. Plant Physiol 137: 911–920.
- 10. Armbrust EV, Berges JA, Bowler C, Green BR, Martinez D, et al. (2004) The genome of the diatom Thalassiosira pseudonana: ecology, evolution, and metabolism. Science 306: 79–86.
- 11. Maheswari U, Montsant A, Goll J, Krishnasamy S, Rajyashri KR, et al. (2005) The Diatom EST Database. Nucleic Acids Res 33 Database Issue: D344–D347.
- 12. Zaslavskaia LA, Lippmeier JC, Kroth PG, Grossman AR, Apt KE (2000) Transformation of the diatom Phaeodactylum tricornutum (Bacillariophyceae) with a variety of selectable marker and reporter genes. J Phycol 36: 379–386.
- 13. Fischer H, Robl I, Sumper M, Kröger N (1999) Targeting and covalent modification of cell wall and membrane proteins heterologously expressed in the diatom Cylindrotheca fusiformis. J Phycol 35: 113–120.
- 14. Poulsen N, Kröger N (2005) A new molecular tool for transgenic diatoms: control of mRNA and protein biosynthesis by an inducible promoter-terminator cassette. FEBS J 272: 3413–3423.
- 15. Patron NJ, Rogers MB, Keeling PJ (2004) Gene replacement of fructose-1,6-bisphosphate aldolase supports the hypothesis of a single photosynthetic ancestor of chromalveolates. Eukaryot Cell 3: 1169–1175.
- 16. Kilian O, Kroth PG (2005) Identification and characterization of a new conserved motif within the presequence of proteins targeted into complex diatom plastids. Plant J 41: 175–83.
- 17. Gruber A, Vugrinec S, Hempel F, Gould SB, Maier U-G, et al. (2007) Protein targeting into complex diatom plastids: functional characterisation of a specific targeting motif. Plant Mol Biol 64: 519–530.
- 18. Gibbs SP (1981) The chloroplast endoplasmic reticulum: structure, function, and evolutionary significance. Int Rev Cytol 72: 49–99.
- 19. Behrenfeld MJ, Bale AJ, Kolber ZS, Aiken J, Falkowski PG (1996) Confirmation of iron limitation of phytoplankton photosynthesis in the equatorial Pacific Ocean. Nature 383: 508–511.
- 20. Gervais F, Riebesell U, Gorbunov MY (2002) Changes in primary productivity and chlorophyll a in response to iron fertilization in the Southern Polar Frontal Zone. Limnol Oceanogr 47: 1324–1335.
- 21. Reinfelder JR, Kraepiel AML, Morel FMM (2000) Unicellular C-4 photosynthesis in a marine diatom. Nature 407: 996–999.
- 22. Reinfelder JR, Milligan AJ, Morel FM (2004) The role of the C4 pathway in carbon accumulation and fixation in a marine diatom. Plant Physiol 135: 2106–2111.
- 23. Cabello-Passini A, Swift H, Smith GJ, Alberte RS (2001) Phosphoenolpyruvate carboxykinase from the marine diatom Skeletonema costatum and the phaeophyte Laminaria setchellii. II. Immunological characterization and subcellular localization. Bot Marina 44: 199–207.
- 24. Wittpoth C, Kroth PG, Weyrauch K, Kowallik KV, Strotmann H (1998) Functional characterization of isolated plastids from two marine diatoms. Planta 206: 79–85.
- 25. Edwards GE, Franceschi VR, Voznesenskaya EV (2004) Single-cell C(4) photosynthesis versus the dual-cell (Kranz) paradigm. Annu Rev Plant Biol 55: 173–196.
- 26. Long JJ, Wang JL, Berry JO (1994) Cloning and analysis of the C4 photosynthetic NAD-depedent malic enzyme of Amaranth mitochondria. J Biol Chem 269: 2827–2833.
- 27. Burnell JN, Hatch MD (1988) Photosynthesis in phosphoenolpyruvate carboxykinase-type C4 plants: pathways of C4 acid decarboxylation in bundle sheath cells of Urochloa panicoides. Arch Biochem Biophys 260: 187–199.
- 28. Kanai R, Edwards GE (1999) The biochemistry of C4 plants. In: Sage RF, Monson RK, editors. C4 plant biology. London, UK: Academic Press. pp. 49–87.
- 29. von Caemmerer S, Furbank RT (2003) The C(4) pathway: an efficient CO2 pump. Photosynth Res 77: 191–207.
- 30. Lee RE, Kugrens P (1998) Hypothesis: The ecological advantage of chloroplast ER - The ability to outcompete at low dissolved CO2 concentrations. Protist 149: 341–345.
- 31. Kaplan A, Reinhold L (1999) The CO2-concentrating mechanism of photosynthetic microorganisms. Annu Rev Plant Physiol Plant Mol Biol 50: 539–570.
- 32. Ogawa T, Kaplan A (2003) Inorganic carbon acquisition systems in cyanobacteria. Photosynthesis Res 77: 105–115.
- 33. Badger MR, Price GD, Long BM, Woodger FJ (2006) The environmental plasticity and ecological genomics of the cyanobacterial CO2 concentrating mechanism. J Exp Bot 57: 249–65.
- 34. Drechsler Z, Sharkia R, Cabantchik ZI, Beer S (1993) Bicarbonate uptake in the marine macroalga Ulva sp. is inhibited by classical probes of anion exchange by red blood cells. Planta 191: 34–40.
- 35. Park H, Song B, Morel FM (2007) Diversity of the cadmium-containing carbonic anhydrase in marine diatoms and natural waters. Environ Microb 9: 403–413.
- 36. Satoh D, Hiraoka Y, Colman B, Matsuda Y (2001) Physiological and molecular biological characterization of intracellular carbonic anhydrase from the marine diatom Phaeodactylum tricornutum. Plant Physiol 126: 1459–1470.
- 37. Tanaka Y, Nakatsuma D, Harada H, Ishida M, Matsuda Y (2005) Localization of soluble beta-carbonic anhydrase in the marine diatom Phaeodactylum tricornutum. Sorting to the chloroplast and cluster formation on the girdle lamellae. Plant Physiol 138: 207–17.
- 38. Szabo E, Colman B (2007) Isolation and characterization of carbonic anhydrases from the marine diatom Phaeodactylum tricornutum. Physiol Plantarum 129: 484–492.
- 39. Harada H, Nakatsuma D, Ishida M, Matsuda Y (2005) Regulation of the expression of intracellular beta-carbonic anhydrase in response to CO2 and light in the marine diatom Phaeodactylum tricornutum. Plant Physiol 139: 1041–1050.
- 40. Mitra M, Lato SM, Ynalvez RA, Xiao Y, Moroney JV (2004) Identification of a new chloroplast carbonic anhydrase in Chlamydomonas reinhardtii. Plant Physiol 135: 173–182.
- 41. Harada H, Nakajima K, Sakaue K, Matsuda Y (2006) CO2 sensing at ocean surface mediated by cAMP in a marine diatom. Plant Physiol 142: 1318–1328.
- 42. Harada H, Matsuda Y (2005) Identification and characterization of a new carbonic anhydrase in the marine diatom Phaeodactylum tricornutum. Can J Bot 83: 909–16.
- 43. Tortell PD, Reinfelder JR, Morel FM (1997) Active uptake of bicarbonate by diatoms. Nature (London) 390: 243–244.
- 44. Igamberdiev AU, Bykova NV, Lea PJ, Gardenstrom P (2001) The role of photorespiration in redox and energy balanc of photosynthetic plant cells: a study with a barley mutant deficient in glycine decarboxylase. Physiol Plantarum 111: 427–438.
- 45. Park YI, Chow WS, Osmond CB, Anderson JM (1996) Electron transport to oxygen mitigates against photoinactivation of Photosystem II in vivo. Photosynthesis Res 50: 23–32.
- 46. Wu J, Neimanis S, Heber U (1991) Photorespiration is more effective than the Mehler reaction in protecting the photosynthetic apparatus against photoinhibition. Bot Acta 104: 283–291.
- 47. Wingler A, Lea PJ, Quick WP, Leegood RC (2000) Photorespiration: metabolic pathways and their role in stress protection. Phil Trans R Soc Lond B 355: 1517–1529.
- 48. Yamaguchi K, Nishimura M (2000) Reduction to below threshold levels of glycolate oxidase activities in transgenic tobacco enhances photoinhibition during irradiation. Plant Cell Physiol 41: 1397–1406.
- 49. Kozaki A, Takeba G (1996) Photorespiration protects C3 plants from photooxidation. Nature 384: 557–560.
- 50. Tortell PD (2000) Evolutionary and ecological perspectives on carbon acquisition in phytoplankton. Limnol Oceanogr 45: 744–750.
- 51. Paul JS, Volcani BE (1976) Photorespiration in diatoms. 4. 2 pathways of glycolate metabolism in synchronized cultures of Cylindrotheca fusiformis. Arch Microbiol 110: 247–252.
- 52. Colman B, Rotatore C (1988) Uptake and accumulation of inorganic carbon by a freshwater diatom. J Exp Bot 39: 1025–1032.
- 53. Beardall J (1989) Photosynthesis and photorespiration in marine phytoplankton. Aquat Bot 34: 105–130.
- 54. Beardall J, Raven JA (1990) Pathways and mechanisms of respiration in microalgae. Mar Microb Food Webs 4: 7–30.
- 55. Husic DW, Husic HD, Tolbert NE (1987) The oxidative photosynthetic carbon cycle or C2 cycle. CRC Crit Rev Plant Sci 5: 45–100.
- 56. Norman EG, Colman B (1991) Purification and characterization of phosphoglycolate phosphatase from the cyanobacterium Coccochloris peniocystis. Plant Physiol 95: 693–698.
- 57. Tolbert NE (1997) The C-2 oxidative photosynthetic carbon cycle. Ann Rev Plant Physiol Plant Mol Biol 48: 1–23.
- 58. Tolbert NE (1980) Photorespiration. In: Stumpf P, Conn E, editors. The Biochemistry of Plants. Academic NY. pp. 2, 488–525.
- 59. Merrett MJ, Lord JM (1973) Glycolate formation and metabolism by algae. New Phytol 72: 751–767.
- 60. Leboulanger C, Martin-Jézéquel V, Descolas-Gros C, Sciandra A, Jupin HJ (1998) Photorespiration in continuous culture of Dunaliella tertiolecta (Chlorophyta): relationships between serine, glycine, and extracellular glycolate. J Phycol 34: 651–654.
- 61. Kaplan A, Berry JA (1981) Glycolate excretion and the O2/CO2 net exchange ratio during photosynthesis in Chlamydomonas reinhardtii. Plant Physiol 67: 229–232.
- 62. Fogg GE (1983) The ecological significance of extracellular products of phytoplankton photosynthesis. Botanica Marina 26: 3–14.
- 63. Parker MS, Armbrust EV, Piovia-Scott J, Keil RG (2004) Induction of photorespiration by light in the centric diatom Thalassiosira weissflogii (Bacillariophyceae): Molecular characterization and physiological consequences. J Phycol 40: 557–567.
- 64. Leboulanger C, Oriol L, Jupin H, Descolas-Gros C (1997) Diel variability of glycolate in the eastern tropical Atlantic Ocean. Deep Sea Res 44: 2131–2139.
- 65. Eisenhut M, Hasse D, Kahlon S, Lieman-Hurwitz J, Ogawa T, et al. (2006) The plant-type C2 glycolate pathway and the bacterial-like glycerate cycle cooperate in the metabolism of phosphoglycolate in cyanobacteria. Plant Physiol 142: 333–342.
- 66. Parker MS, Armbrust EV (2005) Synergistic effects of light, temperature, and nitrogen source on transcription of genes for carbon and nitrogen metabolism in the centric diatom Thalassiosira pseudonana (Bacillariophyceae). J Phycol 41: 1142–1153.
- 67. Winkler U, Stabenau H (1995) Isolation and characterization of peroxisomes from diatoms. Planta 195: 403–407.
- 68. Suzuki K, Iwamoto K, Yokoyama S, Ikawa T (1991) Glycolate-oxidizing enzymes in algae. J Phycol 27: 492–498.
- 69. Noctor G, Foyer CH (1998) Ascorbate and glutathione: Keeping active oxygen under control. Annu Rev Plant Physiol 49: 249–279.
- 70. Raven JA, Beardall J Platt , editor. (1981) Respiration and photorespiration.Physiological Bases of Phytoplankton Ecology. Can Bull Fish Aquat Sci 210: 55–82.
- 71. Martin W, Scheibe R, Schnarrenberger C (2000) The Calvin cycle and its regulation. In: Leegood RC, Sharkey TD, von Caemmerer S, editors. Photosynthesis: Physiology and Metabolism. Dordrecht: Kluwer Academic Publishers. pp. 9–51.
- 72. Sun N, Ma L, Pan D, Zhao H, Deng XW (2003) Evaluation of light regulatory potential of Calvin cycle steps based on large-scale gene expression profiling data. Plant Mol Biol 53: 467–478.
- 73. Fey V, Wagner R, Bräutigam K, Pfannschmidt T (2005) Photosynthetic redox control of nuclear gene expression. J Exp Bot 56: 1491–1498.
- 74. Ruelland E, Miginiac MM (1999) Regulation of chloroplast enzyme activities by thioredoxins: activation or relief from inhibition? Trends Plant Science 4: 136–141.
- 75. Jacquot J-P, Lancelin J-M, Meyer Y (1997) Thioredoxins: structure and function in plant cells. New Phytol 136: 543–570.
- 76. Oudot-Le Secq M-P, Grimwood J, Shapiro H, Armbrust EV, Bowler C, et al. (2007) Chloroplast genomes of the diatoms Phaeodactylum tricornutum and Thalassiosira pseudonana: comparison with other plastid genomes of the red lineage. Mol Gen Genomics 277: 427–439.
- 77. Tamoi M, Ishikawa T, Takeda T, Shigeoka S (1996) Molecular characterization and resistance to hydrogen peroxide of two fructose-1,6-bisphosphatases from Synechococcus PCC 7942. Arch Biochem Biophys 334: 27–36.
- 78. Kroth PG, Schroers Y, Kilian O (2005) The peculiar distribution of class I and class II aldolases in diatoms and in red algae. Curr Genet 48: 389–400.
- 79. Michelet L, Zaffagnini M, Marchand C, Collin V, Decottignies P, et al. (2005) Glutathionylation of chloroplast thioredoxin f is a redox signaling mechanism in plants. Proc Natl Acad Sci USA 102: 16478–16483.
- 80. Liaud MF, Lichtle C, Apt K, Martin W, Cerff R (2000) Compartment-specific isoforms of TPI and GAPDH are imported into diatom mitochondria as a fusion protein: Evidence in favor of a mitochondrial origin of the eukaryotic glycolytic pathway. Mol Biol Evol 17: 213–223.
- 81. Scheibe R, Backhausen JE, Emmerlich V, Holtgrefe S (2005) Strategies to maintain redox homeostasis during photosynthesis under changing conditions. J Exp Bot 56: 1481–1489.
- 82. Geigenberger P, Kolbe A, Tiessen A (2005) Redox regulation of carbon storage and partitioning in response to light and sugars. J Exp Bot 56: 1469–1479.
- 83. Zhang N, Portis AR (1999) Mechanism of light regulation of Rubisco: A specific role for the larger Rubisco activase isoform involving reductive activation by thioredoxin-f. Proc Natl Acad Sci USA 96: 9438–9443.
- 84. Wedel N, Soll J (1998) Evolutionary conserved light regulation of Calvin cycle activity by NADPH-mediated reversible phosphoribulokinase/CP12/glyceraldehyde-3-phosphatedehydrogenase complex dissociation. Proc Natl Acad Sci USA 95: 9699–9704.
- 85. Graciet E, Lebreton S, Camadro JM, Gontero B (2002) Thermodynamic analysis of the emergence of new regulatory properties in a phosphoribulokinase-glyceraldehyde 3-phosphate dehydrogenase complex. J Biol Chem 277: 12697–12702.
- 86. Harper JT, Keeling PJ (2003) Nucleus-encoded, plastid-targeted glyceraldehyde-3-phosphate dehydrogenase (GAPDH) indicates a single origin for chromalveolate plastids. Mol Biol Evol 20: 1730–1735.
- 87. Grauvogel C, Brinkmann H, Petersen J (2007) Evolution of the glucose-6-phosphate isomerase: the plasticity of primary metabolism in photosynthetic eukaryotes. Mol Biol Evol 24: 1611–1621.
- 88. Huppe HC, Turpin DH (1994) Integration of Carbon and Nitrogen Metabolism in Plant and Algal Cells. Ann Rev Plant Physiol Plant Mol Biol 45: 577–607.
- 89. Cárdenas ML, Cornish-Brown A, Ureta T (1998) Evolution and regulatory role of the hexokinases. Biochim Biophys Acta 140: 242–264.
- 90. Handa N (1969) Carbohydrate metabolism in the marine diatom Skeletonema costatum. Mar Biol 4: 208–14.
- 91. Hama J, Handa N (1992) Diel variation of water-extractable carbohydrate composition of natural phytoplankton populations in Kinu-ura Bay. J Exp Mar Biol Ecol 162: 159–176.
- 92. Van Oijen T, Van Leuwe MA, Gieskes WWC, De Baar HJW (2004) Effects of iron limitation on photosynthesis and carbohydrate metabolism in the Antarctic diatom Chaetoceros brevis (Bacillariophyceae). Eur J Phycol 39: 161–171.
- 93. Vårum KM, Myklestad S (1984) Effects of light, salinity and nutrient limitation on the production of β-1,3-D-Glucan and Exo-D-Glunanase activity in Skeletonema costatum (Grev.) Cleve. J Exp Mar Biol Ecol 83: 13–25.
- 94. Vårum KM, Østgaard K, Grimsrud K (1986) Diurnal rhythms in carbohydrate metabolism of the marine diatom Skeletonema costatum (Grev.) Cleve. J Exp Mar Biol Ecol 102: 249–256.
- 95. Granum E, Kirkvold S, Myklestad SM (2002) Cellular and extracellular production of carbohydrates and amino acids by the marine diatom Skeletonema costatum: diel variations and effects of N depletion. Mar Ecol Prog Ser 242: 83–94.
- 96. Ford CW, Percival E (1965) The carbohydrates of Phaeodactylum tricornutum. Part I. Preliminary examination of the organism, and characterization of low molecular weight material and of a glucan. J Chem Soc 7035–7041.
- 97. Smestad Paulsen B, Myklestad H (1978) Structural studies of the reserve glucan produced by the marine diatom Skeletonema costatum (Grev.) Cleve. Carbohydr Res 62: 386–388.
- 98. McConville MJ, Bacic A, Clarke AE (1986) Structural studies of chrysolaminaran from the ice diatom Stauroneis amphioxys (Gregory). Carbohydr Res 153: 330–333.
- 99. Chiovitti A, Molino P, Crawford SA, Teng R, Spurck T, et al. (2004) The glucans extracted with warm water from diatoms are mainly derived from intracellular chrysolaminaran and not extracellular polysaccharides. Eur J Phycol 39: 117–128.
- 100. Alexseeva SA, Shevchenko NM, Kusaykin MI, Ponomorenko LP, Isakov VV, et al. (2005) Polysaccharides of diatoms occurring in Lake Baikal. Appl Biochem Microbiol 41: 185–191.
- 101. Størseth TR, Hansen K, Reitan KI, Skjermo J (2005) Structural characterization of β-D-(1→3)-glucans from different growth phases of the marine diatoms Chaetoceros mülleri and Thalassiosira weissflogii. Carbohydr Res 340: 1159–1164.
- 102. Størseth TR, Kirkvold S, Skjermo J, Reitan KI (2006) A branched β-D-(1→3,1→6)-glucan from the marine diatom Chaetoceros mülleri characterized by NMR. Carbohydr Res 341: 2108–2114.
- 103. Waterkeyn L, Bienfait A (1987) Localisation et role des β-1,3-glucanes (callose et chrysolaminarine) dans le genre Pinnularia (Diatomées). La Cellule 74: 198–226.
- 104. Roessler PG (1987) UDP-Glucose Pyrophosphorylase activity in the diatom Cyclotella cryptica. Pathway of Chrysolaminarin synthesis. J Phycol 23: 494–498.
- 105. Rayner JC, Pelham HR (1997) Transmembrane domain-dependent sorting of proteins to the ER and plasma membrane in yeast. EMBO J 16: 1832–1841.
- 106. Wiese A, Gröner F, Sonnewald U, Deppner H, Lerchl J, et al. (1999) Spinach hexokinase I is located in the outer envelope membrane of plastids. FEBS Lett 461: 13–18.
- 107. Weber AP, Schneidereit J, Voll LM (2004) Using mutants to probe the in vivo function of plastid envelope membrane metabolite transporters. J Exp Bot 55: 1231–1244.
- 108. Roach PJ (2002) Glycogen and its metabolism. Curr Mol Med 2: 101–120.
- 109. Ball SG, Morell MK (2003) From bacterial glycogen to starch: understanding the biogenesis of the plant starch granule. Annu Rev Plant Biol 54: 207–233.
- 110. Pilkis SJ, Weber IT, Harrison RW, Bell GI (1994) Glucokinase: structural analysis of a protein involved in susceptibility to diabetes. J Biological Chem 269: 21925–21928.
- 111. Sugiyama J, Boisset C, Hashimoto M, Watanabe T (1999) Molecular directionality of β-chitin biosynthesis. J Mol Biol 286: 247–255.
- 112. Tsekos I (1999) The sites of cellulose synthesis in algae: diversity and evolution of cellulose-synthesizing enzyme complexes. J Phycol 35: 635–655.
- 113. Brett CT (2000) Cellulose microfibrils in plants: biosynthesis, deposition, and integration into the cell wall. In: Jeon KW, editor. International Review of Cytology: a Survey of Cell Biology, Volume 199. San Diego, USA: Academic Press. pp. 161–199.
- 114. Lesage G, Bussey H (2006) Cell wall assembly in Saccharomyces cerevisiae. Microbiol Mol Biol Rev 70: 317–343.
- 115. Montijn RC, Vink E, Müller WH, Verkleij AJ, van den Ende H, et al. (1999) Localization of synthesis b1,6-glucan in Saccharomyces cervisiae. J Bacteriology 181: 7414–7420.
- 116. Drew ED (1984) Physiology and metabolism of cyclitols. In: Lewis DH, editor. Storage Carbohydrates in Vascular Plants. Cambridge: Cambridge University Press. pp. 133–155.
- 117. Loewus FA, Loewus MW (1983) Myo-inositol: its biosynthesis and metabolism. Annual Review of Plant Physiology 34: 137–161.
- 118. Stein R, Schnarrenberger C, Gross W (1997) Myo-Inositol dehydrogenase from acido- and thermophilic red alga Galdieria sulphuraria. Phytochemistry 46: 17–20.
- 119. Gross W, Meyer A (2003) Distribution of myo-inositol dehydrogenase in algae. Eur J Phycol 38: 191–194.
- 120. Hipps PP, Eveland MR, Laird MH, Sherman WR (1976) The identification of myo-inositol:NAD(P)+ oxidoreductase in mammalian brain. Biochim Biophys Res Commun 68: 1133–1138.
- 121. Yoshida KI, Aoyama D, Ishio I, Shibayama T, Fujita Y (1997) Organization and transcription of the myo-inositol operon, iol, of Bacillus subtilis. J Bacteriol 179: 4591–4598.
- 122. Lavaud J (2007) Fast Regulation of Photosynthesis in Diatoms: Mechanisms, Evolution and Ecophysiology. Funct Plant Sci Biotechnol 1: 267–287.
- 123. Merchant SS, Prochnik SE, Vallon O, Harris E, Karpowicz SJ, et al. (2007) The Chlamydomonas genome reveals the evolution of key animal and plant functions. Science 318: 245–251.
- 124. Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, et al. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402.
- 125. Matsuzaki M, Misumi O, Shin-i T, Maruyama S, Takahara M, et al. (2004) Genome sequence of the ultrasmall unicellular red alga Cyanidioschyzon merolae 10D. Nature 428: 653–657.
- 126. Weber APM, Oesterhelt C, Gross W, Bräutigam A, Imboden LA, et al. (2004) EST-analysis of the thermo-acidophilic red alga Galdieria sulphararia reveals potential for lipid A biosynthesis and unveils the pathway of carbon export from rhodoplasts. Plant Mol Biol 55: 17–32.
- 127. Barbier G, Oesterhelt C, Larson MD, Halgren RG, Wilkerson C, et al. (2005) Comparative Genomics of Two Closely Related Unicellular Thermo-Acidophilic Red Algae, Galdieria sulphuraria and Cyanidioschyzon merolae, Reveals the Molecular Basis of the Metabolic Flexibility of Galdieria sulphuraria and Significant Differences in Carbohydrate Metabolism of Both Algae. Plant Physiol 137: 460–474.
- 128. Bendtsen JD, Nielsen H, von Heijne G, Brunak S (2004) Improved prediction of signal peptides: SignalP 3.0. J Mol Biol 340: 783–95.
- 129. Nielsen H, Engelbrecht J, Brunak S, von Heijne G (1997) A neural network method for identification of prokaryotic and eukaryotic signal peptides and prediction of their cleavage sites. Int J Neural Syst 8: 581–99.
- 130. Nielsen H, Krogh A (1998) Prediction of signal peptides and signal anchors by a hidden Markov model. Proc Int Conf Intell Syst Mol Biol 6: 122–30.
- 131. Emanuelsson O, Nielsen H, von Heijne G (1999) ChloroP, a neural network-based method for predicting chloroplast transit peptides and their cleavage sites. Protein Sci 8: 978–84.
- 132. Marchler-Bauer A, Anderson JB, Cherukuri PF, DeWeese-Scott C, Geer LY, et al. (2005) CDD: a Conserved Domain Database for protein classification. Nucl Acids Res 33: D192–196.
- 133. Emanuelsson O, Brunak S, von Heijne G (2000) Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J Mol Biol 300: 1005–1016.
- 134. Emanuelsson O, Brunak S, von Heijne G, Nielsen H (2007) Locating proteins in the cell using TargetP, SignalP and related tools. Nature Protocols 2: 953–971.