PDF

Document technical information

Format pdf
Size 507.9 kB
First found May 22, 2018

Document content analysis

Category Also themed
Language
English
Type
not defined
Concepts
no text concepts found

Persons

Organizations

Places

Transcript

A Model for Carbohydrate Metabolism in the Diatom
Phaeodactylum tricornutum Deduced from Comparative
Whole Genome Analysis
Peter G. Kroth1.*, Anthony Chiovitti2., Ansgar Gruber1., Veronique Martin-Jezequel3., Thomas Mock4., Micaela Schnitzler Parker4., Michele S.
Stanley5., Aaron Kaplan6, Lise Caron7, Till Weber1, Uma Maheswari8,9, E. Virginia Armbrust4, Chris Bowler8,9
1 Fachbereich Biologie, University of Konstanz, Konstanz, Germany, 2 School of Botany, University of Melbourne, Melbourne, Victoria, Australia, 3 EA
2663, Faculty of Science, University of Nantes, Nantes, France, 4 School of Oceanography, University of Washington, Seattle, Washington, United
States of America, 5 Scottish Association of Marine Science, Dunstaffnage Marine Laboratory, Oban, Argyll, United Kingdom, 6 Department of Plants
and Environmental Sciences, The Hebrew University of Jerusalem, Jerusalem, Israel, 7 U533, INSERM, Faculty of Medecine, University of Nantes,
France, 8 Centre National de la Recherche Scientifique (CNRS), UMR8186, Ecole Normale Supérieure, Paris, France, 9 Cell Signalling Laboratory,
Stazione Zoologica, Villa Comunale, Naples, Italy
Background. 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.
Methodology/Principal Findings. 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 b-1,3-glucan) outside of the
plastids. We identified various b-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. Conclusions/
Significance. 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
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).
INTRODUCTION
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 [1].
Recent assessments suggest that diatom-mediated export production can influence climate change through uptake and sequestration of atmospheric CO2 [2,3]. 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
Academic Editor: Juergen Kroymann, Max Planck Institute for Chemical Ecology,
Germany
Competing Interests: The authors have declared that no competing interests
exist.
Received July 13, 2007; Accepted December 11, 2007; Published January 9, 2008
Copyright: ß 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.
PLoS ONE | www.plosone.org
* To whom correspondence should be addressed. E-mail: [email protected]
. These authors contributed equally to this work.
1
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
the localization of enzymes and pathways involved in carbon
assimilation and carbohydrate production and catabolism.
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 [4]. For example
the exact mode of CO2 fixation is largely unsolved. Ribulose-1,5bisphosphate carboxylase/oxygenases (Rubisco) from diatoms have
half-saturation constants for CO2 of 30–60 mM [5] despite the fact
that typical sea water contains about 10 mM CO2 [6]. To prevent
potential CO2 limitation, most diatoms have developed mechanisms
to concentrate dissolved inorganic carbon (DIC) via a CO2
concentrating mechanism (CCM) [7]. 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
[8]. Furthermore, some metabolic pathways appear to be missing
altogether from diatoms [9]. 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 [10] and of expressed sequence tags (ESTs) from the
pennate diatom Phaeodactylum tricornutum [11]. 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 [12–14] and various laboratory-based studies of their physiology [8], 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 nonphotosynthetic eukaryote that domesticated a photoautotrophic
eukaryotic cell phylogenetically close to a red alga [15]. 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
[16,17]. 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 59-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
PLoS ONE | www.plosone.org
RESULTS AND DISCUSSION
Structure of the genome and gene annotation
Following publication of the draft Thalassiosira pseudonana Hasle &
Heimdal (CCMP 1335) genome [10], 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 [16]. Diatom plastids are surrounded by
four membranes, the outermost being studded with ribosomes and
continuous with the endoplasmic reticulum (ER) [18]. 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 [16,17].
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 [5]. Extensive diatom blooms
that occur during large iron fertilization experiments in high
nutrient low chlorophyll regions of the oceans [19,20] 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 [3,4]).
Studies on the biochemistry of photosynthesis in the wellcharacterized marine diatom Thalassiosira weissflogii suggested that a
C4-like pathway could exist whereby a C4 compound such as malate
2
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
or OAA is decarboxylated, typically within the chloroplast, to deliver
CO2 to Rubisco [21,22]. The possibility of a C4-like pathway in the
related species T. pseudonana was examined based on an in silico
analysis of gene content [10]. 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 C4related 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 [3]. Despite the presence of typical C4
enzymes in both Thalassiosira species, short 14CO2 labelling
experiments showed marked differences between them [4]. 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. [4] 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 [17]. 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 mitochondriallocalized 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).
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 HCO32 into cells either via specific transporters or by
diffusion of CO2 and its subsequent conversion to HCO32
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 HCO32 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
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
PLoS ONE | www.plosone.org
3
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
decarboxylate PEP inside the mitochondria (see [25,28–29]).
However, most C4-plants carry out the first carboxylation step in
the cytosol [29]. 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 [30] 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 [7,31–33]. 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 Cl2/HCO32 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 HCO32 uptake as shown in Ulva sp. [34], should
inhibit uptake of HCO32 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 [35]. 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 [36], one of these proteins (51305) is located in the
plastid as has been demonstrated by GFP fusion proteins [37]. 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 [38] 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
costatum [23]. Later, Reinfelder et al. [21] found that PEPCK
activity co-localized with Rubisco activity in isolated plastidenriched 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
[24] 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 [25,26]. 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 HCO32)
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. [3]. A potential NADdependent 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 [25], B.
cycloptera and B. aralocaspica [25]. 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
HCO32 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 HCO32 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 [27,28]. 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 NADmalic enzyme, can be used for amino acid synthesis (Fig. 1) [28] 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 HCO32 fixation by PEPC
(Fig. 1). This pathway is also known from C4 plants that
PLoS ONE | www.plosone.org
4
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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 [39].
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 HCO32 to
CO2, thereby raising its concentration in close proximity to
Rubisco [40]. Mutants impaired in this CA demand high CO2
concentrations for growth (see [7,31]). In P. tricornutum, the plastidlocalized 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 [37], and its expression
is strongly enhanced under low CO2 conditions by a mechanism
involving cAMP [41]. 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 [42]. 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 HCO32 fixation
occurs outside the plastid. Finally, enhanced uptake of inorganic
carbon, both as CO2 or HCO32, would be consistent with both
types of CCM, whereas raising the concentration of Ci within the
cells [43] is more consistent with the biophysical CCM. The high
affinity of PEPC for HCO32 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.
Thus, out of four carbons entering the C2 cycle, three are
converted back to PGA and one is released in the form of CO2.
In microalgae, the cyclic process of PGA recovery is not wellstudied. More often, glycolate metabolism is studied in the context
of its excretion as a waste-product to circumvent unfavourable
growth conditions [57–60]. A large fraction of glycolate produced
by fixation of O2 is released from the cell [61–63] and may serve as
an important source of organic carbon in the water body.
Leboulanger et al. [64] 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
[65]. 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
[5]. 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 [63,66]. 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 [67]. 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 [68]. Winkler and Stabenau
[67] 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 [69].
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. [68] and
Winkler and Stabenau [67], 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
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 [44–47]. Mutants of tobacco
(Nicotiana tabacum L.) defective in enzymes of the photorespiratory
pathway demonstrated enhanced photoinhibition under high light
conditions [48,49]. The specificity factor (t) 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
[50]) 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 [51–54].
When O2 out-competes CO2 for RuBP, one molecule of 2-Pglycolate 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 [55] must be degraded via the photorespiratory
pathway (see Fig. 2). In higher plants, metabolism of 2-P-glycolate
takes place via the C2 cycle [56]. 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 Pglycerate which may then enter the Calvin cycle in the chloroplast.
PLoS ONE | www.plosone.org
5
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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
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 [51] 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.
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 [70]).
Another pathway for glyoxylate metabolism, the tartronate
semialdehyde pathway, is known in cyanobacteria [70]. 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 [65]. 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
PLoS ONE | www.plosone.org
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 [71]. Carbon fixation in
6
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
The only Calvin cycle enzymes encoded on the plastid genome
are the small and the large subunit of the Rubisco (GenBank
AY819643) [76]. 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 [77]. 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
land plant plastids is highly regulated, either by substrates and ions
like Mg2+ or by light-dependent redox regulation either at the
transcriptional [72,73] or the enzymatic level via the ferredoxin/
thioredoxin-system [74,75]. 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 pentosephosphates 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 [9,10] 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
PLoS ONE | www.plosone.org
7
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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 [84]. 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 [85]. 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-phosphate dehydrogenase (TIM-GAPC3, 25308) [80], 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 [80] (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.
diatoms by secondary endocytobiosis [15,75]. 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 [78].
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
[75]. 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 [8]).
Another group of proteins involved in redox-regulation in land
plant plastids are glutaredoxins (Glrx), which are involved in finetuning of the thioredoxin system [79]. 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 [9]. 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 [9]. 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 [80] 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 c 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 redoxregulated via thioredoxin and serves as a valve for excess NADPH
[81]. 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 [82]. 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 [83], is apparently also
PLoS ONE | www.plosone.org
Glycolysis
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 UDPGlucose-Pyrophosphorylase/Phosphoglucomutase fusion 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) [80]. GapC2, assigned to be cytosolic [80] 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 [87]
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
8
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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 [88].
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 [89]. The P. tricornutum cytosolic glucokinase (48774) might additionally be involved in the chrysolaminaran pathways (see below).
accumulating during the daylight and becoming depleted in the
dark [90,94,95]. The structure of chrysolaminaran is fundamentally based on a b-1,3-linked glucan backbone, which is
infrequently branched with mainly b-1,6-linkages [96–102].
Vacuolar localization of chrysolaminaran in several diatom
species, including P. tricornutum and T. pseudonana, was demonstrated by staining with aniline blue [103] and by immunolabeling with
a monoclonal anti-1,3-b-D-glucan antibody [99].
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 [104]. Furthermore, exo-1,3-b-glucanase activity was detected in several planktonic
diatoms and upregulation of this activity coincided with chrysolaminaran degradation in the diatom Skeletonema costatum [94].
We focused on exo- and endo-1,3-b-glucanases and bglucosidases as the primary enzymes involved in digesting
chrysolaminaran. We found four putative exo-1,3-b-glucanases
in P. tricornutum, all belonging to the glycosyl hydrolase family 16
(49294, 56510, 56506, 49610). All orthologues possess an Nterminal signal peptide, except (49610), and all contain a Cterminal transmembrane helix. In addition, one (56510) possesses
a putative C-terminal ER-retention signal (REEL). Of the four
exo-1,3-b-glucanases, only one (49294) was represented in T.
pseudonana (13556). Three putative endo-1,3-b-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
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 [90–92]. 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 [93]. Chrysolaminaran concentrations undergo a diel
rhythm characteristic of an assimilatory and respiratory product,
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
PLoS ONE | www.plosone.org
9
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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 [108,109]. 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; [99–102]) compared with broadspecificity eukaryotic hexokinases (KM = 0.007–2.5 mM; [89,110].
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, UDPglucose likely serves as the substrate for chrysolaminaran synthesis
[104]. 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 [104]. 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 membranebound 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-b-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-b-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-b-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 [10,111] and the cellulose synthases of
terrestrial plants and multicellular algae [112,113]. Extracellular
callose has been reported in diatoms and was suggested to serve as a
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-b-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 b-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 bglucosidase 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 b-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-b-glucans were
identified in P. tricornutum. Presumably, at least one of the exo-1,3-bglucanases and one of the endo-1,3-b-glucanases act complementarily to digest the principle b-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 b-1,6-linkages (e.g. gentiobiose)
derived from surviving chrysolaminaran branch points. A bglucosidase 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 exo1,3-b-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
[105]. 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 [106]. 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 Cterminal transmembrane helix [107]. 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 b-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 b-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
PLoS ONE | www.plosone.org
10
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
permeable seal in the girdle regions during cell division [103], so it is
feasible that the identified 1,3-b-glucan synthase is a plasmamembrane-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-b-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 1,6b-glucans [114]. 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 bglucans. 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
[115]. Branching in terrestrial plant starches and mammalian and
fungal glycogen is achieved by specific enzymes that hydrolyze
internal a-1,4-glycosidic bonds and transfer the released reducing
ends to C-6 hydroxyls of the acceptor polysaccharide chain
[108,109]. If analogous processes occur in diatoms, the putative
diatom glucosylase/transglucosylases could act as chrysolaminaran
branching/debranching enzymes.
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 [119]. Interestingly, neither synthesis nor
catabolism involves scyllo-inosose as a reaction product in the algae
studied, whereas in mammals [120] 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 [119] 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 [121]. Without further
investigation its role in inositol metabolism in both P. tricornutum
and T. pseudonana is unclear.
Inositol and Propanoate pathways
There are elementary differences between green
algae and diatoms
Many different cyclitols occur in plants with the most widespread
and extensively studied being myo-inositol [116,117]. 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 wellrepresented, 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 1.1.1.18). Stein
et al. [118] confirmed the presence of InDH in the red alga Galdieria
sulphuraria and a further study by Gross and Meyer [119] 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
PLoS ONE | www.plosone.org
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 [8,122]). 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 [123] 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 posttranslational regulation of photosynthetic products.
MATERIALS AND METHODS
Sequence analysis
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) [10] and Phaeodactylum tricornutum v2.0 (http://
genome.jgi-psf.org/Phatr2/Phatr2.home.html) using the BLAST
algorithm [124]. Comparison with the genome sequences of the
red algae Cyanidioschyzon merolae (http://merolae.biol.s.u-tokyo.ac.
jp/) [125] and Galdieria sulphuraria (http://genomics.msu.edu/
11
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
galdieria) [126,127] 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/) [128]. 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 [16,17]. We screened for signal peptides using SignalP. For
cleavage site predictions the results of SignalP’s Neuronal networks
(NN) [129] or Hidden Markov Models (HMM) [130] were used.
For prediction of chloroplast transit peptide-like domains, the
program ChloroP (http://www.cbs.dtu.dk/services/ChloroP/)
[131] 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) [132] 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/) [133]. 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 [17]. Proteins which (i)
possess a signal peptide but no ER retention signal (ii) possess a Nterminal 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 [133]. 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 [134].
SUPPORTING INFORMATION
Table S1 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/) [133] and SignalP’s (http://www.cbs.dtu.dk/
services/SignalP/) Neuronal networks (NN) [129] or Hidden
Markov Models (HMM) [130]. For a detailed description of
protein localization prediction see also [134]. The Protein IDs and
genomic coordinates are directly linked to the genomic database.
Found at: doi:10.1371/journal.pone.0001426.s001 (0.31 MB
XLS)
ACKNOWLEDGMENTS
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.
Author Contributions
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.
REFERENCES
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 promoterterminator cassette. FEBS J 272: 3413–3423.
15. Patron NJ, Rogers MB, Keeling PJ (2004) Gene replacement of fructose-1,6bisphosphate 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.
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 NO3depletion 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.
PLoS ONE | www.plosone.org
12
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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, eds (1980) 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 (1981) Respiration and photorespiration. In: Platt, ed
(1981) 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, eds (2000)
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.
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, eds (1999) 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.
PLoS ONE | www.plosone.org
13
January 2008 | Issue 1 | e1426
Diatom Carbon Metabolism
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 b-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, ed (2000) 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, ed
(1984) 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 ThermoAcidophilic 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 networkbased 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.
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-phosphate dehydrogenase 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-6phosphate 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 b-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. pp 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 b-D-(1R3)-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 b-D(1R3,1R6)-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 b-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.
PLoS ONE | www.plosone.org
14
January 2008 | Issue 1 | e1426

Similar documents

×

Report this document