Research
Low CO2 results in a rearrangement of carbon metabolism to
support C4 photosynthetic carbon assimilation in Thalassiosira
pseudonana
Adam B. Kustka1, Allen J. Milligan2, Haiyan Zheng3, Ashley M. New1, Colin Gates1, Kay D. Bidle4 and
John R. Reinfelder5
1
Earth and Environmental Sciences, Rutgers University, 101 Warren Street, Newark, NJ 07102, USA; 2Department of Botany and Plant Pathology, Oregon State University, 2082 Cordley
Hall, Corvallis, OR 97331, USA; 3Biological Mass Spectrometry Facility, Rutgers University, 174 Frelinghuysen Road, Piscataway, NJ 08854, USA; 4Institute of Marine and Coastal Sciences,
Rutgers University, 71 Dudley Road, New Brunswick, NJ 08901, USA; 5Department of Environmental Sciences, Rutgers University, 14 College Farm Road, New Brunswick, NJ 08901, USA
Summary
Author for correspondence:
Adam B. Kustka
Tel: +1 973 353 5509
Email: [email protected]
Received: 28 November 2013
Accepted: 28 May 2014
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doi: 10.1111/nph.12926
Key words: C4 metabolism, fatty acid
metabolism, glycine decarboxylase, marine
diatoms, quantitative proteomics, pentose
phosphate pathway, pyruvate phosphate
dikinase (PPDK), pyruvate carboxylase.
The mechanisms of carbon concentration in marine diatoms are controversial. At low CO2,
decreases in O2 evolution after inhibition of phosphoenolpyruvate carboxylases (PEPCs), and
increases in PEPC transcript abundances, have been interpreted as evidence for a C4 mechanism in Thalassiosira pseudonana, but the ascertainment of which proteins are responsible for
the subsequent decarboxylation and PEP regeneration steps has been elusive.
We evaluated the responses of T. pseudonana to steady-state differences in CO2
availability, as well as to transient shifts to low CO2, by integrated measurements of photosynthetic parameters, transcript abundances and quantitative proteomics.
On shifts to low CO2, two PEPC transcript abundances increased and then declined on
timescales consistent with recoveries of Fv/Fm, non-photochemical quenching (NPQ) and
maximum chlorophyll a-specific carbon fixation (Pmax), but transcripts for archetypical decarboxylation enzymes phosphoenolpyruvate carboxykinase (PEPCK) and malic enzyme (ME)
did not change. Of 3688 protein abundances measured, 39 were up-regulated under low
CO2, including both PEPCs and pyruvate carboxylase (PYC), whereas ME abundance did not
change and PEPCK abundance declined.
We propose a closed-loop biochemical model, whereby T. pseudonana produces and subsequently decarboxylates a C4 acid via PEPC2 and PYC, respectively, regenerates phosphoenolpyruvate (PEP) from pyruvate in a pyruvate phosphate dikinase-independent (but glycine
decarboxylase (GDC)-dependent) manner, and recuperates photorespiratory CO2 as oxaloacetate (OAA).
Introduction
Ambient CO2 levels in seawater are vastly under-saturating for
carboxylase activity by ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO). This leads to the possibility of significant
oxygenase activity, which can compete with carboxylation
(Cooper et al., 1969; Badger et al., 1998), and results in photorespiration at the expense of carbon fixation (Riebesell et al., 1993;
Ogren, 1994). To maintain high rates of photosynthesis with
these constraints, marine diatoms have evolved a CO2-concentrating mechanism, or CCM (Rotatore et al., 1995; Fielding
et al., 1998; Burkhardt et al., 2001; Colman et al., 2002; Tortell
et al., 2002), the underlying biochemistry of which remains
uncertain (Reinfelder et al., 2004; Roberts et al., 2007). CO2
concentrations can be elevated in the proximity of RuBisCO
through means commonly referred to as biophysical or biochemical. The biophysical pathway relies on an increased intracellular
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bicarbonate concentration (through active transport), followed
by conversion to CO2 (by carbonic anhydrase) in the vicinity of
RuBisCO, and is well documented in cyanobacteria and green
algae (Amoroso et al., 1998; Badger et al., 2006). Alternatively,
CO2 can be concentrated biochemically through single-cell C4
photosynthesis (Reinfelder et al., 2000; Roberts et al., 2007;
McGinn & Morel, 2008), although it is important to recognize
that a biochemical CCM does not preclude the utility of pumping bicarbonate into the cell.
In the most general terms, C4 photosynthesis is characterized
by the fixation of bicarbonate and phosphoenolpyruvate (PEP)
by PEP carboxylase (PEPC) to form a C4 compound in one
‘compartment’, followed by transport of the C4 compound or its
derivative to other compartments for subsequent decarboxylation. These steps serve to fix CO2 and subsequently decarboxylate a C4 acid in close proximity to RuBisCO, and are segregated
to avoid futile cycling between C3 and C4 compounds. This
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segregation was first (Hatch & Slack, 1966) and is most often
(Sage, 2004) described among different types of cells in multicellular, vascular plant systems, but is also achieved by organelles
within individual cells of multicellular plants (Sage, 2002;
Voznesenskaya et al., 2002; Edwards et al., 2004; Bowes, 2011).
The C4 pathway originated with the appropriation and regulatory modification of existing carbon metabolism enzymes, often
followed by the selection for kinetic properties more suitable for
C4 metabolism (Svensson et al., 2003; and references therein).
Because there have been multiple and evolutionarily independent
origins for C4 photosynthesis (Sage, 2004; Christin et al., 2008),
some degree of flexibility, in which enzymes have been co-opted
for C4 metabolism, could be expected. In virtually every known
example of C4-assisted photosynthesis, the initial step involves
the carboxylation of PEP to oxaloacetate (OAA) by PEPC
(Fig. 1). A notable exception occurs in the marine macroalga
Udotea flabellum, where the C4 pathway starts with the carboxylation of PEP by phosphoenolpyruvate carboxykinase (PEPCK;
Reiskind & Bowes, 1991). In all known systems that begin with
PEPC-mediated carboxylation, the three known decarboxylation
pathways involve either one of two isoforms of malic enzyme
(NAD-ME or NADP-ME) or PEPCK. In pathways mediated by
a malic enzyme (ME) isoform, OAA is rapidly converted to
malate through malate dehydrogenase (MDH), transported to a
separate compartment and decarboxylated by ME to form pyruvate (PYR) and CO2. The cycling of PYR back to PEP is
Fig. 1 Two abridged models of single-cell C4 photosynthesis. Solid arrow
indicates the first step of C4 photosynthesis mediated by
phosphoenolpyruvate carboxylase (PEPC). The first step (indicated by solid
lines) of almost all known C4 pathways begins with the carboxylation of
phosphoenolpyruvate (PEP) via PEPC to form oxaloacetate (OAA).
Although there are some variations on the identity of the C4 acid (OAA,
malate or aspartate) transported to the organelle (most often chloroplast),
the second step is characterized by one of two archetypical pathways that
involve decarboxylation by malic enzyme (ME; dashed lines) or
phosphoenolpyruvate carboxykinase (PEPCK; dotted lines). In the ME
pathway, OAA is converted to malate by malate dehydrogenase (MDH),
and PEP is regenerated from pyruvate (PYR) by pyruvate phosphate
dikinase (PPDK). Carbonic anhydrase (CA) catalyzes the interconversion
between carbon dioxide and bicarbonate.
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accomplished by pyruvate phosphate dikinase (PPDK) and other
enzymes. In a PEPCK-mediated pathway, OAA is decarboxylated
to PEP and CO2.
Recent research on single-cell C4 photosynthesis in marine diatoms has yielded disparate results. On the one hand, some biochemical and molecular data are suggestive of C4-assisted
photosynthesis in T. pseudonana. For example, O2 evolution is
dramatically reduced by the PEPC inhibitor, 3,3-dichloro-2-dihydroxyphosphinoylmethylpropenoate (DCDP), for both
T. weissflogii and T. pseudonana grown at CO2 concentrations
that are subsaturating with respect to RuBisCO carboxylase
(Reinfelder, 2011), and can be rescued by the addition of
HCO3 (McGinn & Morel, 2008). In addition, McGinn &
Morel (2008) observed c. three-fold greater PEPC transcript
abundances for T. pseudonana under low CO2. However,
although Roberts et al. (2007) showed that malate is among the
first products of carbon fixation in T. weissflogii (supporting the
earlier findings of C4 photosynthesis in this species; Reinfelder
et al., 2000), similar external CO2 conditions resulted in minimal
malate labeling in T. pseudonana. In addition, in contrast with
McGinn & Morel (2008), Roberts and colleagues found similar
PEPC transcript abundances for T. pseudonana grown under low
and high CO2 (380 and 100 ll l1, respectively). These findings
led Roberts and co-workers to conclude that T. pseudonana does
not exhibit C4 photosynthesis.
Carbon-concentrating mechanisms help to minimize photorespiratory losses in vascular plants. In diatoms, low CO2 induces
canonical biomarkers of photorespiration, but this is not manifested in decreases in growth rate as might be expected, raising
questions about both the roles and biochemistry of photorespiration in diatoms. All the required genes for photorespiration, apart
from glycerate kinase, are readily identified in both
T. pseudonana and P. tricornutum genomes (Kroth et al., 2008).
In diel experiments with T. pseudonana, Granum et al. (2009)
observed light-dependent increases in glycine decarboxylase
(GDC) transcript abundances under low (100 ll l1) but not
ambient (360 ll l1) CO2. These transcripts, as well as relative
cellular glycolate concentrations, also increased with increasing
light intensity at ambient CO2 (Parker et al., 2004; Parker &
Armbrust, 2005). Despite these indicators of photorespiration,
manipulations of CO2/O2 stoichiometry that should lead to elevated rates of photorespiration do not lead to discernible changes
in growth rate (Beardall, 1989). These apparent discrepancies
between markers of the photorespiratory pathway and the negligible photorespiratory losses themselves suggest the adoption of
some mechanism to recuperate the terminal products of photorespiration, CO2 and NH4+, or that these marker proteins may
have biochemical functions other than photorespiration at low
CO2. It is important to recognize that these enzymes are essential
at both low and ambient CO2 in other phototrophs (Nakamura
et al., 2005; Eisenhut et al., 2008; Zelitch et al., 2009).
We hypothesized that T. pseudonana, like T. weissflogii, exhibits C4-assisted photosynthesis in response to low CO2. We also
hypothesized that C4 metabolism might not entirely eliminate
oxygenase activity, and explored whether compensatory mechanisms might be at play. These hypotheses were tested by
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evaluating the transcriptional, photo-physiological and proteomic
responses of these diatoms under conditions of carefully controlled CO2 availability. From these data, we develop a new
model for carbon fixation and other physiological responses
under low-CO2 conditions.
Materials and Methods
Culture conditions and photo-physiology on CO2 shift
Thalassiosira pseudonana (Cleve 1873), strain CCMP1335, was
grown in medium prepared from synthetic ocean water according
to the Aquil recipe (Sunda et al., 2005) and maintained at 18°C
and 200 lmol photons m2 s1. Macronutrients were added at
100 lmol l1 NO3, 100 lmol l1 Si(OH)4 and 10 lmol l1
PO43. All medium preparation and sample handling were carried out in a class 100 laminar flow hood. All plastic ware was
acid cleaned by soaking in 10% trace metal grade HCl for 5–10 d
and rinsed with 18.2 MΩ cm deionized water. Media and culture
flasks were microwave sterilized according to Keller (1988) in
polycarbonate bottles. Cell density was determined using a Coulter counter (Beckman-Coulter, Fullerton, CA, USA), and growth
rates were computed from linear regressions of ln(cell density) vs
time. Cells were acclimated to a constant CO2 (1780 ll l1 or
61 lM aqueous CO2) and then shifted to 210 ll l1 (7.1 lM)
CO2 within c. 15 s.
Our pre-shift conditions were designed to provide a saturating
supply of CO2, at about twice the half-saturation concentration for
diatom RuBisCO (c. 30 lM; Badger et al., 1998), to greatly reduce
the demand for a CCM relative to that for ambient CO2 levels.
This was performed by maintaining cultures in house-built pH
stats set at pH 7.61 (National Bureau of Standards scale) for > 10
generations. At the shift, 1 M NaOH was added whilst mixing the
culture bottle to achieve a pH of 8.48. Before and following the
shift, the pH was continuously monitored. CO2 was calculated
using these pH values and the TCO2 (total CO2) content of Aquil
(2.38 mM) using CO2SYS (http://cdiac.ornl.gov/oceans/co2rprt.
html). As we control pCO2 by varying the pH at constant dissolved
inorganic carbon concentrations, these experiments resemble
ocean acidification experiments, except that here the concentration
of aqueous CO2 is decreased, as occurs during bloom progression.
The pH stats were assembled as follows. A gel-filled combination
pH electrode (9106BNWP; Thermo, Waltham, MA, USA) was
mounted through the wall of a 1-l polycarbonate bottle using a
bulkhead mount. The electrode potential was monitored using a
pH relay (pH200 controller; Eutech Inst., Singapore). When the
pH increased above a set threshold, the relay switched on a peristaltic pump (Master Flex C/L; Cole Parmer, Vernon Hills, IL, USA)
and an aquarium air pump to deliver weak acid (0.03 M trace metal
grade HCl) whilst mixing the culture.
Before and following a shift to low-CO2 conditions, samples
were collected for the determination of chlorophyll a (Chl
a)concentration, cell concentration, growth rate and photosynthesis–irradiance (PE) relationships. Samples (20–30 ml) for Chl a
concentration were collected on glass fiber filters (GFF Whatman,
GE Healthcare, Piscataway, NJ, USA). Chl was extracted
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overnight with 90% acetone and determined spectrophotometrically following the methods of Jeffrey & Humphrey (1975). Cell
concentrations and growth rate were determined as above.
The PE relationships for samples were determined from
10 min incubations with 14C bicarbonate in a photosynthetron
(Lewis & Smith, 1983). Ten irradiances (0–1750 lmol photons m2 s1) were used during each incubation, with the resulting data fitted to the equation of Platt & Gallegos (1980) using
non-linear regression:
PB ¼ Ps ð1 e a Þe b ;
Eqn 1
where PB is the photosynthetic rate normalized to Chl a and Ps is
the maximum photosynthetic rate in the absence of photoinhibition. For the exponents of Eqn 1, a = aE/Ps and b = bE/Ps, where
a is the initial slope of the P vs E curve, b is the descending slope
at high light and E is the scalar irradiance. There was no evidence
of photoinhibition in our P vs E data, and so b was set to zero for
fitting purposes.
Variable fluorescence (Fv/Fm = (Fm F0)/Fm) was measured
using a fast-repetition-rate fluorometer (FRRf; Kolber et al.,
1998). Samples were removed from the culture vessel and immediately measured (no dark acclimation period) for F0 (Chl fluorescence yield when the photosystem II, PSII, reaction centers are
mostly oxidized) and Fm (Chl fluorescence yield when all functional PSII reaction centers are reduced). We purposely did not
provide an opportunity for acclimation to capture the state of
variable fluorescence in the culture vessel.
The capacity to induce non-photochemical quenching (NPQ)
was determined in a temperature-controlled (18°C) pulse amplitude-modulated (PAM) fluorometer (DUALPAM-100; Heinz
Walz GmbH, Effeltrich, Germany) fitted with a photomultiplier
detector (DUAL-DPM). NPQ was determined using a PAM
protocol entailing an initial low-light (16 lmol photons m2 s1)
acclimation period of 15 min, followed by a dark period of
5 min. It has been shown that diatoms require a low-light exposure to allow NPQ mechanisms to fully revert to a non-quenching state (Grouneva et al., 2008; Miloslavina et al., 2009;
Milligan et al., 2012). The relaxation period was followed by an
actinic light exposure (300 lmol photons m2 s1) for 10 min to
induce NPQ, and finally a second dark period of 21 min. The
maximum fluorescence (Fm) in the NPQ relaxed state was determined before actinic light exposure and during actinic light exposure (Fm0 ). NPQ was calculated as (Govindjee, 2004):
NPQ ¼ ðFm =Fm0 Þ 1
Eqn 2
Quantitative PCR analysis of transcript abundances on CO2
shift
Cultures were maintained under steady-state high CO2 at
350 lmol photons m2 s1 and subject to CO2 shifts as in the
photo-physiology experiments, except that Aquil (Sunda et al.,
2005) was assembled using coastal seawater rather than synthetic
ocean water. In this case, the high- and low-CO2 pH values of
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7.61 and 8.48 correspond to CO2 values of 184 and 1500 ll l1
(54 and 6.6 lM) based on a coastal seawater total alkalinity of
2.04 mM (18°C, salinity = 25; Lee et al., 2006).
Gene transcript abundances for seven potentially important
enzymes in C4 metabolism were quantified using quantitative
reverse transcription-polymerase chain reaction (Q-RT-PCR
hereafter) over an eight-point time course corresponding to 30,
20, 10, + 10, + 20, + 30, + 60 and + 90 min relative to the
time of shift to low CO2, using actin as the housekeeping gene.
RNA extraction, complementary DNA (cDNA) synthesis and QRT-PCR procedures generally followed those of Kustka et al.
(2007) with some modifications. RNA was extracted using a
modification of the Trizol method; cells were scraped from a 47mm polycarbonate filter into 60–65°C Trizol, passed through a
25-gauge needle four to six times to disperse clumps, frozen in
liquid nitrogen and stored at 80°C for < 30 d. On thawing, solids were cleared via centrifugation and the supernatant was transferred to a new vial. Chloroform extraction was performed, and
the aqueous phase was removed, mixed with 1 : 1 v/v 67% ethanol and loaded onto an ‘Absolutely RNA’ spinkit (StratageneAgilent, La Jolla, CA, USA). To remove genomic DNA, two
sequential 30-min reactions were performed with Turbo-DNAse
(Ambion-Life Technologies, Grand Island, NY, USA). Q-RTPCRs were performed with a Stratagene Mx3000P thermal cycler
using Power SYBR mastermix. Equal amounts of template (either
10 or 25 ng of RNA equivalent) were used in triplicate 20-ll
reactions. The absence of interfering genomic DNA contamination was validated with PCRs containing ‘no reverse transcription’ RNA as template, using identical conditions.
Copy numbers of genes of interest (GOI), relative to that of
actin, were estimated as 2[1(CtGOI – Ctactin)] [shorthand as
2(DCt)] according to Pfaffl (2001). This quantity was determined from replicate analyses for each observation, within each
experiment. Melt curves, inspected with each quantitative PCR
analysis for each reaction well, indicated single peaks with melting temperatures consistent with the absence of primer dimers.
This finding was additionally confirmed by gel electrophoresis
for each primer from one treatment once from each of two biological replicates. Standard templates were generated by gel purification of PCR products using gene-specific primers and
quantified with a NanoDrop 1000 (Wilmington, DE, USA)
spectrophotometer. For each primer pair, real-time PCR efficiency was calculated by plotting Ct vs log(template copy number) from the analysis of a four-point serial dilution spanning a
range of 256-fold template concentration. The slope was used to
calculate the efficiency, Efficiency = 10(1/slope). The average efficiency for all quantitative PCR primers in this study was
1.90 0.023 (SD), close to the theoretical value of 2. The primers used in this study are given in Table 1.
Validation of housekeeping genes and limits on
quantification
The suitability of a housekeeper depends on the lack of a discernible trend in cycle threshold value along gradients of treatments
(or time in this case). The analysis of residual Ct values vs time
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showed no trend (Supporting Information Table S1), suggesting
that this housekeeping gene is suitable for studies of transcript
abundance with shifting CO2. The minimum level of differential
gene expression that can be detected (below which apparent shifts
in transcript abundance may be caused by some small variation
in measured actin transcript) was calculated with a 5% type I
error rate by analyzing a population of standard deviations in
actin Ct from all pairwise combinations of pre-shift time points.
This threshold equaled a standard deviation of 0.26 Ct, which
translates into a 20% copy number difference in actin transcripts
between any two identical samples. For transcripts which exhibited shifts above these copy number difference thresholds (PEPC1
and PEPC2), differences between pre- and post-shift transcript
abundances relative to actin (TRA) were evaluated using a
one-tailed t-test on pooled data from both biological replicates.
Quantitative proteomic analyses at steady state
Cultures were grown under steady-state conditions of low and
high CO2 (210 and 1780 ll l1) in pH stats with Aquil medium,
as described above, except that either low- or high-CO2 cells were
grown with 15N- nitrate (> 98%) or nitrate with a natural isotopic composition (i.e. 99.6% 14N). Two sets of independent biological replicates from each condition were processed. For each
condition, c. 900 ml were harvested at (4–5) 9 105 cells ml1
and flash frozen in liquid nitrogen. Protein samples were
extracted in 4% sodium dodecylsulfate (SDS), 7.5% glycerol, in
0.1 M Na2CO3 and protease inhibitor (Sigma-Aldrich P-2714),
and quantified before the addition of 0.1 mM dithiothreitol
Table 1 Primers used in quantitative PCR. Each gene name is as given in
the text, followed by the protein identification number (in parentheses)
and genome locus (underscored)
Gene name locus forward and reverse primers (50 –30 )
Phosphoenolpyruvate carboxylase (3830) chr_3:1899971–1903147
CAGCGTCTCATGGCAGTTAGAAATC
CTCCGAGCTTCCCTCACTTTCAC
Phosphoenolpyruvate carboxylase (5027) chr_5:200957–204623
GGCGACCCCTGAGTTGGAACTT
CACGGAATAGCTCTCAAACTGTCAACAC
Malate dehydrogenase; cytoplasmic (41425) chr_7:1677378–1678600
CATGGAGAAGCTGCAAGTCACTGA
GCCAACGAAGCAGTGGATACAGTAG
Malate dehydrogenase; mitochondrial (175) chr_1:985790–987134
ATGGCGTATGCTGGCTATGTATTTACAG
GCAAAATCTCCTCTACACCTCCCTTAC
Malic enzyme (5100) chr_5:366187–368316
GCGAAACACTGCGAAAGACCAATC
ACGGGCTGCCACTAGCAAAGATAC
Phosphoenolpyruvate carboxykinase (5186) chr_5:566357–568410
GGTGGGAGCTTCCTGCCTCTATT
AATCGACATCTCCTTTCCCCTTGTAC
Pyruvate phosphate dikinase (5500) chr_5:1377134–1380283
GAGGCTGAGCGCATTAAAGTTGAAC
TGGCTTGCTGTCTTCGTCGTCTA
Actin (25772) chr_22:804575–806436
ACTGGATTGGAGATGGATGG
CAAAGCCGTAATCTCCTTCG
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(DTT). Fifty micrograms of protein from each condition were
mixed, yielding a 1 : 1 ratio of 15N- to 14N-labeled protein mixture. Subsequent processing was carried out at the Biological
Mass Spectrometry Facility of the UMDNJ-Robert Wood Johnson Medical School, New Brunswick, NJ, USA. The protein
mixture was digested by trypsin. Digested peptides were solubilized in buffer A (20 mM ammonium formate, pH10) and subjected to high-pH reverse-phase high-performance liquid
chromatography (HPLC) (Gilson 306 pumps, 805 manometric
module and UV/VIS 155 detector) equipped with a XbridgeTM
C18 column (3.5 lm, 2.1 9 150 mm, Waters, Milford, MA,
USA). The gradients used for separation of the peptides were 2%
buffer B (20 mM ammonium formate, 90% acetonitrile, pH 10)
for 2 min, then 2–45% B in 43 min, 45–100% B in 5 min;
1-min fractions were collected and vacuum dried before being
combined or individually analyzed by nano-LC-MS/MS.
Nano-LC-MS/MS was performed using an RSLC system
interfaced with an LTQ Orbitrap Velos (ThermoFisher,
Waltham, MA, USA). Samples were loaded onto a self-packed
100 lm 9 2 cm trap packed with Magic C18AQ, 5 lm 200 A
(Bruker Auburn, Auburn, CA, USA), and washed with buffer A
(0.2% formic acid) for 5 min with a flow rate of 10 ll min1.
The trap was brought in-line with the home-built analytical column (Magic C18AQ, 3 lm 200 A, 75 lm 9 50 cm) and peptides were fractionated at 300 nl min1 with a multistepped
gradient (4–15% buffer B (0.16% formic acid, 80% acetonitrile)
in 25 min and 15–25% B in 65 min and 25–50% B in 55 min).
The mass spectrometer acquisition cycled through one MS in Orbitrap (resolution 60 000), followed by 20 MS/MS (CID) in
LTQ with dynamic exclusion (two repeat counts within 30 s and
exclusion time of 60 s). The LC-MS/MS data were analyzed
using Proteome Discoverer software v1.3 (ThermoFisher). The
data were first searched against T. pseudonana (composed of
sequences queried from Uniprot) using a Sequest search engine
through a ‘light’ search assuming normal nitrogen isotope distribution and a ‘heavy’ search assuming all amino acids were labeled
with 15N. For both ‘heavy’ and ‘light’ searches, carbamidoethyl
on cysteine was used as a fixed modification. For the ‘light’
search, oxidation of methionine was included as a variable modification. For the ‘heavy’ search, N-terminal modification of
0.997 Da was included as a variable modification. The identified peptides were quantified with a custom-built precursor ion
quantification method within the software. The peptide quantification results of each HPLC fraction were combined into one list
(MUDPIT). Peptides were filtered to include only top ranked
identification with confidence above medium. The heavy/light
ratio of each peptide was normalized to the median ratios of all
peptides. The heavy/light ratio of protein was calculated as the
median value of all peptides belonging to this protein. The protein ratio was accepted if three or more peptides were quantified.
Detection limits for the responsive proteome
The minimum difference in 15N : 14N ratios for differential protein abundances was determined by evaluating the distribution of
relative protein abundances for T. pseudonana grown under
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(a)
(b)
Fig. 2 Thalassiosira pseudonana photosynthetic parameters before and
following a shift (at t = 0) from high to low CO2. (a) The initial slope of the
photosynthesis–irradiance (PE) relationship (ab, circles) and chlorophyllspecific maximum carbon fixation rate (Pbmax, squares) for PE relationships
determined in 10-min 14C-bicarbonate assimilation assays. (b) Light
reaction parameters: variable to maximum fluorescence (Fv/Fm, circles)
determined immediately (no dark acclimation period) and nonphotochemical quenching (NPQ, rectangles) determined on samples in
which NPQ was permitted to relax. The widths of the symbols for each
parameter are equal to the time required to make the measurement,
except for Fv/Fm, which is instantaneous. Error bars, SD.
(a)
(b)
Fig. 3 Relative gene expression for phosphoenolpyruvate carboxylase 1 (a)
and phosphoenolpyruvate carboxylase 2 (b) in Thalassiosira pseudonana.
At time zero, cells were shifted from high to low CO2. Gene expression
relative to that for actin was determined in two replicate analyses per
experiment. Experiments one and two are indicated as filled and hollow
symbols, respectively. For each of two replicate experiments, error
bars SD of replicate analyses.
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identical conditions (CO2 at c. 1780 ll l1), except that the N
source was provided as either 14N-NO3 or 15N-NO3. This
serves to integrate the error caused by the variability in mass spectrometer ionization efficiencies as well as biological variability.
The raw 15N : 14N peak area ratio for each protein was normalized to the global median 15N : 14N value and then log2 transformed. The standard deviation of these log2-transformed
15
N : 14N ratios was 0.167 (Table S2). Setting the type I error
rate at 1% corresponded to a threshold log2-normalized
15
N : 14N ratio of 0.388. Proteins from experimental treatments (grown in 14N and 15N under low and high CO2, respectively) were considered to be differentially expressed when the
log2-transformed value in both biological replicates was either
< 0.388 (corresponding to a median-normalized 15N : 14N
ratio of 0.7641 or less; up-regulated under low CO2) or > 0.388
(corresponding to a median normalized 15N : 14N ratio of 1.309;
up-regulated under high CO2). We applied this criterion to the
15
N : 14N ratios for each protein from both biological replicates.
Assuming that the error of the 15N : 14N ratio measured for any
protein is random and represented by the standard deviation
measured here, this criterion translates to a false positive rate of
0.01%.
For proteins differentially expressed in both biological replicates, we report the average fold change (geometric mean) in protein abundance, with an exception for proteins that are extremely
abundant under one condition relative to the other. Isotopic
ratios are difficult to constrain for such instances (i.e. those with
Subcellular protein targeting
Targeting was assessed using the predicted localization of gene/
Uniprot models (Table S3). In some cases, gene and Uniprot
models were re-evaluated and models were extended upstream.
We followed the general approaches outlined in Gruber et al.
(2007), Kroth et al. (2008) and Bender et al. (2012). Specific criteria to assess targeting are provided in more detail in Table S3.
Results
Photo-physiology and transcripts in response to low - CO2
shift
Maximum Chl-specific carbon fixation rates (Pbmax) declined by
54% within the first 10 min after the shift to low CO2, whereas
the initial slope of the irradiance vs fixation curve (ab) was unaffected (Fig. 2a). Before Pbmax recovery, the variable fluorescence
(a)
(d)
(b)
(e)
(c)
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ratios < 1 : 12 or > 12 : 1; Huttlin et al., 2007). For this reason,
we caution on the over-interpretation of the quantitative nature
of such extreme peptide ratios, and rather support the interpretation that such peptides have abundance ratios much less (or
greater) than 1 : 1. In cases in which the apparent fold increase in
protein abundance for one replicate was > 12, we indicate the
fold change in protein abundance as at least as great as the lesser
fold change measured.
Fig. 4 Relative gene expression for proteins
potentially involved in either of the two
archetypical C4 decarboxylation pathways in
Thalassiosira pseudonana. At time zero, cells
were shifted from high to low CO2. (a)
Cytosolic malate dehydrogenase, (b) NADdependent malic enzyme, (c) pyruvate
phosphate dikinase, (d) mitochondrial malate
dehydrogenase, (e) phosphoenolpyruvate
carboxykinase. Gene expression relative to
that for actin was determined in two replicate
analyses per experiment. Experiment one
and two are indicated as filled and hollow
symbols, respectively. For each of two
replicate experiments, error bars SD of
replicate analyses. These values were all
below the 20% transcript relative abundance
threshold (Supporting Information Table S1).
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(Fv/Fm) dropped (11%) and the capacity to induce NPQ rose by
37% (Fig. 2b). These responses are consistent with a more
reduced photosynthetic electron transport chain under acute
CO2 limitation. Within 30–50 min after their initial deviation,
NPQ and Fv/Fm both recovered to values obtained under
steady-state high CO2. The culture pH remained at 8.48
throughout the recovery period (data not shown).
When cultures were shifted from high to low CO2, mitochondrial targeted PEPC1 TRA increased by 5.7-fold within
30 min of the shift (Fig. 3a). By 90 min, PEPC1 TRA had
declined to 2.2-fold of pre-shift values. Similarly, TRA of
PEPC2 (targeted to the chloroplastic endoplasmic reticulum
(CER) or periplastidic space (PPS)) increased by 6.5-fold within
30 min of the shift (Fig. 3b), and decreased to c. 2.3-fold of
pre-shift values by 90 min. These increases were above the minimum threshold of 20% (see the Materials and Methods section
and Table S1). Transcript abundances were significantly higher
for PEPC1 (P < 0.003) and PEPC2 (P < 0.005) after exposure
to low CO2. Transcript abundances for other genes potentially involved in pathways for C4 acid transport and decarboxylation were also evaluated (Fig. 4). The average differences in
TRA between pre- and post-shift conditions for two MDHs,
NAD-ME, PPDK and PEPCK were below the 20% TRA
threshold (Table S1).
Quantitative proteomic analyses at steady state
Under steady-state growth conditions, growth rates during
these experiments did not differ at the two pH/CO2 levels
investigated (2.06 0.014 d1 at pH 7.61 and 2.03 0.071 d1
at pH 8.48), which is consistent with other findings that ocean
acidification (lower pH) does not have an obvious direct effect on
T. pseudonana growth (Crawfurd et al., 2011). Using a nitrogen
isotope labeling approach, we were able to quantify 3688 proteins
Table 2 Proteins up-regulated under low CO2 in both replicates
Uniprot ID
Description
Replicate 1
Replicate 2
Average fold change
B8BTB4
B8C083
B8C250
B8LE18
B8LE19
B8C0D9
B8CG97
B8C4I8
B8C0D8
B8LE17
B8BSW5
B8C8T5
B8BQZ0
B8CDL6
B8BY90
B8BZ35
B8BSI9
B8CE42
B8BPY6
B8BVD1
B8C3W3
B5YMF5
B8C1R7
B8LDH1
B8BX31
B8C596
B8C017
B8C1C4
B8C5V3
B8BXJ5
B8BWX9
B8BZM5
B8BYW8
B8BYN5
Delta carbonic anhydrase
Putative DNA-binding heat shock factor protein
Carbonic anhydrase
Mucin-like protein
HNKH
Bestrophin 2, similar to low-CO2 inducible membrane protein
Cadmium carbonic anhydrase
Putative glycoprotein
Bestrophin 1; similar to low-CO2 inducible membrane protein
HNKH
WAX2-like protein
Glutamine-dependent carbamoyl phosphate synthase-like protein
Ornithine cyclodeaminase
AMP yield acyl-CoA synthetase
Formate–tetrahydrofolate ligase
Alanine glyoxylate aminotransferase/serine pyruvate transaminase
HNKH
Pyruvate carboxylase
HNKH
Related to biotin carboxylase and acetyl-CoA carboxylase
HNKH
Acetyl-CoA carboxylase
Phosphoenolpyruvate carboxylase
Putative SAM-dependent methyltransferase
Glycine decarboxylase p-protein
HNKH
Alanine aminotransferase
Cold-shock DNA-binding domain-containing protein
Biotin synthase
L-Lactate dehydrogenase
Microsomal omega-6 fatty acid desaturase
BolA-like protein
Phosphoenolpyruvate carboxylase
HNKH
580.00
140.23
506.71
15.18
7.89
4.75
77.83
2.36
3.51
3.19
1.36
2.28
1.88
2.37
1.93
1.60
1.49
1.99
1.95
1.75
1.83
1.62
1.71
1.64
1.58
1.50
1.48
1.50
1.59
1.55
1.46
1.47
1.32
1.39
11.37
14.68
9.08
8.67
5.36
3.78
4.04
4.74
3.13
2.30
2.59
1.63
1.95
1.37
1.66
1.95
2.06
1.54
1.51
1.63
1.34
1.45
1.35
1.40
1.45
1.49
1.48
1.46
1.34
1.35
1.42
1.39
1.46
1.38
≥ 11
≥ 15
≥9
≥9
6.50
4.24
≥4
3.34
3.31
2.71
1.88
1.93
1.91
1.80
1.79
1.77
1.75
1.75
1.72
1.69
1.57
1.53
1.52
1.52
1.51
1.49
1.48
1.48
1.46
1.45
1.44
1.43
1.39
1.38
The fold increase in protein abundance within each biological replicate was calculated as the inverse of the normalized 15N : 14N ratio. Averages were
calculated as the geometric mean from biological replicates. When apparent fold increases for one replicate were > 12, the average was substituted with a
range equal to or greater than the lesser value (see text for more details). Protein descriptions listed as HNKH were hypothetical proteins with no known
described homologs according to BLASTp search (National Center for Biotechnology Information, NCBI) with a cut-off of E > 1030. Up-regulated but
incomplete protein models without plausible N termini (Supporting Information Table S3; B8C5S5, B8C235, B8C333, B8LDG9, B8CEV2) are not shown
here.
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Table 3 Proteins down-regulated under low CO2 in both replicates
Uniprot ID
Description
Replicate 1
Replicate 2
Average fold change
B8C274
B8LE04
B8BW41
B5YNT2
B8LEL7
B8BW74
B8BVA0
B8C8L8
B8BXS2
B8LD81
B8LBQ0
B8C5Q7
B5YNK8
B8BSB3
B8BRH9
B8BT47
B8C7H4
B8LE89
Phosphoenolpyruvate carboxykinase
ISIP3-like protein
Similar to cobalamin synthesis protein
Glutathione reductase
Nickel ABC transporter
HNKH
Transaldolase
HNKH
Fructose-1,6-bisphosphatase
K exchanger-like protein
Putative NAD-dependent epimerase/dehydratase
HNKH
HNKH
Putative triose phosphate translocator
HNKH
Similar to Lipid A export msbA protein
HNKH
HNKH
321
5.04
1724
4.00
579
2.34
2.50
2.21
2.31
2.11
1.80
19
1.37
1.71
1.59
1.61
1.37
1.36
5.08
1.84
2.71
1.36
2.50
1.86
1.44
1.55
1.40
1.36
1.67
1.72
2.00
1.52
1.39
1.34
1.36
1.33
≥5
3.05
≥ 2.71
2.33
≥ 2.5
2.09
1.90
1.85
1.80
1.69
1.73
≥ 1.72
1.66
1.61
1.49
1.47
1.36
1.34
The fold decrease in protein abundance within each biological replicate is the normalized 15N : 14N ratio. Averages and protein descriptions are given as in
Table 2. Down-regulated but incomplete protein models without plausible N termini (Supporting Information Table S3; B8BWI1, B8C0W2, B8C443,
B8C231) are not shown here.
from the T. pseudonana proteome from both replicate experiments (Table S4). Of these, 39 and 22 proteins were significantly
up-regulated (Tables 2, S3) and down-regulated (Tables 3, S3)
under low CO2, respectively. Targeting predictions and relative
abundances for select proteins are shown in Tables S3 and S5,
respectively. The most highly up-regulated proteins at low CO2
included three carbonic anhydrases, CA-4, CA-6 and CA-3 (Uniprot IDs B8BTB4, B8C250 and B8CG97, respectively), following the nomenclature of Tachibana et al. (2011). CA-4 and CA-3
may be localized to the cytosol, whereas CA-6 may be targeted to
the cell surface or ER membrane. Two chloroplast-targeted proteins in the bestrophin family of anion channels (B8C0D8 and
B8C0D9) were also present in higher abundance at low CO2.
Among the candidate C4 metabolism proteins, both PEPC paralogs were more abundant under low-CO2 conditions. PEPC2
(B8C1R7) exhibited SignalP targeting scores suggestive of targeting to the CER or PPS, whereas PEPC1 (B8BYW8) seemed to be
mitochondrial. ME (B8C1Z0) abundances did not change in
response to CO2 (Table S5), whereas PEPCK (B8C274) abundances decreased by at least five-fold. Notably, PPDK (B8C332)
was not detected in these proteomes. The abundance of a chloroplast-targeted pyruvate carboxylase (PYC; B8CE42) was 1.8-fold
higher under low CO2. The expression patterns of other proteins
include the 1.6-fold down-regulation of a putative triose phosphate translocator (B8BSB3), which belongs to a family of proteins capable of triose phosphate or 3-phosphoglycerate (3-PGA)
transport (Fl€
ugge, 1999), and has ER targeting and possible
anchoring to the ER membrane. In addition, cytoplasmic fructose
1,6-bisphosphatase (B8BXS2), transaldolase (B8BVA0) and
epimerase (B8LBQ0), components of the non-oxidative portion
of the pentose phosphate pathway, were down-regulated at low
CO2 (Table 3). Core proteins in the oxidative phase of the pentose phosphate pathway were not differentially expressed, but
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glutathione reductase (B5YNT2), which participates in a side
reaction consuming NADPH, was down-regulated by c. 2.3-fold
under low CO2 (Fig. 5; Table 3). Some photorespiratory proteins, including GDC P protein (B8BX31) and alanine glyoxylate
aminotransferase (AGAT; B8BZ35), were up-regulated (by 1.5and 1.8-fold, respectively) under low-CO2 conditions (Table 2).
Proteins involved in ornithine and carbamoyl phosphate metabolism were up-regulated under steady-state low CO2. Ornithine
cyclodeaminase (B8BQZ0) was 1.9-fold more abundant under
low CO2, whereas glutamine-dependent carbamoyl phosphate
synthetase (B8C8T5) was 1.9-fold more abundant. However, no
proteins involved in the ornithine–urea cycle (OUC) were differentially expressed (Table S5).
Discussion
These experiments were designed to examine the physiological
responses to decreasing CO2 which are known to occur during
blooms. Our pre-shift conditions provided a supply of CO2 at
about twice the half-saturation concentration for diatom RuBisCO (c. 30 lM; Badger et al., 1998) to greatly reduce the demand
for a CCM relative to that at ambient CO2. Following the shift to
low CO2 (7.1 lM), carbon fixation (Pbmax) dropped immediately.
The concomitant drop in variable fluorescence and increase in
NPQ capacity are consistent with the limitation of photosynthetic
electron transport by CO2 availability. The recovery of all photophysiological parameters was complete within 100 min.
Both PEPC transcripts were up-regulated on shifting to low
CO2 and, after 90 min, these transcripts were c. two-fold greater
than pre-shift values (Fig. 3). The slightly greater response (c.
three-fold) observed by McGinn & Morel (2008) may reflect the
lower CO2 values (at pH 8.9 vs pH 8.48 here) in their experiments. In contrast, Roberts et al. (2007) did not observe
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Fig. 5 Effects of low CO2 on cytoplasmic pentose phosphate pathway
relative protein abundances in Thalassiosira pseudonana (determined as
described in the text). Red and black arrows indicate down-regulated and
equally represented proteins at low CO2 relative to high CO2, whereas
gray arrow indicates protein was not quantified in both biological
replicates. The upper and lower portions represent the oxidative and nonoxidative phases of the pentose phosphate pathway, respectively.
Enzymes highlighted here – and their corresponding Uniprot IDs – include
NAD-dependent epimerase (EP; B8LBQ0), fructose bisphosphatase (FBP;
B8BXS2), fructose bisphosphate aldolase (ALDO; B8CFH1), glucose-6phosphate dehydrogenase (G6PD; B8C3E7), glucose-6-phosphate
isomerase (GPI; B8BYC8 or B8LDA5), 6-phosphogluconate
dehydrogenase (6PGD; B8BZH6), transketolase (TK; B8BTR4),
transaldolase (TA; B8BVA0), triose phosphate isomerase (TPI; B5YLS7,
B5YNQ0 or B8C5E1) and ribose-5-phosphate isomerase (RPI; B8BUF5).
Abbreviations for substrates: Ery4P, erythrose-4-phosphate; F6P, fructose6-phosphate; GADP, glyceraldehyde phosphate; G6P, glucose-6phosphate; PEP, phosphoenolpyruvate; 6PG, 6-phosphogluconate; R5P,
ribose-5-phosphate, Ru5P, ribulose-5-phosphate; S7P, sedoheptulose-7phosphate; Xu5P, xylulose-5-phosphate. The down-regulation of
cytoplasmic PPP enzymes may serve to diminish the drawdown of GADP
in the cytoplasm and retain more GADP in the chloroplast to ensure
sufficient regeneration of Ru5P lost to photorespiration. Plastid PPP
enzyme abundances did not change between treatments (Supporting
Information Table S4).
significant changes in transcript abundances for either PEPC.
Two factors may explain these disparate results. Steady-state differences in transcript abundances are expected to be minimal,
whereas, during the rapid shifts from high to low CO2, we
expected (and observed) a transient state in which photosynthesis
was limited by CO2 availability, followed by recovery after CO2responsive genes were transcribed. Therefore, the results of Roberts et al. (2007) may reflect the conditions of the cultures, rather
than the absence of C4-assisted photosynthesis. In addition, their
CO2 concentrations (380 and 100 ll l1 CO2) vastly undersaturate RuBisCO carboxylase activity (Badger et al., 1998), raising
the possibility that the CCM would be operating at similar rates
under both conditions.
Short-term changes in PEPC transcription observed in our
CO2 shift experiments are corroborated by steady-state proteomic results (Table 2), where abundances of CER- or PPS-targeted
PEPC2 increased c. 1.5-fold under low CO2. Increased PEPC2
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protein levels may elevate OAA production rates in the CER (or
PPS). Given the filamentous distribution of the ER, this carboxylation is functionally analogous to the cytoplasmic OAA production in other single-cell C4 systems (Voznesenskaya et al.,
2002; Edwards et al., 2004). These increases in PEPC abundance
may also be accompanied by other adaptations to low CO2 that
serve to increase PEPC activity. For example, the down-regulation of an ER membrane-anchored PGA/glyceraldehyde phosphate (GADP) transporter (B8BSB3) under low CO2 would help
to decrease 3-PGA export to the cytosol and promote the production of PEP in the CER (described in detail below).
Our results do not support either ME- or PEPCK-mediated
decarboxylation for C4 photosynthesis in T. pseudonana. Intracellular targeting analysis suggests that neither protein is localized in
the chloroplast, inconsistent with the most parsimonious model
of where C4 decarboxylation should occur. Moreover, various
transcripts for proteins involved in the putative ME-mediated C4
pathway (Fig. 1) did not increase after the shift to low CO2
(Fig. 4). This suggests that, if ME is responsible for decarboxylation here, it is not regulated at the transcriptional level. The
indistinguishable protein abundances for ME (Table S5) further
suggest that ME may not be involved in C4-assisted photosynthesis in this species. The absence of transcript abundance differences (Fig. 4) and the five-fold or greater decrease in PEPCK
protein abundances (Table 3) at low CO2 are consistent with the
decrease in P. tricornutum PEPCK activities at low CO2 (Cassar
& Laws, 2007), but not with a role for PEPCK-mediated decarboxylation in C4-assisted photosynthesis here.
The identity of the proteins responsible for the decarboxylation
step of C4 photosynthesis in these diatoms is unresolved. The
independent origins for C4 photosynthesis (Sage, 2004) leave
open the possibility that some other enzyme altogether may have
been co-opted for the decarboxylation step in diatoms. The chloroplast-localized and reversible PYC (B8CE42) was c. 1.8-fold
more abundant under low CO2. The decarboxylation of OAA by
PYC, coupled to the synthesis of ATP from ADP, is exothermic at
neutral pH, an ATP/ADP ratio of 2.5 and an OAA concentration
> 1 mM (calculations not shown), which raises the possibility that
this protein carries out the decarboxylation reaction (as has been
observed in model systems; Attwood & Cleland, 1986). However,
a forward-acting PYC in T. pseudonana would seem to compete
with RuBisCO for HCO3/CO2 under the very conditions in
which plastid CO2 must be elevated to support rapid growth.
Four conditions should be satisfied to support the hypothesized role for PYC. First, OAA formed in the CER or PPS needs
to be transported to the chloroplast. Two chloroplast-targeted
bestrophin family anion channels were among the most up-regulated proteins under low CO2. One plausible role for these bestrophins is to facilitate diffusion of CER/PPS-localized OAA to
the chloroplast, particularly given the recent discovery that some
bestrophins are permeable to carboxylated organic anions (Roberts et al., 2011). Our current ability to characterize the substrate
specificity of bestrophins in silico is admittedly very limited
(Hagen et al., 2005), leaving open the possibility of an altogether
different function for these proteins. Second, this mechanism of
PYC-mediated decarboxylation also requires the participation of
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a plastid-localized CA. Tachibana et al. (2011) showed clear
experimental evidence of plastid targeting of CA1-green fluorescent protein (CA1-GFP) fusion proteins in T. pseudonana. Yet,
bioinformatics analysis suggests that this may be targeted to the
CER/PPS (Table S3), as it lacks the canonical signal-transit peptide boundary motif (Kilian & Kroth, 2005; Gruber et al.,
2007). This example illustrates a shortcoming of our in silico
knowledge of diatom physiology brought about by uncertainties
in protein targeting. Third, the regeneration of PEP from PYR is
required to sustain the C4 pump. This is most often thought to
be achieved by PPDK (Kroth et al., 2008), but transcription of
PPDK did not respond to CO2, and PPDK was not detected in
our proteome or by probing with antibodies raised towards Zea
mays (courtesy of Chris Chastain, Minnesota State University,
Moorhead, MN, USA; data not shown). In addition, PPDK has
been shown recently to have little effect on growth or photosynthesis in P. tricornutum (Haimovich-Dayan et al., 2013). Fourth,
plastid PYR concentrations should be drawn down relative to
those of OAA to promote the back-reaction.
The proteomic response of T. pseudonana suggests that a
PPDK-independent mechanism of PEP regeneration may be at
play (Fig. 6), which also satisfies the third and fourth criteria.
Greater abundances of cytoplasmic alanine aminotransferase
(ALAT), the bifunctional mitochondrial AGAT/serine pyruvate
transaminase and mitochondrial GDC at low CO2 suggest an
alternative pathway to recover PYR through its transfer to the
mitochondria as alanine and its sequential conversion to 3-hydroxypyruvate, glycerate and 3-PGA (Kroth et al., 2008). A
potential complete pathway for the regeneration of PEP from
PYR is outlined in Fig. 6(b). The unchanged and lower abundances of chloroplast membrane- and CER-localized triose
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phosphate translocators (TPT3; B8BSB3 and TPT4; B5YLS2),
respectively, at low CO2 would support the retention of 3-PGA
in the CER for subsequent conversion to 2-PGA by phosphoglycerate mutase (PGM) (Fig. 6), although the topologies of these
translocators are unknown. PEP formed by cytosolic enolase may
be imported to the CER by a PEP/inorganic phosphate (PEP/Pi)
antiporter (PPT), which can accept either Pi or 2-PGA as a
counter-exchange substrate (Fischer et al., 1997). These proteins
have been functionally verified and best described in vascular
plants (Fischer et al., 1994, 1997; Fl€ugge et al., 2011), but there
has been no heterologous expression work in any photosynthetic
chromalveolates, and there are currently no clear candidates in
the T. pseudonana genome. Three proteins from our limited list
of those up-regulated at low CO2 would serve to draw down
PYR concentrations and facilitate the net export of PYR from the
plastid. These up-regulated proteins are also integral to the regeneration of PEP from PYR.
Malate is the principal C4 acid isolated from low-CO2-acclimated T. weissflogii (Roberts et al., 2007), which, at first blush,
might suggest that malate is the decarboxylation substrate.
However, both malate and OAA equally compete for 14CO2 fixation (Reinfelder et al., 2004), suggesting that diatoms rapidly
interconvert these substrates. This proposed model of OAA decarboxylation in T. pseudonana is consistent with these observations,
considering that the CER/PPS-localized MDH may serve to
buffer the CER/PPS pool of OAA. Without some mechanism to
buffer the OAA supply, the PEPC2-mediated production and flux
of OAA into the plastid would result in futile CO2 production
during, for example, periods of transient low light. The production of malate by MDH when CER/PPS OAA concentrations
become transiently elevated, as OAA production exceeds demand,
Fig. 6 Model of C4 photosynthetic carbon assimilation and associated rearrangement of carbon metabolism to support growth at low CO2 in Thalassiosira
pseudonana. (a) Core components of the model. Red, green and black arrows indicate down-regulated, up-regulated and equally represented proteins at
low CO2 relative to high CO2, respectively, whereas gray arrows indicate that protein was not quantified in both biological replicates. In cases in which
more than one candidate protein may be responsible, multiple arrows indicating CO2-responsive status are shown. In the chloroplastic endoplasmic
reticulum CER or periplastidic space PPS, PEPC2 converts bicarbonate and PEP to OAA. PEP concentrations are enhanced by the down-regulation of a PEP
transporter (TPT3) anchored to the ER membrane. Elevated mitochondrial production of CO2 by photorespiration (only GDC is shown here for simplicity)
may be scavenged by mitochondrial CA and PEPC1 (forming OAA), whereas diminution of mitochondrial PEPCK-catalyzed C4 decarboxylation would
prevent a futile CO2–C4 cycle in the mitochondrion. The down-regulation of PEPCK should favor a greater flux of OAA decarboxylation through pyruvate
carboxylase. Greater abundances of one or both anion channel bestrophin proteins (BEST1/2) may be responsible for facilitating OAA diffusion into the
chloroplast at low CO2. Malate dehydrogenase may serve to minimize the futile decarboxylation of OAA under transient conditions of low light by
minimizing any accumulation of OAA from PEPC2 under these conditions. OAA decarboxylation by PYC produces bicarbonate, which is dehydrated to
CO2 in the proximity of RuBisCO by a chloroplast-localized CA, such as CA1 (reported by Tachibana et al., 2011). Elevated mitochondrial production of
CO2 by photorespiration (only GDC is shown here for simplicity) may be scavenged by mitochondrial CA and PEPC1 (forming OAA), whereas PEPCK is
down-regulated to prevent a futile cycle and promote the flux of OAA to the chloroplast. NH4+ derived from photorespiration may be sequestered as
cytoplasmic carbamoyl phosphate through GS and glutamine-dependent CPScyto, presumably for pyrimidine synthesis. (b) Proposed PPDK-independent
pathway to regenerate phosphoenolpyruvate from pyruvate. Plastidic pyruvate may be used to regenerate PEP through the coordinated reaction of ALAT,
AGAT/SPT, HPR, GK, PGM and ENO and others (as shown in simplified form in a, where proposed reactions that regenerate glyoxylate and serine are
omitted for clarity). Currently, no known homologs for GK and PPT have been identified in the T. pseudonana genome. Enzymes highlighted in (a) – and
their corresponding Uniprot IDs – include alanine aminotransferase (ALAT; B8C017), alanine glyoxylate aminotransferase (AGAT; B8BZ35), bestrophins 1
and 2 (BEST1, BEST2; B8C0D8, B8C0D9), carbamoyl phosphate synthetase (CPScyto; B8C8T5), carbonic anhydrase (CA; possibly B8C025), cytosolic CA
(CAcyto, CA3 or CA4; B8C250 or B8BTB4), CA6 (B8C250), mitochondrial CA (CAmito, CA8, CA9 or CA13; B8C2H3, B8BRC3 or B8C215), enolase (ENO,
B8C355), glutamine synthetase (GS; B8BVI3), glycine decarboxylase (GDC P-protein; B8BX31), hydroxypyruvate reductase (HPR; B8BVI2), malate
dehydrogenase (MDH; B8BQC2), PEP carboxykinase (PEPCK; B8C274), PEP carboxylase (PEPC1; B8BYW8 and PEPC2; B8C1R7), phosphoglycerate
mutase (PGM; B8C354), pyruvate carboxylase (PYC; B8CE42), RuBisCO (rbcL; A8DP73 and rbcS; A0T0N5), serine pyruvate transaminase (SPT; B8BZ35)
and triose phosphate transporters (TPT3; B8BSB3 and TPT4; B5YLS2). Abbreviations for substrates: CP, carbamoyl phosphate; GADP, glyceraldehyde
phosphate; GLC, glycerate; GLX, glyoxylate; MAL, malate; OAA, oxaloacetate; OXO, 2-oxoglutarate; 2PGA, 2-phosphoglycerate; PEP,
phosphoenolpyruvate; PGA, phosphoglycerate; PYR, pyruvate; RuBP, ribulose bisphosphate. Adenosine phosphates and reductants not shown for
simplicity.
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would prevent this futile CO2 production, potentially by both
buffering the OAA pool and by inhibiting PEPC2 activity (Budde
& Chollet, 1986). As bestrophins only facilitate diffusion,
unutilized chloroplast OAA levels would lead to the net conversion of OAA in the CER/PPS to malate. On return to high light,
(a)
ALAT
Ala
OXO
Ala
PYR
OAA drawdown in the plastid will lead to the net conversion of
malate back to OAA in the CER/PPS (and possibly the
de-repression of PEPC2). This model would be entirely consistent
with the ecology of marine diatoms, which are particularly
adapted to turbulent conditions in which their incident light
cytoplasm
Glu
RuBisCO
PGA
CO2 + RuBP
AGAT
PYR
Ser
S
Gly P
GLX
T Ala
OH-PYR
HPR
GLC
CO2+
OAA
PEP
PEPCK
GLC
PGA
GK
PEP + HCO3–
OAA
PEPC1
CAmito
2PGA
Gly
GDC
PYR + HCO3–
mitochondrion
NH4+ GSI
Glu
PYC
OAA
BEST1/2
MAL
TPT4
PEPC2
PGA
MDH
–
PGM PEP + HCO3 OAA
CO2
PEP
PPT
CO2 + NH4+
plastid
CA
2PGA PEP
ENO
CPScyto
Gln
HCO3–
CA6
CAcyto
CO2
HCO3–
CP
pyrimidine
synthesis
(b)
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518 Research
regime rapidly changes as cells transit throughout the mixed layer
(Cerme~
no et al., 2008).
If in vivo RuBisCO oxygenase activities represent a substantial
sink for fixed carbon in marine diatoms at low CO2, a highly efficient compensatory recycling mechanism must be at play, as
growth rates are independent of CO2. Others have suggested that
diffusive losses of NH4+ and CO2 may be avoided by scavenging
and recycling through the OUC (Parker et al., 2008; Allen et al.,
2011). However, of the five OUC proteins quantified, none were
up-regulated. Higher levels of mitochondrial C4 carboxylation
catalyzed by PEPC1 and various CAs may serve to recover CO2
released from photorespiration, whereas the diminution of PEPCK-catalyzed C4 decarboxylation would prevent a futile CO2–C4
cycle in the mitochondria. Cytoplasmic carbamoyl phosphate
synthetase (CPS) was up-regulated at low CO2, suggesting a possible recuperation of NH4+ through glutamine synthetase and
CPS activity (Fig. 6).
The present results support a model of diatom metabolism
in which high carbon fixation and growth rates are maintained
at low CO2 by elevated OAA synthesis in the CER, brought
about by elevated PEPC2 abundance and elevated PEP concentrations derived from the antiport of PGA out of and PEP
into the CER. Our results further indicate that the decarboxylation of OAA may be catalyzed by plastidic PYC (Fig. 6). It
should be noted that this model would require and is entirely
consistent with the presence of bicarbonate transporters in the
plasmalemma (Nakajima et al., 2013) and ER membranes.
The up-regulation of ALAT, AGAT and GDC at low CO2
presents a mechanism to recycle pyruvate and complete the
proposed C4 pathway. Validation of this model will require
further verification of the intracellular localizations of the various enzymes involved, as well as the quantification of their
kinetic and regulatory control. These proteomic data provide
some insight into both the mechanisms of carbon concentration and compensation for photorespiration in diatoms, which
may contribute to their success during blooms when aqueous
CO2 is drawn down to levels far below atmospheric equilibrium and well below the levels that saturate the carboxylase
activity of RuBisCO.
Acknowledgements
This work was supported by National Science Foundation (NSF)
grants to J.R.R. and K.D.B. (OCE 0526365), A.J.M. (OCE
0526188) and A.B.K. (OCE 0927733).
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Supporting Information
Additional supporting information may be found in the online
version of this article.
Table S1 Analysis of actin transcript abundance and intra-treatment cycle threshold variability
Table S2 Thalassiosira pseudonana proteome 15N : 14N ratios
grown under high CO2 with either 14N or 15N nitrate
Table S4 Thalassiosira pseudonana proteome
grown under low and high CO2
15
N : 14N ratios
Table S5 Relative quantification of proteins from pathways of
interest
Please note: Wiley Blackwell are not responsible for the content
or functionality of any supporting information supplied by the
authors. Any queries (other than missing material) should be
directed to the New Phytologist Central Office.
Table S3 Targeting predictions and assigned cellular localization
for proteins of interest
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