Ulva prolifera, a typical green-tide-forming alga, can accumulate a large biomass in a relatively short time period, suggesting that photosynthesis in this organism, particularly its carbon fixation pathway, must be very efficient. Green algae are known to generally perform C3 photosynthesis, but recent metabolic labeling and genome sequencing data suggest that they may also perform C4 photosynthesis, so C4 photosynthesis might be more wide-spread than previously anticipated. Both C3 and C4 photosynthesis genes were found in U. prolifera by transcriptome sequencing. We also discovered the key enzymes of C4 metabolism based on functional analysis, such as pyruvate orthophosphate dikinase (PPDK), phosphoenolpyruvate carboxylase (PEPC), and phosphoenolpyruvate carboxykinase (PCK). To investigate whether the alga operates a C4-like pathway, the expression of rbcL and PPDK and their enzyme activities were measured under various forms and intensities of stress (differing levels of salinity, light intensity, and temperature). The expression of rbcL and PPDK and their enzyme activities were higher under adverse circumstances. However, under conditions of desiccation, the expression of rbcL and ribulose-1, 5-biphosphate carboxylase (RuBPCase) activity was lower, whereas that of PPDK was higher. These results suggest that elevated PPDK activity may alter carbon metabolism and lead to a partial operation of C4-type carbon metabolism in U. prolifera, probably contributing to its wide distribution and massive, repeated blooms in the Yellow Sea.
Citation: Xu J, Fan X, Zhang X, Xu D, Mou S, Cao S, et al. (2012) Evidence of Coexistence of C3 and C4 Photosynthetic Pathways in a Green-Tide-Forming Alga, Ulva prolifera. PLoS ONE 7(5): e37438. https://doi.org/10.1371/journal.pone.0037438
Editor: Vasu D. Appanna, Laurentian University, Canada
Received: February 14, 2012; Accepted: April 22, 2012; Published: May 16, 2012
Copyright: © 2012 Xu et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by Shandong Science and Technology plan project (2011GHY11528), the Specialized Fund for the Basic Research Operating expenses Program (20603022012004), National Natural Science Foundation of China (41176153), Natural Science Foundation of Shandong Province (2009ZRA02075), Qingdao Municipal Science and Technology plan project (11-3-1-5-hy), Qingdao Municipal Science and Technology plan project (10-3-4-11-1-jch), National Marine Public Welfare Research Project (200805069). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Carbon fixation is an important biological process in all photosynthetic organisms. C4 plants are characterized by high rates of photosynthesis and efficient use of water and nitrogen resources . High photosynthetic rates are achieved by addition of a new metabolic pathway, the C4 cycle, in which the initial product of CO2 fixation is a four-carbon (C) organic acid rather than a three-carbon (C) acid. C4 plants show drastically reduced rates of photorespiration because CO2 is concentrated at the site of Rubisco and is able to outcompete molecular oxygen, which, when used by Rubisco, results in photorespiration . The C4 photosynthetic carbon cycle is an elaborated addition to the C3 photosynthetic pathway, which ensures high rates of photosynthesis even when CO2 concentrations are low. C4 photosynthesis evolved several times independently during the evolution of higher plants. It originated at least 32 times in eudicots and 16 times in monocots . It had evolved from ancestral C3 plants via a series of anatomical and physiological adaptations to high light intensities, high temperatures, low pCO2, and dryness .
In aquatic environments, [CO2] can be a primary limitation for photosynthesis because of the low capacity of water to hold gaseous CO2 and the slow diffusion rate of dissolved molecules , . It has been demonstrated that many aquatic photosynthetic organisms can take up both CO2 and HCO3− from the surrounding media, and this capacity is greatly strengthened under CO2-limiting conditions, including the atmospheric pressure of CO2. This system is generally known as the inorganic carbon-concentrating mechanism (CCM) . Cyanobacteria, algae, and some angiosperms evolved multiple mechanisms to actively accumulate inorganic carbon around Rubisco by use of membrane transporters and carbonic anhydrases . The aquatic environment is home to as great a diversity of photosynthetic pathways as terrestrial environments, and there exist C3, C4, CAM, and C3–C4 photosynthetic pathways . Although apparently lacking Kranz anatomy, aquatic Orcuttia californica (an aquatic embryophyte) could also conduct C4 photosynthesis . Some species, such as Chara contraria (a charophyte green algae), Marsilea vestita (an embryophyte), Eleocharis acicularis (an embryophyte) and Pilularia Americana (an embryophyte), have both C3 and C4 fixation in aquatic habitats . Alterations of photosynthetic pathways under environmental stress have been suggested to contribute to the adaptation of plants to environmental stress . For example, Hydrilla verticillata, a submerged aquatic plant, changes its photosynthetic pathway from C3 to C4 under conditions of CO2 deficiency . Therefore, environmental factors are of critical importance in the change of photosynthetic pathways.
From many studies on primary photosynthetic carbon metabolism, it is believed that the operation of the Calvin–Benson cycle (C3 cycle) is predominant in algae , . However, recent papers have reported evidence for the operation of C4 photosynthesis as an alternative CCM in the marine diatom Thalassiosira weissflogii –. The case for C4 photosynthesis has been further strengthened by the occurrence of relevant genes in recently sequenced marine phytoplankton genomes, including the diatoms Thalassiosira pseudonana and Phaeodactylum tricornutum and the green alga Ostreococcus tauri and Micromonas –. Ostreococcus has all the machinery necessary to perform C4 photosynthesis. This includes a plastid-targeted NADP(1)-dependent malic enzyme and a phosphoenolpyruvate carboxylase . However, conflicting experimental data shedding doubt on C4 photosynthesis in diatoms have been reported , , and genomic data do not fully clarify the presence and localization of the enzymes that may drive this mechanism , . No clear evidence for such C4-like processes have been found in the marine diatoms P. tricornutum and T. pseudonana, for which whole genome sequences are available . The general occurrence of C4-like mechanisms in diatoms is therefore still in question , .
As a special type of harmful algal blooms (HABs), green tides have been increasing in severity and geographic range and are now of growing concern globally. Green tides are vast accumulations of unattached green macroalgae usually associated with eutrophied marine environments , . The great majority of green tides are reported to consist of members of just one genus, Ulva (some of the species formerly known as Enteromorpha) , . Ulva prolifera, a representative green-tide-forming macroalga , is the dominant Ulva species along the coastline of the Yellow Sea between June and August , . U. prolifera, as an intertidal macroalga, can tolerate various kinds of abiotic stresses, including desiccation, changes in temperature and salinity, and exposure to high levels of solar radiation during low tide . Furthermore, the evolutionary status of intertidal pluricellular green algae is between the unicellular green algae and lower land plants, which is an important stage during evolution .
It has been proved that marine algae contain C4-Pathway, including Ulva species . Kremer and Küppers (1977) found that the percentage of malate and aspartate usually accounts for distinctly less than 10% of the total 14C-labelling in three Ulva species, and these findings were consistent with data from enzymatic analyses, since 86–90% of the carboxylation capacity was due to ribulose-l.5-biphosphate carboxylase in those green algae . Moreover, the occurrence of PEP-C besides RubP-C has been reported from Ulva using 14C-labelling technique , . One of the most standard comparisons of differences in isotopic ratios is the comparison of 13C to 12C in plants to determine photosynthetic pathway of plants. C3 and C4 plants have different δ13C values, −28.1±2.5‰, −13.5±1.5‰ respectively . Among C3 and C4 plants, δ13C variation can range from 2–5‰. Previous research approved that Ulva are C4 species since there δ13C values are in the range of −14±4‰ , .
In this study we used next generation sequencing (NGS) technology confirmed the existence of genes necessary for a C4 pathway in U. prolifera, and we then chose to compare transcript abundance of U. prolifera with that of the closest relative, U. linza, which has been confirmed to possess the C4 pathway (unpublished data). Subsequently, we focused on the expression profile of two key enzymes, namely RuBPCase and PPDK. Ribulose-1, 5-biphosphate carboxylase, a key enzyme of the C3 pathway, catalyzes the first major step in carbon fixation. Pyruvate orthophosphate dikinase, a cardinal enzyme of the C4 pathway, catalyzes the regeneration of phosphoenolpyruvate (PEP), the primary carboxylation substrate from pyruvate, Pi, and ATP . The rate of PEP formation by PPDK is the lowest in the C4 pathway; therefore, this reaction is considered to be the rate-limiting step in the C4 pathway . Our results demonstrate that U. prolifera may be either a C3–C4 intermediate species or a C3 species displaying C4 metabolic characteristics. The involvement of C4 metabolism in U. prolifera might account for the boom of green tide.
Materials and Methods
Sample collection and culture conditions
Floating specimens of U. prolifera were collected in the Yellow Sea during the green tide bloom in 2011. In the laboratory, the intact samples were washed several times with sterile seawater, sterilized with 1% sodium hypochlorite for 2 min, and then rinsed with autoclaved seawater. The sterilized material was then placed into an aquarium (d = 40 cm, h = 30 cm) containing enriched and continually aerated seawater (500 µM NaNO3 and 50 µM NaH2PO4) and maintained at 15°C under a 12∶12 h LD photoperiod with 50 µmol photons m−2 s−1 provided by cool-white fluorescent tubes.
U. prolifera was exposed to different kinds of stress, namely desiccation and differing levels of salinity, light intensity, and temperature. For desiccation stress, the alga were cultured at 50 µmol photons m−2 s−1 for different durations (0, 1, 2, 3, 4, and 5 h). Salinity stress consisted of subjecting the organism for 3 h to different salt concentrations (0‰, 15‰, 30‰, 45‰, and 60‰); In light intensity treatment, the samples were exposure to 0, 50, 100, 300, 600, 1000, and 2000 µmol photons m−2 s−1 for 3 h. For the three forms of stress, temperature was constant at 15°C, and light intensity during the salinity treatment and the temperature treatment was maintained at 50 µmol photons m−2 s−1. For temperature stress, the materials were cultured at 5, 10, 15, 20, 25, 30 and 35°C for 3 h. Following each stress treatment, rbcL and PPDK mRNA expression level was measured using qPCR, RuBPCase and PPDK activity assessed, and Fv/Fm and Y(II) determined using Dual-PAM-100 (Walz GmbH, Germany).
Light and transmission electron microscopy
The sample preparation was finished according to the methods mentioned by Chen et al.  It consisted of the following steps: collecting the algal; fixing with 1% (v/v) glutaraldehyde and postfixing with 1%(v/v) osmium tetroxide both in sterilizing seawater; dehydration in a series of acetone solutions; suspension in the mixture of epoxy resin (Epon812) and acetone; embedded in 100% Epon812; polymerized and sectioned using a LeicaUC6 ultra microtome; picked up on 200-mesh copper grids and post-stained with urinal acetate. Finally, the sections were examined under an optical microscope (Nikon Eclipse 80i) and a transmission electron microscopy (Hitachi H-7650) at an accelerating voltage of 60 kv.
The alga were treated with different stress conditions, such as low temperature (6°C, 2 h), high temperature (42°C, 1 h), high light (1000 µmol photons m−2 s−1, 1 h), high salt (93‰, 3 h) and UV-B stress (60 µw cm−2, 3 h). Total RNA of all treated samples was extracted and purified, followed by synthesis and purification of double-stranded cDNA and sequencing of cDNA using a Roche GS FLX Titanium platform. To reconstruct the metabolic pathways in U. prolifera, high-quality reads were assigned to the Kyoto Encyclopedia of Genes and Genomes (KEGG) using the software package MEGAN (version 4.0) .
The partial rbcL cDNA sequence acquired from GenBank and the cDNA open reading frame (ORF) sequence of PPDK obtained from transcriptome sequencing, were examined for homology with other known sequences using the BLAST X program available at the website of the National Center for Biotechnology Information <www.ncbi.nlm.nih.gov/blast>. We used the Six Frame Translation of Sequence system <http://searchlauncher.bcm.tmc.edu/seq-util/Options/sixframe.html> analyzing deduced amino acid sequence. Multiple sequence alignments were generated using the program CLUSTAL X and then analyzed using the program BioEdit , . A phylogenetic tree was constructed using the neighbor-joining algorithm of the MEGA 4.0 program , .
Real-time quantitative PCR
Total RNA of U. prolifera exposed to each form and level of stress was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) as specified in the user manual and dissolved in diethypyrocarbonate (DEPC)-treated water. The cDNA used for real-time quantitative PCR was synthesized from the total RNA using Moloney murine leukemia virus reverse transcriptase (Promega Biotech Co., Madison, Wisconsin, USA).
The real-time quantitative PCR reactions were performed with the ABI StepOne Plus Real-Time PCR System (Applied Biosystems, USA) using SYBR Green fluorescence (TaKaRa) according to the manufacturer's instructions. To normalize the relative expression of the selected genes, an 18S rDNA gene was used as reference. Three pairs of gene-specific primers (Table 1) were designed according to the rbcL cDNA, PPDK cDNA, and 18S rDNA sequences using Primer Express 3.0. For each selected gene, three biological replicates were assayed independently. The qPCR amplifications were carried out in a total volume of 20 µL containing 10 µl of 2× SYBR Premix Ex TaqTM II (TaKaRa Biotech Co., Dalian, China), 0.6 µl (10 µM) of each primer, 2.0 µl of the diluted cDNA mix, and 6.8 µl de-ionized water. The qPCR amplification profile was obtained as follows: 95°C for 30 s followed by 40 cycles of 95°C for 5 s, 60°C for 10 s, and 72°C for 40 s. The 2−ΔΔCT method  was used to analyze the quantitative real-time PCR data.
The activity of RuBP carboxylase and PPDK in U. prolifera exposed to the treatments was measured, RuBP carboxylase activity by the method described by Gerard and Driscoll and PPDK activity by that described by Sayre et al. , ; both methods were modified as required.
For measuring RuBP carboxylase activity, each sample was ground to a fine powder in liquid nitrogen and homogenized in pre-cooled rubisco extraction solution (1 ml g−1 fresh weight), pH 7.6, containing 40 mM Tris-HCl buffer with 10 mM MgCl2, 0.25 mM EDTA, and 5 mM reduced glutathione. The homogenate was centrifuged at 10 000 g for 10 min at 4°C. The activity was measured in a 4.5 ml cuvette by adding 3 ml of a reaction mixture containing 0.2 ml NADH (5 mM), 0.2 ml ATP (50 mM), 0.1 ml enzyme extract, 0.2 ml creatine phosphate (50 mM), 0.2 ml NaHCO3 (0.2 mM), 1.4 ml reaction buffer (0.1 M Tris-HCl buffer, pH 7.8, with 12 mM MgCl2 and 0.4 mM EDTA), 0.1 ml creatinephosphokinase (160 units ml−1), 0.1 ml phosphoglycerate kinase (160 units ml−1), 0.1 ml glyceraldehyde-3-phosphate dehydrogenase (160 units ml−1), and 0.3 ml distilled water. The reaction was initiated by adding 0.1 mL ribulose-1, 5-bisphosphate (RuBP) to the reaction cuvette and OD values were recorded every 20 seconds for 3 min by a spectrophotometer at 340 nm. The enzyme activity was expressed in terms of micromoles per gram of fresh weight per minute (µmol g−1 FW min−1).
For measuring PPDK activity, the samples were ground to a fine powder in liquid nitrogen and homogenized in pre-cooled PPDK extraction solution at pH 8.3 (1 ml g−1 fresh weight) containing 100 mM Tris-HCl buffer with 5 mM mercaptoethanol and 2 mM EDTA. The homogenate was centrifuged at 10 000 g for 10 min at 4°C. The activity was measured in a 4.5 ml cuvette by adding 3 ml of a reaction mixture containing 0.1 ml Tris-HCl buffer (150 mM, pH 8.3, with 18 mM MgSO4), 0.1 ml DTT (300 mM), 0.1 ml PEP (30 mM), 0.1 ml NADH (4.5 mM), 0.1 ml AMP (30 mM), 0.1 ml lactic dehydrogenase (60 units ml−1), 0.1 ml enzyme extract, and 1.3 ml distilled water. The reaction was initiated by adding 0.1 mL pyrophosphate natrium to the reaction cuvette and the OD values were recorded every 20 seconds for 3 min at 340 nm. The PPDK activity was also expressed in terms of micromoles per gram of fresh weight per minute (µmol g−1 FW min−1).
Chlorophyll uorescence measurements
Photosynthetic performance of U. prolifera subjected to the different treatments was measured using Dual-PAM-100. The maximal photochemical efficiency of PS II (Fv/Fm) and the effective PS II quantum yield (Y II) were measured by the method of Fleming et al. . Before measurement, samples were dark adapted for 20 min. Optimal chlorophyll fluorescence quantum yield was calculated according to the following equation: Fv/Fm = (Fm−F0)/Fm. Fo and Fm refer to the minimal fluorescence and the maximal fluorescence from dark adapted samples, respectively. Fv is the difference between Fm and Fo. The culture experiments were repeated four times.
We analyzed the carbon fixation pathway in detail and discovered some key genes of enzymes involved in the carbon fixation pathway in U. prolifera, such as phosphoenolpyruvate carboxylase, aspartate aminotransferase, ribulose bisphosphate carboxylase, phosphoglycerate kinase, phosphoribulokinase, phosphoenolpyruvate carboxykinase, alanine transaminase, malate dehydrogenase (NADP+), pyruvate orthophosphate dikinase, and pyruvate kinase (Fig. 1), which provided unequivocal molecular evidence that most of the C3 pathway, C4 pathway, and CAM pathway genes were actively transcribed in U. prolifera. Figure 1 shows that both U. linza (unpublished) and U. prolifera have most of the genes that are indispensable to C3 and C4 pathways, and the relative enzymes are all the same in both algae. However, the abundances of C3 and C4 pathway genes in U. linza and U. prolifera are different. The results suggest the possibility of the existence of two photosynthetic pathways in U. prolifera, the Calvin cycle (C3) and the Hatch-Slack (C4) carbon fixation pathway.
cDNA Sequence Analysis
The partial rbcL cDNA sequence (FJ042888) was acquired from GenBank with a 1305 bp sequence encoding 435 amino acid residues. The PPDK cDNA sequence (JN936854) of ORF was obtained from the U. prolifera transcriptome database with a 2700 bp sequence encoding 889 amino acid residues. Phylogenetic analysis was conducted using the amino acid sequences of rbcL and PPDK (Fig. 2). The phylogenetic tree of rbcL indicated a species clustering that was basically consistent with the evolution of the species, and that of PPDK revealed that the C4 pathway had multiple independent origins. In the phylogenetic tree of rbcL, the clade of green algae diverged into two clusters: a C3–C4 cluster including both U. prolifera and O. tauri, which have all the genes involved in the C4 pathway, and a C3 cluster including C. reinhardtii and V. carteri. However, PPDK of O. tauri was clustered with the genes from land plants, and PPDK of O. tauri and E. vivipara appears to be more ancient than that of higher land plants. PPDK in U. prolifera was clustered with the genes found in the C3 green algae (C. reinhardtii and V. carteri.) and in the C3–C4 brown alga T. pseudonana, and PPDK in T. pseudonana appears to be more ancient than that in green algae. Overall, PPDK in green algae also has multiple independent origins as that in land plants.
The phylogenetic tree was constructed by the neighbor-joining (NJ) method using Mega (version 4.0). Bootstrap analysis was computed with 1000 replicates and bootstrap values below 50% were omitted. C3–C4 refers to species that possessed the genes for both C3 and C4 photosynthesis with C3 photosynthesis being the primary pathway. (A) Phylogenetic analysis of rbcL. GenBank accession numbers of the sequences used for constructing the phylogenetic tree of rbcL were as follows: Ulva prolifera (FJ042888), Thalassiosira pseudonana (YP_874498), Flaveria bidentis (ADW80649), Flaveria trinervia (ADW80661), Flaveria pringlei (ADW80648), Zea mays (NP_043033), Sorghum bicolor (ABK79504), Oryza sativa (CAG34174), Saccharum officinarum (YP_054639), Arabidopsis thaliana (AAB68400), Volvox carteri (ACY06055), Chlamydomonas reinhardtii (ACJ50136), Ostreococcus tauri (YP_717262), Ectocarpus siliculosus (CBH31935), Populus tremula (CAD12560), and Eleocharis vivipara (CAQ53780). (B) Phylogenetic analysis of PPDK. GenBank accession numbers of the sequences used for constructing the phylogenetic tree of PPDK were as follows: Ulva prolifera (JN936854), Thalassiosira pseudonana (XP_002290738), Flaveria bidentis (AAA86941), Flaveria trinervia (CAA55703), Flaveria pringlei (CAA53223), Zea mays (ADC32810), Sorghum bicolor (AAP23874), Oryza sativa (CAA06247), Saccharum officinarum (AAF06668), Arabidopsis thaliana (AEE83621), Volvox carteri (XP_002955807), Chlamydomonas reinhardtii (XP_001702572), Ostreococcus tauri (XP_003075283), Ectocarpus siliculosus (CBN74442), Populus tremula (CAX83740), and Eleocharis vivipara (BAA21654).
Analysis of rbcL and PPDK gene expression under various forms of stress
Relative quantitative PCR were carried out to determine the differences in expression levels of rbcL and PPDK genes under the different stress treatments. Figures 3A and 3B show the profiles of expression of rbcL and PPDK as affected by desiccation for varying lengths of time. The expression levels of rbcL and PPDK under normal conditions were taken as 1. The expression levels of rbcL decreased slowly with time, whereas those of PPDK increased steadily at first, peaking (a 4.9-fold increase) at 2 h, and decreased thereafter. Levels of salinity affected the expression markedly compared to that under normal salinity (30‰), which was taken as 1. The transcript levels of both rbcL and PPDK increased at lower and higher levels of salinity but then decreased at very high and very low salinity (Fig. 3C and 3D). Changes in expression levels under different light intensities are shown in Figures 3E and 3F. For each gene, the expression under 50 µmol m−2 s−1 was taken as 1. The expression level of rbcL in the dark was similar to that under normal light intensity, whereas that of PPDK was up-regulated 1.5-fold in the dark. The expression level of rbcL peaked at 300 µmol photons m−2 s−1, while that of PPDK peaked at 600 µmol photons m−2 s−1. Although the expression of PPDK decreased under high light intensity, it was still higher than it was under normal light intensity. Moreover, the effect of light intensities on PPDK was significantly higher than it was on rbcL. The expression of rbcL and PPDK at normal temperature (15°C) was taken as 1. The expression levels of rbcL reached the lowest point at 20°C, whereas those of PPDK were reached at 25°C. The expression of both rose at both higher and lower temperatures (Fig. 3G and 3H).
Data are means of three independent experiments (±SD). Relative mRNA expression of rbcL and PPDK exposed to different stress conditions: (A, B) desiccation for different durations up to 5 h, (C, D) different salt concentrations for 3 h, (E, F) different light intensities for 3 h, (G, H) different temperatures for 3 h.
Activity of RuBP carboxylase and PPDK
The activity of RuBP carboxylase decreased significantly with the duration of desiccation, whereas that of PPDK increased with the duration up to 2 h, the peak value being 1.4 times the normal value, and decreased thereafter (Fig. 4A). The effects of salinity level on RuBP carboxylase activity and PPDK activity were consistent (Fig. 4B): enzyme activity increased at low and high levels of salinity but then decreased at very low and very high values. Different light intensities clearly influenced the activity of both enzymes in a similar direction: the activity began to rise initially, peaked at 300 or 600 µmol photons m−2 s−1, and decreased thereafter as light intensity increased further (Fig. 4C). There was almost no difference in the activity of RuBP carboxylase and PPDK between the level under darkness and that under normal light intensity. Temperature also affected both enzymes significantly and similarly (Fig. 4D): RuBP carboxylase reached minimum activity at 20°C and PPDK at 25°C. The activity of both rose with increasing and decreasing temperatures.
Assay of photosynthetic rate
The optimum quantum yield (Fv/Fm) and effective PSII quantum yield (Y II) reached higher levels under normal conditions (15°C, 50 µmol photons m−2 s−1) and achieved the maximum values at 25°C, 100 µmol photons m−2 s−1 (Fig. 5). Neither was markedly affected by salinity or temperature, but both decreased rapidly under prolonged desiccation and high light intensities.
Studies of photosynthetic pathways of marine macroalgae are scanty, and we have a very limited understanding of the mechanisms controlling the altered cell biology and morphology associated with C4 Ulva species. In the present study, we found that almost all transcripts encoding the proteins required for the core C4 cycle have higher steady-state mRNA levels, suggesting that the C4 pathway does exist and that the activity of the C4 cycle enzymes is controlled at least partially at the level of transcript abundance (Fig. 1). The different expression profiles and product accumulations of rbcL and PPDK indicated that these two genes had respectively taken part in C3 and C4 core cycles under different conditions. We acquired a full-length cDNA sequence of PPDK, a key enzyme of the C4 pathway, to gain insights into the evolutionary optimization of C4 biochemistry in Ulva. The combination of photosynthetic, anatomical, and molecular datasets enabled us to isolate some of the steps in C4 evolution and provides fertile new ground for developing hypotheses about anatomical and ecological conditions that promote the evolution of this complex trait.
C4 photosynthesis is a series of anatomical and biochemical modifications that concentrate CO2 around the carboxylating enzyme Rubisco, thereby increasing photosynthetic efficiency in conditions promoting high rates of photorespiration. C4 plants are believed to have evolved gradually from C3 plants through several intermediate stages of C3–C4 plants . However, the evolutionary processes giving rise to C3–C4 intermediates and C4 plants are yet to be elucidated. Phylogenetic analysis of PPDK revealed that C4-like photosynthesis in green algae has multiple independent origins (Fig. 2), a result that is consistent with the results from diatoms , , . Relative studies on diatoms reveal that they have obtained a redundant set of carboxylation and decarboxylation enzymes during complicated endosymbiosis events, which could potentially constitute C4-type pathways including lateral-gene transfer (LTG) . Higher plants were exposed to much higher pCO2 at the beginning of evolutional history but then became starved for CO2 by a steep decrease of CO2 and increase of O2. These changes were a major driving force for land plants to develop C4 metabolism for suppression of photorespiration. Analogous evolutionary events might have taken place in the marine environment without loss of biophysical CCM .
Information about C4-related enzyme variations under various treatments is considerable. In Egeria densa, transfer from low temperature and light to high temperature and light conditions induced increases in the activities and amounts of both PEPC and NADP-ME. After 3 d of treatment, PEPC specific activity increased about 1.7 times relative to values in plants at LTL, whereas NADP-ME activity increased 1.26 times . The submersed monocot Hydrilla verticillata is a facultative C4 NADP-malic enzyme (NADP-ME) plant in which the C4 and C3 cycles co-exist in the same cell. The transcript expression of PEPC in H. verticillata was substantially up-regulated during light stress . In U. prolifera, both C3 and C4 pathway enzymes exist under normal conditions (Fig. 4). The expression levels of rbcL and PPDK increased under stress conditions, such as high salinity, low salinity, high temperature, and low temperature, but the levels of PPDK were higher than those of rbcL by 3.25, 4.25, 2.8 and 4.5 times, respectively (Fig. 3). The expression levels of rbcL decreased slowly with desiccation time, whereas those of PPDK increased steadily at first and decreased thereafter. These results indicate that both C3 and C4 cycles may function under normal conditions in U. prolifera, while C4 photosynthesis may play a more significant role under stress conditions.
Ulva prolifera is a green macroalga with single-layered tubular thalli (Fig. 6A). It differs from most other multi-cellular C4 land plants, in which, with few exceptions –, the assimilation of CO2 is distributed over two cell types, the mesophyll cells (MCs) and the bundle sheath cells (BSCs) . The distribution of CO2 assimilation over two distinct cell types requires a massive flux of metabolites between MCs and BSCs , . Bienertia sinuspersici, a land plant, is a recently discovered species with a unique form of C4 photosynthesis. In this single-cell C4 species (SCC4), the carbon concentrating mechanism does not depend on cooperation between M and BS cells, as it does in Kranz-type C4 species. Rather, it possesses a unique chlorenchyma with two functional and biochemically different chloroplast types within photosynthetic cells. Peripheral chloroplasts are spatially separated by a large vacuole from chloroplasts clustered in a central compartment (C-CP). This structural arrangement allows for enrichment of CO2 in the Rubisco-containing C-CP, ultimately repressing photorespiration, similar to the mechanism in Kranz-type C4 plants. In U. prolifera, chloroplasts aggregate lucipetally along the outer side of the layer, and there are apparently no functionally or biochemically different chloroplast types (Fig. 6B), so the chloroplast differentiation mechanism is not fit for this species. Indeed, information about the mechanisms controlling the altered cell biology and morphology associated with C4 photosynthesis is very limited. The C4 cycle likely affects not only the relatively small number of enzymes and transport proteins needed to perform the core reactions but, given the consequences to the ecological performance of the plants, also a range of other processes .
A, Transmission electron microscopy of transverse section. EX, external of cavity; IN, inner of cavity. Bar, 5 µm. B, Longitudinal and transverse section view with an optical microscope. TS, transverse section; LS, longitudinal section. Bar, 20 µm.
In the present study, the results showed that the expression of PPDK in U. prolifera was higher under some daily-encountered stress conditions, such as desiccation, high light intensity, high temperature, and low temperature (Figs. 3, 4). High temperature is a major environmental requirement for C4 evolution because it directly stimulates photorespiration and dark respiration in C3 plants , . The availability of CO2 as a substrate also declines at elevated temperature because of the reduced solubility of CO2 relative to O2 . Aridity and salinity are important because they promote stomatal closure and thus reduce intercellular CO2 levels, again stimulating photorespiration and aggravating a CO2 substrate deficiency . C4 photosynthesis has been found in some marine algae. The implications of marine C4 photosynthesis are very significant. The presence of the C4 pathway is likely to influence algal sensitivity to changes in CO2 concentrations. As in terrestrial ecosystems, C4 photosynthesis may therefore be a factor that is shaping species distribution and succession if it occurs in only some members of the phytoplankton. It could operate both on geological timescales and in response to the present rise in atmospheric CO2 concentrations. If C4 photosynthesis can account for a significant portion of marine carbon fixation in some species, it will affect various aspects of marine ecology and biogeochemistry . C4 photosynthesis is a complex biological trait that enables plants to either accumulate biomass at a much faster rate or live in adverse environments compared with “ordinary” plants , . Our results suggest that photosynthetic organisms may have evolved a unique mechanism for coping with environmental transition, before losing CCM, and the C4 pathway may have first formed in intertidal pluricellular green algae before plants colonized terrestrial habitats. An added benefit of the C4 syndrome is improved nitrogen- and water-use efficiencies that have likely contributed to their global distribution and high rates of productivity –. Therefore, the manmade environmental changes, such as CO2 rise and eutrophication, stimulate the expression of the C4 pathway, while the cooperation of CCM and the C4 pathway may enhance the capacity of photosynthesis, which may be one of the most important factors leading to the rapid accumulation of the vast biomass of U. prolifera in the green tide that has occurred in the Yellow Sea in four consecutive years since 2008 , .
Conceived and designed the experiments: JX NY XZ. Performed the experiments: JX XF SM SC. Analyzed the data: JX DX ZZ JM. Contributed reagents/materials/analysis tools: XF XZ NY. Wrote the paper: JX.
- 1. Gowik U, Brautigam A, Weber KL, Weber APM, Westhoff P (2011) Evolution of C4 photosynthesis in the genus Flaveria: how many and which genes does it take to make C4? Plant Cell 23: 2087–2105.
- 2. Weber APM, von Caemmerer S (2010) Plastid transport and metabolism of C3 and C4 plants – comparative analysis and possible biotechnological exploitation. Curr Opin Plant Biol 13: 257–265.
- 3. Sage RF (2004) The evolution of C4 photosynthesis. New Phytologis 161: 341–370.
- 4. Gowik U, Westhoff P (2011) The Path from C3 to C4 Photosynthesis. Plant Physiol 155: 56–63.
- 5. Beer S, Israel A, Drechsler Z, Cohen Y (1990) Photosynthesis in Ulva fasciata V. Evidence for an Inorganic Carbon Concentrating System, and Ribulose-1,5-Bisphosphate Carboxylase/Oxygenase CO2 Kinetics. Plant Physiol 94: 1542–1546.
- 6. Drechsler Z, Beer S (1991) Utilization of Inorganic Carbon by Ulva lactuca. Plant Physiol 97: 439–1444.
- 7. Tachibana M, Allen AE, Kikutani S, Endo Y, Bowler C (2011) Localization of putative carbonic anhydrases in two marine diatoms, Phaeodactylum tricornutum and Thalassiosira pseudonana. Photosynth. Res 109: 205–221.
- 8. Edwards GE, Franceschi VR, Voznesenskaya EV (2004) Single-cell C4 photosynthesis versus the dual-cell (Kranz) paradigm. Annu Rev Plant Biol 55: 173–196.
- 9. Keeley JE (1999) Photosynthetic pathway diversity in a seasonal pool community. Funct Ecol 13: 106–118.
- 10. Ehleringer JR, Cerling TE, Helliker BR (1997) C4 photosynthesis, atmospheric CO2, and climate. Oecologia 112: 285–299.
- 11. Reiskind JB, Madsen TV, Vanginkel LC, Bowes G (1997) Evidence that inducible C4-type photosynthesis is a chloroplatic CO2-concentrating mechanism in Hydrilla, a submersed monocot. Plant, Cell and Environment 20: 211–220.
- 12. Tsuji Y, Suzuki I, Shiraiwa Y (2009) Photosynthetic carbon assimilation in the coccolithophorid Emiliania huxleyi (Haptophyta): evidence for the predominant operation of the C3 cycle and the contribution of β -carboxylases to the active anaplerotic reaction. Plant Cell Physiol 50: 318–329.
- 13. Beer S, Israel A (1986) Photosynthesis of Ulva sp. III. 02 effects, carboxylase activities, and the CO2 incorporation pattern. Plant Physiol 81: 937–938.
- 14. Reinfelder JR, Kraepiel AML, Morel FMM (2000) Unicellular C4 photosynthesis in a marine diatom. Nature 407: 996–999.
- 15. Reinfelder JR, Milligan AJ, Morel FMM (2004) The role of the C4 pathway in carbon accumulation and fixation in a marine diatom. Plant Physiol 135: 2106–2111.
- 16. 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.
- 17. McGinn PJ, Morel FMM (2008) Expression and Inhibition of the carboxylating and decarboxylating enzymes in the photosynthetic C4 pathway of marine diatoms. Plant Physiol 146: 300–309.
- 18. 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.
- 19. Montsant A, Jabbari K, Maheswari U, Bowler C (2005) Comparative genomics of the pennate diatom Phaeodactylum tricornutum. Plant Physiol 137: 500–513.
- 20. Derelle E, Ferraz C, Rombauts S, Rouze P, Worden AZ, et al. (2006) Genome analysis of the smallest free-living eukaryote Ostreococcus tauri unveils many unique features. Proc Nati Acad Sci USA 103: 11647–11652.
- 21. Worden AZ, Lee JH, Mock T, Rouze P, Simmons MP, et al. (2009) Green evolution and dynamic adaptations revealed by genomes of the marine picoeukaryotes Micromonas. Science: 324: 268–272.
- 22. Leliaert F, Smith DR, Moreau H, Herron MD, Verbruggen H, et al. (2012) Phylogeny and molecular evolution of the green algae. Crit Rev Plant Sci 31: 1–46.
- 23. 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: e1426.
- 24. Parker MS, Mock T, Armbrust EV (2008) Genomic insights into marine microalgae. Annu Rev Genet 42: 619–645.
- 25. Bowler C, Vardi A, Allen AE (2010) Oceanographic and biogeochemical insights from diatom genomes. Ann Rev Mar Sci 2: 333–365.
- 26. Charlier RH, Morand P, Finkl CW, Thys A (2007) Green tides on the Brittany coasts. Environ Res Eng Manage 3: 52–59.
- 27. Ye NH, Zhang XW, Mao YZ, Liang CW, Xu D, et al. (2011) ‘Green tides’ are overwhelming the coastline of our blue planet: taking the world's largest example. Ecol Res 26: 477–485.
- 28. Leliaert F, Zhang X, Ye N, Malta EJ, Engelen AH, et al. (2009) Research note: Identity of the Qingdao algal bloom. Phycol Res 57: 147–151.
- 29. Hayden H, Blomster J, Maggs CA, Silva PC, Stanhope MJ, et al. (2003) Linnaeus was right all along: Ulva and Enteromoropha are not distinct genera. Eur J Phycol 38: 277–294.
- 30. Dong M, Zhang X, Zhuang Z, Zou J, Ye N, et al. (2011) Characterization of the LhcSR gene under light and temperature stress in the green alga Ulva linza. Plant Mol Biol Rep. https://doi.org/10.1007/s11105-011-0311-8
- 31. Zhang X, Xu D, Mao Y, Li Y, Xue S, et al. (2011) Vegetative fragments of Ulva prolifera in settlement confirmed as an important seed source for succession of a large-scale green tide bloom. Limnol Oceanogr 56:
- 32. Bischof K, Krabs G, Wiencke C, Hanelt D (2002) Solar ultraviolet radiation affects the activity of ribulose-1,5-bisphosphate carboxylase-oxygenase and the composition of photosynthetic and xanthophyll cycle pigments in the intertidal green alga Ulva lactuca L.. Planta 215: 502–509.
- 33. Aquino RS, Grativol C, Mourao PAS (2011) Rising from the Sea: correlations between sulfated polysaccharides and salinity in plants. PLoS One 6: e18862.
- 34. Kremer BP, Küppers U (1977) Carboxylating enzymes and pathway of photosynthetic carbon assimilation in different marine algae - Evidence for the C4-Pathway? Planta 133: 191–196.
- 35. Karekar MD, Joshi GV (1973) Photosynthetic carbon metabolism in marine algae. Bot Marina 16: 216–220.
- 36. Joshi GV, Karekar MD, Gowda CA, Bhosale L (1974) Photosynthetic carbon metabolism and carboxylating enzymes in algae and mangrove under saline conditions. Photosynthetica 8: 51–52.
- 37. Troughton JH, Card KA, Hendy CH (1974) Photosynthetic pathways and carbon isotope discrimination by plants. Carnegie Institution of Washington Yearbook 73: 768–80.
- 38. Wang WL, Yeh HW (2003) δ13C values of marine macroalgae from Taiwan. Bot. Bull. Acad. Sin. 44: 107–112.
- 39. Mercado JM, Santos CB, Perez-Llorens JL, Vergara JJ (2009) Carbon isotopic fractionation in macroalgae from Cadiz Bay (Southern Spain): Comparison with other bio-geographic regions. Estuar Coast Shelf S 85: 449–458.
- 40. Hatch MD (1987) C4 photosynthesis: a unique blend of modified biochemistry, anatomy and ultrastructure. Biochem. Biophys. Acta 895: 81–106.
- 41. Ishimaru K, Ohkawa Y, Ishige T, Tobias DJ, Ohsugi R (1998) Elevated pyruvate orthophosphate dikinase (PPDK) activity alters carbon metabolism in C3 transgenic potatoes with a C4 maize PPDK gene. Physiol Plantarum 103: 340–346.
- 42. Chen M, Liu T, Chen X, Chen L, Zhang W, et al. (2011) Subcritical co-solvents extraction of lipid from wet microalgae pastes of Nannochloropsis sp. Eur J Lipid Sci Technol. https://doi.org/10.1002/ejlt.201100120
- 43. Huson DH, Auch AF, Qi J, Schuster SC (2007) Megan Analysis of Metagenome Data. Genome Res 17: 377–386.
- 44. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876–4882.
- 45. Chenna R, Sugawara H, Koike T, Lopez R, Gibson TJ, et al. (2003) Multiple sequences alignment with the clustal series of programs. Nucleic Acids Res 31: 3497–3500.
- 46. Liu XM, Anderson JM, Pijut PM (2010) Cloning and characterization of Prunus serotina AGAMOUS, a putative flower homeotic gene. Plant Mol Biol Rep 28: 193–203.
- 47. Tamura K, Dudley J, Nei M, Kumar S (2007) MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Molecular Biology and Evolution 24: 1596–1599.
- 48. Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods 25: 402–408.
- 49. Gerard VA, Driscoll T (1996) A spectrophotometric assay for RuBisCO activity: application to the kelp Laminaria saccharina and implications for radiometric assays. J Phycol 32: 880–884.
- 50. Sayre RT, Kennedy RA, Pringnitz DJ (1979) Photosynthetic enzyme activities and localization in Mollugo verticillata population differing on the leaves of C3 and C4 cycle operations. Plant Physiol. 64: 293–299.
- 51. Fleming ED, Bebout BM, Castenholz RW (2007) Effects of salinity and light intensity on the resumption of photosynthesis in rehydrated cyanobacterial mats from Baja California Sur, Mexico. J Phycol 43: 15–24.
- 52. Ueno O (2011) Structural and biochemical characterization of the C3–C4 intermediate Brassica gravinae and relatives, with particular reference to cellular distribution of Rubisco. J Exp Bot. pp. 1–9.
- 53. Bowler C, Allen AE, Badger JH, Grimwood J, Jabbari K, et al. (2008) The Phaeodactylum genome reveals the evolutionary history of diatom genomes. Nature 456: 239–244.
- 54. Moustafa A, Beszteri B, Maier UG, Bowler C, Valentin K, et al. (2009) Genomic footprints of a cryptic plastid endosymbiosis in diatoms. Science 324: 1724–1726.
- 55. Matsuda Y, Nakajima K, Tachibana M (2011) Recent progresses on the genetic basis of the regulation of CO2 acquisition systems in response to CO2 concentration. Photosynth. Res 109: 191–203.
- 56. Casati P, Lara MV, Andreo CS (2000) Induction of a C4-like mechanism of CO2 fixation in Egeria densa, a submersed aquatic Species. Plant Physiol 123: 1611–1621.
- 57. Rao S, Reiskind J, Bowes G (2006) Light Regulation of the Photosynthetic Phosphoenolpyruvate Carboxylase (PEPC) in Hydrilla verticillata. Plant Cell Physiol 47: 1206–1216.
- 58. Reiskind JB, Berg RH, Salvucci ME, Bowes G (1989) Immunogold localization of primary carboxylases in leaves of aquatic and a C3–C4 intermediate species. Plant Sci 61: 43–52.
- 59. Keeley JE (1998) C4 photosynthetic modifications in the evolutionary transition from land to water in aquatic grasses. Oecologia 116: 85–97.
- 60. Voznesenskaya EV, Franceschi VR, Kiirats O, Freitag H, Edwards GE (2001) Kranz anatomy is not essential for terrestrial C4 plant photosynthesis. Nature 414: 543–546.
- 61. Voznesenskaya EV, Franceschi VR, Kiirats O, Artyusheva EG, Freitag H, et al. (2002) Proof of C4 photosynthesis without Kranz anatomy in Bienertia cycloptera (Chenopodiaceae). Plant J 31: 649–662.
- 62. Voznesenskaya EV, Edwards GE, Kiirats O, Artyusheva EG, Franceschi VR (2003) Development of biochemical specialization and organelle partitioning in the single-cell C4 system in leaves of Borszczowia aralocaspica (Chenopodiaceae). Am J Bot 90: 1669–1680.
- 63. von Caemmerer S, Furbank RT (2003) The C4 pathway: an efficient CO2 pump. Photosynth. Res 77: 191–207.
- 64. Bräutigam A, Hofmann-Benning S, Weber APM (2008) Comparative proteomics of chloroplast envelopes from C3 and C4 plants reveals specific adaptations of the plastid envelope to C4 photosynthesis and candidate proteins required for maintaining C4 metabolite fluxes. Plant Physiol 148: 568–579.
- 65. Brautigam A, Kajala K, Wullenweber J, Sommer M, Gagneul D, et al. (2011) An mRNA blueprint for C4 photosynthesis derived from comparative transcriptomics of closely related C3 and C4 species. Plant Physiol 155: 142–156.
- 66. Brooks A, Farquhar GD (1985) Effect of temperature on the CO2/O2 specificity of ribulose-1,5-bisphosphate carboxylase/oxygenase and the rate of respiration in the light. Planta 165: 397–406.
- 67. Sharkey TD (1988) Estimating the rate of photorespiration in leaves. Physiol. Plantarum 73: 147–152.
- 68. Jordan DB, Ogren WL (1984) The CO2/O2 specificity of ribulose 1,5-bisphosphate carboxylase/oxygenase. Planta 161: 308–313.
- 69. Riebesell U (2000) Carbon fix for a diatom. Nature 407: 959–960.
- 70. Osborne CP, Freckleton RP (2009) Ecological selection pressures for C4 photosynthesis in the grasses. Proc R Soc Lond B Biol Sci 276: 1753–1760.
- 71. Tilman D, Hill J, Lehman C (2006) Carbon-negative biofuels from low-input high-diversity grassland biomass. Science 314: 1598–1600.
- 72. Edwards EJ, Osborne CP, Strömberg CAE, Smith SA, Bond WJ, et al. (2010) The origins of C4 grasslands: integrating evolutionary and ecosystem science. Science 328: 587–591.
- 73. Brutnell TP, Wang L, Swartwood K, Goldschmidt A, Jackson D, et al. (2010) Setaria viridis: a model for C4 photosynthesis. Plant Cell 22: 2537–2544.