Open access peer-reviewed chapter

Absorption and Transport of Inorganic Carbon in Kelps with Emphasis on Saccharina japonica

Written By

Yanhui Bi and Zhigang Zhou

Submitted: 07 September 2015 Reviewed: 26 January 2016 Published: 30 March 2016

DOI: 10.5772/62297

From the Edited Volume

Applied Photosynthesis - New Progress

Edited by Mohammad Mahdi Najafpour

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Due to the low CO2 concentration in seawater, macroalgae including Saccharina japonica have developed mechanisms for using the abundant external pool of HCO3− as an exogenous inorganic carbon (Ci) source. Otherwise, the high photosynthetic efficiency of some macroalgae indicates that they might possess CO2 concentrating mechanisms (CCMs) to elevate CO2 concentration intracellularly around the active site of ribulose-1, 5-bisphosphate carboxylase/oxygenase (RuBisCo). As the photosynthetic modes of macroalgae are diverse (C3, C4 or a combination of C3 and C4 pathway), CCMs in different carbon fixation pathways should vary correspondingly. However, both in C3 and C4 pathways, carbonic anhydrase (CA) plays a key role by supplying either CO2 to RuBisCO or HCO3− to PEPC. Over the past decade, although CA activities have been detected in a number of macroalgae, genes of CA family, expression levels of CA genes under different CO2 concentrations, as well as subcellular location of each CA have been rarely reported. Based on analysis the reported high-throughput sequencing data of S. japonica, 12 CAs of S. japonica (SjCA) genes were obtained. Neighbor-Joining (NJ) phylogenetic tree of SjCAs constructed using Mega6.0 and the subcellular location prediction of each CA by WoLFPSORT are also conducted in this article.


  • Macroalgae
  • Inorganic carbon uptake
  • C3 and C4 metabolism
  • Carbonic anhydrase
  • Saccharina japonica

1. Introduction

Kelps demonstrate high photosynthetic rates. According to the reports, productivity of large brown algae (e.g., Macrocystis, Laminaria, Ecklonia, Sargassum) ranges from 1000 to 3400 g m−2yr−1C or about 3300 to 11,300 g m−2yr−l dry weight, and red algae show a similar range of production. Cultivated macroalgae can yield even higher values. The projected yield of cultivated Laminaria japonica on an annualized basis is equivalent to 1300 t ha−1 fresh weight or 6.5 times the maximum projected yield for sugarcane, the most productive of land plants under cultivation. In general, 45% yield of the dry weight of plants is accounted by carbon, which is assimilated in plant through Calvin cycle. The high productivities of kelps indicate their higher photosynthetic efficiency than C4 terrestrial plants [1].

The enzyme ribulose-1, 5-bisphosphate carboxylase/oxygenase (RuBisCo) is crucial in CO2 assimilation. This bifunctional enzyme could catalyse the initial steps of photosynthetic carbon reduction and photorespiratory carbon oxidation cycles by combining CO2 and O2 with ribulose-1, 5-bisphosphate (RuBP) [2, 3]. RuBP carboxylation determines the net photosynthetic efficiency of photoautotrophs [4]. However, RuBisCo has a surprisingly low affinity for CO2 and the oxygenase activity is intrinsic to RuBisCo. For kelps, the enzymatic efficiency of RuBisCo is also limited by the low concentration and diffusion coefficient of CO2 in seawater [5]. At a natural pH of about 8, the major part of the dissolved inorganic carbon (DIC) is in the form of bicarbonate (HCO3), and only about 12 μM is present as dissolved CO2 [6], which is much lower than the half-saturation constant (Ks) of RuBisCo for CO2 ranges from 30 μM to 60 μM in marine macroalgae [7, 8]. To support photosynthesis and growth, seaweeds require an exogenous inorganic carbon (Ci), while only CO2 and HCO3 can be used as a CO2 source for photosynthesis. Due to the low CO2 concentration in seawater, it is not surprising that most seaweed have developed mechanisms for using the abundant external pool of HCO3 as an exogenous Ci source [911]. And it seems likely that those macrophytes that are able to use HCO3 would possess advantages compared with that rely solely on diffusive CO2 entry. Here the question is how Ci is absorbed, transported to supply high CO2 concentration around RuBisCo in kelps since unlike CO2, HCO3 cannot diffuse through the lipid bilayer of the plasma membrane [12] and the produced or absorbed CO2 are readily leaked out due to the high CO2 permeability of cytomembrane. Otherwise, different models of photosynthesis such as C3, C4 and CAM might employ different CCMs in kelps. Thus, this review mainly focuses on the mechanisms of Ci absorption, transportation and concentration mechanisms of multicellular marine algae, including representatives of Chlorophyceae, Rhodophyceae and Phaeophyceae with different photosynthetic types.


2. Photosynthetic modes of macroalgae

As with terrestrial angiosperms where a single family may possess species with divergent photosynthetic modes [13], the marine macroalgal divisions also exhibit diversity. The photosynthetic carbon fixation pathways of marine macrophytic algae generally follow that of C3 plants [14]. However, for certain genera, a number of studies have shown photosynthesis to possess C4-like photosynthetic characteristics, including the high phosphoenolpyruvate carboxykinase (PEPCK) activity with low phosphoenolpyruvate carboxylase (PEPC) activity, little photorespiration and the labelling of malate and aspartate as an early product of carbon fixation. Based on this, it has been suggested that these macroalgae are of the C4 type, or a combination of C3 and C4, type [1517], although Kremer and Küppers [18] had contradicted the decision whether a species is a C4 plant or not based only on chromatographic and enzymatic analysis. In recent decades, our understanding of the possible metabolic pathways of macroalgae has been extended with using the available sequencing resources and molecular technologies and applying molecular approaches. Reiskind et al. [19] reported that a limited C4-like system in the green alga Udotea with the high PEPCK activity and low PEPC activity was a novel characteristic. Whereafter, Reiskind and Bowes [20] found that when PEPCK activity was inhibited in vivo with 3-mercaptopicolinic acid, thallus photosynthesis was decreased by 70% and the labelling of early photosynthetic products such as malate and aspartate was reduced by 66% and thus provided new evidences for the existence of C4 acid metabolism in this green alga. In contrast to Udotea, Codium, a macroalga closely related to Udotea, exhibits gas exchange characteristics that resemble terrestrial C3 plants, and neither C4 acids nor PEPCK plays a part in photosynthesis [19]. This demonstrates the diversity of photosynthetic mechanisms in the Chlorophyta. Ulva, a common green seaweed, was previously reported as a typical C3 plant based on some biochemical evidences that 3-phosphoglyceric acid (3-PGA) was the main primary product formed photosynthetically and a high RuBPcase/PEPcase ratio was found in it [21], while, recently, it was reported that Ulva possessed rather comprehensive carbon fixation pathways including C3, C4 and CAM mechanisms because key genes of enzymes involved in these photosynthetic modes were got from the expressed sequence tag (EST) using Kyoto encyclopedia of genes and genomes (KEGG) [22]. Recently, C4-like carbon fixation pathway was also found in representatives of Rhodophyceae and Phaeophyceae based on the analysis of ESTs or transcriptomes. In red algae, Fan et al. [23] speculated that the sporophyte of Pyropia haitanensis most likely possesses a C4-like carbon fixation pathway since genes of the key enzymes in the PCK-type C4 carbon-fixation pathway were abundantly transcribed. Wang et al. [24] assumed that a C4-like carbon-fixation pathway might play a special role in fixing inorganic CO2 in Porphyra yezoensis with the evidence that except pyruvate-phosphate dikinase all genes involved in C4-pathway were discovered from the transcriptome. Xu et al. [25] had reported that PEPCK, an important enzyme in carbon fixation in C4 plants, had very high activity in the sporophyte of L. japonica. Besides, haploid gametophytes and diploid sporophytes of some marine macroalgae with dimorphic life cycles might even employ different photosynthetic mode. Wang et al. [24] found that both the RuBisCo content and the initial carboxylase activity were notably higher in gametophytes than in the sporophytes of four seaweed species — P. yezoensis, P. haitanensis, Bangia fuscopurpurea (Rhodophyte) and L. japonica (Phaeophyceae). They assumed that in the sporophyte of these algae, the major carbon fixation pathway may be a C4-like carbon fixation pathway, and thus a high abundance of RuBisCo would not be necessary for the sporophytes. And for L. japonica, the higher RuBisCo content and activity in gametophyte was corresponding to the lower photosynthetic rate, which implied there might be a greater difference between sporophytes and gametophytes of this alga in their photosynthetic mode. Conclusively, the existence of C4-like pathway in macroalgae has been verified using more evidence, while the distribution between C3 and C4 pathways was unknown during growth of macroalgae with comprehensive carbon fixation pathways including C3 and C4.

In C3 and C4 metabolisms, CO2 is the substrate of RuBisCo and assimilated through the Calvin cycle. In this cycle, CO2, catalysed with RuBisCo, combines with RuBP to form two molecules of 3-PGA. PGA is reduced to triose. RuBisCo, a bifunctional enzyme, may catalyse the combination of RuBP and CO2 for photosynthetic carbon reduction or may combine with O2 for C2 photorespiration [3]. The ratio of CO2 to O2 around RuBisCo is a major factor for the enzyme to choose the photosynthetic carbon reduction or C2 photorespiration carbon oxidation [26]. The low CO2 concentration around RuBisCo may not only impose restrictions on photosynthesis but also cause permanent light injuries to photosynthetic organelle [2729]. The speciation of DIC (Ci) is pH dependent. Above pH 4.5, the proportion occurring as CO2 (aq) decreases and HCO3 increases, while above pH 8.3, the bicarbonate equivalence point, the equilibrium begins to shift towards carbonate (CO32−). In the upper layer of the oceans, HCO3 ions predominate, and the dissolved CO2 represents only about 1% of the total dissolved carbon with a concentration of about 21 μM [30]. The Km (CO2) value of RuBisCO is significantly higher than this, having been reported as being as high as 200 μM in some cyanobacteria [31]. To survive under the selective pressure of low CO2 concentration, high permeability of CO2 for plasma membrane and low affinity of CO2 for RuBisCo, many algae, including macroalgae living in the subtidal zone, have evolved with inorganic CCM that allows them to overcome this potentially limiting shortage of CO2 [9, 3236]. So, the productivity of most macroalgae is not currently considered limited by DIC. Unlike terrestrial C4 plants possessing Kranz anatomy to prevent futile recycling of CO2 by segregating the initial carboxylation and decarboxylation reactions in different cells, macroalgae concentrate CO2 internally, which is mediated by Ci transporters at the plasma membrane or chloroplast envelope and CA. As for carboxylases are different between C3 and C4 metabolism, Ci acquisition, transportation and concentration mechanisms might be diverse.

Based on a series of reports on the presence of CCM in blue-green algae and Chlamydomonas (Chlamydomonas reinhardtii) and some other microalgae [3740], Badger [41] reported that the CCM of algae possess at least three functional elements: (1) the transportation of the Ci dissolved in seawater into cells in the form of CO2 and/or HCO3 ; (2) the accumulation of the Ci in cells in the form of HCO3 , forming pools of the dissolved Ci and (3) the delivery of CO2 to the periphery of RuBisCo from such pools.


3. Inorganic carbon absorption mechanisms of macroalgae

Figure 1.

A schematic diagram on the photosynthetic carbon physiology of some macroalgae revised from [45].

The methods of CO2 and/or HCO3 absorption of macroalgae cells (Figure 1) include the following: (1) non-CCM macroalgae (that do not possess or use CCM) rely exclusively on diffusive uptake of CO2, (2) CCM macroalgae uptake of Ci, as CO2 and/or HCO3 via mechanisms of the external carbonic anhydrase (CAext) mechanism, the anion exchange (AE) transport mechanism, the plasma membrane associated with H+-ATPase mechanism and passive transport of CO2 by diffusion. In the first mechanism, HCO3 in the periplasmic space is converted to CO2 at the presence of CAext, an enzyme that is located in the cell wall in the majority of seaweeds and could be inhibited by the membrane impermeable acetazolamide (AZ), and then the resulting CO2 is readily taken into the cell by passive diffusion. This seems to be the most prevalent for HCO3 utilization among seaweeds [42, 43], but it may be non-functional under high pH (>9.00) [44, 45]. The AE transport mechanism is HCO3 direct uptake through the AE protein in plasma membrane [11, 43, 4648], which is 4,4'-diisothiocyano-stilbene-2,2'-disulfonate (DIDS) sensitive. This operates equally well at pH 8.4 and 9.4 [44, 45]. H+-ATPase mechanism refers to a plasma membrane associated H+-ATPase pump that extrudes the excess cellular H+ to the outside of the plasma membrane facilitating a H + / HCO3 co-transportation or enhancement of the external uncatalysed dehydration of HCO3 to CO2 in the periplasmic space [49]. However, this has only been reported in some Laminariales such as S. latissima and L. digitata. Along with the uptake of CO2 and/or HCO3 , the internal charge balance (OH/H+) will be absolutely changed. To maintain intracellular ion balance, macroalgae employ diverse strategies. In AE mechanism, the active transport of HCO3 into the cell might result in an outward flux of OH [5053, 45] as this mechanism is involved in a one-for-one exchange of anions across the plasma membrane. The OH efflux can increase H+ in the cell [52]. To maintain the intracellular OH/H+ balance, H+ extrusion might be required. In macroalgae possessing H+-ATPase mechanism, their plasma membrane associated with H+-ATPase pump might extrude excess cellular H+ to the outside of the plasma membrane, while in macroalgae that do not have H+-ATPase pump in their plasma membrane, the regulation of intracellular ion balance might be related to a high activity of internal carbonic anhydrase (CAint), including the CA in cytoplasm, chloroplast stroma, thylakoid lumen and mitochondria [45].

The extent to which marine macroalgae are able to acquire HCO3 for photosynthesis varies among taxa and/or species, and the special strategies by which the alga acquire Ci is closely related to habitat including pH and depth, conferring as adaptation advantage to the alga [9, 33, 36, 5456]. Cornwall et al. [57] reported when light is low, CCM activity of macroalgae is reduced in favour of diffusive CO2 uptake and the proportion of non-CCM (diffusive uptake of CO2) species increased with depth. Otherwise, pH might also control Ci use by macroalgae. In U. lactuca, the CAext-mediated mechanism is the main method of HCO3 utilization under normal pH conditions, whereas when they were grown at high pH, direct uptake of HCO3 via a DIDS-sensitive mechanism can be induced [44]. Similar HCO3 utilizing mechanisms were found in another green macroalgae Enteromorpha intestinalis [54]. For the red alga Gracilaria gaditana, the HCO3 use is also carried out by the two DIC uptake mechanisms, in which the indirect use of HCO3 by an external CA activity being the main pathway and the potential contribution to HCO3 acquisition by the DIDS-sensitive AE mechanism was higher after culturing at a high pH [58]. However, these two mechanisms do not occur simultaneously, and the DIDS-sensitive mechanism is induced only under high pH. Solieria filiformis, another red marine macroalgae, in which the general form of Ci transported across the plasma membrane is CO2, but HCO3 acquisition takes place simultaneously between CAext mechanism and direct uptake [59]. CAext mechanism is also the main pathway for DIC acquisition for the species of Phaeophyta. S. latissima mainly uses CAext mechanism for HCO3 absorption, since when AZ is used to treat S. latissima, its photosynthetic efficiency drops by 80% [11]. Otherwise, S. latissima also has a H+-ATPase mechanism, of which the proton pump may support the antiport of H + / HCO3 or the discharge of H+, creating an acid environment in the periplasmic space and causing the dehydration of HCO3 into CO2 with CA to quickly diffuse into cells [49]. Similar to S. latissima, L. digitata also has a CAext mechanism of absorbing HCO3 and a P-H+-ATPase mechanism [49]. Gametophytes of Ectocarpus siliculosus utilize the CAext mechanism and the HCO3 transport protein [60] on the cell membrane to absorb HCO3 . Macrocystis pyrifera utilizes the CAext mechanism and the AE protein mechanism to absorb HCO3 , in which the main mechanism of HCO3 uptake is via AE protein and CAext contributes little [45]. For Sargassum henslowianum, like most seaweed, the main Ci acquisition strategy is also CAext metabolism, since its photosynthetic O2 evolution could be drastically depressed by AZ at pH 8.1 (i.e., the normal seawater pH value) and at pH 9.0. And direct uptake for HCO3 via DIDS-sensitive AE protein mechanism was unlikely to be present in Ci acquisition of this kelp, because the photosynthesis in either blade or receptacle tissue of this alga was not affected by DIDS [61]. For Hizikia fusiformis, CAext+ diffusive uptake of CO2 could support its metabolic requirements sufficiently since there is no known other active Ci transport mechanisms [62]. For S. japonica, Yue et al. [63] found that the Ci absorption of the CAext mechanism in its juvenile sporophytes accounts for 75% of the total Ci absorption in algae cells, whereas free CO2 absorption accounts for 25% only.

Thus, the CAext mechanism plays an important role in the CCM macroalgae absorption and the utilization of the relatively abundant HCO3 in seawater.


4. Ci transition process in CCMs of macroalgae

Ci acquisition mechanisms are extensively studied and well-known in microalgae [44, 38]. For instance, regardless of the Ci form (CO2 or HCO3 ) taken up by the microalga C. reinhardtii, HCO3 is the primary form accumulated into the cell to prevent CO2 leakage [38]. In macroalgae, most Ci use processes are speculated based on some biochemical evidence. For C3 photosynthesis, the CO2 that entered the cytoplasm is transformed into HCO3 under the catalytic action of CA in the cytoplasm and stored in the cytoplasm [38] to maintain the equilibrium of different forms of Ci and to regulate the pH value of the cytoplasm [26, 38]. The HCO3 in the cytoplasm enters the chloroplast stroma via the Ci transport protein on the chloroplast membrane, and the CO2 in the cytoplasm directly enters the stroma via the chloroplast membrane. In diatom Phaeodactylum tricornutum, genes with homology to bicarbonate transporters from SLC4 and SLC6 families, two HCO3 transporters studied thoroughly in human, were got from its genome and one of these SLC4-type HCO3 transporters has recently been confirmed to function as a Na+-dependent HCO3 transporter on the outer membrane [64, 65]. However, the molecular nature of HCO3 transporters of macroalgae is unknown now, and their similarity to those found in diatoms is uncertain. The transportation of Ci from the cytoplasm to the chloroplast is the major Ci flux in the cell and the primary driving force for the CCM. This flux drives the accumulation of Ci in the chloroplast stroma and generates a CO2 deficit in the cytoplasm, inducing CO2 influx into the cell. Given that the pH value of the chloroplast stroma is closer to 8, the stroma Ci is mostly enriched in the form of HCO3 , forming Ci pools [66]. In macroalgae, which have pyrenoids, HCO3 is putatively carried into the thylakoid by the Ci transport protein on the thylakoid membrane, forming CO2 in the thylakoid space under the catalytic action of thylakoid CA [67, 68]. The thylakoid membrane partially sinks into the pyrenoids [69], where the diffused CO2 is quickly fixed by the RuBisCo in the pyrenoids. The diffused CO2 from the thylakoid space outside the pyrenoids or the unfixed CO2 leaked from the pyrenoids is transformed into HCO3 under the action of CA in the starch sheath on the periphery of the pyrenoid, thus increasing the number of HCO3 pools in the matrix [70]. For macroalgae without pyrenoids, such as L. japonica, HCO3 entered the chloroplast stroma after being dehydrated under the action of chloroplast stroma CA and provided CO2 for the RuBisCo in the matrix (Figure 1).

For C4 photosynthesis, CA is required to convert CO2 to HCO3 in the cytosol, and thus supply PEPC with substrate. HCO3 will be fixed into malate. For non-PEPC algae with PEPCK, the CO2 entering the cytoplasm will be directly fixed in the form of four-carbon acid [71]. The produced four-carbon acid may be transported into the mitochondria, forming pyruvate after decarboxylation and CO2 release, which is fixed in the form of carbohydrate in the Calvin cycle. In fact, the presence of CA in C4 plants has been suggested to accelerate the rate of photosynthesis in C4 plants 104-fold over what it would be if this enzyme were absent [72].

In conclusion, CA (CAext+CAint) is essential for the reversible HCO3 C O 2 conversion both in the cell and in the periplasm. They participate in photosynthesis by supplying either CO2 to RuBisCO or HCO3 to PEPC for C4 type.


5. Carbonic anhydrase

CAs are metalloenzymes that catalyse the reversible interconversion of CO2 and HCO3 [73]. They are encoded by six evolutionary divergent gene families and the corresponding enzymes are designated as α, β, γ, δ, ε and ζ-CA [39]. These six types of CAs share no sequence similarity in their primary amino acid sequences and seem to have evolved independently [26, 74]. In macroalgae, almost all known CAs belong to α, β and γ classes, with the β class predominating [26, 39]. The δ, ε and ζ classes of CA are found only in some diatoms [75], bacteria [76] and marine protists [77, 78]. The active site of CA contains a zinc ion (Zn2+), which plays a critical role in the catalytic activity of the enzyme. The ζ and γ classes of CAs represent exceptions to this rule since they can use cadmium (ζ), iron (γ) or cobalt (γ) as cofactors [7981]. CA plays an important role in photosynthesis by supplying either CO2 to RuBPCO or HCO3 to PEPC. They also participate in some other physiological reactions such as respiration, pH homeostasis, ion transport and catalysis of key steps in the pathways for the biosynthesis of physiologically important metabolites [41]. The CA synthesis in the cytoplasm [82] is located in the periplasmic space, mitochondria, chloroplast stroma and chloroplast thylakoid lumen, carboxysome and pyrenoid [66, 70, 83, 84]. Different subcellular localizations make different CA functions in CCM. Periplasmic CA (CAext) can catalyse the conversion of HCO3 into CO2 to promote the diffusion of CO2 at the cell surface across the plasma membrane [85, 86]. Therefore, CAext has been postulated to be part of the CCM in most macroalgae. The cytoplasm CA stores Ci in the form of HCO3 to avoid leakage of CO2 and to regulate the pH value of cytoplasm by maintaining the equilibrium of different forms of Ci, which is important for algal CCM [39]. CAs on the chloroplast membrane and in the stroma mainly provide CO2 for RuBisCo [26, 38, 87]. In cyanobacteria, CAs in the carboxysomal shell function to convert accumulated HCO3 into CO2 and pass it to RuBisCo inside the cytoxysome [88]. CA in the thylakoid lumen was proposed to function to create an efficient CO2 supply to RuBisCo by taking advantage of the acidity of the lumenal compartment [69]. Stromal CA is also thought to operate by converting leaking CO2 into HCO3 [70]. Recently, data provided by various genome sequencing studies have revealed the multiplicity of CA isoforms in algae. For example, in the model microalga C. reinhardtii, there are at least 12 genes that encode CA isoforms, including three α, six β and three γ or γ-like CAs [39]. For marine diatom, nine and thirteen CA sequences were found in the genomes of P. tricornutum and Thalassiosira pseudonana, respectively [89]. P. tricornutum contains two β-CA genes, five α and two γ CA genes, whereas T. pseudonana has three α-, five γ-, four δ- and one ζ-CA genes [89]. As for macroalgae, CA genes have only been reported in few species. Six full-length CA of P. haitanensis (PhCA) genes were reported, which include two α-CAs, three β-CAs and one γ-CA [90]. Besides, one β-CA and one α-CA were reported in P. yezoensis [91] and S. japonica [92, 93]. Otherwise, although the activity of CAext and CAint has been detected in many macroalgae, the subcellular localization and functions of CAext and CAint remain unclear [71, 93].

Conclusively, CAs, including CAext and CAint (Figure 1), play an important role in the transportation or concentration process of the Ci. And as for C3 and C4 metablisms have different carboxylase, CAs might play different roles in CCMs of macroalgae with different photosynthetic mode. Thus, isolating of the CA genes, studies on their expression levels in different CO2 concentrations, in different life phase, and under different environmental stress, as well as studies on subcellular locations of CAs should be conducted in macroalgae to help reveal their Ci assimilation processes.


6. Studies of S. japonica CCM

S. japonica is an economically important brown seaweed. It has been cultivated extensively for food and industrial alginate in East Asia, such as in China, Japan and South Korea. China is by far the largest producer, and in 2009, its production in China rose sharply to 4.14×109 kg wet weight [94], accounting for approximately 80% of the global production, over several decades. This has been attributed to both its large-scale farming and high kelp yield per unit area. Production of this kelp in China under natural conditions is within the range of 3,300 to 11,300 g dry matter m−2·year−1, whereas that under artificial conditions is higher [1]. For example, its production during the 7-month cultivation is 15,000 g dry matter m−2 area (equivalent to 150 t per ha), which is 2.8 times higher than the maximum productivity of sugarcane in the United States (fresh weight about 95 t per ha·year) [1], which indicates that S. japonica has higher photosynthetic efficiency than sugarcane and other C4 plants. In fact, the photosynthetic efficiency of macroalgae (e.g., kelp) is 6%–8%, which is 1.8%–2.2% higher than that of land plants [95]. In seawater, the dominant species of Ci is HCO3 [11]. Since there is a fairly high photosynthetic rate in these kelps [34], a CCM involving an efficient HCO3 utilization mechanism is expected to exist. Indeed, 75% of the total Ci absorption in the juvenile sporophytes of this kelp is via the CAext mechanism [63], whereas CO2 diffusion accounts for 25% only. By analysis of genome annotation data of S. japonica [96], all the essential genes related to C3-pathway (23 unigenes) were discovered (Table 1), which provided the unequivocal molecular evidence that there existed C3-pathway in S. japonica. Otherwise, 16 enzyme-encoding unigenes involved in C4-pathway were found, covering almost all enzymes needed for C4-carbon fixation except the malic enzyme (Table 1). The results helped us to understand the carbon fixation process of this species.

Photosynthesis modes Enzyme names Unigenes
C3-pathway 23
Glyceraldehyde-3-phosphate dehydrogenase (phosphorylating) (GAPDH) 4
Transketolase 1
Phosphoribulokinase 2
Phosphoglycerate kinase (PGK) 5
Fructose-1,6-bisphosphatase (FBPase) 1
Sedoheptulose-bisphosphatase (SBPase) 3
Fructose-bisphosphate aldolase 1
Ribulose-phosphate 3-epimerase 2
Triose-phosphate isomerase (TIM) 1
Ribose-5-phosphate isomerase 1
Ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCo), small 1
Ribulose-1,5-bisphosphate carboxylase/oxygenase(RuBisCo), large 1
C4-pathway 16
Malate dehydrogenase 4
Aspartate aminotransferase (AST) 4
Pyruvate kinase 4
Phosphoenolpyruvate carboxylase (PEPC) 1
Phosphoenolpyruvate carboxykinase (PEPCK) 1
Pyruvate phosphate dikinase 1
Arginine/alanine aminopeptidase 1
Total 39

Table 1.

Statistics of C3/C4-pathway related enzymes of S. japonica.

Considering CAs play key roles in CCMs of macroalgae, it is important to determine the numbers and characterizations of CA genes of S. japonica. Herein, based on unigene sequences [96], the high-throughput sequencing data of S. japonica [97, 98] and S. latissima [99], as well as combined with the preparatory work of our group [92, 93], 12 CAs of S. japonica (SjCA) genes were obtained. Among them, we have cloned the full-length complementary DNA (cDNA) sequences of SjαCA1, SjβCA1 and SjβCA2 using rapid amplification of cDNA ends, which are 2804 [94], 1291 and 1261 nucleotides, respectively. The encoded proteins were 290, 314 and 307 amino acids. For further analysis the gene subtypes of CAs, a phylogenetic tree was constructed by using the neighbour-joining algorithm of the MEGA6.0 software [100] with Poisson correction and pairwise deletion parameters. A total of 1000 bootstrap replicates were performed. On the basis of conserved motifs and phylogenetic tree analysis (Figure 2), the SjCAs were divided into three CA classes: from SjαCA1 to SjαCA7 are α-CA; SjβCA1 and SjβCA2 are β-CA; SjγCA1, SjγCA2 and SjγCA3 are γ-CA. Among them, only one α-CA (SjαCA1) has been localized in the chloroplast and thylakoid membrane of the gametocytes of S. japonica under immunogold electron microscopy [93]. To get a general idea of functions of each SjCA, herein, the subcellular localizations of SjCAs were predicted using WoLFPSORT ( Based on the predicted results (Table 2), SjαCA2 might be an external CA and exist in periplasmic space, SjαCA3; SjαCA4, SjαCA6, SjαCA7 and

Figure 2.

Phylogenetic tree constructed using SjCA amino acid sequences.

SjγCA1 might be cytoplasmic CA; SjαCA5, SjβCA2 and SjγCA2 might present in mitochondria; SjβCA1 and SjγCA3 might exist in chloroplasts. However, most of the SjCAs’ subcellular localizations are predicted, which need to be verified by further studies. Otherwise, sporophyte and gametophyte of this kelp might employ different carbon fixation process since the content and activity of RuBisCo enzyme in gametophyte are significantly higher than those in sporophyte implying they may have different types of photosynthetic metabolism [24]. As for CA might play different role in CCMs of C3 and C4 pathway, full-length cDNA as well as DNA sequences of each SjCA should be cloned from sporophytes and gametophytes of this kelp in the future studies. CA gene expression levels under different CO2 concentrations and the subcellular location of each CA should also be conducted to help reveal Ci assimilation process of S japonica.

Enzyme Gene IDa AA no. Full length (Y/N) Subcellular location prediction
SjαCA1 JF827608 290 Y Chloroplast and thylakoid membrane [93]
SjαCA2 SJ07762 205 N Secreted
SjαCA3 SJ07765 160 N Cytoplasmic
SjαCA4 SJ13238 151 N Cytoplasmic
SjαCA5 SJ13240 294 N Mitochondrial inner membrane
SjαCA6 SJ18135 257 N Cytoplasmic
SjαCA7 SJ18141 189 N Cytoplasmic
SjβCA1 SJ12311 314 Y Chloroplast thylakoid membrane
SjβCA2 SJ17783 307 Y Mitochondrial
SjγCA1 SJ07587 305 N Cytoplasmic
SjγCA2 SJ22175 161 N Mitochondrial
SjγCA3 SJ21158 246 N Chloroplast

Table 2.

Prediction of subcellular locations of SjCAs.

Abbreviation: AA, amino acid.

a JF827608 is the NCBI gene accession number; ‘SJ’ in the table stands for the gene IDs for S. japonica.

The completion of the CCM modelling of sporophyte and gametophyte in S. japonica will give a solid foundation for further exploring its highly efficient photosynthetic mechanism. In addition, conducting studies on the inorganic carbon metabolism of macroalgae is of positive significance on developing the biomass energy from kelp and other algae and slowing down seawater acidification and global warming.


  1. 1. Gao K, McKinley KR. Use of macroalgae for marine biomass production and CO2 remediation: a review. Journal of Applied Phycology. 1994; 6(1): 45–60. DOI: 10.1007/BF02185904
  2. 2. Berry JA, Osmond CB, Lorimer G. Fixation of CO2 during photorespiration. Plant Physiology. 1978; 62: 954–967.
  3. 3. Mizohata E, Matsumura H, Okano Y. Crystal structure of activated ribulose-1,5-bisphosphate carboxylase/oxygenase from green alga Chlamydomonas reinhardtii complexed with 2-carboxyarabinitol-1,5-bisphosphate. Journal of Molecular Biology. 2002; 316: 679–691. DOI: 10.1006/jmbi.2001.5381
  4. 4. Jensen GR, Bahr TJ. Ribulose 1,5-bisphosphate carboxylase-oxygenase. Annual Review of Plant Physiology. 1977; 28: 379–400. DOI: 10.1146/
  5. 5. Reinfelder JR. Carbon concentrating mechanisms in eukaryotic marine phytoplankton. Annual Review of Marine Science. 2011; 3: 291–315. DOI: 10.1146/annurev-marine-120709-142720
  6. 6. Stumm W, Morgan JJ. Aquatic Chemistry: Chemical Equilibria and Rates in Natural Waters. 3rd ed. New York: Wiley; 1996. 1022 p.
  7. 7. Kerby NW, Raven JA. Transport and fixation of inorganic carbon by marine algae. Advances in Botanical Research. 1985; 2: 71–123.
  8. 8. Cook CM, Colman B. Some characteristics of photosynthetic inorganic carbon uptake of a marine macrophytic red alga. Plant, Cell and Environment. 1987; 10: 275–278. DOI: 10.1111/1365-3040.ep11602301
  9. 9. Maberly SC. Exogenous sources of inorganic carbon for photosynthesis by marine macroalgae. Journal of Phycology. 1990; 26(3): 439–449. DOI: 10.1111/j.0022-3646.1990.00439.x
  10. 10. Larsson C, Axelsson L. Bicarbonate uptake and utilization in marine macroalgae. European Journal of Phycology. 1999; 34: 79–86. DOI: 10.1080/09670269910001736112
  11. 11. Axelsson L, Mercado JM, Figueroa FL. Utilization of HCO3 at high pH by the brown macroalga Laminaria saccharina. European Journal of Phycology. 2000; 35(1): 53–59.
  12. 12. Gutknecht JM, Bisson A, Tosteson FC. Diffusion of carbon dioxide through lipid bilayer membranes. Effects of carbonic anhydrase, bicarbonate, and unstirred layers. Journal of General Physiology. 1977; 69: 779–794. DOI: 10.1085/jgp.69.6.779
  13. 13. Edwards GE, Huber SC. The C4 pathway. In: Hatch MD, Boardman NK, editors. The Biochemistry of Plants. A Comprehensive Treatise. Vol. 8, Photosynthesis. New York: Academic Press; 1981. p. 237–281.
  14. 14. Kremer BP. Aspects of carbon metabolism in marine macroalgae. Oceanography and Marine Biology: An Annual Review. 1981; 19: 41–94.
  15. 15. Johsi GV, Karekar MD, Gowda CA. Photosynthetic carbon metabolism and carboxylating enzymes in algae and mangrove under saline conditions. Photosynthetica. 1974; 8: 51–52.
  16. 16. Karekar MD, Joshi GV. Photosynthetic carbon metabolism in marine algae. Botanica Marina. 1973; 16(4): 216–220. DOI: 10.1515/botm.1973.16.4.216
  17. 17. Patil BA, Joshi GV. Photosynthetic studies in Ulva lactuca. Botanica Marina. 1970; 13(2): 111–115. DOI: 10.1515/botm.1970.13.2.111
  18. 18. Kremer BP, Küppers U. Carboxylating enzymes and pathway of photosynthetic carbon assimilation in different marine algae—Evidence for the C4-pathway? Planta. 1977; 133(2): 191–196. DOI: 10.1007/BF00391918
  19. 19. Reiskind JB, Seamon PT, Bowes G. Alternative methods of photosynthetic carbon assimilation in marine macroalgae. Plant Physiology. 1988; 87(3): 686–692. DOI: 10.1104/pp.87.3.686
  20. 20. Reiskind JB, Bowes G. The role of phosphoenolpyruvate carboxykinase in a marine macroalga with C4-like photosynthetic characteristics. Proceedings of the National Academy of Sciences of the United States of America. 1991; 88(7): 2883–2887. DOI: 10.1073/pnas.88.7.2883
  21. 21. Beer S, Israel A. Photosynthesis of Ulva sp III. O2 effects, carboxylase activities, and the CO2 incorporation pattern. Plant Physiology. 1986; 81(3): 937–938. DOI: 10.1104/pp.81.3.937
  22. 22. Niu J, Hu H, Hu S. Analysis of expressed sequence tags from the Ulva prolifera (Chlorophyta). Chinese Journal of Oceanology and Limnology. 2010; 28: 26–36. DOI: 10.1007/s00343-010-9120-4
  23. 23. Fan XL, Fang YJ, Hu SN. Generation and analysis of 5318 expressed sequence tags from the filamentous sporophyte of Porphyra haitanensis (Rhodophyta). Journal of Phycology. 2007; 43(6): 1287–1294. DOI: 10.1111/j.1529-8817.2007.00415.x
  24. 24. Wang C, Fan X, Wang G. Differential expression of RuBisCo in sporophytes and gametophytes of some marine macroalgae. PloS One. 2011; 6(1): e16351. DOI: 10.1371/journal.pone.0016351
  25. 25. Xu ZM, Yao NY, Li JZ. Studies on the activity of PEPck in L. japonica. Marine Science. 1991; 2: 41–45.
  26. 26. Moroney JV, Bartlett SG, Samuelsson G. Carbonic anhydrases in plants and algae. Plant, Cell & Environment. 2001; 24(2): 141–153. DOI: 10.1111/j.1365-3040.2001.00669.x
  27. 27. Krause GH. Photoinhibition of photosynthesis. An evaluation of damaging and protective mechanisms. Physiologia Plantarum. 1988; 74(3): 566–574. DOI: 10.1111/j.1399-3054.1988.tb02020.x
  28. 28. Hanelt D, Huppertz K, Nultsch W. Photoinhibition of photosynthesis and its recovery in red algae. Botanica Acta. 1992; 105(4): 278–284. DOI: 10.1111/j.1438-8677.1992.tb00299.x
  29. 29. Hanelt D, Huppertz K, Nultsch W. Daily course of photosynthesis and photoinhibition in marine macroalgae investigated in the laboratory and field. Marine Ecology Progress Series. 1993; 97(1): 31–37. DOI: 10.3354/meps097031
  30. 30. Roleda MY, Hurd CL. Seaweed responses to ocean acidification//Seaweed Biology. Springer Berlin Heidelberg. 2012; 219: 407–431. DOI: 10.1007/978-3-642-28451-9_19
  31. 31. Moroney J V, Somanchi A. How do algae concentrate CO2 to increase the efficiency of photosynthetic carbon fixation? Plant Physiology. 1999, 119(1): 9–16. DOI: 10.1104/pp.119.1.9
  32. 32. Cook CM, Lanaras T, Colman B. Evidence for bicarbonate transport in species of red and brown macrophytic marine algae. Journal of Experimental Botany. 1986; 37(7): 977–984. DOI: 10.1093/jxb/37.7.977
  33. 33. Surif MB, Raven JA. Exogenous inorganic carbon sources for photosynthesis in seawater by members of the Fucales and the Laminariales (Phaeophyta): ecological and taxonomic implications. Oecologia. 1989; 78(1): 97–105. DOI: 10.1007/BF00377203
  34. 34. Axelsson L, Uusitalo J, Ryberg H. Mechanisms for concentrating and storage of inorganic carbon in marine macroalgae. Seaweed Cellular Biotechnology, Physiology and Intensive Cultivation. COST-48. Universidad de las Palmas de Gran Canaria, España. 1991; 185–198.
  35. 35. Johnston AM, Maberly SC, Raven JA. The acquisition of inorganic carbon by four red macroalgae. Oecologia. 92(3): 317–326. DOI: 10.1007/BF00317457
  36. 36. Mercado JM, Gordillo FJL, Figueroa FL. External carbonic anhydrase and affinity for inorganic carbon in intertidal macroalgae. Journal of Experimental Marine Biology and Ecology. 1998; 221(2): 209–220. DOI: 10.1016/S0022-0981(97)00127-5
  37. 37. Badger MR, Hanson D, Price GD. Evolution and diversity of CO2 concentrating mechanisms in cyanobacteria. Functional Plant Biology. 2002; 29(3): 161–173. DOI: 10.1071/PP01213
  38. 38. Spalding MH. Microalgal carbon-dioxide-concentrating mechanisms: Chlamydomonas inorganic carbon transporters. Journal of Experimental Botany. 2008; 59(7): 1463–1473. DOI: 10.1093/jxb/erm128
  39. 39. Moroney JV, Ma YB, Frey WD. The carbonic anhydrase isoforms of Chlamydomonas reinhardtii: intracellular location, expression, and physiological roles. Photosynthesis Research. 2011; 109(1–3): 133–149. DOI: 10.1007/s11120-011-9635-3
  40. 40. Price GD. Inorganic carbon transporters of the cyanobacterial CO2 concentrating mechanism. Photosynthesis Research. 2011; 109(1–3): 47–57. DOI: 10.1007/s11120-010-9608-y
  41. 41. Badger M. The roles of carbonic anhydrases in photosynthetic CO2 concentrating mechanisms. Photosynthesis Research. 2003; 77(2–3): 83–94. DOI: 10.1023/A:1025821717773
  42. 42. Israel A, Hophy M. Growth, photosynthetic properties and RuBisCo activities and amounts of marine macroalgae grown under current and elevated seawater CO2 concentrations. Global Change Biology. 2002; 8(9): 831–840. DOI: 10.1046/j.1365-2486.2002.00518.x
  43. 43. Fernández PA, Roleda MY, Hurd CL. Effects of ocean acidification on the photosynthetic performance, carbonic anhydrase activity and growth of the giant kelp Macrocystis pyrifera. Photosynthesis Research. 2015; 124: 293–304. DOI: 10.1007/s11120-015-0138-5
  44. 44. Axelsson L, Ryberg H, Beer S. Two modes of bicarbonate utilization in the marine green macroalga Ulva lactuca. Plant, Cell & Environment. 1995; 18(4): 439–445. DOI: 10.1111/j.1365-3040.1995.tb00378.x
  45. 45. Fernández PA, Hurd CL, Roleda MY. Bicarbonate uptake via an anion exchange protein is the main mechanism of inorganic carbon acquisition by the giant kelp Macrocystis pyrifera (Laminariales, Phaeophyceae) under variable pH. Journal of Phycology. 2014; 50(6): 998–1008. DOI: 10.1111/jpy.12247
  46. 46. Drechsler Z, Sharkia R, Cabantchik ZI. Bicarbonate uptake in the marine macroalga Ulva sp. is inhibited by classica probes of anion exchange by red blood cells. Planta. 1993; 191: 34–40. DOI: 10.1007/BF00240893
  47. 47. Axelsson L, Larsson C, Ryberg H. Affinity, capacity and oxygen sensitivity of two different mechanisms for bicarbonate utilization in Ulva lactuca L.(Chlorophyta). Plant, Cell & Environment. 1999; 22(8): 969–978. DOI: 10.1046/j.1365-3040.1999.00470.x
  48. 48. Giordano M, Beardall J, Raven JA. CO2 concentrating mechanisms in algae: mechanisms, environmental modulation, and evolution. Annual Review of Plant Biology. 2005; 56: 99–131. DOI: 10.1146/annurev.arplant.56.032604.144052
  49. 49. Klenell M, Snoeijs P, Pedersén M. Active carbon uptake in Laminaria digitata and L. saccharina (Phaeophyta) is driven by a proton pump in the plasma membrane. Hydrobiologia. 2004; 514(1–3): 41–53. DOI: 10.1023/B:hydr.0000018205.80186.3e
  50. 50. Smith RG, Bidwell RGS. Mechanism of photosynthetic carbon dioxide uptake by the red macroalga, Chondrus crispus. Plant Physiology. 1989; 89(1): 93–99. DOI: 10.1104/pp.89.1.93
  51. 51. Choo K, Snoeijs P, Pedersén M. Uptake of inorganic carbon by Cladophora glomerata (Chlorophyta) from the Baltic Sea. Journal of Phycology. 2002; 38(3): 493–502. DOI: 10.1046/j.1529-8817.2002.01083.x
  52. 52. Beer S. Mechanisms of inorganic carbon acquisition in marine microalgae (with special reference to the Chlorophyta). Progress in Phycological Research. 1994; 10: 179–207.
  53. 53. Beer S. Photosynthetic utilization of inorganic carbon in Ulva. Scientia Marina. 1996; 60(Supl. 1): 125–128.
  54. 54. Larsson C, Axelsson L, Ryberg H. Photosynthetic carbon utilization by Enteromorpha intestinalis (Chlorophyta) from a Swedish rockpool. European Journal of Phycology. 1997; 32(1): 49–54. DOI: 10.1080/09541449710001719365
  55. 55. Snoeijs P, Klenell M, Choo K. Strategies for carbon acquisition in the red marine macroalga Coccotylus truncatus from the Baltic Sea. Marine Biology. 2002; 140(3): 435–444. DOI: 10.1007/s00227-001-0729-x
  56. 56. Murru M, Sandgren CD. Habitat matters for inorganic carbon acquisition in 38 species of red macroalgae (Rhodophyta) from Puget Sound, Washington, USA. Journal of Phycology. 2004; 40(5): 837–845. DOI: 10.1111/j.1529-8817.2004.03182.x
  57. 57. Cornwall CE, Revill AT, Hurd CL. High prevalence of diffusive uptake of CO2 by macroalgae in a temperate subtidal ecosystem. Photosynthesis Research. 2015; 124(2): 181–190. DOI: 10.1007/s11120-015-0114-0
  58. 58. Andría JR, Pérez-Lloréns JL, Vergara JJ. Mechanisms of inorganic carbon acquisition in Gracilaria gaditana nom. prov.(Rhodophyta). Planta. 1999; 208(4): 564–573. DOI: 10.1007/s004250050594
  59. 59. Gómez-Pinchetti JL, Ramazanov Z, García-Reina G. Effect of inhibitors of carbonic anhydrase activity on photosynthesis in the red alga Soliera filiformis (Gigartinales: Rhodophyta). Marine Biology. 1992; 114(2): 335–339. DOI: 10.1007/BF00349536
  60. 60. Gravot A, Dittami SM, Rousvoal S. Diurnal oscillations of metabolite abundances and gene analysis provide new insights into central metabolic processes of the brown alga Ectocarpus siliculosus. New Phytologist. 2010; 188(1): 98–110. DOI: 10.1111/j.1469-8137.2010.03400.x
  61. 61. Zou D, Gao K, Chen W. Photosynthetic carbon acquisition in Sargassum henslowianum (Fucales, Phaeophyta), with special reference to the comparison between the vegetative and reproductive tissues. Photosynthesis Research. 2011; 107(2): 159–168. DOI: 10.1007/s11120-010-9612-2
  62. 62. Zou DH, Gao KS, Xia JR. Photosynthetic utilization of inorganic carbon in the economic brown alga, Hizikia fusiforme (Sargassaceae) from the South China Sea. Journal of Phycology. 2003; 36: 1095–1100. DOI: 10.1111/j.0022-3646.2003.03-038.x
  63. 63. Yue GF, Wang JX, Wang JF. Inorganic carbon acquisition by juvenile sporophyte of Laminarials (L. japonica ×L. longissima). Oceanologia et Limnologia Sinica. 2001; 32(6): 647–652.
  64. 64. Nakajima K, Tanaka A, Matsuda Y. SLC4 family transporters in a marine diatom directly pump bicarbonate from seawater. Proceedings of the National Academy of Sciences of the United States of America. 2013; 110(5): 1767–1772. DOI: 10.1073/pnas.1216234110
  65. 65. Hopkinson BM. A chloroplast pump model for the CO2 concentrating mechanism in the diatom Phaeodactylum tricornutum. Photosynthesis Research. 2014; 121(2–3): 223–233. DOI: 10.1007/s11120-013-9954-7
  66. 66. Karlsson J, Clarke AK, Chen ZY. A novel α-type carbonic anhydrase associated with the thylakoid membrane in Chlamydomonas reinhardtii is required for growth at ambient CO2. EMBO Journal. 1998; 17(5): 1208–1216. DOI: 10.1093/emboj/17.5.1208
  67. 67. Karlsson J, Hiltonen T, Husic HD. Intracellular carbonic anhydrase of Chlamydomonas reinhardtii. Plant Physiology. 1995; 109(2): 533–539. DOI: 10.1104/pp.109.2.533
  68. 68. Hanson DT, Franklin LA, Samuelsson G. The Chlamydomonas reinhardtii cia3 mutant lacking a thylakoid lumen-localized carbonic anhydrase is limited by CO2 supply to RuBisCo and not photosystem II function in vivo. Plant Physiology. 2003; 132(4): 2267–2275. DOI: 10.1104/pp.103.023481
  69. 69. Raven JA. CO2-concentrating mechanisms: a direct role for thylakoid lumen acidification? Plant, Cell & Environment. 1997; 20(2): 147–154. DOI: 10.1046/j.1365-3040.1997.d01-67.x
  70. 70. Mitra M, Lato SM, Ynalvez RA. Identification of a new chloroplast carbonic anhydrase in Chlamydomonas reinhardtii. Plant Physiology. 2004; 135(1): 173–182. DOI: 10.1104/pp.103.037283
  71. 71. Sültemeyer D. Carbonic anhydrase in eukaryotic algae: characterization, regulation, and possible function during photosynthesis. Canadian Journal of Botany. 1998; 76(6): 962–972. DOI: 10.1139/b98-082
  72. 72. Badger MR, Price GD. The role of carbonic anhydrase in photosynthesis. Annual Review of Plant Biology. 1994; 45(1): 369–392. DOI: 10.1146/annurev.pp.45.060194.002101
  73. 73. Badger MR, Price GD. The CO2 concentrating mechanism in cyanobacteria and microalgae. Physiologia Plantarum. 1992; 84(4): 606–615. DOI: 10.1034/j.1399-3054.1992.840416.x
  74. 74. Hewett-Emmett D, Tashian RE. Functional diversity, conservation, and convergence in the evolution of the α-, β-, and γ-carbonic anhydrase gene families. Molecular Phylogenetics and Evolution. 1996; 5(1): 50–77. DOI: 10.1006/mpev.1996.0006
  75. 75. Roberts SB, Lane TW, Morel FMM. Carbonic anhydrase in the marine diatom Thalassiosira weissflogii (Bacillariophyceae). Journal of Phycology. 1997; 33: 845–850. DOI: 10.1111/j.0022-3646.1997.00845.x
  76. 76. So AK, Espie GS, Williams EB.A novel evolutionary lineage of carbonic anhydrase (epsilon class) is a component of the carboxysome shell. Journal of Bacteriology. 2004; 186: 623–630. DOI: 10.1128/JB.186.3.623-630.2004
  77. 77. Lane TW, Morel FMM. Regulation of carbonic anhydrase expression by zinc, cobalt, and carbon dioxide in the marine diatom Thalassiosira weissflogii. Plant Physiology. 2000; 123: 345–352. DOI: 10.1104/pp.123.1.345
  78. 78. Park H, Song B, Morel FMM. Diversity of the cadmium-containing carbonic anhydrase in marine diatoms and natural waters. Environmental Microbiology. 2007; 9(2): 403–413. DOI: 10.1111/j.1462-2920.2006.01151.x
  79. 79. Lane TW, Saito MA, George GN. Biochemistry: a cadmium enzyme from a marine diatom. Nature. 2005; 435(7038): 42. DOI: 10.1038/435042a
  80. 80. Xu Y, Feng L, Jeffrey PD. Structure and metal exchange in the cadmium carbonic anhydrase of marine diatoms. Nature. 2008; 452(7183): 56–61. DOI: 10.1038/nature06636
  81. 81. Ferry JG. The γ class of carbonic anhydrases. Biochimica et Biophysica Acta. 2010; 1804(2): 374–381. DOI: 10.1016/j.bbapap.2009.08.026
  82. 82. Coleman JR, Grossman AR. Biosynthesis of carbonic anhydrase in Chlamydomonas reinhardtii during adaptation to low CO2. Proceedings of the National Academy of Sciences of the United States of America. 1984; 81(19): 6049–6053. DOI: 10.1073/pnas.81.19.6049
  83. 83. Fujiwara S, Fukuzawa H, Tachiki A. Structure and differential expression of two genes encoding carbonic anhydrase in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America. 1990; 87(24): 9779–9783. DOI: 10.1073/pnas.87.24.9779
  84. 84. Eriksson M, Karlsson J, Ramazanov Z. Discovery of an algal mitochondrial carbonic anhydrase: molecular cloning and characterization of a low-CO2-induced polypeptide in Chlamydomonas reinhardtii. Proceedings of the National Academy of Sciences of the United States of America. 1996; 93(21): 12031–12034. DOI: 10.1073/pnas.93.21.12031
  85. 85. Moroney JV, Husic HD, Tolbert NE. Effect of carbonic anhydrase inhibitors on inorganic carbon accumulation by Chlamydomonas reinhardtii. Plant Physiology. 1985; 79(1): 177–183. DOI: 10.1104/pp.79.1.177
  86. 86. Van K, Spalding MH. Periplasmic carbonic anhydrase structural gene (Cah1) mutant in Chlamydomonas reinhardtii. Plant Physiology. 1999; 120(3): 757–764. DOI: 10.1104/pp.120.3.757
  87. 87. Lucas WJ. Photosynthetic assimilation of exogenous HCO3 by aquatic plants. Annual Review of Plant Physiology. 1983; 34(1): 71–104. DOI: 10.1146/annurev.pp.34.060183.000443
  88. 88. Peña KL, Castel SE, de Araujo C. Structural basis of the oxidative activation of the carboxysomal γ-carbonic anhydrase, CcmM. Proceedings of the National Academy of Sciences of the United States of America. 2010; 107(6): 2455–2460. DOI: 10.1073/pnas.0910866107
  89. 89. Tachibana M, Allen AE, Kikutani S. Localization of putative carbonic anhydrases in two marine diatoms, Phaeodactylum tricornutum and Thalassiosira pseudonana. Photosynthesis Research. 2011; 109(1–3): 205–221. DOI: 10.1007/s11120-011-9634-4
  90. 90. Chen C, Dai Z, Xu Y. Cloning, expression, and characterization of carbonic anhydrase genes from Pyropia haitanensis (Bangiales, Rhodophyta). Journal of Applied Phycology. DOI: 10.1007/s10811-015-0646-x
  91. 91. Zhang BY, Yang F, Wang GC. Cloning and quantitative analysis of the carbonic anhydrase gene from Porphyra yezoensis. Journal of Phycology. 2010; 46: 290–296. DOI: 10.1111/j.1529-8817.2009.00801.x
  92. 92. Yu Z, Bi YH, Zhou ZG. Cloning and characterization of carbonic anhydrase (CA) gene from Laminaria japonica gametophytes. Journal of Fisheries of China. 2011; 35: 1343–1353.
  93. 93. Ye RX, Yu Z, Shi WW. Characterization of α-type carbonic anhydrase (CA) gene and subcellular localization of α-CA in the gametophytes of Saccharina japonica. Journal of Applied Phycology. 2014; 26: 881–890. DOI: 10.1007/s10811-013-0221-2
  94. 94. The FAO yearbook of Fishery and Aquaculture Statistics: Aquaculture Production. Rome: Food and Agriculture Organization of the United Nations; 2009. 221 p.
  95. 95. Ross AB, Jones JM, Kubacki ML. Classification of macroalgae as fuel and its thermochemical behaviour. Bioresource Technology. 2008; 99(14): 6494–6504. DOI: 10.1016/j.biortech.2007.11.036
  96. 96. Ye N, Zhang X, Miao M. Saccharina genomes provide novel insight into kelp biology. Nature Communications. 2015; 6: 6986. DOI: 10.1038/ncomms7986
  97. 97. Deng YY, Yao JT, Wang XL. Transcriptome sequencing and comparative analysis of Saccharina japonica (Laminariales, Phaeophyceae) under blue light induction. PLoS One. 2012; 7(6): e39704. DOI: 10.1371/journal.pone.0039704
  98. 98. Wang WJ, Wang FJ, Sun XT. Comparison of transcriptome under red and blue light culture of Saccharina japonica (Phaeophyceae). Planta. 2013; 237(4): 1123–1133. DOI: 10.1007/s00425-012-1831-7
  99. 99. Heinrich S, Valentin K, Frickenhaus S. Transcriptomic analysis of acclimation to temperature and light stress in Saccharina latissima (Phaeophyceae). PloS One. 2012; 7(8): e44342. DOI: 10.1371/journal.pone.0044342
  100. 100. Tamura K, Stecher G, Peterson D. MEGA6: molecular evolutionary genetics analysis Version 6.0. Molecular Biology and Evolution. 2013; 30: 2725–2729. DOI: 10.1093/molbev/mst197

Written By

Yanhui Bi and Zhigang Zhou

Submitted: 07 September 2015 Reviewed: 26 January 2016 Published: 30 March 2016