Skip to main content
PMC home page
Plant Physiology logoLink to Plant Physiology
. 1999 May;120(1):105–112. doi: 10.1104/pp.120.1.105

Extracellular Carbonic Anhydrase Facilitates Carbon Dioxide Availability for Photosynthesis in the Marine Dinoflagellate Prorocentrum micans

Nabil A Nimer 1,*, Colin Brownlee 2, Michael J Merrett 1
PMCID: PMC59241  PMID: 10318688

Abstract

This study investigated inorganic carbon accumulation in relation to photosynthesis in the marine dinoflagellate Prorocentrum micans. Measurement of the internal inorganic carbon pool showed a 10-fold accumulation in relation to external dissolved inorganic carbon (DIC). Dextran-bound sulfonamide (DBS), which inhibited extracellular carbonic anhydrase, caused more than 95% inhibition of DIC accumulation and photosynthesis. We used real-time imaging of living cells with confocal laser scanning microscopy and a fluorescent pH indicator dye to measure transient pH changes in relation to inorganic carbon availability. When steady-state photosynthesizing cells were DIC limited, the chloroplast pH decreased from 8.3 to 6.9 and cytosolic pH decreased from 7.7 to 7.1. Re-addition of HCO3− led to a rapid re-establishment of the steady-state pH values abolished by DBS. The addition of DBS to photosynthesizing cells under steady-state conditions resulted in a transient increase in intracellular pH, with photosynthesis maintained for 6 s, the amount of time needed for depletion of the intracellular inorganic carbon pool. These results demonstrate the key role of extracellular carbonic anhydrase in facilitating the availability of CO2 at the exofacial surface of the plasma membrane necessary to maintain the photosynthetic rate. The need for a CO2-concentrating mechanism at ambient CO2 concentrations may reflect the difference in the specificity factor of ribulose-1,5 bisphosphate carboxylase/oxygenase in dinoflagellates compared with other algal phyla.

Marine phytoplankton species are the major dominant fixers of inorganic carbon in the oceans (Raven, 1994; 1997a), removing about 35 Pg of inorganic carbon per year from the ecosphere (Raven, 1997a). The key enzyme of photosynthetic CO2 fixation, Rubisco (EC 4.1.1.39) (Raven, 1995), catalyzes the initial assimilation reaction of CO2 and also the oxygenation of ribulose bisphosphate, initiating the photorespiratory pathway (Lorimer et al., 1973). The ratio of these reactions determines the specificity factor (τ) of the enzyme, which can indicate enzyme type. The diverse universal type I enzyme found in oxygenic phototrophs has a substantially higher specificity for CO2 than for oxygen (Jordan and Ogren, 1981) compared with type II, the more oxygen-sensitive, homomeric form of the enzyme found in heterotrophic anaerobic proteobacteria and cyanobacteria (Delgado et al., 1995; Tabita, 1995; Raven, 1997a). Recently, among the dinoflagellates, a major component of marine phytoplankton, the species tested were found to possess the oxygen-sensitive homomeric type-II form of the enzyme (Morse et al., 1995; Whitney and Yellowlees, 1995; Rowan et al., 1996; Whitney and Andrews, 1998).

Dinoflagellates are morphologically and physiologically diverse, abundant in the marine ecosystem, and ecologically important; they make a major contribution to the global biological carbon pump (Raven and Johnston, 1991). Relatively little, however, is known about their mechanism of inorganic carbon acquisition. In the dinoflagellate species investigated, the specificity factor was approximately 2-fold greater than other homomeric Rubiscos but was still very unlikely to support photosynthetic rates at ambient CO2 levels (Raven and Johnston, 1991; Whitney and Andrews, 1998). This explains the need for a CCM to elevate the CO2 concentration around the active center of Rubisco, suppressing glycolate formation and enhancing carboxylation (Coleman, 1991; Badger and Price, 1994). An essential component of a CCM is an active influx of inorganic carbon across a membrane (Raven, 1995).

In some algal species, extracellular CA (EC 4.2.1.1) is a major component of the CCM (Badger et al., 1980; Spalding et al., 1983; Aizawa and Miyachi, 1986). Extracellular CA was shown to be important in inorganic carbon transport when HCO3− is available but CO2 is limited external to the plasma membrane (Tsuzuki and Miyachi, 1989). Under these conditions the catalytic dehydration of HCO3 by extracellular CA provides CO2 external to the plasma membrane. Overall, the mechanism of inorganic carbon acquisition in marine phytoplankton is species dependent; some species rapidly acclimate to changes in CO2 and/or HCO3− concentrations (Coleman, 1991; Nimer and Merrett, 1996; Nimer et al., 1997). In the dinoflagellate Prorocentrum micans, the activity of constitutive extracellular CA (Nimer et al., 1997) is modulated by environmental conditions, increasing under conditions of inorganic carbon limitation (Nimer et al., 1997).

In the present study we investigated the potential role of extracellular CA in facilitating CO2 entry and sustaining the photosynthetic rate in P. micans by monitoring transient changes in intracellular pH in relation to internal inorganic carbon concentration and photosynthetic CO2 fixation.

MATERIALS AND METHODS

Growth of Cells

Axenic cultures of the marine dinoflagellate Prorocentrum micans (strain CCAP 1136/8 from the Provasoli-Guillard Centre for Culture of Marine Phytoplanktonoban, Oban, UK) were grown on f/2 medium (Guillard and Ryther, 1962) modified as described previously (Nimer et al., 1997). Cultures were grown at 15° ± 1°C with a PFD of 100 μmol m−2 s−1 at the culture surface provided by cool-white fluorescent tubes. Cells were grown in inorganic carbon-replete conditions (2.0 mm total DIC, pH 8.3) and, when required, cells were resuspended in 1.0 mm total DIC at pH 8.3 or 2.0 mm total DIC at pH 9.0 (i.e. inorganic carbon or CO2-limited conditions).

Measurement of Alkalinity, Total DIC, and the Calculation of Free CO2

We followed procedures described previously (Parsons et al., 1989; Merrett et al., 1996) to measure alkalinity and we measured pH with a digital pH meter (Hanna Instruments, Woonsockett, RI). Total DIC was calculated from carbonate alkalinity and pH (McConnaughey, 1991), and the free CO2 concentration was calculated as described previously (Dong et al., 1993).

Measurement of Extracellular CA Activity

Intact cells were harvested by centrifugation at 1500g, washed once, and resuspended in 25 mm barbitol buffer. Intact cells were assayed for extracellular CA by an electrometric method described previously (Dixon et al., 1987; Nimer et al., 1997).

Inorganic Carbon-Dependent Photosynthetic Oxygen Evolution

Inorganic carbon-dependent, photosynthetic oxygen evolution was measured using a Clark-type oxygen electrode (Hansatech, King's Lynn, UK) as described previously (Nimer and Merrett, 1992, 1996). Intact cells were resuspended in 300 mm sorbitol and 25 mm Hepes (pKa 7.5) to pH 8.3, and Hepes was replaced by 25 mm citric acid/trisodium citrate buffer for pH 5.0. Cells were allowed to deplete all endogenous carbon sources (measured by the cessation of oxygen evolution). We measured the rate of oxygen evolution after the addition of various concentrations of KHCO3.

Inorganic 14C Uptake and Photosynthesis

Intact cells were harvested, washed twice, and resuspended in 300 mm sorbitol and 25 mm Hepes at pH 8.3. We placed the cells in an oxygen electrode chamber and allowed them to deplete all endogenous carbon sources. We measured inorganic carbon uptake and photosynthesis after the addition of the required Na14CO3 (Amersham) at a specific activity of 5.6 × 108 Bq/mol, as described previously (Badger et al., 1980; Nimer et al., 1992; Nimer and Merrett, 1996). We used 5 μm [14C]inulin and 3H2O to estimate the intracellular water space and the free water space taken down with the cells through the silicone oil. Separate incubation times for each were 20 s (time needed for equilibration). We determined the 14C and the 3H in the pellet by liquid scintillation counting (Beckman) and the intracellular space (inulin impermeable) as total water minus the inulin-permeable space volume.

Measurement of Intracellular pH

Cells were loaded for 30 min with the fluorescence indicator SNARF (10 μm carboxy-SNARF-1-acetomethylester, Molecular Probes, Eugene, OR). The fluorescence properties of this dye are such that the ratio of fluorescence emission at 630 and 590 nm is pH dependent (Seksek et al., 1991). Cells were settled on a coverslip pretreated with poly-l-Lys (Anning et al., 1996). Single cells were observed with a confocal laser-scanning microscope (model MRC 1024, Bio-Rad). Fluorescence emission was recorded following excitation at 488 nm. We obtained ratio images (630/590) of dye-loaded cells using time-course image-analysis software (Bio-Rad). The fluorescence intensities of the cytosol and the chloroplast from the ratio images were measured and compared with fluorescence ratio images of an “in vitro” calibration, which we obtained from buffered media containing 10 μm SNARF-free acid at the required pH. A good agreement between “in vivo” and “in vitro” calibrations was established previously (Anning et al., 1996). We located the position of the chloroplast by monitoring its autofluorescence at 515 nm emission. A 50-W ellipsoid halogen-reflector bulb (Philips, Eindhoven, The Netherlands) provided light during the imaging of the cells.

We used an improved Neubauer hemocytometer (Weber Scientific International, Cambridge, UK) to determine cell number. A stock solution of DBS (Synthelic, Lund, Sweden) was prepared in double-distilled water and used at a final concentration of 200 μm. DCMU (Sigma) was dissolved in 96% ethanol to give a stock solution of 200 mm and used at a final concentration of 50 μm.

RESULTS

Photosynthetic Rate and Intracellular DIC Accumulation

The rate of inorganic carbon-dependent photosynthetic oxygen evolution was measured at different DIC concentrations, and the affinity of the cells for DIC was determined by calculating the concentration of DIC required to give the half-maximal rate of photosynthetic oxygen evolution, K0.5[DIC]. The maximum rate of photosynthetic oxygen evolution was unaffected by external pH (Fig. 1, A and B). At pH 8.3 the K0.5[DIC] was 750 μm, whereas at pH 5.0 it was 25 μm. These results are consistent with the belief that CO2 is the species of inorganic carbon that crosses the plasma membrane. We used DBS, a membrane-impermeable inhibitor of CA, to investigate the role of extracellular CA in DIC utilization. At pH 8.3, DIC-dependent photosynthetic oxygen evolution was 90% inhibited by DBS, even at a high external DIC concentration (i.e. 3.0 mm). At pH 5.0 photosynthetic oxygen evolution was unaffected by the presence of DBS.

Figure 1.

Figure 1

Open in a new tab

A and B, Rates of photosynthetic oxygen evolution in response to external DIC concentration at pH 8.3 (A) and pH 5.0 (B) in the absence (•) and presence (○) of DBS. C to F, Internal inorganic carbon concentration by P. micans in the absence (•) and presence (○) of DBS (C and E) and in the absence (•) and presence (○) of DCMU (D and F) and photosynthetic 14CO2 fixation in the absence (▪) and presence (□) of DBS (C and E) and in the absence (▪) and presence (□) of DCMU (D and F). The final cell concentration was 2.5 × 106 cells mL−1. In C and D, the intracellular water space was 0.87 μL and the total water space was 1.75 μL; in E and F, the intracellular water space was 0.53 μL and the total water space was 1.08 μL (n = 3). In A and B, the temperature was 15°C ± 1°C, and the PFD was 500 μmol m−2 s−1; in C and D, the pH was 8.3, the temperature was 15°C ± 1°C, and the PFD was 500 μmol m−2 s−1; in E and F, the temperature was 15°C ± 1°C and the PFD was 500 μmol m−2 s−1.  

These results suggest a major role for extracellular CA at an alkaline pH when the available free CO2 concentration is lower, simulating the conditions in the marine environment. This role of extracellular CA was confirmed by measuring the extracellular CA and the photosynthetic rate at pH 8.3 (2.0 mm total DIC and 9.0 μm free CO2) and pH 9.0 (2.0 mm total DIC and 1.0 μm free CO2) (Table I). Cells of P. micans maintained an almost constant photosynthetic rate at pH 8.3 and 9.0 (Table I), although the free CO2 concentration was much lower at pH 9.0 (Table I). At pH 8.3 and 9.0, DBS effectively inhibited extracellular CA activity and DIC-dependent photosynthetic oxygen evolution (Table I). At pH 9.0 extracellular CA activity was more than 60% greater than at pH 8.3 (Table I).

Table I.

Photosynthetic oxygen evolution in relation to extracellular CA activity in P. micans grown at 2.0 mm DIC, pH 8.3, and resuspended under the conditions indicated at a PFD of 500 μmol m−2 s−1 at 15°C

Condition Photosynthetic Oxygen Evolution External CA
nmol 106 cells−1 h−1 EU (106 cells−1)
pH 8.3
 2.0 mm total DIC 20.47  ± 0.92 0.25  ± 0.01
 2.0 mm total DIC plus DBS 2.31  ± 0.20 0.03  ± 0.00
pH 9.0
 2.0 mm total DIC 19.78  ± 0.53 0.41  ± 0.03
 2.0 mm total DIC plus DBS 2.11  ± 0.13 0.05  ± 0.00
Open in a new tab

Values are ±se; n = 4.

Effect of DBS on Photosynthetic Rate and Inorganic Carbon Uptake

Measurement of DIC uptake at pH 8.3 by the silicone oil centrifugation technique showed that a rapid steady state was achieved between the internal inorganic carbon pool of the cells and DIC in the external medium (Fig. 1, C and D). Over the same period, photosynthetically fixed 14CO2 showed a linear increase with time (Fig. 1, C and D). The inhibitors DBS (Moroney et al., 1985; Nimer and Merrett, 1996; Nimer et al., 1998) and DCMU were equally effective in blocking the transport of inorganic carbon into the cell and the photosynthetic fixation of 14CO2 (Fig. 1, C and D).

The rate of photosynthetic 14CO2 fixation increased over a range of external DIC concentrations until nearby inorganic carbon saturation of photosynthesis was observed at 1.0 mm external DIC (Fig. 1, E and F). DBS and DCMU were equally effective in inhibiting the photosynthetic rate and the accumulation of inorganic carbon within the cell and over a range of external DIC concentrations (Fig. 1, E and F). At all of the external DIC concentrations used, the average intracellular inorganic carbon concentration was 10-fold greater than the DIC concentration outside the cells (Fig. 1, E and F).

Intracellular pH

Confocal images of SNARF-loaded P. micans cells show the distribution of the pH-sensitive dye in subcellular compartments (Fig. 2). The position of the chloroplast was determined by autofluoresence (Fig. 2C). The chloroplast autofluoresence and the distribution of SNARF fluorescence (Fig. 2) were comparable to the chloroplast multilobular arrangement and position observed in transmission electron microscope images of P. micans (Vesk and Jeffrey, 1977; Dodge and Greuet, 1987). Both the chloroplast and the cytosol were successfully loaded with SNARF, but the dye was excluded from the vacuoles. The ratio image (Fig. 2D) shows the ability to determine pH in the chloroplast and the cytosol. In the steady-state cells under inorganic carbon-replete conditions (2.0 mm total DIC and external pH 8.3), the average chloroplast and cytosol pH were 8.3 ± 0.13 and 7.7 ± 0.11, respectively (Fig. 2D).

Figure 2.

Figure 2

Open in a new tab

Confocal microscope image of SNARF-loaded P. micans cell displaying the distribution of the dye in subcellular compartments. A, Emission at 630 nm; B, emission at 590 nm; C, chloroplast autofluoresence at 515 nm; and D, 630/590 florescence ratio. Bar = 10 μm.

Inorganic Carbon Concentration and Intracellular Homeostasis

When cells photosynthesizing under steady state were DIC limited (1.0 mm total DIC, pH 8.3), the average cytosolic and chloroplast pH decreased to 6.9 ± 0.10 and 7.14 ± 0.12, respectively (Fig. 3A). The re-addition of HCO3− (2.0 mm DIC final concentration) to these cells led to the rapid re-establishment of the steady-state pH values in the cytosol and chloroplast (Fig. 3, B–D). This response was abolished in the presence of DBS (Fig. 3, E–H). Pre-incubating the cells with DCMU significantly lowered the pH of the cytosol and chloroplast (Fig. 3I). In the presence of DCMU, the addition of HCO3− to inorganic carbon-limited cells did not result in the recovery of the chloroplast and cytosol pH (Fig. 3, J–L).

Figure 3.

Figure 3

Open in a new tab

Cytosolic and chloroplast pH in relation to DIC uptake and photosynthesis in P. micans. A to D, Addition of DIC to inorganic carbon-limited cells; E to H, addition of DIC to inorganic carbon-limited cells in the presence of DBS; and I to L, addition of DIC to inorganic carbon-limited cells in the presence of DCMU. Bar = 10 μm.

Intracellular pH, Internal Inorganic Carbon Pool, and Photosynthesis

To observe the immediate response of intracellular pH to HCO3− concentration and inhibitors, ratio images were collected at 2-s intervals (Fig. 4). Direct images were recorded for the rapid measurement of transient intracellular pH (in seconds), because the filtering of images was not possible. The addition of DBS by a custom-made diffusion system to photosynthesizing cells resulted in an immediate transient increase of average intracellular pH from 7.7 to almost 8.5, followed by a steady decline (Fig. 4A). We did not observe this transient change in intracellular pH in the presence of DCMU (Fig. 4A). The intracellular inorganic carbon concentration and the photosynthetic 14CO2 were measured in parallel experiments (Fig. 4B). After the addition of DBS, a decrease in the intracellular inorganic carbon pool was measured within 4 s. The photosynthetic 14CO2-fixation rate was maintained for 4 s but had declined to near zero by 8 s (Fig. 4B). The time course for transient intracellular pH during DBS inhibition closely paralleled the time course for DIC depletion, whereas the photosynthetic rate was only briefly maintained.

Figure 4.

Figure 4

Open in a new tab

A, Immediate effect of DBS addition on transient intracellular pH shown by a time-course experiment with ratio images recorded every 2 s in the absence (▪) and presence (□) of DCMU. B, Immediate effect of DBS addition on the internal inorganic carbon concentration (red bars) and photosynthetic 14CO2 fixation (•). Bar = 10 s.

DISCUSSION

Marine phytoplankton species acquire inorganic carbon for photosynthesis from the DIC of seawater. Within the pH range of seawater (8.0–8.3), the bulk of total DIC is HCO3−, with CO2 being less than 1% of the total (Skirrow, 1975). Although physical parameters (particularly pH, temperature, and salinity) in the marine environment affect the free CO2 concentration (Raven, 1995), it is always well below the Km[CO2] of Rubisco in those algal species for whichRubisco has been characterized (Colman and Rotatore, 1995; Raven, 1997b). This suggests that, in marine phytoplankton species that rely solely on the diffusive entry of CO2 from the bulk phase of the medium to Rubisco in the cell, ambient CO2 concentrations may be limiting for photosynthesis (Talling, 1976; Raven and Geider, 1988; Reibesell et al., 1993). This problem is circumvented in many phytoplankton species that have evolved strategies of inorganic carbon acquisition that require either the direct uptake of HCO3− across the plasma membrane (Merrett et al., 1996; Nimer et al., 1997; Tortell et al., 1997) and/or the indirect use of HCO3− through the catalytic production of CO2 by exofacial CA (Nimer et al., 1997, 1998). The inhibition of extracellular CA by DBS and the concomitant reduction of the photosynthetic rate in P. micans (Table I) provides the most unequivocal evidence to date that indirect HCO3− use can provide the bulk of CO2 fixed in photosynthesis by a marine phytoplankton species.

Of the marine phytoplankton species investigated thus far, most possess a CCM (Raven, 1991), the presence of which is characterized by two distinctive features: (a) the intracellular accumulation of DIC being several times that of the external medium during photosynthesis (Badger et al., 1980; Burns and Beardal 1987; Colman and Rotatore, 1995) and (b) the K0.5[CO2] of the cells being substantially lower than the Km[CO2] of Rubisco (Raven and Johnston, 1991). The intracellular accumulation of inorganic carbon in a dinoflagellate relative to the external medium, shown in this study for the first time to our knowledge (Fig. 1), provides convincing evidence for the presence of a CCM in P. micans. The K0.5[CO2] at pH 5.0 for P. micans (Fig. 1) is substantially below the Km[CO2] that would be expected if the cells possessed a type-II Rubisco (Whitney and Andrews, 1998). The increase in average intracellular pH after the addition of HCO3− to inorganic carbon-limited cells could arise in response to the stimulation of photosynthesis caused by increased CO2 availability or by direct HCO3− influx and accumulation; however, both processes would result in an alkaline pH, a prerequisite for the substantial accumulation of DIC.

The measurement of intracellular pH (Fig. 4) suggests that in P. micans the accumulation of inorganic carbon probably occurs mainly in the chloroplast, as shown for Chlamydomonas reinhardtii (Moroney and Mason, 1991); this accumulation can arise via CO2 uptake with “alkaline trapping,” as HCO3− in the plastid. The presence of DBS sharply decreases the accumulation of intracellular inorganic carbon, the photosynthetic rate (Fig. 1), and the ability of cells to re-establish steady-state intracellular pH values (Fig. 3), reinforcing the major role of extracellular CA in facilitating the availability of CO2 at the exofacial surface of the plasma membrane.

The transient increase in intracellular pH after the addition of DBS may result from the use of the intracellular inorganic carbon pool in photosynthesis, causing alkalization of the cytosol and the chloroplast. In the presence of DBS, another contributory factor affecting cytosolic pH is the absence of CO2 transport and conversion to HCO3− in the cytosol (Volokita et al., 1984). When the photosynthetic rate decreases due to depletion of the intracellular inorganic carbon pool, a decrease in intracellular pH may result from excess reducing equivalents in the cell under conditions of inorganic carbon limitation in the chloroplast. Marine phytoplankton species (Moroney et al., 1985; Jones and Morel, 1988) and other cells (Rubinstein and Luster, 1993) possess a plasma membrane redox chain. The rate of transfer of reducing equivalents to the exterior surface of the plasma membrane is dependent on the inorganic carbon status of the photosynthetic cell (Moroney et al., 1985). The plasma membrane redox system may fuel a trans-membrane proton pump (Jones and Morel, 1988), providing an essential component of a CCM through the maintainance of extracellular CA in the form needed to catalyze the dehydration of HCO3− (Coleman, 1984; Nimer et al., 1998), although active transport at the chloroplast envelope remains a possibility (Moroney and Mason, 1991).

Extracellular CA is constitutive in most of the dinoflagellates tested, including P. micans (Nimer et al., 1997), whereas in other marine phytoplankton species, the development of extracellular CA activity is a response to very low concentrations of CO2 (Iglesias and Merrett, 1997; Nimer at al., 1997). This suggests that in a dinoflagellate such as P. micans there is a need for a CCM even at ambient CO2 concentrations. This may reflect a difference in the specificity factor of Rubisco in some dinoflagellates (Whitney and Andrews, 1998) compared with other algal phyla (Delgado et al., 1995; Raven, 1997a). The development of dinoflagellate blooms occurs under conditions of low turbulence and high temperature (Holligan, 1985; Steindinger and Vargo, 1988), which results in low ambient free CO2 concentrations compared with the open-ocean conditions under which a CCM may be a prerequisite. Therefore, the CCM may be one of several factors that contribute to the existence of near-monospecific dinoflagellate blooms over thousands of miles of coastal waters lasting from weeks to months under conditions of CO2 limitation (Steindinger and Vargo, 1988).

Abbreviations:

CA

carbonic anhydrase

CCM

CO2-concentrating mechanism

DBS

dextran-bound sulfonamide

DIC

dissolved inorganic carbon

PFD

photon flux density

Footnotes

1

This research was supported by the Natural Environment Research Council (UK) (grant no. GR3/09963).

LITERATURE CITED

  1. Aizawa K, Miyachi S. Carbonic anhydrase and CO2 concentrating mechanisms in microalgae and cyanobacteria. FEMS Microbiol Rev. 1986;39:215–233. [Google Scholar]
  2. Anning T, Nimer N, Merrett MJ, Brownlee C. Costs and benefits of calcification in coccolithophorids. J Mar Syst. 1996;9:45–56. [Google Scholar]
  3. Badger MR, Kaplan A, Berry JA. Internal inorganic carbon pool of Chlamydomonas reinhardtii. Plant Physiol. 1980;66:407–413. doi: 10.1104/pp.66.3.407. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Badger MR, Price GD. The role of carbonic anhydrase in photosynthesis. Annu Rev Plant Physiol Plant Mol Biol. 1994;45:369–392. [Google Scholar]
  5. Burns BD, Beardal J. Utilization of inorganic carbon by marine microalgae. J Exp Mar Biol Ecol. 1987;107:75–86. [Google Scholar]
  6. Coleman JE. Carbonic anhydrase. Zinc and the mechanism of catalysis. Ann NY Acad Sci. 1984;81:6049–6053. doi: 10.1111/j.1749-6632.1984.tb12313.x. [DOI] [PubMed] [Google Scholar]
  7. Coleman JR. The molecular and biochemical analyses of CO2 concentrating mechanism in cyanobacteria and microalgae. Plant Cell Environ. 1991;14:861–867. [Google Scholar]
  8. Colman B, Rotatore C. Photosynthetic inorganic carbon uptake in two marine diatoms. Plant Cell Environ. 1995;18:919–924. [Google Scholar]
  9. Delgado E, Medrano H, Keys AJ, Parry MAJ. Species variation in Rubisco specificity factor. J Exp Bot. 1995;46:1775–1777. [Google Scholar]
  10. Dixon GK, Patel BN, Merrett MJ. Planta. 1987;172:508–513. doi: 10.1007/BF00393867. [DOI] [PubMed] [Google Scholar]
  11. Dodge JD, Greuet C . . Dinoflagellate ultrastructure and complex organelles. In: Taylor FJR, editor. The Biology of Dinoflagellates, Oxford, UK: Blackwell Scientific Publications; 1987. pp. 92–142. [Google Scholar]
  12. Dong LF, Nimer NA, Okus E, Merrett MJ. New Phytol. 1993;123:679–684. [Google Scholar]
  13. Guillard RRL, Ryther JH. Studies on marine planktonic diatoms. Cyclotella nana Hustedt and Detonula confervacea (Cleve) Gran. Can J Microbiol. 1962;8:220–239. doi: 10.1139/m62-029. [DOI] [PubMed] [Google Scholar]
  14. Holligan PM (1985). Marine dinoflagellate blooms: growth strategies and environmental exploitation. In DM Anderson, AW White, DG Baden, eds, Proceedings of the Third International Conference on Toxic Dinoflagellates. Elsevier Science Publishing, St. Andrews, New Brunswick, Canada, pp 133–139
  15. Iglesias MD, Merrett MJ. Dissolved inorganic carbon utilization and the development of extracellular carbonic anhydrase by the marine diatom Phaeodactylum tricornutum. New Phytol. 1997;135:163–168. doi: 10.1046/j.1469-8137.1997.00625.x. [DOI] [PubMed] [Google Scholar]
  16. Jones GJ, Morel FMM. Plasmalemma redox activity in the diatom Thalassiosira. Plant Physiol. 1988;87:143–147. doi: 10.1104/pp.87.1.143. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Jordan DB, Ogren WL. Species variation in the specificity of ribulose bisphosphate carboxylase/oxygenase. Nature. 1981;291:513–515. [Google Scholar]
  18. Lorimer GH, Andrews TJ, Tolbert NE. Ribulose diphosphate oxygenase. II. Further proof of reaction products and mechanism of action. Biochemistry. 1973;12:18–23. doi: 10.1021/bi00725a004. [DOI] [PubMed] [Google Scholar]
  19. McConnaughey T. Calcification in Chara corallina: CO2 hydroxylation generates protons for bicarbonate assimilation. Limnol Oceanogr. 1991;36:619–628. [Google Scholar]
  20. Merrett MJ, Nimer NA, Dong LF. The utilization of bicarbonate ions by the marine microalga Nannochloropsis oculata (Droop) Hibberd. Plant Cell Environ. 1996;19:478–484. [Google Scholar]
  21. Moroney JV, Husic HD, Tolbert NE. Effect of carbonic anhydrase inhibitors on inorganic carbon accumulation by Chlamydomonas reinhardtii. Plant Physiol. 1985;79:177–183. doi: 10.1104/pp.79.1.177. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Moroney JV, Mason CB. The role of the chloroplast in inorganic carbon acquisition by Chlamydomonas reinhardtii. Can J Bot. 1991;69:1017–1024. [Google Scholar]
  23. Morse D, Salois P, Markovic P, Hastings JW. A nuclear-encoded form II Rubisco in dinoflagellates. Science. 1995;268:1622–1624. doi: 10.1126/science.7777861. [DOI] [PubMed] [Google Scholar]
  24. Nimer NA, Dixon GK, Merrett MJ. Utilization of inorganic carbon by the coccolithophorid Emiliania huxleyi. New Phytol. 1992;120:153–158. [Google Scholar]
  25. Nimer NA, Iglesias-Rodriguez MD, Merrett MJ. Bicarbonate utilization by marine phytoplankton species. J Phycol. 1997;33:625–631. [Google Scholar]
  26. Nimer NA, Merrett MJ. Calcification and utilization of inorganic carbon by the coccolithophorid Emiliania huxleyi Lohamnn. New Phytol. 1992;121:173–177. [Google Scholar]
  27. Nimer NA, Merrett MJ. The development of a CO2 concentrating mechanism in Emiliania huxleyi. New Phytol. 1996;133:383–389. [Google Scholar]
  28. Nimer NA, Warren M, Merrett MJ. The regulation of photosynthetic rate and activation of extracellular carbonic anhydrase under CO2-limiting conditions in the marine diatom Skeletonema costatum. Plant Cell Environ. 1998;21:805–812. [Google Scholar]
  29. Parsons T, Maita YR, Lalli CM (1989) A Manual of Chemical and Biological Methods for Sea Water Analysis. Pergamon Press, London, pp 141–148
  30. Raven JA. Implications of inorganic carbon utilization: ecology, evolution, and geochemistry. Can J Bot. 1991;69:908–924. [Google Scholar]
  31. Raven JA. Carbon fixation and carbon availability in marine phytoplankton. Photosynth Res. 1994;39:259–273. doi: 10.1007/BF00014587. [DOI] [PubMed] [Google Scholar]
  32. Raven JA. Photosynthetic and non-photosynthetic roles of carbonic anhydrase in algae and cyanobacteria. Phycologia. 1995;34:93–101. [Google Scholar]
  33. Raven JA. Putting the C in phycology. Eur J Phycol. 1997a;32:319–333. [Google Scholar]
  34. Raven JA. Inorganic carbon acquisition by marine autotrophs. In: Callow JW, editor. Advances in Botanical Research. London: Academic Press; 1997b. pp. 85–209. [Google Scholar]
  35. Raven JA, Geider R. Temperature and algal growth. New Phytol. 1988;110:441–446. [Google Scholar]
  36. Raven JA, Johnston AM. Mechanisms of inorganic carbon acquisition in marine phytoplankton and their implications for the use of other resources. Limnol Oceanogr. 1991;36:1701–1717. [Google Scholar]
  37. Reibesell U, Wolf-Gladrow DA, Smetacek V. Carbon dioxide limitation of phytoplankton growth rates. Nature. 1993;361:249–251. [Google Scholar]
  38. Rowan R, Whitney SM, Fowler A, Yellowlees D. Rubisco in marine symbiotic dinoflagellates: form II enzymes in eukaryotic oxygenic phototrophs encoded by a nuclear multigene family. Plant Cell. 1996;8:539–553. doi: 10.1105/tpc.8.3.539. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Rubinstein B, Luster DG. Plasma membrane redox activity: components and role in plant processes. Annu Rev Plant Physiol Plant Mol Biol. 1993;44:131–155. [Google Scholar]
  40. Seksek O, Henry-Toulme N, Bolard J. SNARF-1 as an intracellular pH indicator in laser microspectrofluorimetry: a critical assessment. Anal Biochem. 1991;193:49–54. doi: 10.1016/0003-2697(91)90042-r. [DOI] [PubMed] [Google Scholar]
  41. Skirrow G. The dissolved gases carbon dioxide. In: Riley JP, Skirrow G, editors. Chemical Oceanography, Vol 2. London: Academic Press; 1975. pp. 1–192. [Google Scholar]
  42. Spalding MH, Spreitzer RJ, Ogren WL. Carbonic anhydrase-deficient mutant of Chlamydomonas reinhardtii requires elevated carbon dioxide concentration for photoautotrophic growth. Plant Physiol. 1983;73:268–272. doi: 10.1104/pp.73.2.268. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Steindinger KA, Vargo G (1988) Marine dinoflagellate blooms: dynamics and impacts. In C Lembi, JR Waaland, eds, Algae and Human Affairs. Cambridge University Press, New York, pp 373–401
  44. Tabita FR. The biochemistry and metabolic regulation of carbon metabolism and CO2 fixation in purple bacteria. In: Blankenship RE, Madigan MT, Bauer CE, editors. Anoxygenic Photosynthetic Bacteria. Dordrecht, The Netherlands: Kluwer Academic Publishers; 1995. pp. 885–914. [Google Scholar]
  45. Talling JF. The depletion of carbon dioxide from lake water by phytoplankton. J Ecol. 1976;64:79–121. [Google Scholar]
  46. Tortell PD, Reinfelder JA, Morel FMM. Active uptake of bicarbonate by diatoms. Nature. 1997;320:243–244. [Google Scholar]
  47. Tsuzuki M, Miyachi S. The function of carbonic anhydrase in aquatic photosynthesis. Aquat Bot. 1989;34:85–104. [Google Scholar]
  48. Vesk M, Jeffrey W. Effect of blue-green light on photosynthetic pigments and chloroplast structure in unicellular marine algae from six classes. J Phycol. 1977;13:280–288. [Google Scholar]
  49. Volokita M, Zenvirth D, Kaplan A, Reinhold L. Nature of the inorganic carbon species actively taken up by the cyanobacterium Anabaena variabilis. Plant Physiol. 1984;76:599–602. doi: 10.1104/pp.76.3.599. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Whitney SM, Andrews TJ. The CO2/O2 specificity of single-subunit ribulose-bisphosphate carboxylase from the dinoflagellate, Amphidinium carterae. Aust J Plant Physiol. 1998;25:131–138. [Google Scholar]
  51. Whitney SM, Yellowlees D. Preliminary investigations into the structure and activity of ribulose bisphosphate carboxylase from two photosynthetic dinoflagellates. J Phycol. 1995;31:138–146. [Google Scholar]

Articles from Plant Physiology are provided here courtesy of Oxford University Press

RESOURCES