Enhanced production of a lutein-rich acidic environment microalga.

Journal of Applied Microbiology ISSN 1364-5072

ORIGINAL ARTICLE

Enhanced production of a lutein-rich acidic environment microalga zquez, M.C. Ruiz-Domınguez and C. Vılchez I. Vaquero, M. Va Algal Biotechnology Group, International Centre for Environmental Research (CIECEM), Almonte, Spain

Keywords acidic environment microalgae, carbonic anhydrase, Coccomyxa, inorganic carbon utilization, lutein. Correspondence Isabel Vaquero, Algal Biotechnology Group, International Centre for Environmental Research, (CIECEM), Parque Dunar s/n, Mat~as, Almonte, 21760 Huelva, Spain. alascan E-mail: [email protected] 2013/1424: received 18 July 2013, revised 2 December 2013 and accepted 18 December 2013 doi:10.1111/jam.12428

Abstract Aims: This study was aimed at increasing productivity of a novel lutein-rich acidic environment microalga, Coccomyxa onubensis, based on efficient inorganic carbon use. Methods and Results: Productivity was determined based on dry weight data; inorganic carbon concentration mechanisms were determined by means of carbonic anhydrase activity; carotenoids were extracted with methanol and measured by HPLC techniques. The existence of carbon concentration mechanisms and conditions that might lead to use them for addressing increased productivity of C. onubensis was studied. Best growth and carbon uptake capacity occurred at acidic pH, proving acid-tolerant behaviour of C. onubensis. Incubation in air followed by shift to high carbon conditions enhanced carbon-use efficiency in terms of growth rate and biomass productivity, based on the action of both carbonic anhydrase activities. Lutein accumulated in the microalga at high concentrations above 5–6 g kg1 dry weight and did not depend on inorganic carbon conditions. Conclusions: Consequently, repeated cycles of air incubation and high CO2 incubation of C. onubensis might become a suitable tool to perform production processes of lutein-enriched biomass. Significant and Impact of the Study: This study intends to show that acidic environment microalgae can be produced at similar productivities of nonextreme microalgae, with the added advantage of their growth in highly selective culture medium. Particularly, it is applied to C. onubensis which accumulates lutein at commercially relevant concentrations.

Introduction Besides light intensity, one of the major factors limiting algal productivities is CO2 availability and uptake. To achieve high productivities of biomass and high-value products, microalgae cultivation is often performed with overly CO2 added. Coccomyxa onubensis naturally grows under a very low pH range (1–3), so inorganic carbon is mostly present as CO2 at air equilibrium concentration. The Tinto River, C. onubensis natural habitat, is an extreme acidic environment characterized by a very low pH (1–3). As a result of such high acidity, inorganic carbon (Ci) is mostly as CO2 at air equilibrium concentration. The equilibrium between inorganic carbon forms in aqueous media is pH dependent. At normal intracellular

ionic strength, when the pH level is below 64 (first ionization constant for carbonic anhydrase, pK1), CO2 predominates; at pH between 64 and 103 (pK2), HCO3 predominates, and above pH 103, CO32 predominates. Therefore, Ci concentration is substantially lower at acidic pH than in higher pH waters with bicarbonate as main inorganic carbon source. That situation denotes that carbon availability is a limiting factor for the micro-organisms inhabiting acidic waters, limiting its potential for systematic mass production. It is also inferred from the lack of reports on acidophilic microalgae biomass production trials. Algae have adapted to these challenges through the development of CO2 concentration mechanism (CCM). The CCM is a biological adaptation to low carbon dioxide concentrations in the environment. It is a

Journal of Applied Microbiology 116, 839--850 © 2013 The Society for Applied Microbiology

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mechanism which augments photosynthetic productivity in algal cells by increasing internal levels of inorganic carbon many times over the environmental concentration of carbon dioxide. Previous research on carbon concentration mechanisms (CCMs) has revealed that most microalgae and cyanobacteria can import both CO2 and HCO3– through the cell membrane (Giordano et al. 2005; Price et al. 2008; Maberly et al. 2009). Once imported into the cell, CO2 or HCO3 is accumulated mainly as HCO3– due to its neutral internal pH. Compared with cyanobacteria, less information is available on Ci transport systems in eukaryotic algae (Yamano and Fukuzawa 2009). However, it has been reported that, in addition to Ci transporters in the cell membrane, eukaryotic algae such as Chlamydomonas also have chloroplast Ci transporters, because photosynthesis in eukaryote microalgae occurs in the chloroplast (Markelova et al. 2009; Yamano and Fukuzawa 2009). In addition, CA might contribute to the transport of HCO3– into the thylakoid lumen and its conversion into CO2 (Yamano and Fukuzawa 2009). In algae, carbonic anhydrase (CA) has been recognized as one of the essential elements of the CO2 concentration mechanisms (Jordan and Ogren 1981; Morita et al. 1998; Spalding 1998; Spreitzer 1999). Carbonic anhydrase is a zinc metalloenzyme that catalyses the interconversion of CO2 and HCO3 (Khalifah 1971). The enzyme was first discovered in red blood cells but has also been found in most organisms including animals, plants, archaebacteria and eubacteria (Hewett-Emmett and Tashian 1996). Carbonic anhydrase (CA) is important in many physiological functions that involve carboxylation or decarboxylation reactions, including both photosynthesis and respiration. In addition, it is clear that CA also participates in the transport of inorganic carbon to active photosynthesizing cells or away from actively respiring cells (Henry 1996). Main function of internal carbonic anhydrase of microalgae is converting bicarbonate to carbon dioxide in the active site of RuBisCO, therefore enhancing carbon fixation in microalgae. Some microalgae living in acidic environments do hardly display CCM under carbon-limiting conditions (Colman and Balkos 2005). Accordingly, inorganic carbon acquisition (Ci) might take place through the cell membrane by either active or passive diffusion. However, there are other acidophilic and acid-tolerant algae that have been shown to have CCM (Raven et al. 1982). In that respect, knowledge about CO2 concentrating mechanisms in Coccomyxa genus is scarce. Verma et al. (2009) proposed the presence of an external carbonic anhydrase (CAext) in Coccomyxa although with a carbon transport facilitating role rather than a concentrating function. These results indicate that there is species-specific variation in the induction mechanism of CCM depending on 840

physiological and ecological conditions (Giordano et al. 2005; Raven 2010). Coccomyxa onubensis is expected to show high affinity for carbon dioxide due to the low availability of CO2 in such acidic media. Therefore, knowing whether C. onubensis possesses CO2 concentrating mechanisms (CCMs) and the potential factors that may lead to increase its activity is of value to understand how to increase the algal productivity. In that respect, our work was focused on the effect of high/low CO2 availability cycles on the algal productivity, a tool to increase CO2 uptake efficiency. The low external pH theoretically forces acidic environment microalgae to expend energy in order to maintain neutral pH into the cytosol, using appropriate biochemical systems that withstand the proton gradient across the plasma membrane (Gross 2000). The energy demand of such systems in part justifies a remarkable activity of PSII, even in cultures growing under low carbon conditions. Coccomyxa onubensis was isolated and identified in a previous work of our group (Garbayo et al. 2012). That acid-tolerant microalga was found in screened water samples obtained from acidic drainages in the pyritic belt area around the Tinto River, in the Province of Huelva (Spain). The natural habitat of that microalga is highly concentrated in solved metals and poor in nutrients including N and P. Such extreme environment addresses Coccomyxa to express some typical antioxidant responses. One of the most attractive physiological responses in terms of commercial applications is the large accumulation of lutein, above 5 g kg1 dry biomass and accounting for about 80% of the total pigment pool. This makes of C. onubensis an attractive model acid-tolerant microalga to show that production of value compounds of extreme microalgae might be feasible. Production of acidophilic or acid-tolerant microalgae meets the advantage of selective growth in acidic culture medium, which avoids competition from most of nonadapted microorganisms. Accordingly, this work first aims at studying C. onubensis growth at different pH values, assessing its ability to grow in terms of proton gradient in several pH ranges. Moreover, in this work, suitable strategies of carbon supply are chosen for conducting the extremophile algal growth, showing results that evidence similar algal productivities to those of the so-called common microalgae. Materials and methods Micro-organism and culture conditions Coccomyxa onubensis was isolated from acidic waters of the Tinto River (Huelva, Spain). This river has some very


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special features, such as low pH and a high concentration of heavy metals, especially iron, copper, magnesium and aluminium (Ferris et al. 2004). An axenic culture of the algae was obtained by streaking it on basal agar medium at pH 25 and then was transferred to the liquid medium. Coccomyxa has been identified by ribosomal 18S subunit rDNA sequence analysis (Garbayo et al. 2012). Identified 18S subunit rDNA sequence was registered at GenBank with accession number GU265559. Coccomyxa cells were grown in 1-l Erlenmeyer flasks. According to the chemical composition of the natural environment (Ferris et al. 2004; Garbayo et al. 2012), cultures were grown at pH 25 in a culture medium based on K9 medium (Silverman and Lundgren 1959). A modified K9 medium was prepared according to the following composition: 395 g K2SO4, 01 g KCl, 05 g K2HPO4, 041 g MgCl2, 229 g KNO3, 001 g CaCl2, 5 ml Hutner solution (Hutner et al. 1950) added to a final volume of 1 l of distilled water. Hutner solution was prepared as it follows, for 1 l: 10 g EDTA disodium salt, 440 g ZnSO47H2O, 228 g H3BO3, 102 g MnCl24 H2O, 032 g CoCl26H2O, 032 g CuSO45H2O, 032 g (NH4)6Mo7O244H2O and 100 g FeSO47H2O. A volume of 5 ml Hutner solution was added to the final volume of 1 l of distilled water. The cultures were incubated at 27°C and were illuminated at 150 lE m2 s1 with white fluorescent lamps. The cultures were bubbled with air only, under the so-called low CO2 (LC) conditions, or bubbled with air containing 5% (v/v) CO2, the so-called high CO2 (HC) conditions, ‘control cultures’. The pH experiments in this manuscript were performed in 1-l flasks run in semicontinuous mode. To do this, algal cultures were always maintained within a range of biomass concentration (12–15 g l1), which was previously determined as optimal. That was done by diluting the cultures with fresh culture media. The semicontinuous cultures were bubbled with air containing 5% (v/v) CO2. The pH of each culture was monitored and corrected by either HCl or NaOH addition. The algal cultures were incubated in a culture room under controlled temperature (27°C) and continuous white fluorescent light at a fixed irradiance of 150 lE m2 s1. The bubbled gas flow rate was 190 ml min1 per litre throughout all the experiments. In all laboratory experiments, 045-lm air filters were used in the air supply line. Dry weight measurements Whatman glass microfiber filters (Ø 47 mm, pore size 07 lm) were dried at 95°C overnight and placed in a desiccator to cool them at room temperature. The empty filters were weighed. Approximately 10 mg of sample


(triplicate) was filtrated. Each filter was rinsed twice with demineralized water to remove adhering inorganic salts. The wet filters containing the samples were dried at 95°C overnight, allowed to cool at room temperature in a desiccator and weighed. Growth rate calculations Growth rates were calculated on dry weight data. Specific growth rates of cultures were calculated using the following expression: l ¼ LnðC=C0 Þ=t where l is the specific growth rate, C0 is the initial biomass concentration (dry weight) and C is the biomass concentration at any time t. In batch culture experiments, specific growth rates were calculated from the logarithmic growth phase. In semicontinuous cultures, average growth rates were calculated on averaged daily dry weight data. Microalgal optical density The optical density of the sample (triplicate) was measured at 750 nm (OD750nm). Demineralized water served as reference. The microalgal samples were diluted using demineralized water to achieve OD750nm values below 1. The calibration constant for dry biomass concentration versus optical density at 750 nm was calculated. OD value of 10 at 750 nm corresponds to 047 g dry weight per l culture. Measurement of CA activity External carbonic anhydrase (CAextt) activity was determined by the modified potentiometric technique described by Williams and Colman (1993). Cells were harvested by centrifugation at 5000 g for 10 min, three times washed with 30 mmol l1 Hepes buffer, pH 81, and resuspended at a concentration of c. 25 lg chl ml1 in 15 ml of the same buffer. To determine total carbonic anhydrase (CAt) activity, cells were harvested by centrifugation at 5000 g for 10 min, three times washed with 30 mmol l1 Hepes buffer, pH 81, and then homogenized using a sonicator. The disrupted biomass was resuspended in 15 ml of the same buffer, and an aliquot of that homogenate was taken for CA measurement. Icecold, CO2-saturated distilled water (05 ml) was injected into the cell suspension or cell homogenate at 4°C, and the time taken for the pH of the suspension to drop from pH 81 to 76 was measured. Carbonic anhydrase activity was calculated as WilburAnderson units (WAU) using the formula (Tc/Ts)-1,


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where Tc and Ts represent the time taken for the pH change, in the presence and absence of cells or extract, respectively (Wilbur and Anderson 1948). Both external and total CA activities were expressed as WAU per milligram of protein (WAU mg1 prot). To determine the effect on photosynthesis of inhibiting external CA, a sample of cell suspension was treated with acetazolamide (AZA), a CA inhibitor.

Statistics Unless otherwise indicated, tables and figures show means and standard deviations (SD) of three independent experiments. Results Effect of pH on algal growth and lutein content

Quantum yield Fluorescence measurements correspond to maximal quantum yield (QY) of PSII (Fv/Fm). QY was measured to evaluate the viability of the cells. QY was determined using a pulse amplitude modulation (PAM) (Schreiber et al. 1995). Samples of each culture were previously adapted to darkness for 15 min. Carotenoids determination by HPLC Carotenoids were extracted using aliquots (1 ml) of the cultures. Cells were spinned down for 8 min at 13 000 g. The obtained pellet was placed in an ultrasonic bath at 60°C for 5 min. The pellet was resuspended in 1 ml of methanol and the suspension shaken vigorously for 1 min and centrifuged for 8 min at 3000 g. Carotenoids and chlorophylls a and b were separated and identified by HPLC (TermoQuest, Thermo Separation products, Waltham, MA) with a RP-18 column, using a modified method described by Young et al. (1997). In the mobile phase, solvent A was ethyl acetate and solvent B was acetonitrile and water (9 : 1, v/v). External standards (DHI) and their corresponding calibration curves were used to identify and quantify both lutein and b-carotene. 842

The pH of culture medium determines main dissolved inorganic carbon form available and its concentration. Therefore, microalgal growth rate and productivity will be pH dependent. Figure 1 shows growth rates and total carbonic anhydrase (CA) activities of C. onubensis cultures acclimated at the indicated pH values. The cultures were grown under high level of CO2 (5% v/v in air). The highest cell densities were obtained at pH 40. Results also show that the lower intensity of the proton gradient, at neutral pH, does not inhibit C. onubensis growth. Actually, C. onubensis showed growth over the pH range 25–90. This suggests that C. onubensis internal pH should remain neutral, enabling the microalga for performing photosynthesis in the whole pH range (25–90). Particularly, the cultures grown at pH 90 lost their cell viability at early growth stages, although still were able to grow at some extent. Therefore, C. onubensis might be considered as an acid-tolerant microalga rather than an acidophilic microalga. Figure 1 also shows total CA activity, which increased with the pH increase in the culture medium, the maximum being found in cultures at pH 6 1·2

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Lutein content (mg g–1)

The protein content of algal samples was measured in crude extracts. These were prepared as indicated: cells were harvested by centrifugation at 5000 g for 10 min and three times washed with 30 mmol l1 Hepes buffer, pH 81, and the cell pellets were homogenized using a sonicator. The disrupted biomass was resuspended in 15 ml of 30 mmol l1 Hepes buffer, pH 81. The cell suspension was centrifuged, and the protein content was measured in the supernatant. The Bradford method (Bradford 1976) was used to measure the protein content. Protein in algal samples is calculated from spectrophotometric absorbance at 595 nm of unknown protein concentration samples. The protein concentrations are obtained on a calibration curve prepared using standard solutions of bovine serum albumin (BSA).

Growth rate (day–1) and CA activity (WAU mg–1 prot)

Protein measurement

0 10

pH Figure 1 Specific growth rate (●), total CA activity ( ) and lutein content ( ) of Coccomyxa onubensis cultures at different pH values. The cultures were operated in semicontinuous mode, as described in Materials and Methods. Growth rates were calculated by measuring optical density at 750 nm in stabilized cultures. Total carbonic anhydrase activities were expressed as per milligram of protein (WAU mg1 prot.). Growth rate, CA activity and lutein content were measured on the same samples. The pH data represent the pH values of the growing cultures. Values given as means SD (three replicates).


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Effect of CO2 conditions on growth rates, biomass productivities and carotenoids content According to the results above, acidic pH values—from 25 to 4 in this work—would be more suitable to run lutein production processes with C. onubensis. Furthermore, pH 25 would add to the culture medium the advantage of highly acidic character, therefore becoming more selective for growth, then making biological contamination more difficult in the large scale. As already discussed, CO2 availability at acid pH is limited and its uptake by the microalga seems to be enhanced through active carbon concentration mechanisms. CO2 level should therefore influence such carbon concentration activity and consequently the growth rate of C. onubensis.

Cell density evolution (DO750)

Growth rates and carbonic anhydrase activity of C. onubensis were determined in batch cultures which were bubbled with 5% (v/v) CO2-enriched air (HC conditions) or only air (LC conditions) under laboratory culture conditions, continuous light of 150 lmol photons m2 s1 and 27°C (Fig. 2). Optical density at 750 nm was measured as a cell density factor, as mentioned in the Materials and Methods section. Calculated growth rates were c. 023 and 017 day1 under HC and LC conditions, respectively, suggesting that C. onubensis cells grew just slightly faster under HC

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1·6 CA activity (WAU mg–1 prot)

and decreasing at higher pH values. CA activity is mainly aimed at converting bicarbonate into carbon dioxide, therefore acting as carbon concentration mechanism to effectively increase inorganic carbon pool inside the microalga cell. When cultivated under suitable conditions, C. onubensis accumulates lutein at intracellular concentrations above 5 g kg1 dry weight, which are among the highest published for microalgae (Garbayo et al. 2012) and make that microalga attractive for lutein production. Figure 1 shows C. onubensis lutein content, when grown at different pH values. Lutein content (mg g1 DW) reached its maximum value in cultures at pH 7. In previous work, we reported that C. onubensis naturally accumulates a high constitutive pool of lutein (Garbayo et al. 2012), which increases depending on cultivation conditions and accounts for about 60–80% of total pigment pool (Vaquero et al. 2012). According to growth rates and lutein accumulation values obtained in Coccomyxa cells grown on continuous supply of high level of CO2 in air (5% v/v), lutein productivities, in mg l1 day1, tend to be slightly higher in the acidic range 25–40. The trends observed, however, were not significant (Table 1). As shown in that table, biomass productivities are also higher within that pH range. In addition to that, the alga ability to grow at highly acid pH (25) becomes a selective competitive advantage for C. onubensis in continuous production processes, compared to non-acid-tolerant microalgae.

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Figure 2 Cell density evolution (a) and carbonic anhydrase activity (b) of control cultures, HC ( ) and low CO2 cultures, LC ( ) of Coccomyxa onubensis. Solid bars show total carbonic anhydrase activity. Cross-hatched bars show external carbonic anhydrase, in HC and LC cultures. Cell density evolution was measured as optical density at 750 nm (OD750). Carbonic anhydrase activities were expressed as WAU mg1 prot. (milligram of protein). Values given as means SD (three replicates).

Table 1 Biomass productivities (g l1 day1) and lutein productivities (mg l1 day1) of Coccomyxa onubensis cultured at different pH. The cultures were operated in semicontinuous mode and supplied with high level of CO2 in air (5% v/v). Values given as means SD (three replicates)

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Biomass productivity (g l day ) Lutein productivity (mg l1 day1)

pH 25

pH 4

pH 6

pH 7

pH 9

041 013 235 021

042 014 252 023

034 011 221 014

031 012 232 018

027 014 155 017


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Biomass concentration (g l–1)

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Biomass productivity (g l–1 day–1)

conditions than under LC conditions. External and total CA activities were measured along the experiment, and the average data are represented in Fig. 2. Both external and total CA activities per milligram of protein (WAU mg1 prot) were higher under LC conditions than under HC conditions. This result suggests that C. onubensis might show higher photosynthetic affinity for inorganic carbon under LC conditions than under HC conditions. According to the obtained results, pre-incubation of C. onubensis under LC conditions (air only) might enhance the algal CO2 uptake capacity, which might further be used to address increased CO2 uptake by means of transferring LC-grown cultures to HC conditions. In that respect and aimed at enhancing productivity of C. onubensis cultures, experiments addressed to assess CO2 acquisition capacity of C. onubensis were run. By means of a suitable regime of inorganic carbon supply to C. onubensis cultures, the microalga might increase its biomass productivity, which would also become essential to achieve high productivities of high-value products, namely lutein in C. onubensis. Accordingly, aliquots of C. onubensis cultures grew under LC conditions over different specified time periods (6, 12, 24 or 36 h, respectively). After that period under LC conditions, C. onubensis cultures were transferred to HC conditions (5% CO2-enriched air), accounting for a total experiment time of 60 h, all cultures. A control culture was grown for 60 h under HC conditions. Incubation time under LC conditions does influence C. onubensis growth. During early growth stages, a growth rate deceleration was observed in those cultures grown under LC conditions, compared to control cultures under HC conditions. However, Fig. 3 also shows that cultures grown under low carbon conditions do reactivate their growth when transferred to a carbon-rich environment, reaching higher growth rates and productivities. Surprisingly, after 60 h, those cultures first incubated under LC conditions and then transferred to HC conditions reached higher cell densities than control cultures (grown for 60 h under high CO2 conditions), accounting for 40% higher values. This is illustrated in Fig. 3 that shows productivities achieved by those C. onubensis batch cultures transferred to HC conditions, as a function of pre-incubation time in low CO2 (LC conditions). The biomass productivities were calculated during the incubation time of Coccomyxa cultures under HC conditions. Figure 3 shows that biomass productivity of Coccomyxa cultures transferred from only air to high CO2 (LC?HC) increases as a function of the previous residence time of cultures in air. Coccomyxa onubensis reaches its maximum productivity (18 g l1 day1) when grown for 36 h under LC conditions (air bubbling

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Figure 3 Growth kinetics (a) and biomass productivity (b) of Coccomyxa onubensis growth under LC conditions and further transferred to CO2-enriched air (HC) conditions. Growth kinetics and biomass productivities were calculated from dry weight data and were expressed as g l1 and g l1 day1, respectively. ( ) represents C. onubensis cultures grown for 6 h under LC conditions, ( ) shows cultures grown for 12 h under LC conditions, ( ) shows cultures grown for 24 h under LC condition ( ) shows cultures grown for 36 h under LC conditions, and ( ) represents the HC culture (no hours in air). Arrows indicate when cultures were transferred from only air to 5% CO2-enriched air. Values given as means SD (three replicates).

only) followed by 24-h incubation under high CO2. That maximal productivity was reached in the last 12 h under HC conditions (experiment time: from 48 to 60 h). That productivity is 4-fold that value achieved in the control culture, which grew 60 h under 5% CO2-enriched air. It has been shown that pre-incubation of acid-tolerant microalga cultures in air followed by shift to high carbon conditions (LC?HC) by far enhances carbon-use efficiency in terms of growth rate and biomass productivity. In a further experiment, it was probed that there is a limit for such enhanced carbon-use efficiency and that such limit depends on for how long carbon demand is active. As shown in Fig. 4, both external and total CA of C. onubensis become particularly active within the first


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LC/HC ratio of external CA activity

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Figure 4 Ratio of LC cultures to HC cultures external carbonic anhydrase activity (a), and ratio of LC cultures to HC cultures total carbonic anhydrase activity (b). Both external and total carbonic anhydrase activities were calculated per milligram of protein (WAUext mg1). Average external CA of HC cultures during 48-h incubation = 0693 0055 WAUext mg1 prot. Average total CA of HC cultures during 48-h incubation = 0987 0079 WAUext mg1 prot. Values given as means SD (three replicates).

24–48 h of incubation in air only (LC), compared to CA of HC-grown cells (control cultures). However, as expected, longer incubation periods in air address CA of LC-grown cells falling to those values of control cultures under high CO2. Therefore, the maximal carbon uptake activity of C. onubensis cultures can be achieved through air incubation periods of 24–48 h. As previously shown (Fig. 3), air-incubated cultures further transferred to HC conditions are followed by massive carbon incorporation into biomass, which seems to be maximal after a period of 24–36 h under HC conditions. Longer periods under HC conditions do not improve productivity. In that respect, Table 2 shows carbonic anhydrase activity,

growth rate and productivity data at the end of the cultivation period for 48-h air-incubated C. onubensis cultures which were further transferred to HC conditions for 6 days. Observation of trends, however not significant, indicates that highest growth rate and biomass productivity occur in LC?HC cultures. In addition to that, quantum yield of C. onubensis cultures under both cultivation scenarios is slightly higher in LC?HC cultures. Values between 07 and 08 are considered as typical QY values for viable algal cells growing under nonphotoinhibiting conditions (Cuaresma et al. 2011). The slightly higher values in cultures of Coccomyxa transferred from low to high CO2 environments are consistent with a higher use of inorganic carbon expected in these cultures, in good agreement with the enhanced carbonic anhydrase activity. Papazi et al. (2008) reported higher quantum yields in algal cultures incubated under very high CO2 levels. The value xanthophyll lutein is the major carotenoid of C. onubensis and its accumulation in large amounts inside the microalga is commercially interesting, as described just for few other nonextremophile microalgae species (Fernandez-Sevilla et al. 2010). For lutein production, it becomes relevant to know whether C. onubensis shows any variation in its lutein content (mg g1) if grown under low carbon conditions. Figure 5 shows C. onubensis lutein and b-carotene content evolution when cultured for 48 h under LC conditions followed by 6 days under HC conditions. Coccomyxa onubensis control cultures grew at HC conditions during the 8-day experiment. Both carotenoids content were identified and quantified by HPLC as mentioned in Materials and Methods section. Lutein and b-carotene contents (mg g1 DW) seem to remain constant during the whole experiment, and CO2 concentration in the medium (only air or HC conditions) seems not to have any impact in the biosynthesis rate. Average intracellular lutein content is roughly 6 mg g1 dry weight, similar to that of control culture, and b-carotene content accounted for 05–07 mg g1 dry weight throughout the experiment. Accordingly, C. onubensis pre-incubation under LC conditions does not result in a decreased intracellular carotenoid content. As growth rate and biomass productivity are enhanced when cultures are grown with combined periods of low CO2 and high CO2

Table 2 Growth rate, carbonic anhydrase activity and quantum yield of Coccomyxa onubensis cultures subjected to pre-incubation in air (LC conditions) and further grown on high CO2 (HC conditions) for 6 days. Results are compared with those obtained in cultures grown in HC conditions only. Values given as means SD (three replicates) Inorganic carbon conditions

Growth rate (d1)

Productivity (g l1 day1)

Total carbonic anhydrase (WAU mg1 prot)

Quantum yield

Control cultures (HC) 48-h air-incubated cultures (LC?HC)

017 008 021 005

019 009 022 007

065 007 070 004

068 004 077 004


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Lutein and β-carotene (mg g–1)


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Time (days) Figure 5 Lutein and b-carotene content evolution of Coccomyxa onubensis cultures pre-incubated for 48 h in air (LC conditions) followed by 6 days under HC conditions ( ). C. onubensis control cultures grew at HC conditions during the 8-day experiment ( ). Solid bars show lutein content, and cross-hatched bars show b-carotene content. Values given as means SD (three replicates).

levels, lutein productivity therefore results enhanced. Consequently, repeated cycles of air incubation and high CO2 incubation of C. onubensis cultures might become a suitable tool to perform production processes of luteinenriched biomass. Discussion Effect of pH on algal growth and lutein content We aimed at studying C. onubensis growth at different pH values to assess its ability to grow in terms of the proton gradient between the outside and inside the cell membrane and to assess its ability to use the available inorganic carbon source at different pH values. That information should allow us for defining suitable growth strategies to increase biomass productivity of that acidic environment microalga. As explained, inorganic carbon form availability depends on pH of the culture medium. The fact that C. onubensis better grows at acidic pH, but that can also grow at neutral or even slightly basic pH, suggests acid-tolerant behaviour of that microalga. This is opposite to what is observed in stringent acidophilic microalgae as Dunaliella acidophila (Gimmler and Weis 1992), but it is in good agreement with Verma et al. (2009) who reported Coccomyxa sp growth over the pH range 30–90, indicating that the microalga should maintain constant internal pH and photosynthesis over that pH range. The last has indeed been reported for several acidic environment microalgae (Seckbach and Oren 2007), including measurements of neutral intracellular pH. Such large proton gradient is overcome through extra energy expenses, which have been calculated to 846

account for about 7% total ATP produced by the cell (Messerli et al. 2005). Extra energy maintenance costs should make growth rate and productivity lower if compared to ‘common’ microalgae (Gross 2000), due to lessenergy fraction available for anabolism. As energy needs are higher in acidic environment microalgae, efficient growth and therefore enhanced productivity should be achieved by making use of specific advantages of such extreme micro-organisms. One of the main tools, proposed in this manuscript, would consist of profiting from the carbon concentrating activity of these microalgae. Coccomyxa onubensis apparently shows external CA even growing at pH 25 (Fig. 1), perhaps suggesting different adaptation patterns to acidic media within the same microalga genus, depending on the natural habitat of each species. Based on the need for enhancing inorganic carbon uptake, carbonic anhydrase (CA) acts as carbon concentration mechanism. Several studies describe that CA activity is pH dependent in some acid-tolerant microalgae (Balkos and Colman 2007), which external carbonic anhydrase activity is mainly expressed when the microalgae grew at pH up to 5. At neutral pH, HCO3 is the predominant inorganic carbon source. If an algal cell can use only CO2 from the external medium, the rate of photosynthesis will be limited, in the absence of a catalyst, by the rate of spontaneous CO2 formation from HCO3 dehydration. If the photosynthetic rates exceed the calculated uncatalysed dehydration rate, it would indicate that HCO3 is taken up as a source of Ci (Verma et al. 2009). Coccomyxa onubensis growth seems to occur more efficiently at pH below 6, where CO2 is by far the most abundant inorganic carbon source. Therefore, growth should be highly dependent on CO2 availability. The higher CA activity at intermediate pH values (6–7) seems not to compensate the low CO2 level available and used by C. onubensis as main carbon source, as inferred from the lower growth rate values obtained at pH above 6. The fact that cell growth occurs at neutral pH suggests that HCO3 can also be used as carbon source, but much less efficiently. Therefore, acidic pH was selected as most suitable to produce C. onubensis. If inorganic carbon can be used in an efficient way as to enhance productivities of the acid-tolerant microalga, also lutein productivity—a value xanthophylls accumulated in high concentrations in C. onubensis—would also result increased. Effect of CO2 conditions on growth rates, biomass productivities and carotenoids content As explained, results in Fig. 2 showed that both external and total CA activities per milligram of protein were higher under LC conditions than under HC conditions,


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suggesting that C. onubensis might show higher photosynthetic affinity for inorganic carbon under LC conditions than under HC conditions. This seems reasonable considering that generally, eukaryotic microalgae and cyanobacteria have developed efficient CO2 utilization mechanisms and exhibit high affinity for CO2 when grown under CO2-limiting conditions (Miyachi et al. 2003). Particularly for acidic environment microalga from the Tinto River, previous results of our group obtained in cultures of an acidic environment microalga showed much higher affinity of the alga for CO2 than for bicarbonate, measured at acid pH as apparent affinity constant for dissolved inorganic carbon (Cuaresma et al. 2006). Under saturating CO2 conditions, microalgae exhibit low affinity for CO2, as enough CO2 is available for photosynthesis (Miyachi et al. 2003). In C. onubensis, these results reveal a likely existence of an external (in addition to the internal) carbonic anhydrase, which seems to raise its activity when C. onubensis grows under low CO2 levels. For that reason, a suitable regime of inorganic carbon supply to C. onubensis cultures, pre-incubation in air plus different time periods under high CO2, was thought to be effective for driving to increased biomass productivities, essential to achieve high productivities of lutein in C. onubensis. As shown in Fig. 3, that biomass productivity was 4-fold that value achieved in control cultures. Such high productivity data have not been previously described for acidophilic microalgae (Visviki and Palladino 2001; Beamud et al. 2010; Spijkerman et al. 2011), meaning that there might be room for improving acidophilic microalgae cultures productivities. Moreover, such values are comparable to those of microalgae that are commonly used for biomass production. These results suggest that the efficient use of CO2 by C. onubensis significantly increases when cells experience low inorganic carbon availability during a certain time period and that the action of carbon concentrating mechanisms might be efficiently applied to obtain increased yields of biomass production of acidophilic/acid-tolerant microalgae, which are usually considered as low-growth microalgae (Gross 2000; Pulz and Gross 2004). At once, results would suggest that carbon concentrating mechanisms of C. onubensis (i) are overactive in air conditions, as expected from carbonic anhydrase (CA) data, and (ii) that such highly active situation might remain for longer time during the subsequent microalgal incubation under HC. Excess carbon might be more efficiently converted into biomass of acidophilic microalgae if carbon concentrating mechanisms are highly active, which can be achieved by preincubation under LC conditions. These results allow us to design mass production strategies of acidophilic microalgae, where quite elevated algal biomass productivities might be achieved, obtaining a higher CO2-use efficiency.


The higher productivities of air-pre-incubated C. onubensis cultures are consistent with an increased carbon demand. Longer air pre-incubation (LC) times resulted in increased carbonic anhydrase activities (Fig. 4), considered as total carbon demanding activity. This is in good agreement with results reported by other authors (Gehl et al. 1990; S€ ultemeyer et al. 1991; Lane and Morel 2000) and just emphasizes that carbon acquisition mechanisms of microalgae, which are expressed for balancing the internal inorganic carbon pool, are up-regulated in extreme carbon-limiting situations (Moroney and Somanchi 1999; Miyachi et al. 2003; Moroney et al. 2011), as particularly happens in acidic environments. According to the obtained results, the observed spike of LC cultures to HC cultures total CA activity ratio at 3 h seems to be particularly driven by an increase in the internal CA, therefore increasing carbon demand of the microalga and allowing for higher biomass productivities. The existence of external carbonic anhydrase activity in C. onubensis is a significant result considering that C. onubensis lives naturally in a very low pH environment (below pH 3). Under highly acidic conditions, an extracellular carbonic anhydrase might in principle be not necessary (Gross 2000) as CO2, and not HCO3, is the available carbon source. As an example of this, according to Geib et al. (1996), Dunaliella acidophila showed very low extracellular carbonic anhydrase activity compared to Dunaliella species living at neutral pH. In contrast, acidtolerant algae, like Chlorella saccharophila, seem to have an external CCM (Raven et al. 1982). Other two acidtolerant microalgae, Coccomyxa and Chlamydomonas spp., were found to express external carbonic anhydrase when grown in acidic media, but above pH 5 (Balkos and Colman 2007; Verma et al. 2009). Low CO2 conditions should therefore enhance carbon uptake as appears to happen in other microalgae (Miyachi et al. 2003), although the specific function of external CA at low pH in Coccomyxa is still unclear. As described for Verma et al. (2009), the function of external CA at low pH might be aimed at maintaining the equilibrium of CO2 concentration at the outside part of the cell membrane, therefore speeding CO2 uptake. Low CO2 conditions seem to undergo some changes in internal CA activity in C. onubensis, as well as in external CA. This is in good agreement with results reported for other nonextremophile microalgae such as C. reinhardtii (S€ ultemeyer et al. 1991), Phaeodactylum tricornutum (Nimer et al. 1997), Chlorella saccharophila (Gehl et al. 1990), Skeeltonema costatum (Korb et al. 1997) and Thalassiosira weissflogii (Nimer et al. 1997; Lane and Morel 2000). Gardner et al. (2012) refer that when algae, especially green algae, are transferred from high CO2 conditions (1–5%, v/v) to low CO2 conditions (atmospheric,


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0035%, v/v), a number of carbonic anhydrases and bicarbonate-specific transporters are synthesized within a short time period (up to 6 h). It is noteworthy that even though the CA enzyme activity of C. onubensis appears to reach its maximum value just shortly after (hours) incubation under LC conditions, CA seems to maintain this peak of activity for longer. As explained, the consequence is an increase in biomass productivity when C. onubensis is transferred to HC conditions due to higher carbon uptake efficiency and a subsequent more efficient RuBisCO activity, a consequence of the higher internal CA activity (Palmqvist et al. 1995). In order to know whether the observed external carbon concentrating activity was really due to the external carbonic anhydrase, experiments were performed with Coccomyxa cultures added with an external CA inhibitor, acetazolamide (AZA). CA activity of AZA-added cultures always resulted in values lower than 003 WAU g1 dry weight. These results suggest the existence of external carbonic anhydrase activity of C. onubensis. However, CA values obtained for C. onubensis are lower than external CA activities of ‘common’ microalgae (S€ ultemeyer et al. 1991, 1995; Badger and Price 1994; Li et al. 2012). This was predictable, considering that C. onubensis was cultured in low pH and that the inorganic carbon source was mainly supplied in the form of CO2 and not as bicarbonate. Gene expression analysis is required to confirm the more intense carbon concentrating activity in C. onubensis cell cultures grown in air, and the enhanced biomass productivity obtained during a given time period after the cultures were transferred to CO2-enriched air. In that respect, several authors proved differences in the expression level of carbonic anhydrase genes in microalgae cultures subjected to changes in the CO2 levels. There is abundant literature on the expression of carbonic anhydrase of microalgae. In particular, Chlamydomonas has been widely used as model alga for gene expression studies regarding carbon acquisition in microalgae. Several reports show increased CA expression in algal cells exposed to air and show elimination of the CA transcript in a period of time of hours once the cultures were transferred from air to a high CO2 environment (Bailly and Coleman 1988; Rawat and Moroney 1995). These experiments might be in good agreement with the finding of an enhanced biomass productivity of acidophilic microalgae, more active in carbon acquisition at acid pH, grown in air and transfer to high CO2 levels. Coccomyxa onubensis accumulates high concentrations of lutein, a well-known carotenoid in the group of xanthophylls (Garbayo et al. 2012; Vaquero et al. 2012). Lutein has recently gained attention as an additive in food industry and especially as a powerful antioxidant, its value being recognized against oxidative diseases such as 848

preventing age-related macular degeneration (Ziegler et al. 1996; Carpentier et al. 2009). Average intracellular lutein content of C. onubensis is around 6 mg g1 dry weight, which is within the range of the most promising lutein-producing microalgae species (Fernandez-Sevilla et al. 2010). Calculations from the results of maximal biomass productivity and lutein concentration of C. onubensis shown in this manuscript suggest that it should be technically feasible to achieve lutein productivities up to about 10 mg l1 d1, which are among the highest obtained in microalgae (Fernandez-Sevilla et al. 2010). This model for the production of acid-tolerant microalgae is now being tested at larger scale and should in principle have the added value, to our knowledge, of using a highly selective culture media due to the very low pH, which allows robust biomass production outdoor, and the large lutein abundance of that microalga, accounting for up to 80% of total carotenoid content (Garbayo et al. 2012). From the obtained results, it might be concluded that incubation of acid-tolerant microalgae cultures (C. onubensis in this manuscript) in air followed by shift to high carbon conditions by far enhances carbon-use efficiency in terms of growth rate and biomass productivity, based on the action of both external and internal carbonic anhydrase activities. Lutein content of C. onubensis is high and does not seem to depend on carbon level supplied to cultures. Consequently, repeated cycles of air incubation and high CO2 incubation of C. onubensis might become a suitable tool to perform production processes of lutein-enriched biomass. These results should allow us to design mass production strategies of acidophilic microalgae, particularly lutein-rich biomass in the case of C. onubensis. Acknowledgements This work has been supported by Grant AGR-4337 (Proyecto de Excelencia, Junta de Andalucıa). Conflict of interest No conflict of interest declared. References Badger, M.R. and Price, G.C. (1994) The role of carbonic anhydrase in photosynthesis. Ann Rev Plant Physiol Plant Mol Biol 45, 369–392. Bailly, J. and Coleman, J.R. (1988) Effect of CO(2) concentration on protein biosynthesis and carbonic anhydrase expression in Chlamydomonas reinhardtii. Plant Physiol 87, 833–840.


I. Vaquero et al.

Balkos, K.D. and Colman, B. (2007) Mechanism of CO2 acquisition in an acid-tolerant Chlamydomonas. Plant Cell Environ 30, 745–752. Beamud, S.G., Diaz, M.M. and Pedrozo, F.L. (2010) Nutrient limitation of phytoplankton in a naturally acidic lake (Lake Caviahue, Argentina). Limnology 11, 103–113. Bradford, M.M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72, 248–254. Carpentier, S., Knaus, M. and Suh, M. (2009) Associations between lutein, zeaxanthin, and age-related macular degeneration: an overview. Crit Rev Food Sci Nutr 49, 313–326. Colman, B. and Balkos, K.D. (2005) Mechanisms of inorganic carbon acquisition by Euglena species. Can J Bot 83, 865– 871. Cuaresma, M., Garbayo, I., Vega, J.M. and Vilchez, C. (2006) Growth and photosynthetic utilization of inorganic carbon of the microalga Chlamydomonas acidophila isolated from Tinto river. Enzyme Microb Technol 40, 158–162. Cuaresma, M., Janssen, M., Vılchez, C. and Wijffels, R.H. (2011) Horizontal or vertical photobioreactors? How to improve microalgae photosynthetic efficiency. Bioresour Technol 102, 5129–5137. Fernandez-Sevilla, J.M., Acien Fernandez, F.G. and Molina Grima, E. (2010) Biotechnological production of lutein and its applications. Appl Microbiol Biotechnol 86, 27–40. Ferris, F.G., Hallbeck, L., Kennedy, C.B. and Pedersen, K. (2004) Geochemistry of acidic Rio Tinto headwaters and role of bacteria in solid phase metal partitioning. Chem Geol 212, 291–300. Garbayo, I., Torronteras, R., Forjan, E., Cuaresma, M., Casal, C., Mogedas, B., Ruiz- Domınguez, M.C., Marquez, M. et al. (2012) Identification and physiological aspects of a novel carotenoid-enriched metal resistant microalga isolated from an acidic river in Huelva (Spain). J Phycol 47, 607–614. Gardner, R.D., Cooksey, K.E., Mus, F., Macur, R., Moll, K., Eustance, E., Carlson, R.P., Gerlach, R. et al. (2012) Use of sodium bicarbonate to stimulate triacylglycerol accumulation in the chlorophyte Scenedesmus sp and the diatom Phaeodactylum tricornutum. J Appl Phycol 24, 1311–1320. Gehl, K.A., Colman, B. and Sposato, L.M. (1990) Mechanism of inorganic carbon uptake in Chlorella saccharophila: lack of involvement of carbonic anhydrase. J Exp Bot 41, 1385–1391. Geib, K., Golldack, D. and Gimmler, H. (1996) Is there a requirement for an external carbonic anhydrase in the extremely acid-resistant green alga Dunaliella acidophila? Euro J Phycol 31, 273–284. Gimmler, H. and Weis, U. (1992) Dunaliella acidophila- life at pH 1.0. In: Dunaliella: Physiology, Biochemistry and Biotechnology ed. Avron, M. and Ben-Amonz, A. pp. 99–134. Boca Raton, Florid: CRC Press.


Giordano, M., Beardall, J. and Raven, J.A. (2005) CO2 concentrating mechanisms in algae: mechanisms, environmental modulation, and evolution. Annu Rev Plant Biol 56, 99–131. Gross, W. (2000) Ecophysiology of algae living in highly acidic environments. Hydrobiologia 433, 31–37. Henry, R.P. (1996) Multiple roles of carbonic anhydrase in cellular transport and metabolism. Annu Rev Physiol 58, 523–538. Hewett-Emmett, D. and Tashian, R.E. (1996) Functional diversity, conservation, and convergence in the evolution of the a-, b-, and c-carbonic anhydrase gene families. Mol Phylogenet Evol 5, 50–77. Hutner, S.H., Provosoli, L., Schatz, A. and Haskins, C.P. (1950) Some approaches to the study of the role of metals in the metabolism of microorganisms. Proc Am Philos Soc 94, 152–170. Jordan, D.B. and Ogren, W.L. (1981) Species variation in the specificity of ribulose bisphosphate carboxylase/oxygenase. Nature 291, 513–515. Khalifah, R.G. (1971) The carbon dioxide hydration activity of carbonic anhydrase, Stop-flow kinetic studies on the native human isoenzymes B and C. J Biol Chem 246, 2561–2573. Korb, R.E., Saville, P.J., Jonhston, A.M. and Raven, J.A. (1997) Sources of inorganic carbon for photosynthesis by three species of marine diatom. J Phycol 33, 433–440. Lane, T.W. and Morel, F.M.M. (2000) Regulation of carbonic anhydrase expression by zinc, cobalt, and carbon dioxide in the marine diatom Thalassiosira weissflogii. Plant Physiol 123, 345–352. Li, L., Fu, M., Zhao, Y. and Zhu, Y. (2012) Characterization of carbonic anhydrase II from Chlorella vulgaris in bio-CO2 capture. Environ Sci Pollut Res 19, 4227–4232. Maberly, S.C., Ball, L., Raven, J.A. and S€ ultemeyer, D. (2009) Inorganic carbon acquisition by chrysophytes. J Phycol 45, 1052–1061. Markelova, A.G., Sinetova, M.P., Kupriyanova, E.V. and Pronina, N.A. (2009) Distribution and functional role of carbonic anhydrase Cah3 associated with thylakoid membranes in the chloroplast and pyrenoid of Chlamydomonas reinhardtii. Russ J Plant Physiol 56, 761–768. Messerli, M.A., Amaral-Zettler, L.A., Zettler, E., Jung, S.K., Smith, P.J.S. and Sogin, M.L. (2005) Life at acidic pH imposes an increased energetic cost for a eukaryotic acidophile. J Exp Biol 208, 2569–2579. Miyachi, S., Iwasaki, I. and Shiraiwa, Y. (2003) Historical perspective on microalgal and cyanobacterial acclimation to low- and extremely high-CO2 conditions. Photosynth Res 77, 139–153. Morita, E., Abe, T., Tsuzuki, M., Fujiwara, S., Sato, N., Hirata, A., Sonoike, K. and Nozaki, H. (1998) Presence of the CO2-concentrating mechanism in some species of the


849


I. Vaquero et al.

pyrenoid-less free-living algal genus Chloromonas (Volvocales, Chlorophyta). Planta 204, 269–276. Moroney, J.V. and Somanchi, A. (1999) How do algae concentrate CO2 to increase the efficiency of photosynthetic carbon fixation? Plant Physiol 19, 9–16. Moroney, J.V., Ma, Y., Frey, W.D., Fusilier, K.A., Pham, T.T., Simms, T.A., DiMario, R.J., Yang, J. et al. (2011) The carbonic anhydrase isoforms of Chlamydomonas reinhardtii: intracellular location, expression, and physiological roles. Photosynth Res 109, 133–149. Nimer, N.A., Iglesias-Rodriguez, M.D. and Merreet, M.J. (1997) Bicarbonate utilization by marine phytoplankton species. J Phycol 33, 625–631. Palmqvist, K., S€ ultemeyer, D., Baldet, P., Andrews, T.J. and Badger, M.R. (1995) Characterisation of inorganic carbon fluxes, carbonic anhydrase(s) and ribulose-1,5-biphosphate carboxylate-oxygenase in the green unicellular alga Coccomyxa: comparison with low-CO2 cells of Chlamydomonas reinhardtii. Planta 197, 352–361. Papazi, A., Makridis, P., Divanach, P. and Kotzabasis, K. (2008) Bioenergetic changes in the microalgal photosynthetic apparatus by extremely high CO2 concentrations induce an intense biomass production. Physiol Plant 132, 338–349. Price, G.D., Badger, M.R., Woodger, F.J. and Long, B.M. (2008) Advances in understanding the cyanobacterial CO2-concentrating-mechanism (CCM): functional components, Ci transporters, diversity, genetic regulation and prospects for engineering into plants. J Exp Bot 59, 1441–1461. Pulz, O. and Gross, W. (2004) Valuable products from biotechnology of microalgae. Appl Microbiol Biotechnol 65, 635–648. Raven, J.A. (2010) Inorganic carbon acquisition by eukaryotic algae: four current questions. Photosynth Res 106, 123–134. Raven, J.A., Beardall, J. and Johnston, A.M. (1982) Inorganic carbon transport in relation to HC transport at the plasmalemma of photosynthetic cells. In Plasmalemma and Tonoplast: Their Functions in the Plant Cell ed. Marme, D., Marre, E. and Hertel, R., pp. 41–47. Amsterdam: Elsevier Biomedical Press. Rawat, M. and Moroney, J.V. (1995) The regulation of carbonic anhydrase and ribulose-1,5-bisphosphate carboxylase/oxygenase activase by light and CO2 in Chlamydomonas reinhardtii. Plant Physiol 109, 937–944. Schreiber, U., Hormann, H., Asada, K. and Neubauer, C. (1995) O2-dependent electron flow in spinach chloroplasts: Properties and possible regulation of the Mehler-ascorbate peroxidase cycle. In Photosynthesis: from Light to Biosphere, vol II ed. Mathis, P., pp. 813–818. Dordrecht: Kluwer Academic Publishers. Seckbach, J. and Oren, A. (2007) Oxygenic photosynthetic microorganisms in extreme environments. In Algae and

850

Cyanobacteria in Extreme Environments ed. Seckbach, J. pp. 3–25. The Netherlands: Springer. Silverman, M.P. and Lundgren, D.G. (1959) Studies on the chemoautotrophic iron bacterium Ferrobacillus ferrooxidans. J Bacteriol 77, 642–647. Spalding, M.H. (1998) CO2 Acquisition. In Acclimation to Changing Carbon Availability, in the Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas ed. Rochaix, J.D., Goldschmidt-Clermont, M. and Merchant, S. pp. 529–547. Dordrecht: Kluwer Academic Publishers. Spijkerman, E., de Castro, F. and Gaedke, U. (2011) Independent colimitation for carbon dioxide and inorganic phosphorus. PLoS ONE 6, e28219. Spreitzer, R.J. (1999) Questions about the complexity of chloroplast ribulose-1,5-bisphosphate carboxylase/ oxygenase. Photosynth Res 60, 29–42. S€ ultemeyer, D., Miller, A.G., Fock, H.P. and Canvin, D.T. (1991) Characterization of the active CO2 uptake mechanism by low CO2 grown Chlamydomonas reinhardtii. Can J Bot 69, 995–1002. S€ ultemeyer, D.F., Amoroso, G. and Fock, H. (1995) Induction of intracellular carbonic anhydrases during the adaptation to low inorganic carbon concentrations in wild- type and ca-1 mutant cells of Chlamydomonas reinhardtii. Planta 196, 217–224. Vaquero, I., Ruiz-Dominguez, M.C., Marquez, M. and Vilchez, C. (2012) Cu-mediated biomass productivity enhancement and lutein enrichment of the novel microalga Coccomyxa onubensis. Process Biochem 47, 694–700. Verma, V., Bhatti, S., Huss, V.R. and Colman, B. (2009) Photosynthetic inorganic carbon acquisition in an acidtolerant, free-living species of Coccomyxa (chlorophyta). J Phycol 45, 847–854. Visviki, I. and Palladino, J. (2001) Growth and cytology of Chlamydomonas acidophila under acidic stress. Bull Environ Contam Toxicol 66, 623–630. Wilbur, K.M. and Anderson, N.G. (1948) Electrometric and colorimetric determination of carbonic anhydrase. J Biol Chem 176, 147–154. Williams, T.G. and Colman, B. (1993) Identification of distinct internal and external isozymes of carbonic anhydrase in Chlorella saccharophila. Plant Physiol 103, 943–948. Yamano, T. and Fukuzawa, H. (2009) Carbon-concentrating mechanism in a green alga, Chlamydomonas reinhardtii, revealed by transcriptome analyses. J Basic Microbiol 49, 42–51. Young, A., Orset, S. and Tsavalos, A. (1997) Method for carotenoids analysis. In Handbook of Photosynthesis ed. Pessarakli, M. pp. 11:597–622. New York: Marcel Dekker. Ziegler, R.G., Colavito, E.A., Hartge, P., McAdams, M.J., Schoenberg, J.B., Mason, T.J. and Fraumeni, J.F. (1996) Importance of a-carotene, b-carotene, and other phytochemicals in the etiology of lung cancer. J Natl Cancer Inst 88, 612–615.


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