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1 Institute of Plant Physiology, Russian Academy of Sciences, Botanicheskaya Street 35, Moscow, 127276 Russia
2 Umeå Plant Science Centre, Department of Plant Physiology, University of Umeå, S-901 87 Umeå, Sweden
3 Department of Biology, Universidad Autonoma de Madrid, 28049 Madrid
4 Institute of Microbiology, Russian Academy of Sciences, Prospect 60-Letiya Oktyabrya 7/2, Moscow, 117312 Russia
Correspondence
Natalia Pronina
pronina{at}ippras.ru
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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and 
classes of CA. Immunocytochemical analysis revealed putative -CA localized in the glycocalyx. This -CA has a molecular mass of about 34 kDa and a pI of 3.5. External CAs showed two peaks of activity at around pH 10 and 7.5. The possible involvement of extracellular CAs of M. chthonoplastes in photosynthetic assimilation of inorganic carbon and its relationship to CaCO3 deposition during mineralization of cyanobacterial cells are discussed.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). At the present point in geological time, stromatolite formation is rather insignificant, and the mechanisms of their formation are poorly known. It is still unclear whether mineralization occurs due to some metabolic processes in cyanobacteria, or whether the cyanobacteria just serve as a matrix for binding and structuring of carbonates in the natural process of calcium precipitation.
Carbonic anhydrase (CA, EC 4.2.1.1) is a zinc-containing enzyme catalysing a reversible hydration of carbon dioxide: 
. CA operates with forms of inorganic carbon (Ci), including bicarbonate, which participate in calcium precipitation in nature. CAs have been found in all groups of living organisms. According to the accepted classification, CAs are divided into three main classes, , and 
, which have no significant primary sequence identity and, supposedly, are evolutionarily independent (Smith & Ferry, 2000). The existence of additional 
and 
classes of CAs have also been reported (Tripp et al., 2001; So et al., 2004).
Some eukaryotic algae are able to precipitate intracellular calcium carbonate due to the activity of intracellular CAs (Quiroga & Gonzalez, 1993). In contrast, cyanobacteria can deposit calcium only outside the cells, and such precipitation is strictly controlled by pH (Zavarzin, 2002). It is likely that extracellular CAs of cyanobacterial cells might stabilize the pericellular pH and participate in cell mineralization. However, the cyanobacterial CAs of benthic communities have not been investigated so far.
It is well known that cyanobacterial communities are rather conserved and have not changed significantly during the past two billion years, in terms of their physiology and morphology. Thus, today's microbial communities can be used as a model system for the study of ancient mineralization. This is especially true for so-called ‘relict’ communities that are considered to be the analogues of ancient ecosystems. At present, such communities exist in extreme environments where no traces of higher organisms have been detected (Sergeev et al., 2002).
We studied CAs of the benthic cyanobacterium Microcoleus chthonoplastes. It dominates in alkaliphilic and halophilic cyanobacterial mats worldwide (Gerasimenko et al., 2003). The formation of calcium minerals on the outer mucous cover (glycocalyx) of this cyanobacterium has been reported both in natural samples in the mats (Gerasimenko et al., 2003) and also in laboratory culture (Zavarzin et al., 2003). We have identified extracellular CAs of M. chthonoplastes with access to outer Ci substrates and indicated their possible participation in extracellular calcium carbonate precipitation.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The organism was grown in medium S (Castenholz & Waterbury, 1989) (g l–1: KCl, 1.0; NaHCO3, 16.8; K2HPO4, 0.5; NaNO3, 2.5; K2SO4, 1.0; NaCl, 30; MgSO4, 0.1; CaCl2, 0.04; FeSO4, 0.01; EDTA, 0.08, pH 9.8) and micronutrients (Rippka et al., 1979) in batch cultures at 31 °C under continuous irradiance of 50 µmol m–2 s–1 from cool white fluorescent lamps. All experiments were carried out with biomass after 4–5 days cultivation, corresponding to the mid-exponential phase of growth.
Isolation of cell envelopes and their fractions.
Cells were broken in cooled buffer (30 mM HEPES-KOH, pH 8.2) using a French press (5x107 Pa). Unbroken cells were sedimented by centrifugation at 700 g for 10 min at 4 °C. Cell envelopes of M. chthonoplastes were isolated by centrifugation of the cell homogenate at 12 000 g according to Weckesser & Jürgens (1988). To remove chlorophyll and soluble protein contaminations, the cell envelope fraction was washed four to five times with 30 mM HEPES-KOH (pH 8.2).
The isolation of cell walls and other components of the envelope was performed as described by Weckesser & Jürgens (1988). The cell envelope fraction was loaded on a discontinuous sucrose gradient (60, 55, 50, 45 and 40 % sucrose in 30 mM HEPES-KOH buffer, pH 8.2) and centrifuged at 20 000 r.p.m. in a Beckman SW 41 rotor for 4 h at 4 °C. As a result, three main fractions were obtained: (1) two lower fractions of cell walls (at the bottom of the tube and at the 55/60 % sucrose interface); (2) a fraction of some cell envelope fragments at the 50/55 % sucrose interface (supposedly glycocalyx); and (3) a lighter fraction of plasma membranes located above 40 % sucrose. All these fractions were collected, and the two cell-wall fractions were combined. Traces of sucrose were removed by washing in 30 mM HEPES-KOH buffer with centrifugation at 176 000 g for 1 h at 4 °C. Cell walls were further purified as described by Weckesser & Jürgens (1988). During all isolation steps, protease inhibitor mix (Sigma) was added to avoid protein degradation.
Assay of CA activity.
CA activity was measured electrometrically (Wilbur & Anderson, 1948) by monitoring the rate of pH change during carbon dioxide hydration using a fast-response ‘blue glass' microelectrode (MI-710; Microelectrodes) and an 18-bit A/D converter (IOtech). Measurements were performed with intact cells, cell homogenate and cell envelopes in 30 mM HEPES-KOH buffer, pH 8.2. The reaction was carried out at 2 °C and started by a rapid injection of saturated CO2 solution into an equal volume of the sample (fermentative reaction) or buffer (non-fermentative reaction, control). The reaction kinetics were recorded for 100 s. CA activity was calculated as the difference in the initial rate of CO2 hydration between control and samples, and expressed in Wilbur–Anderson units (WAU) per 1 mg chlorophyll (Chl) or protein. One WAU is defined as 10x(t0–t)t–1, where t0 and t are the times required for the pH to change by 
pH in the control and the samples, respectively. The measurements were carried out in three to five replicates.
To study the dependence of M. chthonoplastes extracellular CA activity on external pH, the cyanobacterium was cultured at the optimum pH of 9.8 and then transferred to S medium at different pH values (6.5, 7.5, 8.0, 9.0 and 9.8) and grown under similar culture conditions (light and temperature were not changed). CA activity in intact cells was measured after 1.5 h incubation. CA activity was expressed in relative units compared to the activity at the optimal pH of 9.8.
Estimation of chlorophyll and protein content.
Protein content was estimated in accordance with Bio-Rad Laboratories protocols using commercial kits with standard solutions (Bio-Rad DC Protein Assay Kit). Chlorophyll content was determined spectroscopically after extraction with absolute methanol (Porra et al., 1989).
Electrophoresis and immunodetection.
Proteins were separated by 10 or 12 % SDS-PAGE, following Laemmli (1970), or by two-dimensional (2D) electrophoresis. A mix of protein standards (Bio-Rad) was used as molecular mass markers.
For electrophoresis, a standard protocol and Bio-Rad buffers and solutions were used. The lanes were loaded with 15 µg protein for silver-stained gels, and 20 or 45 µg protein for Coomassie-stained gels and immunoblot analysis, respectively. The samples were solubilized for 5 min at 95 °C in the sample buffer.
2D electrophoresis was performed according to the manufacturer's instructions (Amersham). The protein content was 100 µg for immunodetection or 300 µg for silver-stained gels. Each sample was mixed with a rehydration solution containing 8 M urea, 2 % CHAPS, 20 mM DTT, 0.5 % IPG buffer (pH 3–10; Amersham) and 0.001 % bromophenol blue. IPG dry strips (Immobiline DryStrip gels, 7 cm, with a linear pH range coverage of 3–10; Amersham) were allowed to rehydrate in the presence of samples in the IPGphor Isoelectric Focusing System (Amersham), followed by isoelectric focusing of proteins at 20 °C. The following voltage/time profile was used: rehydration for 12 h; increasing voltage by the ‘step-and-hold’ procedure from 500 to 1000 V over 1 h; final phase of 8000 V for 12 h. After the first dimension run, the individual strip was equilibrated for 15 min in SDS buffer (50 mM Tris/HCl, pH 6.8; 6 M urea; 30 %, v/v, glycerol; 2 % SDS) supplemented with 1 % (w/v) DTT, and then for another 15 min in SDS buffer supplemented with 2.5 % (w/v) iodoacetamide. Thereafter, the strips were put on the top of the second dimension gel (12 %) and covered with 0.5 % agarose in SDS buffer. Each gel was run at 10 mA at the beginning and then at 15 mA per gel after all the proteins were transferred from the strip into the gel.
Immunoblotting was performed as described in the Bio-Rad Laboratories protocol. The primary antibodies were raised against: (1) Chlamydomonas reinhardtii -CA (Cah-3) (Karlsson et al., 1998) (affinity-purified); (2) C. reinhardtii mitochondrial -CA (Eriksson et al., 1996); (3) spinach chloroplast -CA (Fawcett et al., 1990); (4) Coccomyxa sp. intracellular -CA (Hiltonen et al., 1998); and (5) C. reinhardtii D1 protein (PsbA) (Nishiyama et al., 2001). Horseradish peroxidase-labelled secondary antibodies (Amersham Life Science) and chemiluminescence solutions (ECL, Amersham) were used to detect an antibody–antigen conjugate.
Immunoelectron microscopy.
For immunogold labelling experiments, M. chthonoplastes cells or the isolated fraction of glycocalyx were fixed in 4 % paraformaldehyde for 4–10 days at 4 °C. Immunocytochemical reactions were performed after washing the sample in 0.1 M phosphate buffer (pH 7.4). The reaction with primary antibodies against C. reinhardtii -CA (Cah-3) was carried out for 1 h at 24 °C and then for 23 h at 4 °C. Thereafter, the samples were washed three times with phosphate buffer over a 24 h period. The second step of the immunochemical reaction and post-washing were performed under similar conditions using Protein-A-Gold (Sigma). The samples were post-fixed in 1 % OsO4, dehydrated in an alcohol series and embedded in Epoxy resin (Sigma). As a control for immunochemical reaction specificity, the sample treatment step with primary antibodies was omitted. Thin sections of samples were prepared using an ultramicrotome and then analysed with a JEM JEOL X-100 transmission electron microscope (Japan) without any additional contrast.
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RESULTS |
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The highest activity of M. chthonoplastes extracellular CA was detected in intact cells incubated at an alkaline pH (
10) which is optimal for the growth of this alkaliphilic cyanobacterium. Enzyme activity was also recorded at a near neutral pH (7.5) (Fig. 1). There was no detectable CA activity in intact cells incubated at pH 6.7 and 8.2.
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-CA-CA (Cah-3) was carried out. These antibodies cross-reacted with only one specific protein of approximately 34 kDa (Fig. 2b). The protein cross-reacting with the Cah3 antibodies was clearly enriched in the cell envelope fraction when compared to the cell homogenate (Fig. 2b). This enrichment corresponded with the higher enzyme activity seen in the cell envelope fraction (see above).
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; Villarejo et al., 2002), it was necessary to be sure that the signal observed in the cell envelope fraction of M. chthonoplastes did not result from cross-contamination of this fraction with thylakoid membranes from the cyanobacterium. Antibodies against D1 (PsbA), a major protein of PS II, was used as a specific marker for thylakoid membranes (Nishiyama et al., 2001). Fig. 2(c)
shows that cell envelopes did not cross-react with these antibodies, which implies the absence of thylakoid membrane contamination in this fraction.
Immunocytochemical detection of -CA
Localization of -CA in M. chthonoplastes cells was also investigated by immunogold electron microscopy using antibodies against C. reinhardtii -CA (Cah-3) (Fig. 3). The electron-impermeable spots of colloidal gold are clearly visible in the glycocalyx (Fig. 3b). This structure and analogous specific signals are also typical of the fragments located at the interface between 50 and 55 % sucrose during cell envelope fractionation (Fig. 3d). Thus, one can conclude that this fraction consists of glycocalyx.
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). Our attempts to detect CA activity in different fractions obtained after further fractionation of the cell envelopes from M. chthonoplastes failed. Apparently, in spite of the presence of protease inhibitors, enzyme activity decreased drastically during fractionation. However, the presence of a potential -CA in these fractions was tested by Western blot analysis with antibodies against C. reinhardtii -CA (Cah-3). Fig. 4 shows that the signal was specifically enriched in a fraction characterized as glycocalyx, but there was no signal in the cell wall or plasma membrane fractions. It thus appears that the specific signal observed in the cell envelopes (Fig. 2b) was due to the presence of a Cah-3-like enzyme in the M. chthonoplastes glycocalyx.
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-CA (Cah-3) enabled us to estimate the protein composition of the M. chthonoplastes glycocalyx fraction as well as the isoelectric point of extracellular -CA (Fig. 5). The analysis of the silver-stained 2D gel showed the presence of approximately 15–20 various polypeptides in the M. chthonoplastes glycocalyx. Practically all the proteins in this fraction were acidic, with pI values ranging from 3 to 5, as has been reported in other cyanobacteria (Huang et al., 2004). However, the most abundant protein of this fraction was basic, with an estimated pI value of 9.5 and a approximate molecular mass of 18 kDa.
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Identification of M. chthonoplastes extracellular -CAs
For identification of -CAs among M. chthonoplastes cell envelope proteins, primary antibodies against -CAs from various organisms were used, i.e. spinach chloroplasts and C. reinhardtii mitochondria, as well as antibodies against intracellular -CA from the photobiont microalga Coccomyxa sp. This decision to use several types of antibody was determined by the multiplicity of the -CA isoforms.
The cross-reaction between the M. chthonoplastes cell envelope polypeptides and the antibodies against spinach and C. reinhardtii -CAs revealed the presence of several specific signals (Fig. 6). These signals were much stronger than those of the cell homogenate. The molecular masses of polypeptides detected by different antibodies were not identical. No cross-reaction was detected by Western blotting when the antibodies against the intracellular CA of Coccomyxa sp. was probed (not shown).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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- or -classes.
Until now, the presence of external -CA (possibly periplasmic) had been shown in only two species of cyanobacteria, Anabaena sp. PCC 7120 and Synechococcus sp. PCC 7942. However, enzyme activity of the identified -CA (EcaA) was not detected either in whole-cell lysates or in cell fractions of these cyanobacteria (Soltes-Rak et al., 1997). It was postulated that the cause of this failure could be due to the low sensitivity of the electrometrical method used for CA activity measurement. Extracellular localization in the periplasmic space was also assumed for EcaB (-CA) of Synechocystis PCC 6803 (So et al., 1998).
The levels of M. chthonoplastes extracellular CA activity detected are comparable with enzyme activities of some other prokaryotic CAs, for example the intracellular enzyme activity of Anabaena variabilis (Yagawa et al., 1984), as well as with the activities of CAs from microalgal chloroplasts (Katzman et al., 1994). In addition to a potential -class enzyme, at least two putative -CAs were found in the cell envelopes of M. chthonoplastes: one of them is similar to chloroplastic -CAs and the other to mitochondrial -CAs (Fig. 6). Numerous other specific signals probably resulted from incomplete denaturation of native -CAs consisting of several subunits (Smith & Ferry, 2000). The exact topology of these extracellular -CAs within cell envelopes of M. chthonoplastes is still not clear.
Studies of the CAs of alkaliphilic cyanobacteria from soda lakes are important with respect to evolutional theory, because these organisms are supposed to be relicts of ancient microbiota (Sergeev et al., 2002). Genome analysis of many prokaryotic organisms has revealed the prevalence of - and -CAs, which have been proposed to be the most ancient classes of CAs (Smith & Ferry, 2000). It has been assumed that -CAs evolved from a common ancestral gene about 0.5–0.6 billion years ago (Smith & Ferry, 2000). This is consistent with the evidence that only a few prokaryotic genomes encode -class enzymes. However, -CAs have been found in some bacteria (Nafi et al., 1990; Chirica et al., 1997) and cyanobacteria (Soltes-Rak et al., 1997; Dudoladova et al., 2004) where they have been characterized as extracellular enzymes, similar to the -CA of M. chthonoplastes. Thus, the presence of this class of CAs in relict organisms, as well as in other bacteria, indicates that -CAs are as ancient as - and -CAs.
The participation of CAs in the autotrophic assimilation of Ci by cyanobacteria is well known. It is supposed that the function of extracellular forms of the enzyme (-EcaA and -EcaB) is associated with Ci transport from the medium into the cell by CO2 and/or 
substrate formation for carbon transporters located in the plasma membrane. For intracellular -CAs (IcfA and CcaA), which are located in the carboxysome, a role in CO2 generation is presumed in the place of the fixing of CO2 by Rubisco (Smith & Ferry, 2000).
A general scheme of extracellular CA participation in Ci assimilation of M. chthonoplastes is presented in Fig. 7. In soda lakes, where the pH is highly alkaline (near 10), all dissolved Ci is present as carbonate and bicarbonate ions. In cyanobacterial cells, the existence of several 
transport systems has been demonstrated (Badger & Price, 2003). The majority of bicarbonate ions absorbed by the cell enter the carboxysome where they are converted to CO2 by intracellular -CAs, which is then fixed by Rubisco. Some of the absorbed 
can also be converted into CO2 before being fixed by photosynthesis in accordance with the Henderson–Hasselbalch equation because of the lower intracellular pH compared to that of the outer medium. It has been shown previously that the cytoplasmic pH of alkaliphilic cyanobacteria is neutral (Kupriyanova et al., 2003). According to the concentration gradient, there must be some leakage of the CO2 produced in the intracellular space out of the cell. However, - and -CAs, which are located in the cell envelope of cyanobacteria, might prevent this leakage of CO2 by converting it to 
. This is then transported through the membrane back into the cell by carbon transporters. This is in accordance with high enzyme activity of extracellular CA observed under alkaline pH conditions (Fig. 1) typical of the natural habitat of M. chthonoplastes in soda lakes.
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) reflects the ability of M. chthonoplastes to live in the seashore environment with a pH close to this value. The mineralization of cyanobacteria usually occurs under such conditions because seawater has a large reserve of Ca2+ ions (Zavarzin et al., 2003). Stromatolite formation does not occur in soda lakes because calcium ions almost disappear from the medium at high pH, having been precipitated as CaCO3.
The mechanisms of bicarbonate uptake discovered in cyanobacterial cells are associated solely with the alkalinization of the pericellular space. Bicarbonate assimilation could be accompanied by hydroxyl excretion into the outer medium (McConnaughey, 1994) and/or by sodium symport (Badger & Price, 2003). The latter mechanism leads to alkalinization of cytoplasm followed by activation of Na+-ATPase that assists Na+ exchange for H+ from the outer medium (Balnokin et al., 2004). These mechanisms are shown schematically in Fig. 7.
When the pH of the pericellular layers rises to 9.0, calcium carbonate granules are produced in the cyanobacterial glycocalyx if free Ca2+ is available in the medium. The glycocalyx facilitates mineralization because it helps in the formation of a physicochemical/chemical microgradient by decreasing the rate of diffusion flow, and it binds divalent cations, including Ca2+, from the external medium due to the presence of polysaccharide carboxyl groups (Arp et al., 2001). Thus, photosynthetic assimilation of Ci by M. chthonoplastes cells appears to be accompanied by their mineralization under pH conditions found in seawater. CAs possibly stabilize pH in the pericellular space and maintain the substrate (
) concentration essential for both photosynthesis and CaCO3 deposition.
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ACKNOWLEDGEMENTS |
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Edited by: K. Forchhammer
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Received 30 October 2006;
revised 6 January 2007;
accepted 11 January 2007.
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