doi: 10.1371/journal.pbio.1001085. Epub 2011 Jun 21.
A voltage-gated H+ channel underlying pH homeostasis in calcifying coccolithophores
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- PMID:21713028
- PMCID: PMC3119654
- DOI: 10.1371/journal.pbio.1001085
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A voltage-gated H+ channel underlying pH homeostasis in calcifying coccolithophores
Alison R Taylor et al. PLoS Biol.2011 Jun.
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Abstract
Marine coccolithophorid phytoplankton are major producers of biogenic calcite, playing a significant role in the global carbon cycle. Predicting the impacts of ocean acidification on coccolithophore calcification has received much recent attention and requires improved knowledge of cellular calcification mechanisms. Uniquely amongst calcifying organisms, coccolithophores produce calcified scales (coccoliths) in an intracellular compartment and secrete them to the cell surface, requiring large transcellular ionic fluxes to support calcification. In particular, intracellular calcite precipitation using HCO₃⁻ as the substrate generates equimolar quantities of H+ that must be rapidly removed to prevent cytoplasmic acidification. We have used electrophysiological approaches to identify a plasma membrane voltage-gated H+ conductance in Coccolithus pelagicus ssp braarudii with remarkably similar biophysical and functional properties to those found in metazoans. We show that both C. pelagicus and Emiliania huxleyi possess homologues of metazoan H(v)1 H+ channels, which function as voltage-gated H+ channels when expressed in heterologous systems. Homologues of the coccolithophore H+ channels were also identified in a diversity of eukaryotes, suggesting a wide range of cellular roles for the H(v)1 class of proteins. Using single cell imaging, we demonstrate that the coccolithophore H+ conductance mediates rapid H+ efflux and plays an important role in pH homeostasis in calcifying cells. The results demonstrate a novel cellular role for voltage gated H+ channels and provide mechanistic insight into biomineralisation by establishing a direct link between pH homeostasis and calcification. As the coccolithophore H+ conductance is dependent on the trans-membrane H+ electrochemical gradient, this mechanism will be directly impacted by, and may underlie adaptation to, ocean acidification. The presence of this H+ efflux pathway suggests that there is no obligate use of H+ derived from calcification for intracellular CO₂ generation. Furthermore, the presence of H(v)1 class ion channels in a wide range of extant eukaryote groups indicates they evolved in an early common ancestor.
Conflict of interest statement
The authors have declared that no competing interests exist.
Figures
(A) Whole cell currents fromC. pelagicus cells in response to incremental 1 s 10 mV depolarisations from −80 to +60 mV. The recording pipette contained (in mM) K-Glutamate 200, MgCl2 5, EGTA 5, and HEPES 100 (P1a, Table S2). The ASW bathing solution contained (in mM) NaCl 450, KCl 8, MgCl2 30, MgSO4 16, CaCl2 10, NaHCO3 2, and HEPES 20 (E1, Table S2). For clarity only every other trace (Δ+20 mV) is indicated. (B) Corresponding current-voltage relationship showing outward current activation at voltages more positive than EH+ (arrow). (C) Tail currents activated by a 1 s depolarizing pulse to +40 mV before 500 ms test pulses from −75 to +25 mV. The dotted line indicates zero current. Solutions are as in (A). (D) pH gradient (ΔpH = pHo − pHi) dependence of average Erev (± SE,n = 6). Data were fitted by a regression with a slope of −43 mV/pH unit. (E) pHo sensitivity of whole current activation. The pipette solution contained (in mM): MgCl2 5, EGTA 5, TEA-Cl 200, and HEPES 5 (P2, Table S2), and the ASW bath solution was as in (A). (F) Average whole cell currents (± SE) under the same conditions as (E) in pHo 8.0 (n = 10) and 7.0 (n = 4). Outward currents are smaller and activation more positive with decreased pHo (arrows = EH+). (G) Effect of 30 µM free Zn2+ on the outward currents. The internal and external solutions used are the same as in (A) (P1a, E1). (H) Average current voltage curves (± SE) for control (n = 5) and in the presence of 30 µM free Zn2+ (n = 5). Arrow indicates EH+.
Tail current analysis was conducted with a range of pipette solutions with major salt as indicated in the schematic KCl 80 mM (A), KCl 400 mM (B), and K-Glutamate 200 mM (C) and 5 mM HEPES buffer. All other constituents were as P1 (Table S2), which are (in mM) MgCl2 5, EGTA 5. The bath solution was ASW containing (in mM) NaCl 450, KCl 8, MgCl2 30, MgSO4 16, CaCl2 10, NaHCO3 2, and HEPES 20; and adjusted to the pH indicated in each schematic (E1, Table S2). Pipette solutions were adjusted to ∼1,000 mOsmol kg−1 with sorbitol. For each recording condition, outward currents were activated by a depolarising voltage pulse to +110 mV and tail currents subsequently recorded at a range of voltages. The upper panels show representative tail current traces for (A) 80 mM KCl, (B) 400 KCl, and (C) 200 K-glutamate pipette solutions. Dotted lines represent zero current and voltages for representative traces are indicated. The initial activated outward currents are truncated to emphasise the tail currents. Lower plots illustrate average tail current-voltage plots for a minimum of 4 cells for each condition (±SE). Arrows indicate the predicted reversal potential for each of the major ions. The tail current reversal for theC. pelagicus outward currents are consistently close to EH+ regardless of ECl− and EK+, suggesting whole cell currents activated by depolarisation are primarily carried by H+ ions in calcifying cells. The slightly positive reversal indicated may be due in part to the weakly buffered pipette solution used in these experiments or because of a minor contribution by endogenous cation currents carried by Mg2+ Ca2+ or Na+ (predicted equilibrium potentials of +26, +182, and > +300 mV for each ion, respectively).
(A) Multiple sequence alignment of animal and coccolithophore Hv1 proteins. The multiple sequence alignment indicates transmembrane residues conserved between animal and coccolithophore Hv1 proteins. Shading indicates residues that are identical or similar in 80% of the sequences (BLOSUM62 matrix), and boxes represent predicted transmembrane domains in human Hv1. The three arginine residues in S4 required for voltage sensing are conserved (numbers below alignment correspond to R205, R208, and R211 in human Hv1). Many of the acidic residues (aspartate and glutamate) appear conserved (D112, E119, D123, E153, D174), although Asp185 is replaced by glutamate and Glu171 is not conserved. Several of these acidic residues are conserved amongst voltage sensor domain proteins and may function in voltage sensing . In contrast, the other basic residues (lysine) are not conserved in coccolithophores (K125, K157, K169, K221). Polar serine residues (S143, S181) are conserved inE. huxleyi, although Ser181 is replaced by alanine inC. pelagicus. Similar sequences identified in the diatomsP. tricornutum andT. pseudonana are also shown in the alignment. The arrows indicate external histidine residues required for Zn2+ inhibition in human Hv1 and external histidine residues which are conserved in algal Hv1 proteins (the numbers above the alignment indicate their position in EhHv1). (B) Multiple sequence alignment indicating conserved domains in transmembrane regions of voltage sensor proteins. Hv1 proteins from animals (Hs,Homo sapiens; Dr,Danio rerio; Ci,Ciona intestinalis) and coccolithophores (Eh,Emiliania huxleyi; Cp,Coccolithus pelagicus) are aligned to transmembrane regions S1–S4 from K+ channels from humans,Drosophila, plants, and prokaryotes and also to the prokaryote Na+ channel NaChBac. Shading indicates identical residues in 5 out of 10 sequences displayed. In this analysis the only conserved residues that are exclusive to Hv1 proteins correspond to human Hv1 residues D112, D123, and S143. (C) Phylogenetic analysis of known Hv1 proteins. The tree was generated by a maximum likelihood analysis of available Hv1 protein sequences. 100 bootstraps were performed.
(A) Whole cell currents in HEK293 cells transfected withEhHVCN1,CpHVCN1, andGFP only. The pipette solution contained (in mM) NMDG 65, MgCl2 3, EGTA 1, and HEPES 150 glucose 70 at pH 7.0 (P4, Table S2), and the bath solution contained (in mM) N-methyl D-glucamine (NMDG) 75, MgCl2 3, CaCl2 1, glucose 160, and HEPES 100 at pH 7.8 (E4, Table S2). Cells were depolarised in 10 mV increments from −60 to +100 mV. For clarity, every other trace is shown. (B) Average current-voltage curves (± SE,n = 6) of HEK293 cells transfected withEhHVCN1 (red circles),CpHVCN1 (blue squares), andGFP only (black triangles), respectively, using the protocol described in (A). (C) Tail current analysis of HEK293 cells expressingEhHVCN1. Currents were activated by a 500 ms depolarisation to +80 mV followed by repolarisation to between −50 and +70 mV. Dotted line represents zero current level. The solutions used for patch and bath solutions were the same as described in (A) (E4, P4). (D) pH dependence of EhHv1 Erev. (E) Outward currents of EhHv1-expressing HEK 293 cells activated by depolarising pulses from −60 mV to +100 mV before (upper panel) and after (lower panel) perfusion with pHo 6.5. The pipette solution was as in (A) and the external solutions were as described above except that 100 mM MES replaced 100 mM HEPES at pH 6.5 (E5, Table S2). (F) Average current-voltage curves (± SE,n = 5) for pHo 7.8 or 6.5. Activation voltage shifts with EH+ (arrows). (G) Inhibition of EhHv1 currents by 500 µM Zn2+. The recording pipette contained in mM: NaCl 30, KCL 100, MgCl2 3, EGTA 1, and HEPES 100 at pH 7.0 (P3, Table S2). Cells were bathed with (in mM) NaCl 160, KCL 2.0, MgCl2 1, CaCl2 1, Glucose 20, and HEPES 100 at pH 7.8 (E3, Table S2). (H) Average current voltage curves (± SE,n = 5) for control and in the presence of 500 µM Zn2+.
(A) Representative simultaneous whole cell patch clamp and pHi imaging inC. pelagicus cells (300 µM BCECF free acid loaded via the patch pipette). Top panel displays false colour BCECF fluorescence ratio images ofC. pelagicus during the voltage step protocol (+20, −50, and +70 mV). The inset (top right) indicates localization of BCECF (green), chlorophyll autofluorescence (red), and reflectance of an internal developing coccolith (white). Scale bar, 10 µm. Membrane depolarization to +20 mV or +70 mV from a holding potential of −50 mV caused an increase in pHi. The pipette solution contained (in mM) K-Glutamate 200, MgCl2 5, EGTA 5 and Pipes 1.0 (P1b, Table S2), and the external solution was ASW containing (in mM) NaCl 450, KCl 8, MgCl2 30, MgSO4 16, CaCl2 10, NaHCO3 2, and HEPES 20 (E1, Table S2). (B) The effect of pHo on pHi in BCECF-loadedC. pelagicus cells. Changing pHo from 8.0 to 6.5 induced a substantial and reproducible reduction in pHi. Traces from 4 individual cells are superimposed. (C) The effect of Zn2+ on pHi in calcifyingC. pelagicus cells. Cells loaded with BCECF-AM were perfused with either f/2 FSW media pH 8.0 (n = 62) or artificial seawater (ASW) pH 8.0, containing 0 mM or 10 mM Ca2+ (n = 28 and 7, respectively). 30 µM free Zn2+ was perfused for 2.5 min (grey box); averaged traces are shown.
(A) Calcification rate following manipulation of pHi. Coccolith production by decalcifiedC. pelagicus cells in f/2 FSW media (pH 8.2, 15 °C) was monitored by cross-polarised light microscopy (upper right and Figure S6). Upper left shows a scanning electron micrograph of aC. pelagicus cell with an intact coccosphere for reference. Scale bars, 10 µm. pHi was manipulated by perfusion with f/2 media at pH 8.2 (control), at pH 6.5, or at pH 8.2 + 10 mM NH4Cl for 10 min (lower panel). Note the appreciable lag in restoration of the initial calcification rate following acidification of the cytosol. Traces are normalized to initial rate (0–150 min). (B) Mean calcification rate (±SE) was calculated by linear regression for each period (0–150, 150–300, and 300–600 min) and is expressed as a percentage of the initial rate (n = 4 except for pH 6.5, wheren = 3).
The scheme illustrates the requirement for efficient H+ efflux pathways in coccolithophores as a result of intracellular calcification. Mature coccoliths are arranged on the extracellular surface, surrounding the cell to form a coccosphere (A). However, coccolith formation occurs within the intracellular Golgi-derived coccolith vacuole. Calcium carbonate (CaCO3) precipitation requires the production of carbonate (CO32−) from bicarbonate (HCO3−) and results in the net production of H+ (B). H+ must be rapidly removed from the coccolith vacuole in order to maintain a suitable pH for CaCO3 precipitation. Once in the cytosol (C), some H+ may potentially be utilised by photosynthesis in the production of CO2 from HCO3−, however H+ efflux provides an efficient mechanism to prevent cytosolic acidosis during fluctuations in photosynthetic rate. At normal seawater pH 8.2, pHi of 7.2, and Vm ∼−46 mV maintained by a Cl– inward rectifier (D) , a drop in pHi alone or in combination with membrane depolarisation would result in a net outward proton motive force across the plasma membrane. This enables passive H+ efflux via a plasma membrane localised H+ channel (E), providing a rapid mechanism for maintaining constant intracellular pH. Other membrane transporters (yet to be characterised) are likely involved in longer term maintenance of cytoplasmic pH (F). Nevertheless, the pH- and voltage-sensitive gating mechanism of the H+ channel coupled to its high transport capacity suggests it plays a major role in the modulation of intracellular pH in coccolithophores. Patch clamp studies indicate that Cl– and H+ are the dominant transmembrane conductances in coccolithophores.
Comment in
- Will ion channels help coccolithophores adapt to ocean acidification?Mejia R.Mejia R.PLoS Biol. 2011 Jun;9(6):e1001087. doi: 10.1371/journal.pbio.1001087. Epub 2011 Jun 21.PLoS Biol. 2011.PMID:21713029Free PMC article.No abstract available.
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