BGBiogeosciencesBGBiogeosciences1726-4189Copernicus GmbHGöttingen, Germany10.5194/bg-12-6493-2015Phytoplankton calcification as an effective mechanism to alleviate
cellular calcium poisoningMüllerM. N.mnmuller@usp.brBarcelos e RamosJ.SchulzK. G.https://orcid.org/0000-0002-8481-4639RiebesellU.https://orcid.org/0000-0002-9442-452XKaźmierczakJ.GalloF.MackinderL.LiY.NesterenkoP. N.TrullT. W.HallegraeffG. M.https://orcid.org/0000-0001-8464-7343Institute for Marine and Antarctic Studies (IMAS),
University of Tasmania, Private Bag 129, Hobart, TAS 7001,
AustraliaInstitute of Oceanography, University of São Paulo,
Praça do Oceanográfico 191, 05508-120 São Paulo, SP,
BrazilCentre of Climate, Meteorology and Global Change (CMMG),
University of Azores, Rua do Capitão d'Ávila, Pico da Urze 970-0042
Angra do Heroísmo, Açores, PortugalCentre for Coastal Biogeochemistry, School of
Environmental Science and Management, Southern Cross University, P.O. Box
157, Lismore, NSW 2480, AustraliaGEOMAR Helmholtz Centre for Ocean Research Kiel,
Düsternbrooker Weg 20, 24105 Kiel, GermanyInstitute of Paleobiology, Polish Academy of Sciences,
Twarda 51/55, 00-818 Warsaw, PolandDepartment of Plant Biology, Carnegie Institution, 260
Panama Street, Stanford, CA 94305, USAAustralian Centre for Research on Separation Science
(ACROSS), School of Chemistry, University of Tasmania, Private Bag 75,
Hobart, TAS 7001, AustraliaAntarctic Climate and Ecosystems Cooperative Research
Centre, University of Tasmania and CSIRO Oceans and Atmosphere Flagship,
Hobart, TAS 7001, AustraliaM. N. Müller (mnmuller@usp.br)13November201512216493650114July201511August20153November20154November2015This work is licensed under a Creative Commons Attribution 3.0 Unported License. To view a copy of this license, visit http://creativecommons.org/licenses/by/3.0/This article is available from https://bg.copernicus.org/articles/12/6493/2015/bg-12-6493-2015.htmlThe full text article is available as a PDF file from https://bg.copernicus.org/articles/12/6493/2015/bg-12-6493-2015.pdf
Marine phytoplankton have developed the remarkable ability to tightly
regulate the concentration of free calcium ions in the intracellular cytosol
at a level of ∼ 0.1 µmol L-1 in the presence of
seawater Ca2+ concentrations of 10 mmol L-1. The low cytosolic
calcium ion concentration is of utmost importance for proper cell signalling
function. While the regulatory mechanisms responsible for the tight control
of intracellular Ca2+ concentration are not completely understood,
phytoplankton taxonomic groups appear to have evolved different strategies,
which may affect their ability to cope with changes in seawater Ca2+
concentrations in their environment on geological timescales. For example,
the Cretaceous (145 to 66 Ma), an era known for the high abundance of
coccolithophores and the production of enormous calcium carbonate deposits,
exhibited seawater calcium concentrations up to 4 times present-day
levels. We show that calcifying coccolithophore species (Emiliania huxleyi, Gephyrocapsa oceanica and Coccolithus braarudii) are able
to maintain their relative fitness (in terms of growth rate and
photosynthesis) at simulated Cretaceous seawater calcium concentrations, whereas these rates are severely reduced under these
conditions in some non-calcareous phytoplankton species (Chaetoceros sp., Ceratoneis closterium and
Heterosigma akashiwo). Most notably, this also applies to a non-calcifying strain of E. huxleyi which
displays a calcium sensitivity similar to the non-calcareous species. We
hypothesize that the process of calcification in coccolithophores provides
an efficient mechanism to alleviate cellular calcium poisoning and thereby
offered a potential key evolutionary advantage, responsible for the
proliferation of coccolithophores during times of high seawater calcium
concentrations. The exact function of calcification and the reason behind
the highly ornate physical structures of coccoliths remain elusive.
Introduction
Calcium is a versatile and crucial ion in biological systems (Case et al.,
2007), which is, among other functions, essential for cellular signalling,
membrane structure and cell division (Sanders et al., 1999). The
concentrations of cytosolic free Ca2+ in eukaryotes are well
regulated and the maintenance of relatively low levels is essential for fast
signal transduction. An excessive influx of Ca2+ to the cytosol can
be lethal as it disturbs intracellular signalling and irreversibly damages
the cell (Orrenius et al., 1989; Kader and Lindberg, 2010). Homeostasis of
Ca2+ in plant cells is predominantly achieved by Ca2+-binding
proteins, reducing the effective diffusion coefficient of Ca2+ in the
cytosol, and ultimately via sequestration by the endoplasmic reticulum,
mitochondria and cellular vacuoles (Case et al., 2007). Cytosolic free
Ca2+ concentrations in marine phytoplankton are about 105 times
lower than modern seawater concentrations and marine eukaryotes have
developed a remarkable capacity to maintain these low cytosolic Ca2+ levels (Brownlee et al., 1987, 1995). It is, however,
unknown whether the regulating mechanisms of marine phytoplankton to keep this
delicate Ca2+ homeostasis differ between species and between functional
groups. In freshwater environments, for example, calcium ions play an
important role shaping microalgal species composition. Desmid green algae
have a narrow tolerance to calcium (Moss, 1972; Tassigny, 1971) and
thrive in soft-water lakes, while submersed macrophytes (Elodea, Stratiotes, Potamogeton) and benthic
cyanobacteria dominate in hard-water lakes, where they can be heavily
encrusted with CaCO3 precipitates.
An early hypothesis describes the invention and the process of
biomineralization in the form of calcium carbonate by marine organisms as a
potential Ca2+ detoxification mechanism (Simkiss, 1977; Kaźmierczak
et al., 1985; Kempe and Degens, 1985). Ocean calcium concentrations have
changed remarkably throughout the Phanerozoic eon (past 541 Myr) as
documented by fluid inclusions of marine halite (Horita et al.,
2002). Over the past 300 Myr, highest seawater Ca2+ concentrations are
documented for the Cretaceous (145 to 66 Ma; Hönisch et al., 2012),
known for massive deposition of biogenic calcareous material produced in the
pelagic ocean. Calcifying phytoplankton (coccolithophores) are the dominant
planktonic calcifiers in the modern ocean and are responsible for up to half
the pelagic production of calcium carbonate (Broecker and Clark, 2009).
Coccolithophores form minute calcite plates (coccoliths) inside a
specialized cell compartment (coccolith vesicle) from where the coccoliths
are subsequently transported to the cell's surface and released via
exocytosis. The record of nannofossils and coccoliths has its origin in the
Late Triassic (about 225 Ma), coinciding with relatively low seawater
Ca2+ concentrations (Bown et al., 2004). Subsequently, seawater
Ca2+ concentrations increased, potentially linked to changes in the
seafloor spreading rates (Skelton, 2003), and peaked in the
Cretaceous at the highest levels since the past 300 Myr (∼ 3 to
4 times the present seawater concentrations of 10 mmol Ca2+ L-1).
Species diversity and abundance of total nannofossils, including
coccolithophores, have increased in concert with high seawater Ca2+
concentrations (Fig. 1).
We tested two calcifying coccolithophores (Emiliania huxleyi and Gephyrocapsa oceanica), two diatoms (Chaetoceros sp. and
Ceratoneis closterium) and one raphidophyte (Heterosigma akashiwo) to elevated seawater calcium concentrations
simulating changes in oceanic Ca2+ levels over the past 300 Myr.
Representative for a non-calcifying coccolithophore, one non-coccolith-carrying (naked) E. huxleyi strain was tested. Furthermore, a possible stimulation of
coccolith production by increased seawater Ca2+ concentration was
investigated in two under-calcifying E. huxleyi strains. If biogenic calcification
represents a viable mechanism to cope with high external Ca2+
concentrations, a diverging response in physiological parameters would be
expected between calcifiers and non-calcifiers.
Seawater Ca2+ concentration and fossil phytoplankton
diversity over the past 300 Myr. Model-reconstructed seawater Ca2+
concentration (blue line; data retrieved from Hönisch et al., 2012),
fossil species diversity of diatoms (red line; data retrieved from Kooistra
et al., 2007), total nannofossils and coccolithophores (black and grey
line, respectively; data retrieved from Bown et al., 2004).
Materials and methodsCulture conditions
Monospecific cultures of the diploid coccolithophores Gephyrocapsa oceanica (CS-335/03) and
Emiliania huxleyi (calcifying CS-370, non-calcifying SO-6.13 and under-calcifying SO-5.25 and
SO-8.04), the diatoms Chaetoceros sp. (CHsp-TB02) and Ceratoneis closterium (CCMMG-3), and the raphidophyte
Heterosigma akashiwo (CS-169) were grown in sterile artificial seawater (Kester et al., 1967)
with macro- and micronutrient additions according to f/2 and f/20 (Guillard,
1975), respectively, or in the case of G. oceanica according to GSe/20 (Loeblich and
Smith, 1968). The under-calcified populations (strains SO-5.25 and SO-8.04)
consist of cells with no or single attached coccoliths. Cells with no
coccoliths attached in these populations either lost their coccoliths,
lacked the ability to produce coccoliths or did not yet produce coccoliths.
Emiliania huxleyi strain SO-6.13 was isolated by Suellen Cook in February 2007 from the Southern
Ocean (54∘ S, 146∘ E; 65 m depth). Multiple single-cell isolates from this water sample resulted in a number of calcified
ecotype B/C E. huxleyi strains. Strain SO-6.13, however, was naked upon isolation and
throughout the conduct of the current study. Much later, in early 2015,
strain SO-6.13 switched from a non-calcifying to a calcifying stage and
started to produce typical B/C coccoliths.
Calcium concentrations were adjusted by varying additions of CaCl2 with
concomitant additions of NaCl, keeping the ionic strength of the artificial
seawater constant. Gephyrocapsa oceanica, H. akashiwo and E. huxleyi (CS-370) were obtained from the Australian
National Algae Culture Collection. Ceratoneis closterium was obtained from the Centre of Climate,
Meteorology and Global Change at the University of Azores (CMMG). All other
species and strains were obtained from the Algae Culture Collection at the
Institute of Marine and Antarctic Studies at the University of Tasmania,
Australia.
Experimental setup
In the first experiment, cells were acclimated to the experimental
conditions (Ca2+ range from 1 to 52 mmol L-1) for more than 50
generations and allowed to consume a maximum of 10 % (non-calcifiers) or
5 % (calcifiers) of dissolved inorganic carbon to avoid major changes in
the carbonate chemistry. Cultures were incubated in triplicates at
12 ∘C (16 ∘C for G. oceanica), a photon flux density of 100 µmol quanta m-2 s-1 and a 16:8 h light : dark cycle at the University
of Tasmania. Ceratoneis closterium was incubated at 20 ∘C, 250 µmol quanta m-2 s-1 and a 14:10 h light : dark cycle at the University of
Azores. The physiological response of all species (except C. closterium) was examined in
terms of growth rate, particulate organic and inorganic carbon cell quota
and production rate, and maximum quantum yield of the photosystem II
(Fv/Fm). Physiology of C. closterium was only examined in terms of growth rate. Seawater
carbonate chemistry was determined from total alkalinity (AT) and
dissolved inorganic carbon (CT) samples taken at the start and the end
of the experiment.
In the second experiment, two under-calcified E. huxleyi strains (SO-5.25 and SO-8.04)
were cultured at the University of Tasmania in triplicates for 2 months under
dilute semi-continuous batch conditions at the identical conditions as
described above with Ca2+ concentrations adjusted to 10 or 36 mmol Ca2+ L-1. Strain-specific growth rate and the number of coccoliths
per cell were monitored over time via cell counts and scanning electron
microscopy, respectively. Cultures were allowed to grow from ∼ 50 to a maximal cell density of ∼ 80 000 cells mL-1, which
prevented major changes in the seawater carbonate chemistry.
Seawater chemistry analysis
Seawater Ca2+ concentrations at the start of the experiment were
determined via chelation ion chromatography (Meléndez et al., 2013),
using an adjusted method to match the different Ca2+ concentrations
(precision of ±1.4 %). Dissolved inorganic carbon and AT were
analysed as the mean of triplicate measurements with the infrared detection
method using an Apollo SciTech DIC analyser (model AS-C3) and the
potentiometric titration method (Dickson et al., 2003), respectively. Data
were corrected to certified reference materials (Scripps Institution of
Oceanography, USA). Consecutive measurements of the Dickson standard
resulted in an average precision of > 99.8 % for both CT
and AT. The carbonate system was calculated using equations from Zeebe and
Wolf-Gladrow (2001) with dissociation constants for carbonic acid after Roy
et al. (1993), modified with sensitivity parameters for [Na+],
[Mg2+] and [Ca2+] (Ben-Yaakov and Goldhaber, 1973). The calcite
saturation state (Ω) was calculated with regard to the Mg / Ca ratio as
described in Tyrrell and Zeebe (2004). Detailed information on the carbonate
system parameters can be found in the Supplement.
Physiological parameters
Maximum quantum yield of the photosystem II (Fv/Fm) was measured on dark-adapted samples (45 min) using a WATER-PAM fluorometer (Walz GmbH,
Germany). Subsamples for total particulate carbon (TPC) and particulate
organic carbon (POC) were filtered onto pre-combusted (7 h,
450 ∘C) quartz-microfibre filters (pore-size of 0.3 µm) and
stored at -24 ∘C. Filters for POC analysis were fumed with
saturated HCl for 10 h to remove all inorganic carbon. TPC and POC were
measured on an elemental analyser (Thermo Finnigan EA 1112, Central Science
Laboratory of the University of Tasmania). Particulate inorganic carbon
(PIC) was calculated as the difference between TPC and POC. Cell numbers
were obtained by means of triplicate measurements with a Multisizer 4
Coulter Counter (Beckman Coulter, USA) or by light microscopy counts. The
average cell number was used to calculate the growth rate μ (d-1)
as μ= (ln(c1)- ln(c0))/(t1-t0), where
c0 and c1 are the cell concentrations at the beginning (t0)
and the end of the incubation period (t1). POC and PIC production rates
were calculated from cell quota and species-specific growth rates.
Phytoplankton physiological responses to seawater Ca2+
concentration. Displayed are laboratory-cultured strains of diatoms (red
markers), raphidophytes (blue markers), coccolithophores (black markers) and
a non-calcifying coccolithophore (black open marker): (a) species-specific
growth rate, (b) maximum quantum yield of photosynthesis (Fv/Fm), (c) cellular POC and
(d) PIC quotas, (e) cellular POC and (f) PIC production
rates as a function of seawater Ca2+ concentration. Error bars denote
±1 SD (n= 3). Note that the physiological response of Ceratoneis closterium was only
determined via growth rate measurements. POC quota of H. akashiwo could not be
determined at a Ca2+ concentration of 42 mmol L-1 due to lack of
growth.
Scanning electron microscopy
Samples for electron microscopy were filtered gently onto polycarbonate
filters, air-dried at 60 ∘C and afterwards sputter-coated with
gold–palladium. Photographs were taken with a Hitachi SU-70 field emission
scanning electron microscope (SEM) at the Central Science Laboratory of the
University of Tasmania. During SEM sessions, > 50 cells were
visually evaluated and representative pictures were taken.
Relative physiological response of phytoplankton species to
seawater Ca2+ concentration. Relative fitness expressed in terms of
(a) growth rate and (b) POC production of all tested species normalized to
ambient seawater Ca2+ concentration of ∼ 10 mmol L-1, and supplemented with coccolithophore literature data from
Müller et al. (2011) and Herfort et al. (2004) to illustrate the effect
of calcium poisoning on calcifiers and non-calcifiers. Solid lines indicate
regressions through calcifiers: (a)y=-0.0036×+1.0483 (r2= 0.278,
p= 0.035, n= 16) and (b)y=-0.0052×+1.0704 (r2= 0.184,
p= 0.067, n= 19). Dotted lines indicate regressions through
non-calcifiers: (a)y=-0.025×+1.307 (r2= 0.858, p < 0.0001,
n= 20) and (b)y=-0.024×+1.303 (r2= 0.826, p < 0.0001, n= 15).
Representative SEM photographs of the under-calcified E. huxleyi
strain SO-8.04 cultured at modern seawater Ca2+ concentration of 10 mmol L-1, showing no or only single attached coccoliths (a). When cultured
for 2 months at elevated Ca2+ concentration of 36 mmol Ca2+ kg-1, E. huxleyi strain SO-8.04 produced a sufficient number of coccoliths to
cover the whole cell (b).
Results
In the first experiment, at Ca2+ concentrations below 2 mmol L-1,
all species exhibited significantly (t test, p < 0.05) lower growth,
particulate organic carbon (POC) production rates and maximum quantum yield
of photosystem II (Fv/Fm) compared to modern seawater concentrations of
∼ 10 mmol Ca2+ L-1 (Fig. 2). Furthermore, the two
calcifying species displayed decreased particulate inorganic carbon (PIC)
production rates at Ca2+ concentrations below 2 mmol L-1 compared
to ∼ 10 mmol Ca2+ L-1 (t test, p < 0.05). At
elevated Ca2+ concentrations all non-calcifying species exhibited a
severe reduction in growth, POC production and maximum quantum yield (Fig. 2). In the most extreme cases no growth was detected at 42 and
52 mmol Ca2+ L-1 in H. akashiwo and C. closterium, respectively. Both tested coccolithophore
species, on the other hand, were able to maintain their growth, Fv/Fm, POC
and PIC production rates with no substantial change at calcium concentration
expected for Cretaceous seawater (25 to 40 mmol Ca2+ L-1). A
further increase in external Ca2+ concentrations up to 52 mmol L-1
adversely affected POC and PIC production only in E. huxleyi, whereas G. oceanica was not
impaired. The non-calcifying strain of E. huxleyi exhibited a similar response to that of the
diatom and raphidophyte species with reduced physiological rates of up to
84 % at Ca2+ concentrations of 19 mmol L-1 and higher (Fig. 2).
To illustrate the diverging physiological response of calcifying
coccolithophores and non-calcifying phytoplankton, we normalized growth and
POC production rates from the current study and literature data to the
species-specific rates exhibited at modern ocean calcium levels (Fig. 3). A
linear regression fit (from 9 to 52 mmol Ca2+ L-1) through
calcifiers and non-calcifiers resulted in a 6.9 times steeper reduction for
the latter group in terms of growth rate (Fig. 3a) and a 4.6 times steeper
reduction in terms of POC production rates (Fig. 3b).
In the second experiment, the two under-calcified E. huxleyi strains (SO-5.25 and
SO-8.04) cultured at elevated seawater Ca2+ concentrations (36 mmol L-1) displayed no significant change in growth rate (t test,
p > 0.05) compared to strains cultured at modern Ca2+
concentrations of 10 mmol L-1 (0.67 ± 0.01 and 0.72 ± 0.01 d-1 compared to 0.68 ± 0.01 and 0.71 ± 0.01 d-1 for the
strains SO-5.25 and SO-8.04, respectively). The number of coccoliths per
cell, however, increased remarkably from fewer than 2 coccoliths per cell
at 10 mmol Ca2+ L-1 to more than 12 coccoliths per cell, forming a
complete coccosphere, at 36 mmol Ca2+ L-1 (Fig. 4).
Discussion
The results presented here demonstrate the influence of seawater Ca2+
concentrations on marine phytoplankton physiology (in terms of growth and
particulate organic carbon production). Whereas previous studies have already
investigated the effects of elevated seawater Ca2+ concentrations on
calcifying coccolithophore physiology and coccolith formation (Herfort et
al., 2004; Langer et al., 2007; Müller et al., 2011), this study is to
our knowledge the first to investigate the Ca2+ sensitivity of
non-calcifying phytoplankton in the laboratory. Marine phytoplankton
presumably operate several mechanisms which contribute to cellular Ca2+
regulation, such as intra- and extracellular enzymatic binding capacities
and/or the influx regulation via selective channels (Gadd, 2010). Over the past decade progress has been made in the discovery of cellular compartments (e.g. endoplasmic
reticulum, chloroplast, mitochondria) regulating plant Ca homeostasis and signalling (McAinsh and Pittmann, 2009;
Webb, 2008; Brownlee and Hetherington, 2011), as well as in the differences between the Ca channels of eukaryotes, higher plants and mammalian cells (Wheeler and Brownlee,
2008). However, many unknowns remain about phytoplankton
intracellular ion regulation and the homeostasis of the major biological
active cations like Ca2+ and Mg2+ and their interaction and
possible influence on each other. For example, Ca2+ has a higher
ion-exchange capacity than Mg2+ (Harris, 2010) and when present in high
concentrations might interfere with enzymatic reactions where Mg2+ acts
as a cofactor (Moore et al., 1960; Legong et al., 2001). However, it remains
speculative whether this is a possible explanation for the observed reduction in
growth rate and Fv/Fm of non-calcifying phytoplankton species (Fig. 2).
The non-calcifying strain of E. huxleyi showed a comparable response to elevated
seawater Ca2+ concentrations as the diatom and raphidophyte species
(Fig. 3). This indicates that the Ca2+ tolerance of calcifying
coccolithophores compared to non-calcifying phytoplankton is not a
taxon-specific trait but connected to the process of calcification itself
and, furthermore, suggests that coccolithophore biomineralization acts as an
efficient mechanism to cope with high external Ca2+ concentrations.
Reduced overall fitness triggered by high external Ca2+ concentrations
is presumably associated with enhanced transmembrane Ca2+ influx, leading
to higher energetic costs for cytosolic Ca2+ removal and might
ultimately result in a disadvantage in resource competition between
phytoplankton species. Dunaliella, a member of the class Chlorophyceae, is one of the most tolerant
phytoplankton species regarding high external ion concentrations and
regularly blooms in highly saline lakes (Oren, 2002, 2005). However, this
extremophile species is inhibited in growth by high external Ca2+
concentrations and only forms blooms in hypersaline lakes when the upper
water layer becomes sufficiently diluted with regard to its Ca2+
concentrations (Baas-Becking, 1931). This emphasizes the ecological
importance of external Ca2+ concentrations for phytoplankton growth
dynamics.
The remarkable tolerance of calcifying coccolithophores to elevated
Ca2+ concentrations likely results from a tight control on
transmembrane Ca2+ entry, intracellular transport, and deposition.
Seawater Ca2+ presumably enters the coccolithophore cell through
permeable channels into the peripheral endoplasmatic reticulum. Via the
endomembrane transport network it reaches a Golgi-derived organelle, the
coccolith vesicle, where it is precipitated as CaCO3 (Mackinder et
al., 2010). Precipitation of Ca2+ in the form of calcite changes the
ion to a biochemically inert state. Large amounts of Ca2+ can thereby
be sequestered in a finite space and time. For Emiliania huxleyi to sustain a typical rate of
calcification, an uptake of 5 × 106 Ca2+ ions s-1
is required (Mackinder et al., 2010). The fact that this massive intracellular Ca2+
flux needs to be achieved at a cytosolic concentration of only 100 nmol Ca2+ L-1 without disturbing the cell's delicate Ca2+
homeostasis exemplifies the level of cellular control involved in
coccolithophore calcification. It appears reasonable to assume that this
tight cellular control of biogenic calcification (which includes CaCO3
precipitation inside the coccolith vesicle and the regulation of cellular
Ca2+ entrance and distribution) also allows for the observed tolerance
to external Ca2+ concentrations. The absence of Ca2+-stimulated
calcification at levels above modern ocean Ca2+ concentrations (Fig. 2f) is in line with previous findings, which indicate saturation of
calcification in E. huxleyi and C. braarudii at ∼ 10 mmol Ca2+ L-1 (Herfort et al., 2004; Trimborn et al., 2007; Leonardos et al., 2009;
Müller et al., 2011). This suggests that in coccolithophores adapted to
modern ocean conditions, factors other than the Ca2+ concentration may
limit CaCO3 precipitation at higher than ambient Ca2+ levels.
Potentially limiting factors include dissolved inorganic carbon acquisition
and energy supply for the process of calcification (Bolton and Stoll, 2013;
Bach et al., 2015).
Emiliania huxleyi is characterized by three distinct different cell forms: (a) the
coccolith-carrying non-motile diploid form (C cell), (b) the naked non-motile diploid
form (N cell) and (c) the scaly motile haploid form (S cell). The latter
haploid form possesses organic body scales covering the cell and two
flagellates that enable motion (Paasche, 2002). The life cycle of E. huxleyi consists
of C and S cells, whereas N cells are mostly observed in the laboratory
after extended culture periods (Paasche, 2002) or under unfavourable culture
conditions (Müller et al., 2015). This study investigated only the
diploid coccolith-carrying (C cell) and the naked (N cell) cell forms of E. huxleyi.
Our observations and the presence of N and S cells in laboratory cultures
and natural populations (Paasche, 2002; Frada et al., 2012; Müller et
al., 2015) indicate that E. huxleyi cells have the ability to control intracellular
Ca2+ homeostasis at modern Ca2+ concentrations without the need of
biomineralization.
At modern seawater conditions some E. huxleyi strains display an incomplete coccolith
cover (coccosphere) with less than 2 coccoliths per cell (Fig. 4a)
instead of the 10 to 15 that are necessary to form a complete coccosphere
(Paasche, 2002). The results of the second experiment indicate that an
existent but under-saturated calcification mechanism can be stimulated by
increased seawater Ca2+ concentrations (Fig. 4b) and, furthermore,
might prevent cellular Ca2+ poisoning as seen in the non-calcifying E. huxleyi
strain (Figs. 2 and 3). However, benefits of coccolith formation are expected
which evidently outweigh the substantial costs of this energy-consuming
process even under modern ocean Ca2+ concentrations. Although numerous
hypotheses have been proposed concerning the precise function of
coccolithophore calcification, including ballasting and protection from
viruses, grazers and damaging irradiance, so far none of these is
conclusively supported by experimental evidence (Raven and Crawfurd, 2012;
Barcelos e Ramos et al., 2012).
Palaeoecological implications
Palaeoceanographic studies have indicated that the oceanic conditions of the
Cretaceous were quite different from those in the modern ocean (e.g. see
Zeebe, 2001; Hay, 2008). Besides elevated seawater Ca2+
concentrations (Fig. 1), the Cretaceous was marked by a warm greenhouse
environment, elevated sea levels, warm shallow shelf seas and altered
oceanic circulation. Here we tested whether the biomineralization mechanism
in coccolithophores increases their resilience to cellular calcium stress,
which indeed is indicated by the physiologically different responses of the
three calcifying coccolithophore species (E. huxleyi, G. oceanica and C. braarudii) compared to the
non-calcifying species (Fig. 3). Cretaceous seawater Ca2+
concentrations may thus have represented a selective advantage for
coccolithophores during this period of the geological past. This could
explain the proliferation and high productivity of coccolithophores during
the Cretaceous compared to non-calcifying phytoplankton. We cannot exclude
the possibility of other environmental factors that might have supported the
proliferation of coccolithophores or suppressed non-calcifiers in the
Cretaceous (e.g. Stanley et al., 2005), but the seawater Ca2+
concentrations seem to be a major environmental aspect promoting
coccolithophore over non-calcifying phytoplankton growth.
It remains an open question whether the onset of calcification in
coccolithophores (approx. 225 Ma) at relatively low seawater Ca2+
concentrations evolved primarily to efficiently regulate cellular Ca2+
homeostasis or whether calcification had other functions at that time. If
calcification in coccolithophores evolved as a Ca2+ detoxification
mechanism, it was presumably an additional instrument to regulate
intracellular Ca2+ levels because other strategies must have existed in
the ancestors of coccolithophores that did not precipitate calcium
carbonate. It is reasonable to assume that the rising oceanic Ca2+
concentrations represented a selective pressure on phytoplankton populations
and may have provided an evolutionary advantage to coccolithophores over
non-calcareous phytoplankton during the Jurassic and Cretaceous period (Fig. 1). However, secondary benefits of calcification are likely responsible for
its continued operation under modern ocean Ca2+ concentrations.
Interestingly, E. huxleyi and G. oceanica, the dominant coccolithophores in the modern ocean, are
two of the few coccolithophore species that have a non-calcifying haploid
life stage, whereas the haploid life stage of the majority of
coccolithophores is calcified (Billard and Inouye, 2004). This led us to
suggest that these two species in the modern ocean do not rely on cellular
Ca2+ detoxification by biomineralization.
Concluding remarks
The concept of biocalcification as a Ca2+ detoxification mechanism in
marine organisms has been proposed earlier (Simkiss, 1977; Kaźmierczak
et al., 1985) and, based on the results of this study, is supported for
coccolithophores. The occurrence of calcified cyanobacteria in the
geological record during the Phanerozoic also appears to be connected to
elevated seawater Ca2+ concentrations (Arp et al., 2001), suggesting
similarities in the benefits of calcification in fossil cyanobacteria and
coccolithophores. It remains speculative to extend the
“Ca2+-detoxification concept” to other marine calcifying groups or to
the onset of biocalcification in the Precambrian–Cambrian transition (Kempe
and Kaźmierczak, 1994; Brennan et al., 2004). However, in view of the
substantial variability in seawater Ca2+ concentration during Earth's
history and the observed Ca2+ sensitivity of dominant marine
phytoplankton species, the ocean's Ca2+ ion concentration should be
considered a potential factor influencing the evolution of marine life on
Earth.
The Supplement related to this article is available online at doi:10.5194/bg-12-6493-2015-supplement.
Acknowledgements
We thank D. Davis for laboratory assistance and A. McMinn for providing a
PAM fluorometer. We are grateful for the constructive comments of T. Tyrrell, J. Young and one
anonymous reviewer. Additional comments from a
research group meeting (composed of L. Munns, M. Duret, C. Daniels, K. Mayers, A. Poulton and R. Sheward) further increased the quality of the
manuscript. The work was funded by the Australian Research Council (DP
1093801 to G. M. Hallegraeff and T. W. Trull) and the “Conselho Nacional de
Desenvolvimento Científico e Tecnológico Brasil (CNPq, Processo:
405585/2013-6)”. K. G. Schulz is the recipient of an Australian Research
Council Future Fellowship (FT120100384). We thank the data publisher PANGAEA
for hosting and making the data fully available under http://doi.pangaea.de/10.1594/PANGAEA.854719.
Edited by: J.-P. Gattuso
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