Coral reefs are constructed by calcifiers that precipitate calcium carbonate
to build their shells or skeletons through the process of calcification.
Accurately assessing coral calcification rates is crucial to determine the
health of these ecosystems and their response to major environmental changes
such as ocean warming and acidification. Several approaches have been used
to assess rates of coral calcification, but there is a real need to compare
these approaches in order to ascertain that high-quality and intercomparable
results can be produced. Here, we assessed four methods (total alkalinity
anomaly, calcium anomaly, 45Ca incorporation, and 13C
incorporation) to determine coral calcification of the reef-building coral
Stylophora pistillata. Given the importance of environmental conditions for this process, the
study was performed under two starting pH levels (ambient: 8.05 and low:
7.2) and two light (light and dark) conditions. Under all conditions,
calcification rates estimated using the alkalinity and calcium anomaly
techniques as well as 45Ca incorporation were highly correlated. Such a
strong correlation between the alkalinity anomaly and 45Ca
incorporation techniques has not been observed in previous studies and most
probably results from improvements described in the present paper. The only
method which provided calcification rates significantly different from the
other three techniques was 13C incorporation. Calcification rates based
on this method were consistently higher than those measured using the other
techniques. Although reasons for these discrepancies remain unclear, the use
of this technique for assessing calcification rates in corals is not
recommended without further investigations.
Introduction
Calcification is the fundamental biological process by which organisms
precipitate calcium carbonate. Calcifying organisms take up calcium and
carbonate or bicarbonate ions to build their biomineral structures
(aragonite, calcite, and/or vaterite) which have physiological, ecological,
and biogeochemical functions. Moreover, calcium carbonate plays a major role
in the services provided by ecosystems to human societies. The ocean has absorbed large amounts of anthropogenic CO2 since the
start of the industrial revolution and is currently sequestering about
22 % of CO2 emissions (average
2008–2017; Le Quéré et al., 2018). This massive input of CO2 in
the ocean impacts seawater chemistry with a decrease in seawater pH and
carbonate ion concentrations [CO32-] and an increase in CO2
and bicarbonate concentrations [HCO3-]. These fundamental changes
to the carbonate system are referred to as “ocean acidification”
(OA; Gattuso and Hansson, 2011). Models project that the average
surface water pH will drop by 0.06 to 0.32 pH units by the end of the
century (IPCC, 2014).
The effect of OA is currently the subject of intense research with
particular attention to organisms producing CaCO3. For instance, coral
communities have already proven to be particularly vulnerable to rapidly
changing global environmental conditions (e.g., Albright et
al., 2018). In order to help project the future of coral reefs, accurate
estimates of calcification rates during realistic perturbation experiments
are necessary in order to produce high-quality and intercomparable results (Cohen
et al., 2017; Gazeau et al., 2015; Langdon et al., 2010; Riebesell et al.,
2010; Schoepf et al., 2017).
Several methods are available to quantify rates of coral calcification.
Calcification can be measured as the increase in CaCO3 mass
(e.g., the buoyant weight technique; Jokiel et al., 1978)
or following the incorporation of radio-labeled carbon or calcium in the
skeleton (Goreau, 1959), but also through the quantification of
changes in a seawater constituent that is stoichiometrically related to the
amount of CaCO3 precipitated. For instance, the alkalinity anomaly
technique (Smith and Key, 1975) has been widely used to estimate net
calcification of organisms and communities, especially of corals and coral
reef environments (e.g., Smith and Kinsey, 1978; Gazeau et al., 2015;
Albright et al., 2016; Cyronak et al., 2018). Total alkalinity (AT) is
directly influenced by bicarbonate and carbonate ion concentrations together
with a multitude of other minor compounds (Wolf-Gladrow et al., 2007).
Calcification consumes carbonate or bicarbonate, following the reversible
reaction
Ca2++2HCO3-↔CaCO3+CO2+H2O.
Calcification consumes 2 mol of HCO3-, hence decreasing
AT by 2 mol mol-1 of CaCO3 produced (Reaction R1). It is possible
to derive the rate of net calcification (gross calcification – dissolution)
by measuring AT before and after incubating an organism or a community.
This method assumes, however, that calcification is the only biological
process influencing AT (Smith and Key, 1975). Nitrogen
assimilation through photosynthetic activities, nitrification, and
aerobic and anaerobic remineralization of organic matter is known to impact
AT through the consumption or release of nutrients (ammonium, nitrate,
and phosphate) and protons (Wolf-Gladrow et al., 2007).
While for some group of species (e.g., bivalves, sea urchins), corrections
appear necessary to take into account the effect of nutrient release on
AT, changes in nutrient concentrations during incubations of isolated
corals are too low (i.e., several orders of magnitude lower than changes in
AT) to introduce a significant bias in the calculations (Gazeau et al., 2015).
In contrast to AT, the concentration of calcium (Ca2+) in seawater
is only biologically influenced by net calcification, and a 1 : 1 relationship
can be used to derive net calcification rates (Reaction R1). The depletion of
AT and Ca2+ needs to be corrected for gains of AT and Ca2+
resulting from evaporation. These corrections can be applied through the
incubation of seawater in the absence of coral (Schoepf et al.,
2017). Both the alkalinity anomaly and calcium anomaly methods are
nondestructive and typically show a good agreement
(Chisholm and Gattuso, 1991; Murillo et al.,
2014; Gazeau et al., 2015).
The 45Ca incorporation technique has been used since the 1950s
(Goreau and Bowen, 1955; Goreau, 1959). While earlier
techniques showed low reproducibility, methodological improvements led to a
significant reduction of the deviations between replicates (see
Tambutté et al., 1995, for more details). The strength of this method is
that it is extremely sensitive for measuring short-term variations in gross
calcification rates. However, in contrast to the AT and Ca2+
anomaly techniques, it is a sample-destructive method.
Previous studies designed to compare calcification rate estimates using the
45Ca incorporation and AT anomaly methods revealed subtle
discrepancies. For example, Smith and
Kinsey (1978) reported an overestimation of rates based on the 45Ca
method. In contrast, Tambutté et al. (1995) and
Cohen et al. (2017) reported a decrease in AT without
concomitant incorporation of 45Ca, therefore suggesting an
overestimation of calcification derived from AT measurements. However,
during these studies, in order to avoid radioactive contamination of
laboratory equipment, estimates of calcification were not performed during
the same incubations, but rather during incubations performed over 2
consecutive days.
In contrast to the 45Ca incorporation method, to the best of our
knowledge, no studies have used carbon-based incorporation techniques to
estimate coral calcification rates in the framework of ocean acidification.
Past studies that compared carbon and calcium incorporation rates in coral
skeletons based on a double labeling technique with H14CO3 and
45Ca showed that only a minor proportion of the labeled seawater
carbon is incorporated in the skeleton (e.g., Marshall and
Wright, 1998) and that the major source of dissolved inorganic carbon for
calcification is metabolic CO2 (70 %–75 % of the total
CaCO3 deposition; Furla et al., 2000). Consequently, under both light
and dark conditions, the rate of 45Ca deposition appears greater than
the rate of 14C incorporation (Furla et al., 2000). To the
best of our knowledge, only one study estimated calcification rates of a
benthic calcifier (coralline algae) using a stable carbon isotopic technique
through addition of 13C-labeled bicarbonate (McCoy et al., 2016). The
present study aimed at comparing calcification rates measured using the
alkalinity and calcium anomaly methods, as well as the 45Ca and
13C incorporation techniques, under different pH and light conditions.
Material and methods
Colonies of the reef-building coral Stylophora pistillata were incubated in the laboratory, in both the light and dark, under ambient and lowered pH conditions. At ambient
pH (experiment conducted in July–August 2017), two sets of incubations were
performed using either 45Ca or 13C additions, and calcification
rates based on these techniques were compared to those derived, during the
same incubations, by the alkalinity and calcium anomaly techniques. At
lowered pH (experiment conducted in August 2018), no incubations with
13C addition were conducted and only the three other techniques were
compared.
Experimental details for the series of incubations of the coral
Stylophora pistillata performed under ambient and low pH, and in the light and dark following
45Ca or 13C labeling. The ratio Ww:Wc corresponds to
the ratio between seawater weight (g) and skeletal dry weight (g). Values
represent mean ± standard deviation (SD); n is the number of true
replicates considered for each experiment. All incubations were conducted at
25±0.5∘C.
Different types of incubations were conducted. In July–August 2017; one set
of incubations was performed under ambient pH conditions with the addition
of radioactive calcium dichloride (45CaCl2). During the same
period, another set of incubations was performed, under ambient pH
conditions, with the addition of 13C-labeled sodium bicarbonate
(13C-NaHCO3 99 %). Finally, in August 2018, one set of
incubations was performed under lowered pH conditions (see thereafter for
more details) with the addition of 45CaCl2. For all sets of
incubations, organisms were incubated for 5 to 11 h (Table 1), in both
the light and dark, in 500 mL polyethylene beakers equipped with a magnetic
stirrer (Fig. 1). Six and five replicates were used, respectively, at
ambient and low pH. Furthermore, for all sets of incubations, one beaker was
incubated, under the same conditions as the other beakers, without coral and
served as a control.
Scheme of the polyethylene container in which a coral nubbin is
suspended with a nylon line and covered with a transparent film.
For all sets of incubations, samples for the measurements of pHT,
AT (200 mL), and Ca2+ concentrations (50 mL) were taken before
distributing seawater to the experimental beakers. While pHT was
measured immediately after sampling, samples for AT measurements were
poisoned with 40 µL of 50 % saturated HgCl2 and stored in the
dark at 4 ∘C pending analysis less than 2 weeks later. Samples
for [Ca2+] measurements were not poisoned and were stored in the dark at 4 ∘C pending analysis less than 2 weeks after sampling.
Incubations in the light were performed at an irradiance of 200 µmol photons m-2 s-1 during daytime whereas dark incubations were
conducted at night. Incubation times were not fixed based on scientific
considerations and differed between the different incubations due to
practical constrains (i.e., access to the lab). Before the
beginning of the incubations, all beakers (containing corals) were precisely
weighed at ±0.01 g (Sartorius BP 310S).
At the conclusion of the incubations, all beakers were precisely weighed to
evaluate evaporation, and seawater samples were analyzed for pHT,
AT, and [Ca2+] as well as for 45Ca activity or δ13C-CT depending on the type of incubations. pHT was measured
immediately and samples for AT and [Ca2+] determinations were
filtered at 0.2 µm (GF/F, Ø 47 mm), poisoned with saturated
HgCl2 (only for AT), and stored in the dark at 4 ∘C
pending analysis (within 2 weeks). The corals were then removed from the
beakers for the analysis of incorporated 45Ca or 13C. Three
additional corals which were not incubated were processed for carbon
isotopic composition of the previously accreted calcium carbonate (see
Sect. 2.3).
Analytical techniques
Immediately after sampling, pHT was measured on a Metrohm 826 mobile
pH logger, and a glass electrode (Metrohm, Ecotrode Plus) was calibrated on the
total scale using a TRIS buffer of salinity 35 (provided by Andrew Dickson,
Scripps University, USA). AT was determined in triplicate 50 mL
subsamples by potentiometric titration on a titrator Titrando 888 (Metrohm)
coupled to a glass electrode (Metrohm, Ecotrode Plus) and a thermometer
(pt1000). The pH electrode was calibrated before every set of measurements
on the total scale using a TRIS buffer of salinity 35 (provided by Andrew Dickson, Scripps University, USA). Measurements were carried out at a
constant temperature of 25 ∘C and AT was calculated as
described in Dickson et al. (2007). Certified reference material
(CRM; batches 143 and 156) provided by Andrew Dickson (Scripps University, USA)
was used to check precision (standard deviation within measurements of the
same batch) and accuracy (deviation from the certified nominal value). Over
the six series of AT measurements performed during the experiment, mean
accuracy and precision (± SD) were respectively 7.2±1.2 and
1.2±0.2µmol kg-1. [Ca2+] was determined in
triplicate using the ethylene glycol tetra acetic acid (EGTA) potentiometric
titration (Lebel and Poisson, 1976). About 10 g of sampled seawater
and 10 g of HgCl2 solution (ca. 1 mmol L-1) were accurately
weighed out. Then, about 10 g of a concentrated EGTA solution (ca. 10 mmol L-1, also by weighing) was added to completely complex Hg2+ and to
complex nearly 95 % of Ca2+. After adding 10 mL of borate buffer
(pHNBS∼ 10) to increase the pH of the solution, the
remaining Ca2+ was titrated by a diluted solution of EGTA (ca. 2 mmol L-1) using a titrator (Titrando 888, Metrohm) coupled to an
amalgamated silver combined electrode (Ag Titrode, Metrohm). Following
Cao and Dai (2011), the volume of EGTA necessary to titrate the
remaining ca. 5 % of Ca2+ was obtained by manually fitting a
polynomial function to the first derivative of the titration curve using the
function “loess” of the R software (R Development Core Team, 2018). The EGTA
solution was calibrated prior to each measurement series using International
Association for the Physical Sciences of the Oceans (IAPSO) standard
seawater (salinity = 38.005). Mean [Ca2+] precision obtained using
this technique was 2.9 µmol kg-1 (n=40), corresponding to a
coefficient of variation (CV) of 0.026 %.
To determine the specific activity in radio-labeled seawater, the 1 mL
aliquots were transferred to 20 mL glass scintillation vials and mixed in
proportion 1 : 10 (v:v) with scintillation liquid Ultima Gold™ XR.
According to a method adapted from Tambutté et al. (1995), at
the end of incubation sampled nubbins were immersed for 30 min in beakers
containing 300 mL of unlabeled seawater to achieve isotopic dilution of the
45Ca contained in the gastrovascular cavity. Constant water motion was
provided in the efflux medium by magnetic stirring bars. Tissues were then
dissolved completely in 1 mol L-1 NaOH at 90 ∘C for 20 min. The skeleton was rinsed twice in 1 mL NaOH and twice in 5 mL of Milli-Q
water. It was then dried for 72 h at 60 ∘C, precisely weighed at
±0.01 g using a Sartorius BP 310S (referred to thereafter as skeleton
dry weight), and dissolved in 12 N HCl. Three 200 µL aliquots from each
skeleton dissolution were transferred to 20 mL glass scintillation vials and
mixed with 10 mL scintillation liquid Ultima Gold™ XR. Radioactive
samples were thoroughly mixed to homogenize the solution and kept in the
dark for 24 h before counting. The radioactivity of 45Ca was counted
using a Tri-Carb 2900 liquid scintillation counter. Counting time was
adapted to obtain a propagated counting error of less than 5 % (maximal
counting duration was 90 min). Radioactivity was determined by comparison
with standards of known activities, and measurements were corrected for
counting efficiency and physical radioactive decay.
The analyses of seawater δ13C-CT as well as of the 13C
signature of coral calcified tissues were performed at Leuven University.
For δ13C-CT analyses, a helium headspace (5 mL) was created
in the vials and samples were acidified with 2 mL of phosphoric acid
(H3PO4, 99 %). Samples were left to equilibrate overnight to
transfer all CT to gaseous CO2. Samples were injected in the
carrier gas stream of an EA-IRMS (Thermo EA1110 and Delta V Advantage), and
data were calibrated with NBS-19 and LSVEC standards (Gillikin and
Bouillon, 2007). Corals were treated following the same protocol as for
45Ca incorporation measurements and powdered. Triplicate subsamples of
carbonate powder (∼100µg) were placed into gas-tight
vials, flushed with helium, and converted into CO2 with
H3PO4. After 24 h, subsamples of the released CO2 were
injected into the EA-IRMS system as described above. Data were calibrated
with NBS-19 and LSVEC. Carbon isotope data are expressed in the delta
notation (δ) relative to the Vienna Pee Dee Belemnite (VPDB) standard
and were calculated as
Rsample=δ13Csample1000+1⋅RVPDB.
Computations and statistics
The carbonate chemistry was assessed using pHT and AT and the R
package seacarb (Gattuso et al., 2019). Propagation of errors on
computed parameters was performed using the new function “error” of the
package seacarb (Orr et al., 2018) on the R software,
considering errors associated with the estimation of AT as well as errors
on dissociation constants.
Estimates of coral calcification rates based on changes in AT and
[Ca2+] during incubations were computed following Eqs. (2) and
(3), respectively. As shown in these equations, initial levels of AT and
[Ca2+] are not necessary to compute calcification rates and only final
values in the incubations with corals and without corals (controls) were
used:
2GAT=-(AT2-AT1)-(AT2c-AT1)2t⋅WwWc=-(AT2-AT2c)2t⋅WwWc,3GCa=-(Ca2-Ca1)-(Ca2c-Ca1)t⋅WwWc=-(Ca2-Ca2c)t⋅WwWc,
where AT1 and Ca1 are AT and Ca2+ concentrations at the
start of the incubations (µmol kg-1; not used in the
computations); AT2/AT2c and Ca2/Ca2c are AT and
Ca2+ concentrations at the end of the incubations, respectively with
and without corals; t is the incubation duration in hours; and Ww and Wc
are respectively the mass of seawater (average between initial and final
weights) and the coral skeleton dry weight (g; DW). GAT and GCa
are therefore expressed in µmol CaCO3 g DW-1 h-1. Error
propagation was used to estimate errors.
4SEGAT=SEAT22+SEAT2c22t⋅WwWc5SEGCa=SECa22+SECa2c2t⋅WwWc
Here
SEAT2/SEAT2c
and
SECa2c/SECa2c correspond to standard errors associated with the measurement of three
analytical replicates per sample for AT and Ca2+ at the end of the
incubations, respectively with and without corals; t is the incubation
duration in hours; and Ww and Wc are respectively the mass of seawater
(average between initial and final weights) and the coral skeleton dry
weight (g DW).
Coral calcification rates based on 45Ca incorporation were estimated
using measured seawater activity and activity recorded in the skeleton
digest. Rates were then normalized per gram of skeleton dry weight using the
formula
G45Ca=Activitysample⋅CaActivityseawaterWc⋅t,
where Activitysample is the average of counts per minute (CPMs) of three
200 µL aliquots from the dissolved skeleton sample,
Activityseawater is the total CPMs in the 1 mL seawater samples, Ca is
the [Ca2+] measured in the corresponding samples (average between
initial and final values, µmol kg-1) and further converted to µmol L-1 considering a temperature of 25 ∘C and a salinity
of 38, Wc is the skeleton dry weight (in grams), and t is the incubation
duration (in hours). G45Ca is therefore expressed in µmol CaCO3 g DW-1 h-1. The standard errors for these calcification rate
estimates were propagated based on standard errors associated with the
measurements of triplicate samples for both Activitysample and
[Ca2+].
The precipitation of calcium carbonate minerals (G) during the incubation
interval was also estimated using measured δ13C values and
isotope mass balance calculations (Eqs. 7 and 8 below). The CO2
released during phosphoric acid digestion is derived from two sources: new
coral CaCO3 and previously accreted skeletal carbonate mineral. The new
carbon acquired in each measured nubbin (δ13CN) was
assumed to have the same carbon isotope composition as the labeled seawater
CT (average between initial and final level, δ13C-CT∼ 1400 ‰–1700 ‰). The previously accreted
skeletal material was assumed to have a δ13C value equal to
the measured value for the background sample (δ13CP). The
δ13C value (δ13CM), representing the mixture
of new calcified material and previously accreted carbonate mineral, is then
calculated with the following mixing equation:
δ13CM=fG⋅δ13CN+(1fG)⋅δ13CP,
where fG is the fraction of the calcium carbonate mineral precipitated
during the experiment, and δ13CN and δ13CP are the carbon isotope compositions of the newly
precipitated and previously accreted calcium carbonate, respectively.
Equation (7) was solved for fG to determine the calcium carbonate
precipitated during the incubation using
G13C=fGt⋅MCaCO3×106,
where MCaCO3 is the molar mass of calcium carbonate (g mol-1) and
t is the incubation duration in hours. G13C is therefore expressed
in µmol CaCO3 g DW-1 h-1. The standard errors for these
calcification rate estimates were calculated based on standard errors
associated with the triplicate measurements of δ13CP
and δ13CN.
Model II linear regressions (Sokal and Rohlf, 1995) were used to compare net
calcification rates obtained with the different methods. All regressions
were performed using the function “lmodel2” of the package
lmodel2 (Legendre and Oksanen, 2018) with the R software.
Results
Environmental conditions at the start of the different incubations are shown
in Table 2. All values in Table 2 as well as in the text below correspond to
the average between replicates (or incubations) ± standard deviation
(SD). All incubations performed under ambient pHT (∼8.05) were conducted under carbonate chemistry favorable to calcification
with saturation states with respect to aragonite (Ωa) well
above 1 (average of 4.0±0.1 over the four incubations). In contrast,
during experiments at low pHT (initial pHT∼7.2),
seawater was corrosive with respect to aragonite (Ωa∼0.75). However, as pH was not regulated during the
incubations (see previous section), it increased, at lowered pH, to an
average of 7.75±0.03 (n=5) in dark conditions and to an average
of 7.84±0.03 in light conditions (n=5). Evolution of pH in
control beakers (final pHT of 7.78 and 7.48; n=1 in both the
light and the dark, respectively) showed that the observed increase in
beakers with corals was due to the additive effects of biological control
(photosynthesis minus respiration and calcification) and exchanges at the
interface in the light, and mostly due to CO2 exchange with air during
the much longer incubations performed in the dark. Assuming linear
variations with time, the average conditions of the carbonate chemistry in
the lowered pH experiments were slightly favorable to aragonite production
(Ωa=1.4±0.2 in the dark, n=5 and 1.6±0.05 in the light, n=5). Under ambient pH conditions (for both 45Ca
and 13C incubations), pH did not change during incubations in the light
(average final pHT of 8.05±0.03, n=12, data not shown)
while it decreased in the dark, due to respiration and calcification, to
reach an average pHT level of 7.62±0.07, n=12 (data not
shown). In control beakers under ambient pH, pHT slightly increased in
the light (8.09, n=2) and did not change in the dark (8.05, n=2).
Environmental conditions at the start of incubations of the coral
Stylophora pistillata. pH on the total scale (pHT), partial pressure of CO2
(pCO2, µatm), total alkalinity (AT, µmol kg-1),
dissolved inorganic carbon (CT, µmol kg-1), saturation
states with respect to aragonite (Ωa) and calcite (Ωc), and calcium concentrations ([Ca2+], µmol kg-1) are presented. Labeled seawater 45Ca activity
(Activityseawater, Bq mL-1) and the isotopic level, after
enrichment, of the seawater dissolved inorganic carbon pool (δ13C-CT, ‰) are also shown. Means ± standard deviation (SD) of analytical triplicates (duplicates for δ13C-CT) are shown when available. All incubations were conducted
at 25±0.5∘C.
45Ca activities in seawater did not change during the incubations,
reaching a final activity of 16.1±1.2 (n=12) and 28.5±0.6 (n=10) Bq mL-1 under ambient and lowered pH conditions,
respectively (including both dark and light incubations, data not shown).
Furthermore, for all incubations, these values were similar to those
measured in beakers without corals (control, data not shown). Under ambient
pH levels (no incubation at lowered pH), seawater was enriched in 13C
(δ13C-CT) from a background level of 0.26±0.05 ‰ (n=3) to 1740±4.7 ‰ (n=2) and 1634±11 ‰
(n=2) in the light and dark, respectively. During light-condition
incubations, δ13C-CT levels decreased to an average of
1636±10 ‰ (n=6, data not shown) while they
decreased to an average of 1466±24 ‰ in dark
conditions (n=6, data not shown). Incubations in control beakers
(without corals) showed that the majority of δ13C-CT loss
for both types of incubations (light and dark) was due to 13C
incorporation by corals with a minor effect of gas exchanges at the
interface (data not shown).
Both AT and [Ca2+] declined in all incubations as a consequence of
coral calcification (Table 3). Changes in AT during incubations in
control beakers (data not shown) comprised between 0.1 % and 1.1 % of
the initial level. Similar results were observed for [Ca2+] with a
relative change that comprised between 0.05 % and 1.15 % of the initial value.
These minimal changes were corroborated with no measurable changes in
seawater weight between the start and the end of all incubations (data not
shown), showing that evaporation, if any, was minimal using our experimental
setup over the considered incubation times. At ambient pH levels, decreases
in AT and [Ca2+] (average of -380±97 and -194±51µmol kg-1 for both parameters, respectively, n=24 including
both 45Ca and 13C incubations) were roughly similar under light
and dark conditions although coral specimens used for dark incubations were
ca. 166 % heavier (skeleton dry weight; see Table 1). Incubations
performed under lowered pH levels showed much lower AT and [Ca2+]
net consumption rates than under ambient pH levels. Under these pH
conditions, an extremely high AT consumption rate was observed in one
beaker (dark incubation; see Table 3) while no changes in [Ca2+] were
observed in a total of three beakers (see Table 3). These estimates (n=4) have been considered as outliers, marked with an asterisk in Table 3 and
not included in the following analyses.
Changes in total alkalinity (AT) and calcium concentrations
([Ca2+]) during the different types of incubations compared to
control beakers: ΔAT=AT2-AT2c, Δ[Ca2+]=Ca2-Ca2c, both expressed in micromoles per kilogram. Standard errors (SE) have been calculated as SEAT22+SEAT2c2 and SECa22+SECa2c2 for AT and [Ca2+], respectively, where SE corresponds to standard
errors associated with the measurement of three analytical replicates per
sample. 45Ca activity (Activitysample, Bq) and 13C
incorporation (δ13CM, ‰) of
sampled corals are also shown. Values of 45Ca activity and δ13C are mean ± standard error of the mean (SE) associated with
the measurement of three aliquots for each coral. Outliers (n=4; see
text for details) are identified with an asterisk.
45Ca activities in coral skeleton reached maximum levels under ambient
pH and light conditions (average of 87.5±9.1 Bq, n=6). Although
seawater was more enriched in 45Ca at the lower pH levels (see above),
45Ca activity in corals incubated under these conditions was much
lower, with the lowest values measured in the dark (average of 19.6±9.1 Bq, n=5). δ13C levels measured in coral skeletons (-3.69 ‰ to
8.92 ‰) showed significant enrichment compared to
background levels (-3.97±0.35 ‰, n=9).
Calcification rates using the different techniques were higher in the light
than in the dark and much lower rates were estimated at lowered pH (Table S1 in the Supplement, Figs. 2, 3 and 4). The rates measured by alkalinity anomaly (GAT)
and calcium anomaly (GCa) techniques were highly correlated (Fig. 2;
R2=0.98, p<0.01, n=34). No significant difference was
observed between rates measured by the two methods (see Table 4 for the
95 % confidence intervals of the slope and intercept). The 45Ca
method also provided rates very similar to those of the two previous approaches
(Fig. 3; GCa vs. G45Ca not shown), although the slope and the
intercept of the geometric regression between GAT and G45Ca were
significantly different from 1 and 0, respectively. Finally, the only
approach that did not provide similar rates to the others was the 13C
incorporation technique. Calcification rates based on this method were
systematically higher than those measured using the other three techniques
(see Table 4), and rates were not always significantly related (e.g., R2=0.33, p>0.05, n=12 for GAT vs. G13C; see Fig. 4; other relationships not shown).
Model II regression results of the comparison between calcification
rates estimated using the different methods considered in this study: the
alkalinity and calcium anomaly techniques (GAT and GCa,
respectively) as well as the 45Ca and 13C incorporation techniques
(G45Ca and G13C, respectively). The number of samples (n), the
regression coefficient (R2), the slope and intercept (including their
95 % confidence intervals, 95 % CI), and the p value are shown for
each comparison. Few identified outliers (n=4) have been removed from
the analyses; see Tables 3 and S1 in the Supplement.
Methods comparednR2Slope Intercept p valueValue95 % CI Value95 % CI LowHighLowHighGAT vs. GCa320.980.950.901.000.090.000.184.9×10-27GAT vs. G45Ca210.990.940.900.980.090.030.153.9×10-21GCa vs. G45Ca200.971.000.911.09-0.06-0.200.075.9×10-15GAT vs. G13C120.330.490.051.20.77-1.22.10.0506GCa vs. G13C120.320.460.031.10.94-0.92.20.0551
Calcification rates estimated based on the alkalinity anomaly
technique (GAT) as a function of calcification rates estimated based on
the calcium anomaly technique (GCa). The dashed line represents the 1:1
relationship while the full line represents the model II regression
relationship. Horizontal error bars represent standard errors (SE)
associated with the estimation of GCa. Vertical error bars representing
SE associated with the estimation of GAT are too small to be visible.
The corresponding dataset can be found in Table S1.
Calcification rates estimated based on the alkalinity anomaly
technique (GAT) as a function of calcification rates estimated based on
the 45Ca incorporation technique (G45Ca). The dashed line
represents the 1:1 relationship while the full line represents the model II
regression relationship. Horizontal error bars represent standard errors
(SE) associated with the estimation of G45Ca. Vertical error bars
representing SE associated with the estimation of GAT are too small to
be visible. The corresponding dataset can be found in Table S1.
Calcification rates estimated based on the alkalinity anomaly
technique (GAT) as a function of calcification rates estimated based on
13C incorporation technique (G13C). The dashed line represents the
1:1 relationship while the full line represents the model II regression
relationship. Horizontal error bars represent standard errors (SE)
associated with the estimation of G13C. Vertical error bars
representing SE associated with the estimation of GAT are too small to
be visible. The corresponding dataset can be found in Table S1.
Discussion
Under all experimental conditions (ambient pH vs. low pH, light vs. dark),
significant consumption rates of AT and Ca2+ as well as significant
incorporation rates of 45Ca and 13C were observed in the
zooxanthellate coral Stylophora pistillata. For all methods, calcification rates were lower in
dark than in light conditions. Such trends are expected as it has long been
established that calcification rates increase in zooxanthellate corals
during periods in which photosynthesis is occurring (Yonge, 1931), a process
known as light-enhanced calcification (e.g., Gattuso et al., 1999). Even
under lowered pH conditions, at pH levels far below those predicted to occur
in the next decades (starting pHT of ca. 7.2, average pHT during
incubations of ca. 7.5), all corals appeared to produce calcifying
structures under both light and dark conditions. The organisms selected for
this experiment were fully coated with tissues with no exposed calcareous
structures which can explain the absence of observable net dissolution such
as reported by Cohen et al. (2017) in a similar study. Since our
experimental protocol was not designed to address the potential impact of
decreasing pH levels on calcification rates of this species (no control of
carbonate chemistry during incubations, no acclimation of the organisms), we will not discuss further the observed decrease in calcification
rates identified by the three techniques used at these pH levels.
Under all experimental conditions, rates of calcification calculated using
the alkalinity and the calcium anomaly techniques were highly correlated
with a slope of 1 and no significant intercept. These results are consistent
with previously published data on colonies of Pocillopora damicornis (Chisholm and
Gattuso, 1991), Cladocora caespitosa (Gazeau et al., 2015), and several other coral
species (Murillo et al., 2014). Although the precision
obtained on Ca2+ measurements is among the highest reported to date
(Gazeau et al., 2015), the alkalinity anomaly technique appears
as the most appropriate to estimate calcification rates of isolated corals
(better precision, stronger signals). As observed by Murillo
et al. (2014), this is not true when an entire community including sediment
is investigated. The occurrence of several processes in the sediment that
can impact AT prevents the use of this technique. It is therefore
recommended to use the calcium anomaly technique when working in natural
settings, assuming that Ca2+ concentrations are measured with an
analytical technique as precise as the one used in our study (CV < 0.05 %). Similarly, although corrections are possible when applying the
alkalinity anomaly technique on organisms that significantly release
nutrients (echinoderms, bivalves, etc.), the use of the calcium anomaly
technique is highly recommended instead (Gazeau et al., 2015).
Calcification rate estimates based on changes of AT or Ca2+ were
highly correlated with estimates based on 45Ca incorporation in corals.
These results are not consistent with those reported by
Smith and Kinsey (1978), Tambutté et al. (1995), and
Cohen et al. (2017). These studies revealed discrepancies between
the alkalinity anomaly and the 45Ca incorporation techniques. Smith and
Kinsey (1978) found that rates measured with the 45Ca method were higher than
those measured using the alkalinity anomaly technique (significant 45Ca
incorporation at ΔAT=0). Results from both
Tambutté et al. (1995) and Cohen et al. (2017)
suggested the opposite with a decrease in AT consumption without any
concomitant 45Ca incorporation. A number of reasons may explain these
discrepancies. First, the present study is the first one comparing these
techniques in the same incubations, in contrast to the other ones in which
incubations for AT anomaly and 45Ca incorporation were performed
over 2 consecutive days (due to radioactive contamination issues). Second,
calcification expressed as absolute changes in AT during incubations,
measured during our experiment, were at least 1 order of magnitude higher
than measured during these studies (44 200 to 745 600 nmol vs. less than
4000 nmol in previous experiments). Cohen et al. (2017) have shown
that such discrepancies were much higher at very low rates and that the
ratio between rates estimated based on 45Ca incorporation and AT
consumption were getting closer to 1 with increasing calcification rates.
Nevertheless, even at the highest levels of calcification computed during
these studies, 45Ca-based rates were still significantly different from
ΔAT-based rates, which is in contrast with our results.
As already mentioned, although calcification rates of the present study were
lower at lowered pH levels, there was still a close to perfect agreement
between the different techniques. While the 45Ca labeling technique is
thought to provide rates of gross calcification, there is no doubt that both
the AT and Ca2+ anomaly techniques allow the estimation of net
calcification rates (gross calcification – dissolution). A full agreement
of rates computed from these methods further suggests that no dissolution of
previously precipitated CaCO3 structures occurred during our study,
even under lowered pH conditions. The corals used in our experiment were
fully covered with tissues, which is likely the reason why no dissolution was
measured.
Furthermore, we must note that the protocol for 45Ca incorporation
considered in our study differed from the one used in the abovementioned
past studies. A much smaller activity was used (0.025 kBq mL-1)
compared to that of Tambutté et al. (1995; 40 kBq mL-1) and Cohen et al. (2017; 9 kBq mL-1). Moreover, in contrast to Cohen et al. (2017), rates
were not corrected for 45Ca incorporation on the skeleton of dead
corals. This choice was motivated by the absence of detectable radioactivity
on bare skeletons exposed for 7 h and treated with the same protocol as the
one used in our study (Chantal Lanctôt, personal communication, 2018).
To the best of our knowledge, this is the first study comparing
calcification rates measured using the 13C labeling technique to the
more widely used alkalinity and calcium anomaly techniques. It shows that
13C-derived rates were systematically higher and much more variable
(with large uncertainties) than the ones estimated using the two other
techniques. As already mentioned, several studies have shown that most of
the carbon precipitated in the skeleton comes from coral and its symbiotic
zooxanthellae (e.g., Erez, 1978; Furla et al., 2000),
leading to an underestimation of calcification rates based on labeled,
radioactive carbon incorporation. As there is no reason for 13C to
behave differently, our results appear inconsistent with a metabolic source
of carbon. As the nubbins were treated following the same protocol as for
45Ca incorporation measurements, it is unclear why much stronger
13C incorporation was obtained and why variability was so high. Before
better insights into such discrepancies can be developed, we recommend to
avoid this technique to estimate coral calcification rates.
Incubation times (tmin; h) necessary to obtain significant
signals using the three methods: the alkalinity anomaly technique
(AT), the calcium anomaly technique (Ca2+), and the 45Ca
incorporation techniques (45Ca); see text for calculation procedures.
tmax (h) is the maximum incubation time to maintain carbonate chemistry
within an acceptable range (ΔpHT<0.06 and ΔCT<10 % and ΔAT<10 %). The ratios between incubation volume (V, in milliliters) and the size of the
nubbins (S, in centimeters), considered in our study for the different sets of
incubations (ambient pH vs. low pH; light vs. dark), are also shown.
tmin values are noted in bold when higher than tmax.
Ratio V : Stmin (h) tmax (h)ATCa2+45CaAmbient pH – light77–950.261.000.64.7Ambient pH – dark59–690.332.101.51.3Lowered pH – light109–1211.256.151.10.5Lowered pH – dark95–1091.6011.203.41.3
Our study was designed to compare different techniques to estimate
calcification rates and not to define the best experimental approach to
study the effects of ocean acidification on coral species using these
different approaches. As such, the chosen experimental protocol (e.g.,
incubation times) was not optimal and led, in some cases, to significant
changes in the carbonate chemistry during incubations. However, our results
provide some insights that we further discuss in the following section.
Measuring and comparing calcification rates of organisms under varying pH
conditions requires the careful choice of a volume and a time interval such
that the precision of the calcification rate measurement is large enough to
observe significant signals and that the change in carbonate chemistry
parameters between the beginning and end of the incubation is small compared
to the range of these parameters in the different treatments (Langdon et
al., 2010). Table 5 illustrates the incubation time necessary to obtain
measurable changes for each method (tmin) considering the ratio between
incubation volume and coral size chosen for our study. As the 13C
incorporation method did not provide reliable rates, this technique was not
considered in this analysis. The threshold for significant signals was set
at 10-fold the analytical precision of the instruments (Langdon et al.,
2010) for AT and Ca2+ measurements (1.2 and 2.9 µmol kg-1, respectively) and above the detection limit of 15 CPM for
45Ca activity estimated. Maximum incubation times are more difficult to
estimate. Langdon et al. (2010) and Riebesell et al. (2010) recommend
considering incubation times short enough to maintain AT and CT
within an acceptable range (ΔAT and ΔCT<10 %). As it is more difficult to estimate what changes in pH are
acceptable, we have arbitrarily considered a maximal change in pH of 0.06,
corresponding to the lowest change in global surface ocean pH projected for
2100 (IPCC, 2014). Maximal incubation times, as presented in Table 5
(tmax), correspond then to incubation times that should not be exceeded
in order to maintain acceptable conditions of the carbonate chemistry
(ΔpHT<0.06 and ΔAT<10 % and
ΔCT<10 %).
Under light and ambient pH conditions, even if the ratio between incubation
volume and nubbin size is much higher than for previous similar studies
(e.g., Cohen et al., 2017), all methods would allow a precise estimation of
calcification rates over very short incubation times (∼15 min
to 1 h, depending on the method) while leading to moderate changes in
carbonate chemistry. In the dark, and under ambient pH conditions, in the
absence of pH increase due to photosynthesis, the decrease in pH due to
respiration narrows the possible incubation period to 1.3 h. While this is
still larger than the incubation time allowing us to obtain a significant
signal with the alkalinity anomaly technique (∼20 min), the
other two methods necessitate longer incubation times to obtain precise
estimates (>1.5 h). At lower pH, under both light and dark
conditions, and using open systems without a continuous pH regulation as in
our study, it is obvious that all techniques are not well adapted to this
experimental protocol. Indeed, as a consequence of lower calcification rates
at lower pH and significant CO2 degassing, incubation times necessary
to obtain significant signals using these techniques are too large to
maintain the carbonate parameters within an acceptable range. This is not
insurmountable as a continuous regulation of pH using for instance pure
CO2 bubbling or incubations performed in a closed container (i.e.,
without contact with the atmosphere) would alleviate these problems.
In conclusion, the present study is the first one allowing a direct (i.e.,
during the same incubations) comparison of three methods used to estimate
coral calcification rates, the calcium and alkalinity anomaly techniques and
the 45Ca incorporation technique. These methods provided very
consistent calcification rates of the coral Stylophora pistillata independently of the conditions
set for the incubations (light vs. dark, ambient vs. low pH). Among these
three methods, the alkalinity anomaly and the 45Ca incorporation
techniques appear to be the most sensitive, allowing the quantification of
coral calcification rates without significant changes in targeted
environmental conditions. In contrast, the 13C incorporation technique
did not provide reliable calcification rates and its use is not recommended
until further investigations clarify the discrepancies. Finally, this study
was restricted to a single coral species and used nubbins fully covered with
tissues. Conducting similar comparison studies with other coral species as
well as other major calcifying groups widely studied in the context of ocean
acidification (e.g., coralline algae, mollusks) would be
necessary for a better understanding of ocean acidification impacts on
ecosystem services provided by calcifying organisms.
Data availability
All data used in this manuscript are freely available at: https://doi.pangaea.de/10.1594/PANGAEA.912222 (Gómez Batista et al., 2020).
The supplement related to this article is available online at: https://doi.org/10.5194/bg-17-887-2020-supplement.
Author contributions
FG and MM designed and supervised the study. MGB conducted the research, and MGB and FG wrote the paper with contributions from all authors.
Competing interests
The authors declare that they have no conflict of interest.
Acknowledgements
We thank the
Monaco government and the Centre Scientifique de Monaco for propagating and
maintaining the coral nubbins and Samir Alliouane for technical assistance
for total alkalinity and calcium measurements.
The authors are also grateful to the editor and two anonymous reviewers whose comments and suggestions helped improve the paper.
Financial support
This work was supported by the IAEA's Ocean Acidification International Coordination Center (OA-ICC) and the IAEA-ICTP Sandwich Training Educational Programme (STEP) and the project “Strengthening the National System for Analysis of the Risks and Vulnerability of Cuban Coastal Zone Through the Application of Nuclear and Isotopic Techniques” National Program PNUOLU/4-1/2 No. /2017 of the National Nuclear Agency (AENTA).
Review statement
This paper was edited by Lennart de Nooijer and reviewed by two anonymous referees.
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