Introduction
Since the industrial revolution the surface ocean pH has dropped by 0.1 units
and is predicted to drop another 0.3–0.5 units by 2100 (Caldeira and
Wickett, 2005; Orr et al., 2005; IPCC, 2013). This is due to the increasing
amount of atmospheric carbon dioxide (CO2) absorbed by the ocean that
extensively affects seawater carbonate chemistry (e.g. Caldeira and Wickett,
2003, 2005; Orr et al., 2005; Feely et al., 2004). Increased concentrations
of anthropogenic CO2 are reflected in an elevated concentration of
hydrogen ions, which lowers the pH and the availability of carbonate ions.
Effects on marine organisms are of great scientific interest for
understanding the geological past and the consequences in the immediate
future (e.g. Ries et al., 2009), since the decrease in calcium carbonate
saturation potentially threatens marine organisms forming biogenic calcium
carbonate (e.g. Orr et al., 2005; Guinott et al., 2006; McCulloch et al.,
2012; Jantzen et al., 2013a, b). This applies to calcium carbonate
shell-forming species, such as brachiopods and mollusks, because they are
considered excellent archives documenting changes in environmental conditions
affecting marine organisms (e.g. Kurihara, 2008; Comeau et al., 2009; Hahn et
al., 2012, 2014; Watson et al., 2012; Cross et al., 2015, 2016, 2018; Crippa
et al., 2016a; Milano et al., 2016; Garbelli et al., 2017; Jurikova et al.,
2019).
Recently, several experiments were performed to investigate whether a change
in seawater pH may affect growth rate, shell repair and oxygen consumption of
calcifying organisms, and how they respond, in general, to ocean
acidification (Supplement Table S1). However, despite the great number of
studies, the consequences for biomineral formation remain not well
understood, as most studies focused mainly on growth, metabolic rates, shell
dissolution and shell repair (Table S1, and references therein). Only a few
studies deal with the effect of acidification on microstructure (Beniash et
al., 2010; Hahn et al., 2012; Stemmer et al., 2013; Fitzer et al., 2014a, b;
Milano et al., 2016), and most of them focused on bivalves and show that
neither microstructure nor shell hardness seem to be affected by seawater pH.
The few studies that examined brachiopods or brachiopod shells suggest that
the latter suffered increased dissolution under lower seawater pH. In other
studies, the organism either exhibited no changes or an increase in shell
density (calculated as dry mass of the shell (g)/shell volume (cm3)),
but otherwise no changes in shell morphology and trace chemistry (Table 1).
Cross et al. (2018) found that punctae became narrower over the past 120
years, which partially explained the increase in shell density over this
period. Overall, there appears to be little to no apparent effect on
brachiopod morphology or chemistry with lower seawater pH (Cross et al.,
2015, 2016, 2018).
Culturing, dissolution experiments and natural variation on several
brachiopod species and shells.
Species n (number of sample)
Growth parameters
Shell repair/microstructure/oxygen consumption/dissolution of shell/microstructure
Method and material
Environment/conditions T: temperature (∘C) S: salinity (PSU) pCO2 (µatm)
Duration of experiment
Source
Calloria inconspicua (Sowerby, 1846) n= 123
(1) > 3 mm in length undamaged individuals were not affected by lower pH; (2) < 3 mm in length undamaged individuals grew faster at pH 7.62 than the control conditions
Shell growth rates and shell repair frequencies were not affected by low pH (> 80 % of all damaged individuals repaired after 12 weeks)
Culture experiment
(a) pH 8.16, T 16.5, S 33.9, pCO2 465, Ωcalcite 3.5 (b) pH 7.79, T 16.9, S 33.9, pCO2 1130, Ωcalcite 1.6 (c) pH 7.62, T 16.6, S 33.9, pCO2 1536, Ωcalcite 1.3
12 weeks
Cross et al. (2016)
Calloria inconspicua (Sowerby, 1846) nadult= 389 for shell morphology analyses*
Punctae width decreased by 8.26 %, shell density increased by 3.43 %, no change in shell morphology, punctae density, shell thickness, and shell elemental composition (Ca, Mg, Na, Sr and P) No changes were found in shell dissolution over the last 120 years.
Collected every decade from one locality
Last 2 decades pH reduced 0.1 unit Temperature varied from 10.7 to 13.0 ∘C pCO2 varied from 320 to 400 Salinity and Ωcalcite not provided
120-year record
Cross et al. (2018)
Liothyrella uva (Broderip, 1833) n=156
Not affected by lower pH and temperature
Shell repair frequencies were not affected by low pH and temperature (> 83 % of individuals repaired after 7 months)
Culture experiment
(a) pH 7.98, T-0.3, S 35, pCO2 417, Ωcalcite 1.20 (b) pH 8.05, T 1.7, S 35, pCO2 365, Ωcalcite 1.49 (c) pH 7.75, T 1.9, S 35, pCO2 725, Ωcalcite 0.78 (d) pH 7.54, T 2.2, S 35, pCO2 1221, Ωcalcite 0.50
7 months
Cross et al. (2015)
Liothyrella uva (Broderip, 1833) npost-mortem=5
Not applicable
Higher dissolution in gastropods and brachiopods at lower pH after 14 days
Empty shells
(a) pH 7.4, T 4, S 35, Ωcalcite 0.74 (b) pH 8.2, T 4, S 35, Ωcalcite 4.22 pCO2 not provided
14 to 63 days
McClintock et al. (2009)
* A subsample of 40 brachiopods (two to five specimens per
decade over the last 120 years) were used for further shell analysis of shell
density, punctal width, punctal density, shell dissolution, shell thickness
and shell elemental composition.
Brachiopods possess a low-magnesium calcite shell, which should be more
resistant compared to the more soluble forms of CaCO3 such as aragonite
and high-Mg calcite (Brand and Veizer, 1980; Morse et al., 2007). The shell
microstructure of Rhynchonelliformean brachiopods has been used as a
powerful tool to understand the biomineral's response to modern global ocean
acidification and similar events in the geologic past (Payne and Clapham,
2012; Cross et al., 2015, 2016; Garbelli et al., 2017). A comprehensive
study focusing on fossil brachiopods during the end-Permian mass extinction
showed that brachiopods produce shells with increased organic matter content
during ocean acidification events (Garbelli et al., 2017).
Here, we describe the microstructure and carbon and oxygen isotopic
composition of brachiopod shells belonging to the cold–temperate water
species Magellania venosa (Dixon, 1789) grown in natural
environments as well as under pH-controlled culturing conditions. M. venosa represents the largest recent brachiopod species, is often locally
abundant (e.g. in Chile; Försterra et al., 2008), and has the highest
growth rate recorded for recent brachiopods (Baumgarten et al.,
2014). Its low-magnesium calcite shell consists of a microgranular primary
layer and a fibrous secondary layer (Smirnova et al., 1991; Baumgarten et
al., 2014; Casella et al., 2018; Romanin et al., 2018) crossed by
perforations – endopunctae.
Since little is known about the microstructural and geochemical responses of
brachiopods to increased ocean acidification, the main goal of this study is
to document any changes in this highly important archival marine organism. We
will describe whether and how shell microstructural features such as the
primary layer thickness, density of endopunctae and fibre morphology,
together with their stable carbon (δ13C) and oxygen
(δ18O) isotope compositions, respond to low seawater pH
conditions.
Materials and methods
Brachiopod samples and culturing set-up
A full description of the brachiopod sampling and culturing procedure is
provided in Jurikova et al. (2019), but we provide an abbreviated
version and reiterate the key points. Nine adult individuals of M. venosa (Dixon, 1789) were chosen for microstructure investigation and
evaluation of their δ13C and δ18O values. All
specimens were collected by scientific SCUBA divers alive from 20 m water
depth of Comau Fjord (Chile) at different localities (Fig. 1). Specimens no. 158
and no. 223 did not experience any treatment after collection from Comau Fjord.
The other specimens, no. 43, no. 63, no. 8004, no. 8005, no. 9004, no. 9005
and no. 9006, were cultured under different pH conditions at either AWI in
Bremerhaven or GEOMAR (at KIMOCC – Kiel Marine Organisms Culture Centre) in
Kiel, Germany (Tables 2 and 3).
Culture and sensor systems used in M. venosa culturing
(specimens: no. 43, no. 63, no. 8004, no. 8005, no. 9004, no. 9005 and no.
9006). Operated under controlled experimental settings in climate-controlled
laboratories at Alfred-Wegener-Institut Helmholtz-Zentrum für Polar- und
Meeresforschung, Bremerhaven, Germany, and at GEOMAR Helmholtz-Zentrum
für Ozeanforschung Kiel, Germany.
Culture system at AWI
Automated sensor systems at AWI
Culture system at GEOMAR
Automated sensor systems at GEOMAR
Aquarium (150 L/each pHtreatment)
Aquarium (150 L/each pH treatment)
Supplied from a reservoir tank (twice a week 20 % water was replaced)
Supplied from a reservoir tank (twice a month 10 % water was replaced)
Temperature
Controlled in temperature constant room
Controlled using heaters or coolers
Temperature sensor pond
pCO2
Bubbling of CO2
COMPORT, Dennerle, Vinningen; IKS aquastar Aquarium computer V2.xx with Aquapilot 2011
Bubbling of CO2-enriched air
CONTROS HydroC® underwater CO2 sensor
Salinity
Mixing Reef commercial sea salt (until October: Aqua Medic, Bissendorf, Germany, thereafter Dupla Marin Reef Salt, Dohse Aquaristik, Grafschaft-Gelsdorf, Germany) with deionized water
Conductivity electrode
Mixing Tropic Marin Pro–Reef commercial sea salt with deionized water
Conductivity electrode
Filtering
Biofilter, protein skimmer and UV sterilizer
Biofilter, protein skimmer and UV sterilizer
Food
Regularly fed (typically five times per week) with Dupla Rin, Coral Food, Reef Pearls 5–200 µm, alive Thalassiosira weissflogii, and 1-day old nauplii of Artemia salina
Regularly fed (typically five times per week) with Rhodomonas baltica
Substrate
Sabia Corallina, 7–8 mm, Dohse Aquaristik, Grafschaft-Gelsdorf, Germany
No
Specimens of M. venosa sampled from Comau
Fjord, Chile, and natural and experimental culturing conditions.
Sample ID
Sample locality at Comau Fjord (Chile)1
Sample seawater conditions2
Date of collection
Length of ventral valve (mm)
Duration of experiment
Experimental conditions
No. 43
Lilliguapi
pH: ∼7.9 T:∼13 S: ∼32
D: 20
Feb 2012
37
214 days3
pCO2: 1390, pH: 7.66±0.04 T: 11.6±0.5, S: 32.6 Ωcal: 2.0
No. 63
Lilliguapi
pH: ∼7.9 T: ∼13 S: ∼32 D: 20
Feb 2012
23
214 days3
pCO2: 2600, pH: 7.44±0.08 T: 11.7±0.5, S: 32.7 Ωcal: 1.4
No. 158
Huinay Dock
pH: ∼7.9 T: ∼13 S: ∼32 D: 20
Dec 2011
36
no
No. 223
Cahuelmó
pH: ∼7.9 T: ∼13 S: ∼32 D: 23
Feb 2012
30
no
No. 8004
Comau Fjord
pH: ∼7.9 T: ∼13 S: ∼32 D: 21
Apr 2016
31
335 days4
pCO2: 600 pH: 8.00 to 8.15±0.05 T: 10±1, S: 30 Ωcal: 2.0–3.5
No. 8005
Comau Fjord
pH: ∼7.9 T: ∼13 S: ∼32 D: 21
Apr 2016
46
335 days4
pCO2: 600 pH: 8.00 to 8.15±0.05 T: 10±1, S: 30 Ωcal: 2.0–3.5
No. 9004
Comau Fjord
pH: ∼7.9 T: ∼13 S: ∼32 D: 21
Apr 2016
41
335 days4
pCO2: 2000–40005 pH: 7.60 to 7.35±0.05 T: 10±1, S: 30 Ωcal: 0.6–1.1
No. 9005
Comau Fjord
pH: ∼7.9 T: ∼13 S: ∼32 D: 21
Apr 2016
25
335 days4
pCO2: 2000–40005 pH: 7.60 to 7.35±0.05 T: 10±1, S: 30 Ωcal: 0.6–1.1
No. 9006
Comau Fjord
pH: ∼7.9 T: ∼13 S: ∼32 D: 21
Apr 2016
43
335 days4
pCO2: 2000–40005 pH: 7.60 to 7.35±0.05 T: 10±1, S: 30 Ωcal: 0.6–1.1
Note: D: depth (m), T: temperature (∘C), S:
salinity (PSU – practical salinity units), pCO2 (µatm).
1 Cahuelmó 42∘15′23′′ S, 72∘26′42′′ W,
Cross–Huinay 42∘23′28′′ S, 72∘27′27′′ W, Jetty
(Huinay Dock) 42∘22′47′′ S, 72∘24′56′′ W,
Lilliguapi 42∘9′43′′ S, 72∘35′55′′ W; sample nos.
8004, 8005, 9004, 9005, and 9006 were harvested from three sites in Comau
Fjord (Cross-Huinay, Jetty, and Liliguapy), Chile. 2 Reference: Laudien
et al. (2014) and Jantzen et al. (2017). 3 Culture experiments conducted
at Alfred-Wegener-Institut Helmholtz-Zentrum für Polar- und
Meeresforschung, Bremerhaven, Germany. 4 Culture experiments conducted
at GEOMAR Helmholtz-Zentrum für Ozeanforschung Kiel, Germany (Jurikova et
al., 2019). 5 CO2 concentration was changed during the
experiment: to 2000 µatm from 4 August 2016 to 18 April 2017 and to
4000 µatm from 18 April to 5 July 2017.
Map of Comau Fjord. (a) Overview of Chilean Patagonia.
(b) Gulf of Ancud with connections in the north and south to the
Pacific Ocean. (c) Comau Fjord with brachiopod sample collection
localities. In both maps the rectangle marks the location of Comau Fjord.
In summary, individuals of M. venosa were collected alive in Chile
and transported to GEOMAR, Germany, in plastic bags filled with seawater, and
maintained under controlled conditions in a climate laboratory. The
brachiopods were first acclimatized under control conditions for 5 weeks, and
prior to the start of the experiment were labelled with a fluorescent dye –
calcein (Sigma, CAS 1461–15–0; 50 mg L-1 for 3 h) (e.g. Baumgarten
et al., 2014; Jurikova et al., 2019). As a culture medium we used artificial
seawater, which was prepared by mixing a commercial salt with deionized water
until the desired salinity and chemical composition were achieved (Atkinson
and Bingman, 1998). An overview of the culturing set-up at both laboratories
is available in Table 2. Specimens no. 43 and no. 63 were cultured at AWI at
pH = 7.66 (pCO2 = 1390 µatm) and pH = 7.44
(pCO2 = 2610 µatm) from 29 August 2013 to
31 March 2014, respectively. Specimens nos. 8004, 8005, 9004, 9005 and 9006 were
cultured concurrently at GEOMAR under control or low-pH conditions. Specimens
no. 8004 and no. 8005 were maintained under pH settings of 8.0–8.15 from
4 August 2016 to 5 July 2017, conditions similar to those of their fjord
habitat. In contrast, specimens nos. 9004, 9005 and 9006 were cultured under
low-pH artificial seawater conditions. Low-pH conditions were achieved by
bubbling of CO2 through the tanks at AWI, and by bubbling
CO2-enriched air through the tanks at GEOMAR (Table 2). The
acidification experiment at GEOMAR was performed in two phases; the first one
from 4 August 2016 to 18 April 2017 during which the pCO2 was set
to 2000 µatm (corresponding to a pH of 7.60), and the second one
during which the pCO2 was set to 4000 µatm (corresponding
to a pH of 7.35) from 18 April 2017 to 5 July 2017 (Table 3). In order to
distinguish between the shell parts precipitated under the specific pH
conditions as well as to allow exact comparison to shells of the control
treatment, calcein marking was also carried out prior to the second low-pH
experiment of 4000 µatm. Parts of the shell grown under specific pH
conditions are indicated in Fig. 2. In addition to the calcein marking, newly
grown shell parts may be distinguished from visible growth lines on the
surface of the shell (Fig. 2). The total length (maximum distance from the
blue line to the anterior margin) of the curved dorsal and ventral valves
grown during 11 months of culturing (Fig. 2) varied from < 5 to
15.6 mm (Table 4).
Growth tracked with calcein and marked by blue and red lines
on the surface of the brachiopod specimens (no. 9006).
Shell length of specimens of M. venosa
before and during the culture.
Sample ID
Valve
Initial length
New shell
pH
New shell
pH
before culturing
growtha
growthb
(mm)
257 days (mm)
78 days (mm)
No. 8004
ventral
15.4
14
8.00
1.6
8.15
No. 8005
ventral
40
5
8.00
< 1
8.15
No. 8005
dorsal
36
4
8.00
< 1
8.15
No. 9004
ventral
26.8
13
7.60
1.2
7.35
No. 9005
ventral
11.2
12
7.60
1.8
7.35
No. 9006
ventral
33
9
7.60
< 1
7.35
No. 9006
dorsal
29
8
7.60
< 1
7.35
Note: a culturing from 4 August 2016 to
18 April 2017; b culturing from 18 April 2017 to 5 July 2017.
Microstructural analysis
This study followed the sample preparation method shells suggested by Crippa
et al. (2016b). In order to obtain more detailed data on microstructural
changes, the samples were cut with a diamond blade along different axes and
directions (Fig. 3a). Subsequently, the samples were immersed in 36-volume
hydrogen peroxide (H2O2) for 24/48 h to remove the organic
tissue. The sectioned surfaces were manually smoothed with 1200 grit
sandpaper, then quickly (3 s) cleaned with 5 % hydrochloric acid (HCl),
immediately washed with water and air-dried. The time of acid etching was
kept short so as not to affect the microstructure (Crippa et al., 2016b).
Finally, the valve sections were gold-coated and analysed by a Cambridge
S-360 scanning electron microscope with a lanthanum hexaboride (LaB6)
source operating at an acceleration voltage of 20 kV (Dipartimento di
Scienze della Terra “A. Desio”, Università di Milano).
Brachiopod shell sample cut along different axes.
(a) Longitudinal and transverse sections; (b) transverse
sections at the anterior margin of the shell; (c) plane grinding of
the external surface of the shell.
The methods described by Ye et al. (2018a) were followed to investigate the
basic microstructural units (fibres) in SEM images. We focused primarily on
the anterior margin of the valves, the part that was produced during
culturing (hereinafter referred to as during-culturing) under different pH conditions.
Therefore, additional transverse sections along the growth lines were
obtained in the most anterior part (black lines in Fig. 3b) by manually
smoothing with 1200 grit sandpaper. Plane grinding was performed on the
external surface of the shell (Fig. 3) to investigate the distribution of
endopunctae.
Measurement methods used for the thickness of the primary
layer (a) and the density of the endopunctae (b). Note that
for the latter, endopunctae were counted when included for more than their
half-diameter inside the square. dv: dorsal valve.
The thickness of the primary layer was measured on the SEM images of
specimens no. 8005 and no. 9006 (Fig. 4a) in different positions along the
longitudinal growth axis (posterior, central and anterior regions). In the
vicinity of the transition from natural growth to cultured growth, the region
was further subdivided into four sub-zones.
To calculate and measure the density and diameter (max) of endopunctae,
squares (800 µm × 800 µm) were located randomly
over the smoothed external surface of the anterior shell (Fig. 4b). Four
sub-zones (C2, A1, A2, A3) were defined according to their position along the
posterior–anterior direction (Fig. 4), while distinguishing the part of the
shell produced before-culturing and that produced
during-culturing.
For morphometric analyses, fibres were manually outlined using polygonal
lasso in Adobe Photoshop CS6, and size and shape parameters were measured
with Image-Pro Plus 6.0 and ImageJ (for convexity). In particular, following
Ye et al. (2018a, b) we measured/calculated the Feret diameter (max), area,
roundness (4area/π× Feret diameter (max)2) and convexity
(convex perimeter / perimeter). The width of an individual fibre roughly
corresponds to its Max Feret diameter, whereas its height corresponds to the
Min Feret diameter (see Fig. 6 in Ye et al., 2018a).
As individual fibres are irregular in shape in the most anterior section of
brachiopods, the morphometric measurement method proposed by Ye et
al. (2018a, b) is not always suitable. Thus, modifications had to be made to
the Ye et al. (2018a, b) measurement method to make the comparative
morphometric analysis of the fibres from the anterior part (Fig. 5a, b).
First, all SEM images were oriented in the same direction with the base of
the primary layer facing upwards. Then a uniformly sized zone
(20 µm × 20 µm) was selected for additional
measurements with the upper side of the square always placed at the boundary
between the primary and secondary layers (Fig. 5c). Two new methods were
developed and applied: for Method 1, the width of fibres crossed by two
standard lines was measured, which were always located in the same position
and at the same distance in all the selected zones (yellow and orange lines
in Fig. 5, Method 1). For Method 2, we calculated the number of boundaries based on the
number of fibres crossed by the two standard lines (Fig. 5, Method 2).
Sub-zones were named according to the following nomenclature: the most
anterior transection zone of the ventral valve was named Z1, the second most
anterior transection zone of the ventral valve Z2, and so on; the most
anterior transection zone of the dorsal valve was named Z4. The standard line
facing the primary layer was named “1” and the second standard line “2”
(example: “Z1-1” is the sample of the standard line facing the primary
layer at the most anterior transect zone of the ventral valve).
Methods of measurements used in the anterior transverse sections.
All SEM images are oriented in the same direction: base of the primary layer
facing upwards. A square (20 µm × 20 µm) was
analysed with its upper side just overlapping the boundary between the
primary and secondary layers. Method 1 refers to the measurement of the width
of the fibres crossed by two standard lines, which were located in the same
position and at the same distance in all 194 squares (yellow and orange
lines); Method 2 involved the calculation of the numbers of boundaries
between the fibres that are crossed by two standard lines. vv: ventral
valve.
Stable isotope analyses of shells
Cleaned shells of specimens nos. 8004, 8005, 9004, 9005 and 9006 were chosen for
carbon and oxygen isotope analyses. For specimens no. 8005 and no. 9006, surface
contaminants and the primary layer were first manually and then chemically
removed by leaching with 10 % HCl, rinsed with distilled water and
air-dried. As the primary layer is not secreted in equilibrium with ambient
seawater (e.g. Carpenter and Lohmann, 1995; Brand et al., 2003, 2013), it is
important to chemically remove it in order to avoid cross-contamination of
results. Individual growth increments exclusively come from the secondary
layer, and were separated from the shell in both dorsal and ventral valves
using a WECHEER (WE 248) microdrill at low speed with a tungsten–carbide
milling bit. Shell increment fragments, of similar width, were then powdered
using an agate mortar and pestle. For carbon and oxygen isotope analyses
about 250 µg of powdered calcite of each sample was analysed using
an automated carbonate preparation device (GasBench II) connected to a Delta
V Advantage (Thermo Fisher Scientific Inc.) isotopic ratio mass spectrometer
at the Earth Sciences Department, University of Milan, Italy. The carbon and
oxygen isotope compositions are expressed in the conventional delta notation
calibrated to the Vienna Pee-Dee Belemnite (V-PDB) scale by the international
standards IAEA 603 (International Atomic Energy Agency 603;
δ18O: -2.37±0.04 ‰, δ13C: +2.46±0.01 ‰) and NBS 18 (δ18O: -23.2±0.1 ‰, δ13C: -5.014±0.035 ‰).
Analytical reproducibility (1σ) for these analyses was better than
±0.04 ‰ for δ13C and ±0.1 ‰ for
δ18O (Appendix A). Another set of shells, no. 8004, no. 9004
and no. 9005, were gently rinsed with ultra pure water (Milli-Q) and dried
for a few days on a hotplate at 40 ∘C in a clean flow hood. Targeted
parts of the shell were sampled for powder under binoculars using a precision
drill (Proxxon) with a mounted dental tip. Stable isotope analyses of powders
of these specimens were performed at GEOMAR, Kiel, on a Thermo Finnigan MAT
252 mass spectrometer coupled online to an automated Kiel carbonate
preparation line. The external reproducibility (1σ) of in-house
carbonate standards was better than ±0.1 ‰ and ±0.08 ‰ for δ13C and δ18O,
respectively (Appendix A).
Stable isotope analyses of water samples
In addition to carbon and oxygen isotope analyses of shells, analyses were
also carried out on seawater samples collected from the culturing tanks.
Measurements of δ13CDIC and δ18OH2O were
performed using Thermo Scientific™ Delta Ray™ IRIS
with URI Connect.
Isotope values (δ13C, δ18O) are reported as per
mil (‰) deviations of the isotopic ratios (13C/12C,
18O/16O) calculated to the VPDB scale for δ13C
and VSMOW scale for δ18O values. Analytical reproducibility
(1σ) on three aliquots of each water sample was ≤0.03 ‰
for both δ13C and δ18O values (Appendix B).
Discussion
Microstructure and organic component relationship
Before discussing whether and how acidification may affect the microstructure
of the brachiopod shell, it is important to examine the relationship between
the microstructure and the number of organic components within the shell. It
has already been stated that, in fossil and recent brachiopods, different
shell microstructures have different numbers of shell organic components
(Garbelli et al., 2014; Garbelli, 2017; Casella et al., 2018; Ye et al., 2018a).
This holds true for most rhynchonelliformean brachiopods: the primary layer
of M. venosa consists of acicular and granular calcite (Williams,
1968, 1973, 1997; MacKinnon and Williams, 1974; Williams and Cusack, 2007;
Casella et al., 2018). Analyses of electron backscatter diffraction show that
the primary layer is a thin nanocrystalline layer with higher micro-hardness
and smaller-sized calcite crystallites compared to those of the secondary
layer (Griesshaber et al., 2004). In addition, each spherical and small unit
is coated by a mixture of organics and amorphous calcium carbonate (Cusack et
al., 2010). This, per se, may suggest a higher amount of organic
components associated with the primary layer in contrast to other shell
layers (i.e. secondary or in some species tertiary layer), but it has never
been proven.
In fossils, the primary layer is likely absent or, if present, diagenetically
altered, and it will luminesce (Grossman et al., 1991), suggesting that
higher amounts of organics may be present. However, this has also been
ascribed to the incorporation of magnesium into the lattice (Popov et al.,
2007; Cusack et al., 2008). A report of higher sulfur concentration
in the primary layer of the brachiopod Terebratulina retusa may
suggest the presence of a sulfur-rich organic component, but backscatter
electron imaging revealed contradictory results (England et al., 2007).
Cusack et al. (2008) showed that, in the same species, the sulfate
concentration is higher in the primary layer than in the secondary layer.
Since there is no conclusive evidence for this observation, we cannot relate
the increase in thickness of the primary layer to changes in organics within
the shell. With respect to previous findings (Williams, 1966; Parkinson et
al., 2005), our results show that the thickness of the primary layer of
M. venosa is much less uniform and shows an increase with growth,
which is more evident during-culturing under low-pH conditions. However,
disturbances (stress condition with handling before and at the start of the
culturing) may also cause an abrupt change in thickness.
Endopunctae, which during life are filled with mantle expansions, are widely
distributed in the shell of M. venosa and show the superficial
hexagonal close-packing pattern documented by Cowen (1966). The biological
function of endopunctae is still controversial, with some suggesting that,
generally in living organisms, they serve as support and protection
structures (Williams, 1956, 1997), as sensors, or as storage and respiration
features (Pérez-Huerta et al., 2009). With more endopunctae filled by
mantle expansions, the amount of organic tissue would increase in the same
volume of shell. The density of endopunctae has been related to temperature,
as species living at higher temperatures have greater endopunctae density
(Campbell, 1965; Foster, 1974; Peck et al., 1987; Ackerly et al., 1993). The
present analyses support the concept that the increase in endopunctae density
may be related in part to ontogeny and to low-pH conditions. This may be
expected, as organisms living under low-pH conditions have to up-regulate
their internal pH to be able to calcify, as demonstrated in M. venosa by Jurikova et al. (2019) and also observed in other calcifiers such
as corals (McCulloch et al., 2012; Movilla et al., 2014). This would require
a higher energy cost and a larger respiration/storage surface would satisfy
this requirement.
In addition to the thickness of the primary layer and the density of the
endopunctae, the size changes in the individual fibres within the fibrous
secondary layer may also contribute to the variability in organic components.
Most of the recent rhynchonelliformean brachiopods, and M. venosa in
particular, possess a shell mainly made of a fibrous secondary layer
(Williams, 1997; Parkinson et al., 2005; Williams and Cusack, 2007). Each
fibre of this layer is secreted by the mantle and it is ensheathed by an
organic membrane (e.g. Jope, 1965; Williams, 1968; MacKinnon, 1974; Williams
and Cusack, 2007; Cusack et al., 2008; Casella et al., 2018). Thus, with a
decrease in size but within the same shell volume the surface area increases
and with it the number of organic components. Recently, the relationship
between the size of fibres and shell organic components was discussed in
detail (Garbelli, 2017; Garbelli et al., 2017; Ye et al., 2018a). The main
conclusion is that the smaller the calcite fibres, the higher the organic
component in the shell (cf. Fig. 11). Thus, smaller fibres and a greater
endopunctae density may lead to higher organic content per shell volume
(Fig. 11).
Relationship between the microstructure and the organic components
of calcified shells of brachiopods. Position information: see Figs. 6 and 7;
dv: dorsal valve; vv: ventral valve; CM: central middle part; AM: anterior
middle part.
Low pH and brachiopod microstructure
Several studies tried to understand how marine carbonate-shelled animals
respond to ocean acidification, such as brachiopods (McClintock et al., 2009;
Cross et al., 2015, 2016, 2018; Jurikova et al., 2019), bivalves (e.g. Berge
et al., 2006; McClintock et al., 2009; Beniash et al., 2010; Parker et al.,
2010; Melzner et al., 2011; Talmage and Gobler, 2011; Amaral et al., 2012;
Hiebenthal et al., 2013; Coleman et al., 2014; Gobler et al., 2014; Milano et
al., 2016), cold-water scleractinian corals (e.g. Form and Riebesell, 2011;
McCulloch et al., 2012; Jantzen et al., 2013b; Büscher et al., 2017) and
sea urchins (Suckling et al., 2015) (Table S1). The results of these studies
show that, in general, seawater acidification reduces the growth rate of
marine calcifiers (Michaelidis et al., 2005; Shirayama and Thornton, 2005;
Berge et al., 2006; Bibby et al., 2007; Beniash et al., 2010; Nienhuis et
al., 2010; Thomsen and Melzner, 2010; Fernández-Reiriz et al., 2011;
Melzner et al., 2011; Mingliang et al., 2011; Parker et al., 2011, 2012;
Talmage and Gobler, 2011; Liu and He, 2012; Navarro et al., 2013; Milano et
al., 2016).
For brachiopods, in the Liothyrella uva (Antarctic) and
Calloria inconspicua (New Zealand), no ocean acidification effects
on shell growth were detected by Cross et al. (2015, 2016, 2018), although
the shells of the former species may rapidly dissolve in acidified waters
(McClintock et al., 2009). However, C. inconspicua from the same
locality in New Zealand (Paterson Inlet, Stewart Island) laid down a denser
shell over the last 120 years, with nearby environmental conditions
increasing by 0.6 ∘C from 1953 to 2016 and slightly increasing by
35.7 µatm in pCO2 from 1998 to 2016 (Cross et al., 2018).
These changes are in line with global trends of ocean pH and temperature
since the industrial revolution (Caldeira and Wickett, 2005; Orr et al.,
2005; IPCC, 2013). The present experiment showed that growth of specimens was
not affected by the low-pH conditions; instead, their growth was similar to
that of specimens cultured under control conditions (no. 9006, ∼0.9 cm
in the ventral valve, ∼0.8 cm in the dorsal valve; no. 8005, ∼0.5 cm in the ventral valve, ∼0.4 cm in the dorsal valve). Based on
the von Bertalanffy growth function, Baumgarten et al. (2014) calculated an
expected growth increment and rate, and we compared those parameters with the
measured ones under control and low-pH conditions. The results in Fig. 12
demonstrate that the measured individual growth rates are within the range of
the ones of naturally growing individuals.
Projection of shell length of ventral valves on the von Bertalanffy
growth function (grey line) Lt= 71.53 (1-e-0.336(t-t0)),
source from Baumgarten et al. (2014); Lb: shell length at the
beginning of culturing; Lm: measured shell growth at the end of
culturing; Le: expected shell growth.
A limiting factor is the small database, but in general, the present
observations agree with studies that show no or little impact of
acidification on the growth rates of marine calcifiers (cf. Marchant et al.,
2010; Thomsen et al., 2010; Range et al., 2011, 2012; Talmage and Gobler,
2011; Dickinson et al., 2012; Fernández-Reiriz et al., 2012; Liu and He,
2012; Hiebenthal et al., 2013; Cross et al., 2015, 2016, 2018), or even an
increase in respiration, shell growth or metabolic rates after having
experienced low-pH conditions (Wood et al., 2008; Cummings et al., 2011;
Parker et al., 2012). We note however that a combined effect of multiple
stressors, such as low pH, lower dissolved oxygen and higher temperature or
scarce food availability is more complex and potentially detrimental. For
instance, Steckbauer et al. (2015) reported that hypoxia and increased
pCO2 could significantly reduce the respiration rate of some marine
invertebrates (Anthozoa, Gastropoda, Echinoidea and Crustacea). On the other
hand, the highest growth rate in the bivalve Macoma balthica
(Limecola balthica (Linnaeus, 1758)) was observed in seawater with
low O2 and high pH (Jansson et al., 2015). Gobler et al. (2014)
reported that juveniles of the bivalves Argopecten irradians
(Lamarck, 1819) and Mercenaria mercenaria (Linnaeus, 1758) were not
affected by hypoxia or acidification being applied individually, but the
growth rate decreased when juveniles were exposed to both conditions
simultaneously.
To explore the effects of acidification on brachiopod biomineralization, the
microstructures of the specimens cultured for 214 days (no. 43,
pH = 7.66±0.04; and no. 63, pH = 7.44±0.08) and the other
population cultured for 335 days (no. 8005, pH = 8.0 to 8.15±0.05;
and no. 9006, pH = 7.6 to 7.35±0.05) were investigated in detail.
No conclusive consideration can be carried out on the specimens cultured for
214 days (no. 43 and no. 63), but in the other culturing experiments
conducted for 335 days, the microstructure produced by the specimen cultured
under low-pH conditions was different from that produced under control
conditions: (1) the thickness of the primary layer increased with culturing
(Fig. S1a–d); (2) the density and size of the endopunctae were higher
(Fig. 1e–h); and (3) the fibres of the secondary layer were smaller. The
punctal pattern detected here is different from that observed by Cross et
al. (2018), who recorded no change in the punctal density of the ventral
valve of C. inconspicua on specimens from the last 120 years. Also
different is the trend in size of the endopunctae, which measured in the
dorsal valve by Cross et al. (2018) seems to decrease. However, the slight
environmental changes in the natural environment (references in Cross et al., 2018) are very different
from those of our culturing experiments. Furthermore, the size of the
endopunctae was measured from the dorsal valve only by Cross et al. (2018),
whereas the increase in size we report was observed only from the ventral
valve of M. venosa. A potential factor controlling this could be the
duration of culturing under low-pH conditions. We note, however, that during
the second phase of this acidification experiment (pH = 7.35), the
seawater was strongly undersaturated with respect to calcite
(Ωcal=0.6), suggesting that the observed structural
changes could also be linked to the saturation state. Conversely, the
duration of low-pH conditions as a controlling factor is also in line with
the few data available in the literature on microstructural changes during
acidification. Milano et al. (2016) reported no significant difference in the
prismatic microstructure of the cockle Cerastoderma edule when
cultured under low-pH conditions for about 2 months, except for dissolution
of ontogenetically younger parts of the shell. Similarly, a study by Stemmer
et al. (2013) on the clam Arctica islandica revealed that there was
no effect on the shape and size of the crystals in the homogeneous
microstructure after 3 months of culturing at low pH (Table S1). However, the
experiments conducted by Fitzer et al. (2014a, b) for 6 months on the blue
mussel Mytilus edulis showed that the animals exposed to low pH and
high pCO2 tend to produce less organized, disorientated calcite
crystals and an unordered layer structure.
Thus, in bivalves, and similar to our observations, the duration of culturing
may be crucial in recording significant effects. The present results lend
support to the microstructure variation observed in brachiopods during the
end-Permian extinction event and concomitant ocean acidification (Garbelli et
al., 2017). During this event, both Strophomenata and Rhynchonellata produced
more organic-rich shells to cope with the long-term and protracted seawater
acidification effects (Garbelli et al., 2017).
Stable isotope variation under low-pH conditions
Brachiopod shells are commonly used as archives for deep-time
paleoenvironmental reconstructions as they potentially record the original
geochemical composition of the seawater they lived in (Grossman et al., 1993;
Banner and Kaufman, 1994; Mii and Grossman, 1994; Mii et al., 2001; Brand et
al., 2003, 2011, 2016; Jurikova et al., 2019). Several studies suggest that
carbon and oxygen isotope compositions of the secondary layer of brachiopod
shells, especially slow-growing species – and particularly the innermost
shell parts – tend to be close to equilibrium with the ambient seawater
temperature (e.g. Popp et al., 1986; Carpenter and Lohmann, 1995; Parkinson
et al., 2005; Brand et al., 2013, 2015, 2016; Takayanagi et al., 2013;
Yamamoto et al., 2013). Recently, Bajnai et al. (2018) documented that
brachiopods do not incorporate oxygen isotopes in thermodynamic equilibrium
with ambient seawater, and appear to be subjected to taxon-specific
growth-rate-induced kinetic effects. The documented isotopic offset appears
to be relatively constant throughout the range of brachiopod shell production
from cold to warm environments. Thus, the brachiopod oxygen isotope
composition, when corrected for the seawater-18O contribution,
records ambient water temperatures close to those observed for their ambient
environment (Brand et al., 2013). Overall, the δ18O values of
brachiopods remain a mainstay and robust proxies of paleoenvironmental
temperature conditions.
In general, the measured δ13C (between -8.05 ‰ and
+0.45 ‰) and δ18O (between -3.04 ‰ and
+0.21 ‰) values of the secondary layer produced during growth in
the natural environment (Fig. 10) are similar to previous results from the
shells of M. venosa (Penman et al., 2013; Ullmann et al., 2017;
Romanin et al., 2018). Furthermore, the present results show that there are
no significant differences in δ13C and δ18O
values between the dorsal and ventral valves (p-values in
δ13C and δ18O of no. 8005 are 0.437 and 0.491,
respectively, and p-values in δ13C and δ18O
of no. 9006 are 0.862 and 0.910, respectively), which is in agreement with
previous findings (e.g. Parkinson et al., 2005; Brand et al., 2015; Romanin
et al., 2018).
In the naturally grown shell before-culturing, the
δ13C and δ18O values are relatively stable
along the ontogenetic direction (Table S2), except for the depleted values at
approximately mid-shell length in both no. 8005 and no. 9006. In particular,
in no. 9006, in this part of the shell values drop to about -6 ‰
for δ13C and -2 ‰ for δ18O values
(Fig. 10). Since the samples were taken from the mid-shell layer and not from
the shell interior, we can exclude the isotope negative shift being produced
by shell material added during the during-culturing shell
thickening. While this drop may be an artefact of both sampling and
analytical uncertainties, a possibility also exists that it could be linked
to shell repair processes. Brachiopods are well known to show a remarkable
shell repair ability (Cross et al. 2015, 2016), and thus it cannot be ruled
out that this shell part, although originally formed early in life under
natural conditions, also contains a contribution from material precipitated
in the culture seawater later in life, in particular under low-pH conditions.
Also, negative isotope excursions of a similar magnitude were recorded in
M. venosa specimens from the South America shelf by Ullmann et
al. (2017) and Romanin et al. (2018). Ullmann et al. (2017) implied that
these variable δ13C and δ18O values indicate
isotope disequilibrium with ambient waters in Terebratellids. In contrast,
Romanin et al. (2018), who also analysed specimens collected from Comau
Fjord, attributed the negative isotope excursion to environmental
perturbations, in particular, to changes in seawater productivity and
temperature, and/or to anthropogenic activities. Negative shifts in both
δ13C and δ18O values during ontogeny have also
been observed in the brachiopod Terebratella dorsata, which
co-occurs with M. venosa and which has been explained by the effect
of resorption in corresponding muscle scar areas (Carpenter and Lohmann,
1995). Here, we follow the interpretation of Romanin et al. (2018) to explain
the mid-shell excursion observed in our specimens.
Calculated carbon and oxygen fractionation factors for brachiopods
based on cultured M. venosa and culture seawater.
Sample
Treatment
Avg.
Avg.
Growth
ID
Δ13Ccal-DIC
Δ18Ocal-sw
temperature
No. 8004
Control
-4.06
29.99
10 ∘C
No. 9005
Acidification pH 7.35
-1.21
30.92
10 ∘C
No. 9004
Acidification pH 7.35
-2.23
30.70
10 ∘C
The most prominent change in δ13C values was observed in the
secondary layer produced during-culturing under low-pH
conditions (δ13C VPDB: ∼-25 ‰), reflecting the
composition of the δ13CDIC (δ13C
VPDB: -24 ‰ for the low-pH/high-pCO2 conditions). The
δ13C values were significantly depleted by more than
20 ‰ in the specimens cultured under low-pH/high-pCO2
conditions (pH 7.60 and pH 7.35; no. 9004, no. 9005 and no. 9006) (Fig. 10,
Appendix A, Table S2), whereas the depletion was lower and only a few per mil
(about 0.9 ‰–1.2 ‰) in the control specimens (pH 8.00 and
8.15; no. 8004 and no. 8005). This demonstrates that the δ13C
values of M. venosa to a large extent reflect the composition of the
CO2 source and thus present a valuable geochemical archive. Similar
observations have also been reported for other calcifiers cultured under
controlled experimental settings with pH mediated by CO2-bubbling.
For a comparison, Hahn et al. (2014) reported a decreasing trend of about
10 ‰ in δ13C values in the blue mussel
Mytilus edulis when exposed to seawater conditions of pH 8.03
(pCO2 612 µatm) and pH 7.21 (pCO2
4237 µatm). In corals, a species-specific δ13C
response to high-pCO2 conditions was reported by Krief et
al. (2010) of more negative 2.3 ‰ and 1.5 ‰
δ13C values in Porites sp. after 14 months of
culturing under low-pH conditions (pH 7.49, pCO2 1908 µatm
and 7.19 pCO2, 3976 µatm), whereas no significant
difference was found in other coral species, such as Stylophora pistillata (Esper, 1797).
In our culturing experiments, oxygen isotope compositions of the shells
record only a minor depletion during-culturing under different
pH conditions (δ18O (VPDB): -6.4 ‰ to -7.9 ‰/(VSMOW): ∼+23.6 ‰ to +24.3 ‰) in
comparison to the values observed in the shell parts grown under natural
conditions, following the changes in δ18OH2O.
The fractionation of carbon and oxygen isotopes between phases – brachiopod
calcite and culture seawater – is defined as
Δ13Ccal-DIC
or Δ18Ocal-sw=1000×lnαcal-DIC/sw,
where αcal-DIC/sw=
[13C/12C]cal/[13C/12C]DIC or
[18O/16O]cal/[18O/16O]sw, respectively. The
calculated values based on our culture measurements are presented in Table 11.
For carbon isotopes, we observe variable Δ13Ccal-DIC
between the different specimens and culturing treatments, and it is
inconclusive whether this is linked to culturing conditions, differences
between individuals or an ontogenetic component. It appears that there is
about a 2 ‰ difference between the control specimen and samples from
the acidification treatments (pH 7.35), with the last one being, strikingly,
closer to the equilibrium with seawater DIC. Possibly, this illustrates the
variability in kinetic effects (Bajnai et al., 2018), but may also be linked
to changes in the source δ13CDIC in the control
treatment. More studies are needed to fully answer this question.
Similarly, for oxygen isotopes, we find variable
Δ18Ocal-sw with an apparent trend with pH. These
values are offset from the equilibrium Δ18Ocal-sw
(Δ18Ocal-sw = 32.9 at 10∘C) determined
by Watkins et al. (2013, 2014). This suggests that M. venosa present
non-equilibrium growth-rate-related isotope effects up to about
-2.9 ‰, larger than the approx. -1.5 ‰ previously
recorded by Bajnai et al. (2018). Provided that this offset can be
constrained, brachiopods continue to present robust archives for
palaeo-temperature reconstructions.
In summary, although it appears that variable growth rates present the most
prominent confounding parameter complicating the interpretation of carbon and
oxygen data, provided that we account for them, our results support the
notion that brachiopods present robust geochemical archives, even when
stressed by ocean acidification.