Introduction
Biomineralization is a process through which living organisms produce a
protective, mineralized hard tissue. The considerable diversity of
biomineralizing species contributes to high variability in terms of shape,
organization and mineralogy of the structures produced (Lowenstam and Weiner,
1989; Carter et al., 2012). Different architectures at the micrometer and
nanometer scale and different biochemical compositions determine material
properties that serve specific functions (Weiner and Addadi, 1997; Currey,
1999; Merkel et al., 2007). Besides these differences, all mineralized
tissues are hybrid materials consisting in hierarchical arrangements of
biomineral units surrounded by organic matrix, also known as
“microstructures” (Bøggild, 1930; Carter and Clark, 1985;
Rodriguez-Navarro et al., 2006), “ultrastructures” (Blackwell et al.,
1977; Olson et al., 2012) or overall “fabrics” (Schöne, 2013;
Schöne et al., 2013). The carbonate and organic phases represent the
fundamental level of the organization of biomaterials (Aizenberg et al.,
2005; Meyers et al., 2006). The mechanisms of microstructure formation and
shaping, especially in mollusks, has attracted increasing attention during
recent decades. At present, it is commonly accepted that the organic
compounds play an important role in the formation of the inorganic phases of
biominerals (Weiner and Addadi, 1991; Berman et al., 1993; Dauphin et al.,
2003; Nudelman et al., 2006). However, the identification of the exact
mechanisms driving biomineralization is still an open research question.
Previous studies conducted on mollusks show that environmental parameters can
influence microstructure formation (Lutz, 1984; Tan Tiu and Prezant, 1987;
Tan Tiu, 1988; Nishida et al., 2012). These results set the stage for
research that focuses on the use of shell microstructures as proxies for
reconstructing environmental conditions (Tan Tiu, 1988; Tan Tiu and Prezant,
1989; Olson et al., 2012; Milano et al., 2017).
Mollusks are routinely used as climate and environmental proxy archives
because they can record a large amount of environmental information in their
shells (Richardson, 2001; Wanamaker et al., 2011a; Schöne and Gillikin,
2013). Whereas structures at nanometric levels are still underexplored as
potential environmental recorders, shell patterns at lower magnification,
such as individual growth increments, are commonly used for this purpose
(Jones, 1983; Schöne et al., 2005; Marali and Schöne, 2015; Mette et
al., 2016). Mollusks deposit skeletal material on a periodic basis and at
different rates (Thompson et al., 1980; Deith, 1985). During periods of fast
growth, growth increments are formed, whereas during periods of slower
growth,
growth lines are formed (Schöne, 2008; Schöne and Gillikin, 2013).
The periodicity of such structures ranges from tidal to annual (Gordon and
Carriker, 1978; Schöne and Surge, 2012). By cross-dating time series with
similar growth patterns it is possible to construct century- and
millennia-long master chronologies (Marchitto et al., 2000; Black et al.,
2008, 2016; Butler et al., 2013). This basic approach, in combination with
geochemical methods, has great potential in reconstructing past climatic
conditions (Wanamaker et al., 2011b). At present, the most frequently used
and well-accepted geochemical proxy is oxygen isotopic composition of shell
material (δ18Oshell) (Epstein, 1953; Grossman and Ku,
1986; Schöne et al., 2004; Wanamaker et al., 2007), which may serve as a
paleothermometer and/or paleosalinometer (Mook, 1971; Andrus, 2011); however,
δ18Oshell value is influenced by both seawater
temperature and the isotopic composition of seawater (δ18Owater; related to salinity). Thus, δ18Oshell-based temperature reconstructions are particularly
challenging in habitats with fluctuating δ18Owater
conditions such as estuaries or restricted basins (Gillikin et al., 2005).
Because of the multiple impacts on δ18Oshell values,
there have been substantial efforts to develop alternative techniques to
reconstruct environmental variables from mollusk shells (Schöne et al.,
2010; Milano et al., 2017).
This study investigates the possibility using shell microstructure properties
to serve as a new environmental proxy. For this purpose, the effects of
seawater temperature (grown at 10 and 15 ∘C) and dietary regime on
the microstructural units of Arctica islandica cultured under
controlled conditions were analyzed and quantified. A. islandica was
chosen as model species because of its great potential in paleoclimatology
and paleoceanography (see Schöne, 2013; Wanamaker et al., 2016). Its
extreme longevity of up to more than 500 years makes this species a highly
suitable archive for long-term paleoclimate and environmental reconstructions
(Schöne et al., 2005; Wanamaker et al., 2008, 2012; Butler et al., 2013).
Materials and methods
The analyses were conducted on 11 A. islandica shells. Three
juvenile A. islandica shells, sampled for the seawater temperature
experiment, were collected alive on 21 November 2009 aboard the F.V.
Three of a Kind off Jonesport, Maine, USA
(44∘26′9.829′′ N, 67∘26′18.045′′ W), in 82 m water
depth. From 2009 to 2011, all animals were kept in a flowing seawater
laboratory at the Darling Marine Center, Walpole, Maine, USA (see Beirne et
al., 2012, for additional details). In 2011, clams were grown at two different
temperature regimes for 16 weeks (Table 1). At the completion of the
experiment, shells were estimated to be between 4 to 5 years old. Eight
1-year old juveniles were collected in July 2014 from Kiel Bay, Baltic Sea
(54∘32′ N, 10∘42′ E; Fig. 1) and kept alive in tanks at
7 ∘C for 6 months at the Alfred Wegener Institute (AWI) for Polar and
Marine Research, Bremerhaven, Germany. During this time interval, the
animals were fed with an algal mix of Nannochloropsis sp.,
Isochrysis galbana and Pavlova lutheri. Then, they were
transferred to the Royal Netherlands Institute for Sea Research (NIOZ),
Texel, the Netherlands, and cultured in tanks at three different dietary
conditions for 11 weeks (Table 1).
Shell of adult Arctica islandica used in the temperature
experiment (top) and juvenile from the Baltic Sea used in the food
experiment (bottom). The map indicates the localities where the two sets of
shells were collected: Jonesport, Maine (circle), and Kiel Bay (square).
Seawater temperature experiment
The seawater temperature experiment started on 27 March and ended on
21 July 2011. Prior to the start of the experiment the animals were marked
with calcein. The staining leaves a clear fluorescent marker in the shells
that can be used to identify which shell material has formed prior to and
during the experiment. Initially, the animals were kept at
10.3 ± 0.2 ∘C for 48 days. Then, they were briefly removed
from the tanks and marked again. Subsequently, the clams were cultured for 69
more days at 15.0 ± 0.3 ∘C. Ambient seawater was pumped from
the adjacent Damariscotta River estuary and adjusted to the desired temperature.
The salinity was measured with a Hydrolab®
MiniSonde. It ranged between 30.2 ± 0.7 and 30.7 ± 0.7, in the
two experimental phases, respectively. During the entire culture period, all
clams were exposed to ambient food conditions. At the end of the experiment
the soft tissues were removed.
Food experiment
The food experiment was carried out from 9 February to 29 April 2015. The
animals were placed in aquaria inside a climate room at 9 ∘C. Water
temperature in the tanks ranged between 8 and 10 ∘C. Water salinity
was measured by using an ENDECO 102 refractometer and ranged between 29.6 and
29.9 ± 0.1 in each aquarium. The 15 L tanks were constantly supplied
with aerated water from the Wadden Sea. The clams were acclimated for 3
weeks before the start of the experiment. Three dietary regimes were chosen.
One treatment consisted of feeding the animals with Microalgae Mix (food
type 1), a ready-made solution of four marine microalgae (25 %
Isochrysis, 25 % Tetraselmis, 25 %
Thalassiosira and 25 % Nannochloropsis) with a particle
size range of 2–30 µm. A second treatment was based on PhytoMaxx
(food type 2), a solution of living Nannochloropsis algae with a
2–5 µm size range. A third treatment served as control; i.e., the
animals were not fed with any additional food. In treatments with food type 1
and 2, microalgae were provided at the constant optimum concentration of
20 × 106 cells L-1 (Winter, 1969). A dispenser equipped with a timer was used to
distribute the food 5 times per day. At the end of the experiment the soft
tissues were removed. A distinct dark line in the shells indicated the
transposition to the NIOZ aquaria and the associated stress. This line marks
the beginning of the tank experiment.
List of the studied specimens of Arctica islandica and
experimental conditions.
Sample ID
Locality
Age
Experiment
Treatment
A2
Maine
5
Temperature
10 ∘C + 15 ∘C
A4
Maine
4
Temperature
10 ∘C + 15 ∘C
A5
Maine
4
Temperature
10 ∘C + 15 ∘C
S12
Kiel Bay
1
Diet
Food 1
S14
Kiel Bay
1
Diet
Food 1
S15
Kiel Bay
1
Diet
Food 1
G11
Kiel Bay
1
Diet
Food 2
G12
Kiel Bay
1
Diet
Food 2
G15
Kiel Bay
1
Diet
Food 2
N13
Kiel Bay
1
Diet
No additional food
N15
Kiel Bay
1
Diet
No additional food
Sample preparation
The right valve of each specimen was cut into two 1.5 mm thick sections
along the axis of maximum growth. For this purpose, a low-speed precision saw
(Buehler Isomet 1000) was used. Given the small size and fragility of the
juvenile shells used in the food experiment, the valves were fully embedded
in a block of Struers EpoFix (epoxy) and air-dried overnight prior the
sectioning. Sections of the clams used in the temperature experiment were
embedded in epoxy after the cutting. All samples were ground using a Buehler
Metaserv 2000 machine equipped with Buehler silicon carbide papers of
different grit sizes (P320, P600, P1200, P2500). In addition, the samples
were manually ground with Buehler P4000 grit paper and polished with a
Buehler diamond polycrystalline suspension (3 µm). Sample surfaces
were rinsed in demineralized water and air-dried. In the samples of the
temperature experiment, the calcein marks were located under a fluorescence
light microscope (Zeiss Axio Imager.A1m microscope equipped with a Zeiss
HBO100 mercury lamp and filter set 38: excitation wavelength, ca.
450–500 nm; emission wavelength, ca. 500–550 nm).
Sketch showing the microstructures characterizing the different
shell layers of Arctica islandica. The oOSL is formed by homogenous
microstructure (HOM), whereas the oOSL and ISL are composed of crossed
acicular structure (CA); dog is direction of growth.
A. islandica shell organization
The shell of A. islandica consists of pure aragonite and it is
divided in two major layers, an outer shell layer (OSL) and the inner shell layer (ISL).
The OSL is further subdivided in outer portion (oOSL) and inner portion (iOSL)
(Schöne, 2013). These layers are characterized by specific
microstructures (Ropes et al., 1984; Fig. 2). The oOSL largely consists of
homogenous microstructures with a granular aspect (Schöne et al., 2013).
This type of architecture is characterized by approximately equidimensional
units of about 5 µm in width. The unit shape tends to be irregular
with a bulky aspect. The organization lacks of specific structural
arrangement typical of other types of microstructures such as the
crossed-lamellar
and crossed-acicular microstructures. The latter are the main
component characterizing the iOSL and ISL (Dunca et al., 2009). Here,
elongated units are arranged with two main dip directions, resulting in a
relative oblique alignment. As shown in Fig. 2, the elongation of the
structures becomes more evident in the ISL.
The present study focuses on ventral margin of the shells. Analyses were
carried out exclusively in the OSL.
Similar to other mollusks, the shell of A. islandica contains
pigment polyenes, which are obviously visible when using CRM (Hedegaard et
al., 2006). Polyenes are organic compounds containing single (C-C) and
double (C=C) carbon–carbon bonds forming a polyenic chain. Their
distribution across the shell is not homogenous. The pigments are abundant in
the oOSL, whereas they become scarce in the iOSL. Furthermore, an enrichment
in polyenes has been observed in the growth lines, potentially indicating
their involvement in the biomineralization process (Stemmer and Nehrke,
2014). However, the specific functions of these organic compounds have not
been disclosed yet (Hedegaard et al., 2006; Karampelas et al., 2009). Given
the high phenotypic variation in pigmentation among and within mollusk
species, it has been proposed that coloration does not have a primary
function as an adaptive tool (i.e., camouflage, warning signaling) as in other
animals (Seilacher, 1972; Evans et al., 2009). This, in turn, can indicate a
certain degree of influence of the environment on the pigments, in particular
by diet (Hedegaard et al., 2006; Soldatov et al., 2013). In the current
study, the effect of different dietary regimes was tested in order to explore
the potential of polyenes as environmental proxy.
Confocal Raman microscopy and image processing
Shells were mapped with a WITec alpha 300 R (WITec GmbH, Germany) confocal
Raman microscope. Scans of 50 × 50 µm,
100 × 50 µm and 150 × 50 µm were
performed using a piezoelectric scanner table. All Raman measurements were
carried out using a 488 nm diode laser. A spectrometer (UHTS 300, WITec,
Germany) was used with a 600 mm-1 grating, a 500 nm blaze and an
integration time of 0.03 s. On each sample two to six scans were made,
depending of the thickness of the shell. For instance, in juvenile shells
(food experiment), two scans of each sample were made. On larger shells used
in the temperature experiment, six maps were completed, i.e., two maps in the
oOSL, two in the middle of the iOSL and two in the inner portion of the iOSL.
Each scan contained between 40 000 and 120 000 data points, depending on
the map size. The spatial resolution equaled 250 nm. Half of the maps were
performed on the shell portion formed before the experiments. The other half
were made on the shell portion formed under experimental conditions. In order
to avoid areas affected by transplantation or marking stress, the scans were
located far off the calcein and stress lines. Raman maps on food experiment
shells were performed 300 µm away from the stress line. In the
shells from the temperature experiment, the scans were made 1 mm away from
the calcein mark.
Raman spectrum of Arctica islandica showing the typical
aragonite peaks (gray line). The exact position of the polyene peaks R1
and R4 was determined by using a peak fitting routine based on a
Gaussian function (black line).
Polarized Raman microscopy is known to provide comprehensive information
about the crystallographic properties of the materials (Hopkins and Farrow,
1985). The aragonite spectrum is characterized by two lattice modes
(translation mode Ta, 152 cm-1 and librational mode La,
206 cm-1) and the two internal modes (in-plane band ν4,
705 cm-1 and symmetric stretch ν1, 1085 cm-1). The ratio
(Rν1/Ta) between peak intensities belonging to ν1and
Ta is caused by different crystallographic orientations of the
aragonitic units (Hopkins and Farrow, 1985; Nehrke and Nouet, 2011). Within
each scan, Rν1/Ta was calculated for each data point. New spectral
images were generated using WITecProject software (version 4.1, WITec GmbH,
Germany). These images were then binarized by replacing all values above 2.5
with 1 and the others with 0. The orientation was quantified by calculating
the area formed by pixels of value 1 over the total scan area. The imaging
software Gwyddion (http://gwyddion.net/; last access: June 2016) was
used for this purpose. The results were expressed in percentage.
The Raman scans of the food experiment shells were analyzed to investigate
the pigment composition. Polyene peaks have definite positions in the
spectrum according to the number of the C-C and C=C bonds of the chain,
which are specific for certain types of pigments. The two major polyene peaks
at ∼ 1130 (R1) and 1520 cm-1 (R4) were identified by
using the “multipeak fitting 2” routine of IGOR Pro (version 7.00,
WaveMetrics, USA). Their exact position was determined adopting a Gaussian
fitting function (Fig. 3). The number of single (N1) and double carbon
bonds (N4) was calculated by applying the equations by Schaffer et
al. (1991):
N1=476(R1-1082),N4=830(R4-1438).
Spectral images of the R4 band were used to locate the polyenes in the
shell and measure the thickness of the pigmented layer. The images were
analyzed using the software Panopea (© Schöne and Peinl).
The thickness of the pigmented layer was calculated as distance between the
outer shell margin and the point where the concentration of polyenes suddenly
declined. The measurements were taken perpendicular to the shell outer
margin. This analysis was conducted only on the shells of the food
experiment. Given the larger size of the shells used in the temperature
experiment, the spectral maps were not sufficient for a correct localization
of the pigmented layer boundaries and estimation of its thickness.
To quantify changes of the orientation of individual biomineral units of the
juvenile shells (food experiment), the spectral maps were subdivided into
two portions. The outermost shell portion (oOSL) was enriched in pigments
whereas the iOSL showed a decrease in polyene content.
Arctica islandica shell growth under controlled conditions.
(a) Total growth and (b) instantaneous growth rate during
the temperature experiment. (c) Total growth during the food
experiment.
Effect of temperature increase on biomineral orientation.
(a) Position of the Raman maps of the three specimens reared at 10
and 15 ∘C. Dotted lines indicate the location of the calcein marks;
dog is direction of growth. (b) Raman spectral maps of Rν1/Ta. Left images of each column represents shell portion formed at
10 ∘C, right images represent shell portions formed at
15 ∘C. First row of pairs refers to oOSL, the other two represent
the iOSL; scale bars = 10 µm. (c) Proportions of
biominerals with Rν1/Ta>2.5 a.u. with respect to the total map
area. Asterisks indicate significant difference between the orientation of
iOSL microstructures formed at 10 and 15 ∘C (p<0.05).
Scanning electron microscopy
After performing Raman measurements, the samples were prepared for SEM
analysis. Each shell slab was ground with a Buehler Metaserv 2000 machine and
Buehler silicon F2500 grit carbide paper. To reduce the impact of grinding on
the sample surface of juvenile shells, extra grinding was done by hand. Then,
the slabs were polished with a Buehler diamond polycrystalline suspension
(3 µm). Afterward, shell surfaces were etched in 0.12 N HCl
solution for 10 (food experiment samples) to 90 s (temperature experiment
samples) and subsequently placed in 6 vol % NaClO solution for 30 min.
After being rinsed in demineralized water, air-dried samples were
sputter coated with a 2 nm thick platinum film by using a Low Vacuum Coater Leica EM ACE200.
A scanning electron microscope (LOT Quantum Design 2nd generation Phenom Pro
desktop SEM) with backscattered electron detector and 10 kV accelerating
voltage was used to analyze the shells. Images were taken at the same
distances from the calcein and stress lines as in the case of the Raman
measurements to assure comparability of the data.
In addition, stitched SEM images of the ventral margins were used to
accurately determine the shell growth advance during the culturing
experiments. Growth increment widths were measured with the software Panopea.
Given the difference in duration of the two phases of the temperature
experiment, the measurements were expressed as total growth and instantaneous
growth rate (Fig. 4a, b). The latter was calculated using the following
equation (Brey et al., 1990; Witbaard et al., 1997):
Instantaneous growth rate=(ln(yt/y0)/a),
where y0 represents the initial shell height, yt is the final shell
height and a is the duration of the experiment. In the case of the food
experiment, only the total growth was calculated (Fig. 4c).
Results
Effect of seawater temperature and diet on A. islandica shell growth
When exposed to a water temperature of 10 ∘C, the shells grew
between 11.67 and 14.17 mm during a period of 48 days. During a period of
69 days at 15 ∘C, the growth ranged between 2.32 and 5.77 mm
(Fig. 4a). The instantaneous growth rate showed a decrease between the two
experimental phases. At 10 and 15 ∘C, the average instantaneous
growth per day was 0.0091 and 0.0013, respectively (Fig. 4b). The decrease in
total growth and growth rate between the two temperatures was statistically
significant (t test, p<0.01).
During the food experiment, shells grew between 0.37 and 3.71 mm with large
differences due to the different food types. Growth of specimens exposed to
food type 1 ranged between 1.87 to 3.71 mm, whereas those cultured with food
type 2 grew between 0.55 to 0.96 mm. Both control specimens added 0.37 mm
of shell during the experimental phase (Fig. 4c). ANOVA and Tukey's HSD post
hoc tests showed significant differences between specimens cultured with food
type 1 and 2 (p<0.05) and between food type 1 and control shells (p<0.05).
Effect of seawater temperature on A. islandica
microstructure
At a water temperature of 10 ∘C, the area occupied by
microstructural units oriented with Rν1/Ta higher than 2.5 a.u.
(= arbitrary units) ranged between 31.3 and 50.6 % in the oOSL and
between 21.3 and 33.5 % in the iOSL. When exposed to 15 ∘C,
values ranged between 25.6 and 48.7 and between 45.7 and 55.9 % in the
oOSL and iOSL, respectively (Fig. 5). Whereas the slight difference of area
with Rν1/Ta>2.5 in the oOSL was not significant between the two
water temperatures (t test, p=0.62), the area with Rν1/Ta
>2.5 in the iOSL significantly increased at 15 ∘C
(t test, p=0.02). Under the SEM, no difference was visible between
units formed at 10 and 15 ∘C (Fig. 6).
SEM images of Arctica islandica shell microstructures
formed at 10 ∘C (left column) and at 15 ∘C (right column).
The sketch indicates the position of the images 1 mm away from the calcein
mark (gray line). The first row of images refers to the oOSL, the other two
row refers to the iOSL. Scale bars if not otherwise
indicated = 5 µm.
Effect of different diets based on (a) food type 1,
(b) food type 2 and (c) ambient food on biomineral
orientation. The optical microscope images indicate the position of the Raman
scans. Dotted line marks the start of the experiment. The portion of shell
prior the line was formed during the acclimation phase; dog is direction
of growth. The Raman spectral maps indicate the ratio Rν1/Ta for
each data point of the scan. For each shell, maps on the left represent shell
portions during the experiment, maps on the right represent shell portions
formed during the acclimation phase. In the acclimation portion of the sample
reared with ambient food, a significant change in the microstructure
orientation is visible. The respective area of the Raman map was not
considered in further calculations because it was influenced by the emersion
and transportation stress at the start of the experiment. Scale
bars = 10 µm. The graphs show the proportions of biominerals of
oOSL and iOSL with Rν1/Ta>2.5 a.u. with respect to the total map
area.
Details of the pigment composition of the Arctica islandica shells used in the food
experiment. The position of the major polyene peaks R1 and R4 in
the Raman spectrum is indicated together with the number of single and
double carbon bonds of the pigment molecular chain (N1 and N4).
Each shell was analyzed in the portions formed before and during the
experimental phase.
Sample ID
Shell portion
R1 (cm-1)
R4 (cm-1)
N1
N4
S12
Acclimation
1130.9
1515.2
9.7
10.8
Food 1
1121.4
1515.3
12.1
10.7
S14
Acclimation
1133.2
1519.4
9.3
10.2
Food 1
1132.2
1518.6
9.5
10.3
S15
Acclimation
1129.5
1516.5
10.0
10.6
Food 1
1132.1
1519.8
9.5
10.1
G11
Acclimation
1132.6
1518.4
9.4
10.3
Food 2
1129.5
1517.0
10.0
10.5
G12
Acclimation
1131.7
1518.7
9.6
10.3
Food 2
1132.1
1518.2
9.5
10.4
G15
Acclimation
1132.4
1519.5
9.4
10.2
Food 2
1128.0
1520.9
10.3
10.0
N13
Acclimation
1130.2
1515.6
9.9
10.7
Ambient food
1131.4
1514.1
9.6
10.9
N15
Acclimation
1117.9
1516.0
13.3
10.6
Ambient food
1130.7
1517.0
9.8
10.5
Average
1129.7 ± 4.2
1517.5 ± 2.0
10.1 ± 1.1
10.4 ± 0.3
Effect of food on A. islandica microstructure and
pigments
In the shells cultured with food type 1, the area occupied by biomineral
units oriented with Rν1/Ta higher than 2.5 a.u. during the
experiment ranged between 24.8 % (oOSL) and 43.0 % (iOSL). In the
shell portion deposited during the acclimation phase, the ratio varied
between 19.4 % (oOSL) and 36.2 % (iOSL). Although a trend was
recognized, these variations were not statistically different
(t tests OSL: p=0.43; ISL: p=0.57;
Fig. 7a). On the contrary, in the clams exposed to food type 2, the area
occupied by units oriented with Rν1/Ta>2.5 ranged between
11.7 % (oOSL) and 20.4 % (iOSL). Before the experiment, the
proportions were higher, i.e., 18.1 % (oOSL) and 26.3 % (iOSL)
(Fig. 7b). As for the other treatment, the difference was not significant
(t tests oOSL: p=0.34; iOSL: p=0.28). In the control shells
grown with no extra food supply, the area with Rν1/Ta>2.5 ranged
between 24.6 % (oOSL) and 44.8 % (iOSL) during the experiment and
21.2 % (oOSL) and 44.5 % (iOSL) before the experiment (Fig. 7c).
Hence, no trend was visible and the two portions did not show significant
differences (t tests oOSL: p=0.59; iOSL: p=0.99). As for the
temperature experiment, under the SEM, the microstructure of the shells from
the food experiment did not show any change (Fig. 8).
SEM images of Arctica islandica shell microstructures
formed during the acclimation phase at AWI (left column) and during the food
experiment (right column). Scale bars = 4 µm.
Effects of diet on shell pigment distribution. (a) Raman
spectral maps of the 1524 cm-1 band representing the distribution of
the polyenes in the shell cultured with food type 2. Dotted line marks the
start of the experiment; dog is direction of growth. (b) The
graph shows the thickness of the pigmented layer over the whole shell
thickness before and during the food experiments.
All treatments showed a slightly thicker pigmented layer formed during the
experiment than during the acclimation phase (Fig. 9a). During the
experiment, clams cultured with food type 1 showed, on average, a thickening
by 6.4 %. In the food type 2 specimens, the layer thickness increased by
9.9 %. Control shells showed an increase of 10.4 % (Fig. 9b).
However, none of these differences was statistically significant (t test.
Food type 1: p=0.43; food type 2: p=0.39; control: p=0.10).
According to the position of the polyene peaks, the number of single carbon
bonds in the pigment chain did not change between the acclimation and
experimental phase (N1=10.1±1.3 and N1=10.0±0.9,
respectively). Likely, no significant variation was observed in the number of
double carbon bonds (N4=10.5±0.2 and N4=10.4±0.3,
respectively; Table 2).
Discussion
According to the results, variations of both food type and water temperature
can influence the shell production rate of A. islandica. However,
the shell microstructure and pigmentation react differently to these two
environmental variables. Whereas changes of the dietary conditions do not
affect the shell architecture and pigment composition, the crystallographic
orientation of the biomineral units responds to seawater temperature
fluctuations.
Environmental influence on shell microstructure
The environmental conditions experienced by mollusks during the process of
biomineralization appear to influence shell organization (Carter, 1980).
Among the different environmental variables, water temperature is the most
studied driving force of structural changes of the shell. For instance, shell
mineralogy can vary depending on water temperature (Carter, 1980). According
to the thermal potentiation hypothesis, nucleation and growth of calcitic
structural units is favored at low temperatures by kinetic factors (Carter et
al., 1998). As a consequence, bivalve species living in cold water
environments exhibit additional or thicker calcitic layers compared to the
corresponding species from warm waters (Lowenstam, 1954; Taylor and Kennedy,
1969). Changes in the calcium carbonate polymorph also affect the type of
microstructures (Milano et al., 2016a).
However, architectural variations often occur without mineralogical impact
(Carter, 1980).
The present results indicate that temperature induces a change in the
crystallographic orientation of the biomineral units of A. islandica. Although water temperature was previously shown to have an impact
on microstructure formation, the attention has been mainly focused on the
effects on the morphometric characteristics (e.g., size and shape) or on the
type of microstructure. Milano et al. (2017) demonstrated that size and
elongation of prismatic structural units of Cerastoderma edule were
positively correlated to seawater temperature variation throughout the
growing season. Likely, low temperatures induced the formation of small nacre
tablets in Geukensia demissa (Lutz, 1984). Seasonal changes of the
microstructural type were reported in the freshwater bivalve
Corbicula fluminea (Prezant and Tan Tiu, 1986; Tan Tiu and Prezant,
1989). During the warm months, crossed-acicular structure was produced,
whereas simple crossed-lamellar were formed during the winter period. So far,
variations of the crystallographic properties of bivalve biominerals have
been exclusively investigated as a response to hypercapnic (acidified)
conditions. Mytilus galloprovincialis and Mytilus edulis
showed a significant change in the orientation of the prisms forming shell
calcitic layer when subjected to hypercapnia (Hahn et al., 2012; Fitzer et
al., 2014a). Altered crystallographic organization may derive from the animal
exposure to suboptimal conditions. These findings together with the present
results suggest that thermal- and hypercapnic-induced stress are likely to
affect the ability of the bivalves to preserve the orientation of their
microstructural units (Fitzer et al., 2014b).
Different food sources do not significantly influence the orientation of the
biomineral units or the composition and distribution of pigments in shells of
A. islandica. In previous studies, the relationship between
microstructure and diet was virtually overlooked resulting in a lack of data
in the literature. As suggested by Hedegaard et al. (2006), however, the type
of polyenes is influenced by food. The ingestion of pigment-enriched
microalgae potentially leads to an accumulation of pigments in mollusk
tissues and the shell (Soldatov et al., 2013). On the other hand, it has been
argued that polyenes do not generate from food sources like other pigments
(i.e., carotenoids), but they are locally synthesized (Karampelas et al.,
2009). In accordance to Stemmer and Nehrke (2014), the results presented here
support the view that the specific diets on which the animals rely on do not
influence shell pigment composition. The chemical characteristics of the
polyenes are likely to be species-specific and independent from the habitats.
Confocal Raman microscopy as tool for microstructural analysis
From a methodological perspective, the present study represents an innovative
approach in the investigation of shell microstructural organization. Electron
backscatter diffraction (EBSD) has been previously used to determine the
crystallographic orientation of gastropod (Fryda et al., 2009;
Pérez-Huerta et al., 2011) and bivalve microstructural units (Checa et
al., 2006; Frenzel et al., 2012; Karney et al., 2012). Whereas, CRM on
mollusk shells is generally applied within studies on taphonomic
mineralogical alteration and pigment identification (Stemmer and Nehrke,
2014; Beierlein et al., 2015). Both techniques provide considerably high
spatially resolved analysis up to 250 nm, allowing for the identification of
individual structural units at micrometer and nanometer scale (Cusack et al.,
2008; Karney et al., 2012). CRM offers important advantages supporting a
broader application of this methodology in the biomineralization research
field. For instance, samples do not require any pre-treatment. Unlike EBSD,
there is no need for preparing thin sections (∼ 150 µm thick)
or etching the shell surface (Griesshaber et al., 2010; Hahn et al., 2012).
Therefore, further structural and geochemical analyses can be easily
performed on the same sections (Nehrke et al., 2012). In addition, the size
of CRM scans can be remarkably large (∼ 7–8 mm2) without
compromising the achievable resolution. By overlapping adjacent scans, it is
possible to produce stitched scans allowing one to further increase the region of
interest on the shell surface. On the other side, EBSD provides a relevant
advantage to take into consideration. It allows for absolute measures of the
crystallographic orientation of the carbonate structures. The CRM, instead,
determines the relative change in the orientation between the single units
without providing absolute values.
SEM has previously been demonstrated to provide a convenient approach for
the identification of individual structural units and the quantification of
potential changes occurring within them (Milano et al., 2017, 2016b).
However, SEM exclusively provides information about the morphometric
characteristics of the microstructural units. As highlighted by the present
study, to achieve an exhaustive examination, it is suggested to combine SEM
with techniques assessing crystallographic properties of the biomaterials.
For instance, our results show that the effect of water temperature is
detectable in crystallographic orientation but not in morphometric features
of the biomineral units.
Environmental influence on shell growth
Numerous previous studies demonstrated that the growth rate of A. islandica is linked to environmental variables (e.g., Witbaard et al., 1997,
1999; Schöne et al., 2004; Butler et al., 2010; Mette et al., 2016).
However, the relative importance of the main factors, temperature and food
supply/quality driving shell formation are still not well understood.
Positive correlations between shell growth and water temperature have been
identified (i.e., Schöne et al., 2005; Wanamaker et al., 2009; Marali et
al., 2015), but the relationship between shell growth and environment is more
complex (Marchitto et al., 2010; Stott et al., 2010; Schöne et al., 2013)
and likely dependent on the synergic effect of food availability and water
temperature (Butler et al., 2013; Lohmann and Schöne, 2013; Mette et al.,
2016). Tank experiments were run in order to precisely identify the role of
these two parameters on the shell growth of A. islandica (Witbaard et
al., 1997; Hiebenthal et al., 2012). A 10-fold increase in instantaneous
growth rate was observed between 1 and 12 ∘C, with the greatest
variation occurring below 6 ∘C (Witbaard et al., 1997). On the
contrary, a temperature increase between 4 and 16 ∘C was shown to
produce a slowdown of shell production (Hiebenthal et al., 2012). Our results
are in agreement with the latter study and show a decrease in the
instantaneous growth rate between 10 and 15 ∘C. High temperatures
are often associated with an increase of free radical production (Abele et
al., 2002). A large amount of energy then has to be allocated to limit
oxidative cellular damage (Abele and Puntarulo, 2004). This translates into a
higher accumulation of lipofuscin and slower shell production rate
(Hiebenthal et al., 2013). The contrasting results of previous studies may be
explained by individual differences in the tolerance toward temperature
change (Marchitto et al., 2000).
Along with water temperature, food availability was also shown to influence
A. islandica shell growth (Witbaard et al., 1997). At high algal
cell densities, the siphon activity increased. This, in turn, was positively
correlated to shell growth. Previous experiments used different combinations
of algae, such as Isochrysis galbana and Dunaliella marina
(Witbaard et al., 1997), or Nannochloropsis oculata, Phaeodactylum tricornutum and Chlorella sp. (Hiebenthal et al., 2012) to grow
the clams. However, there are still uncertainties about the composition of
the primary food source for this species (Butler et al., 2010). Even though
it is challenging to determine the preferred algal species, our results show
that the use of a mixture of different algal species results in significantly
faster shell growth than the used of just one algal species. In the natural
environment, suspension feeders such as A. islandica preferentially
ingest certain particle sizes (Rubenstein and Koehl, 1977; Jorgensen, 1996;
Baker et al., 1998). The exposure to a limited algal size range, as in the
case of food type 2, may affect shell growth. Furthermore, multispecific
solutions contain a higher variability of biochemical components that better
meet the nutritional requirements of the animal (Widdows, 1991). Our results
are in good agreement with previous findings. For instance, it has been shown
by Strömgren and Cary (1984) that Mytilus edulis shell growth
increased as a result of a diet based on three different algal species.
Furthermore, Epifanio (1979) tested the differences on the growth of
Crasssostrea virginica and Mercenaria mercenaria of a mixed
diet composed by Isochrysis galbana and Thalassiosira pseudonana and diets consisting of the single species. Faster growth was
measured in the mixed diet treatment, indicating a synergic effect of the
relative food composition (Epifanio, 1979). Likely, Mytilus edulis
grew faster when reared with different types of mixed diets as opposed to
monospecific diets (Galley et al., 2010).