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Benthic diatoms from Potter Cove, 25 de Mayo (King George) Island, Antarctica: Mucilage and glucan storage as a C-source for limpets
Accepted Manuscript Benthic diatoms from Potter Cove, 25 de Mayo (King George) Island, Antarctica: Mucilage and glucan storage as a C-source for limpets Yasmin Daglio, Hernán Sacristán, Martín Ansaldo, María C. Rodríguez PII:
S1873-9652(17)30056-7
DOI:
10.1016/j.polar.2018.01.004
Reference:
POLAR 370
To appear in:
Polar Science
Received Date: 5 May 2017 Revised Date:
15 December 2017
Accepted Date: 26 January 2018
Please cite this article as: Daglio, Y., Sacristán, Herná., Ansaldo, Martí., Rodríguez, Marí.C., Benthic diatoms from Potter Cove, 25 de Mayo (King George) Island, Antarctica: Mucilage and glucan storage as a C-source for limpets, Polar Science (2018), doi: 10.1016/j.polar.2018.01.004. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Benthic diatoms from Potter Cove, 25 de Mayo (King George) Island,
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Antarctica: mucilage and glucan storage as a C-source for limpets.
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Yasmin Daglioa,Hernán Sacristánb, Martín Ansaldoc, María C. Rodrígueza,d*
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a
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Departamento de Química Orgánica, Consejo Nacional de Investigaciones
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Científicas y Técnicas- Centro de Investigación de Hidratos de Carbono
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(CIHIDECAR, CONICET-UBA), Ciudad Universitaria-Pabellón 2, Intendente
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Universidad de Buenos Aires, Facultad de Ciencias Exactas y Naturales,
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Güiraldes 2160, C1428EGA Buenos Aires, Argentina
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b
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V9410CAB Ushuaia, Argentina.
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c
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Martín, Buenos Aires, Argentina.
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d
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Departamento de Biodiversidad y Biología Experimental, Ciudad Universitaria-
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Pabellón 2, Intendente Güiraldes 2160, C1428EGA Buenos Aires, Argentina.
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Instituto Antártico Argentino, Avenida 25 de Mayo 1151, B1650HML San
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Universidad de Buenos Aires, Facultad de Ciencias Exactas y Naturales,
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Centro Austral de Investigaciones Científicas, CONICET, Houssay 200,
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* Corresponding author:
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E-mail address: [email protected]
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Abstract Biofilms were allowed to develop on ceramic tiles placed in closed containers on the shore of Potter Cove, 25 de Mayo (King George) Island. Water pumping
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from the cove inside the containers extended for 25 days. Diatoms were the
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dominant microalgae in these biofilms, which were removed from a set of tiles to
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a) characterize the extracellular mucilage, b) carry out floristic determination and
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c) perform grazing experiments with the limpet Nacella concinna. Biofilms
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mucilaginous matrix consisted of proteins and carbohydrates. Room temperature
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aqueous extraction of the freeze-dried material rendered a fraction enriched in the
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storage glucano chrysolaminarin, its identity confirmed by methylation structural
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analyses. Hot water extracted products showed greater heterogeneity in
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monosaccharide composition, including glucose, mannose, galactose, fucose,
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xylose and rhamnose. Diatom identification revealed that Pseudogomphonema
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kamtschaticum was the dominant species followed by several Navicula species,
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Nitzschia pellucida and Synedra kerguelensis. Photographical survey of colonized
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tiles placed in glass flasks together with a specimen of Nacella concinna exhibited
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between 5-30% removal of the biofilms coverage after 24 h of exposure to the
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limpet, suggesting that EPS and chrysolaminarin constitute a C-source for the
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gastropod.
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Keywords Benthic diatoms, Feeding, Gastropods, Nacella concinna, β-1,3-
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glucan, Antarctica, Potter Cove, 25 de Mayo (King George) Island.
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1. Introduction In the last 50 years, especially during austral winter, the Western Antarctic Peninsula (WAP) experienced rapid increases in temperature together with fast
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glacial retreat and strong sea ice decreases (Turner et al., 2009; Ducklow et al.,
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2013). In Potter Cove (25 de Mayo/King George Island) the retreat and melting of
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the Fourcade Glacier has left newly ice-free areas (Rückamp et al., 2011)
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available for benthic colonization (Quartino et al., 2013). These areas are exposed
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to high loads of sediment input from subglacial waters (Eraso and Domíınguez,
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2007), a fact particularly relevant for macroalgal and microalgal communities due
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to the reduction in light penetration within the water column (Schloss et al., 2012;
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Wiencke and Amsler, 2012; Quartino et al., 2013; Deregibus et al., 2016). On the
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other hand, nutrient income into the cove due to glacier underwater run-off has
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been reported to promote benthic microfilm blooms together with a proliferation
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of associated common benthic megafauna (Ahn et al., 2016).
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Diatoms frequently dominate in natural benthic biofilms which can be defined
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as consortia of autotrophic and heterotrophic organisms embedded in a matrix of
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polymers and particles (Decho, 2000).Within these biofilms, benthic diatoms are
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propelled in their gliding movement by mucilage secretion (Molino and
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Wetherbee, 2008). As the cells move on, they leave behind a trail of mucilage,
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which in time is hydrated and partially solubilized (Daglio et al., 2016).
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EPS (extracellular polymeric substances) are biopolymers of microbial origin
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in which biofilms microorganisms are embedded. The nature of EPS presents a
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complex biochemical profile comprising, among others, proteins, glycoproteins,
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sulfated and acid polysaccharides and glycolipids (for reviews see Underwood
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and Paterson, 2003; Delattre et al., 2016; Xiao and Zheng, 2016). This highly
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hydrated matrix attaches biofilms to the surfaces and allows the sequestering of
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dissolved and particulate substances on which biofilms organisms nourish
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(Fleming et al., 2007 and literature therein). Low molecular weight EPS are
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soluble (s-EPS), whereas high molecular weight polymers are associated with the
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cell surface or the substratum, constituting a jelly-like to a solid interface, termed
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bound EPS (b-EPS) (Nielsen and Jahn, 1999, Shniukova and Zolotareva, 2015).
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In addition, diatoms accumulate the storage polysaccharide chrysolaminarin.
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Its structure consists of a β-1,3-glucan backbone chain with different degree of
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substitution in C-2 and/or C-6 (Alekseeva et al., 2005; Storseth et al., 2006; Xia et
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al., 2014), resembling the laminaran of brown seaweeds, except for the lack of
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guluronic and mannuronic acid terminal groups.
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Regardless the chemical nature of the EPS, these polymers together with chrysolaminarin can constitute a carbon source for heterotrophic organisms, such
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as microscopic bacteria, viruses, eukaryotic protists (Pierre et al., 2014) and
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macroscopic grazers. Among the latter, the limpet Nacella concinna, a frequent
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invertebrate of the Antarctic and Sub-Antarctic nearshore fauna has been reported
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to feed on microbial films, on the microepiflora, calcareous rhodophytes, brown
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and red seaweed, bryozoans and sessile spirorbids (polychaetes) (Kim, 2001 and
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references therein; Ansaldo et al., 2007; Suda et al., 2015). N. concinna has a
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worldwide circumpolar distribution and lives in the intertidal up to depths ca. 100
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m. Its ubiquitous distribution in polar regions, together with its feeding habits
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throughout the year, turn this limpet a suitable model organism for histological,
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physiological, biochemical and environmental studies (Ansaldo et al., 2007 and
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references therein). In this work we analyzed the monosaccharide composition of the mucilage and
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intracellular fractions and the possible role of chrysolaminarin and EPS as a
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carbohydrate source in grazing experiments with the limpet Nacella concinna.We
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also present a floristic list of the epipelic and epilithic diatom taxa grown on an
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artificial substrate during the austral summer of 2012-2013 in the Potter Cove
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area, 25 de Mayo (King George) Island, Antarctica. We compare their diversity and relative abundance with previous reports on natural substrates.
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2. Materials y methods
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2.1. Study area
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Potter Cove is located on the southwest coast of 25 de Mayo (King George) Island (62°14'4.23" S, 58°39'53.90" W) in the South Shetland Archipelago of the
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Antarctic Peninsula (Fig. 1a).
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2.2. Diatom colonization on artificial substrate
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Two polypropylene containers (1 m3 capacity each) were located on the shore
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(Fig 1b). The containers received a continuous flow of seawater from the cove by pumping and they were sealed with a dark cover in order to prevent saturating
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natural light intensities. Inside, at the bottom of each of these containers, 30
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ceramic tiles with a rough face (5 x 5 cm) were used as artificial substrate (Fig
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1c). The rough side was placed up to be colonized by diatoms. Colonization was
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carried out during 25 days in January 2013.
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2.3. EPS fractionation and structural analysis Ten randomly selected tiles, colonized as described above, were gently scraped
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with a toothbrush to remove the biofilm and placed in Eppendorf vials for
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preservation at -20°C. Part of this material was used in floristic analysis (see
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below).
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In the laboratory, biofilms material was freeze-dried. The lyophilized material (980 mg) was resuspended and agitated in distilled water at room temperature.
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Cells were separated by centrifugation (10000 ×g for 30 min), and the supernatant
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lyophilized to obtain fraction RTW (room temperature extract, 617 mg). The
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cellular residue was freeze-dried and subjected to extraction in boiling water twice
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(100°C, 1 h), to obtain products HW1 (first hot water extract, 40.8 mg) and HW2
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(second hot water extract, 16.8 mg). Carbohydrate content was dosed by the
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phenol-sulfuric acid method (Dubois et al., 1956) using galactose as standard.
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Protein was determined according to Lowry et al. (1951) referring to bovine
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serum albumin.
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The extract RTW was desalted by gel permeation chromatography in Bio-Gel
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P-2 (Bio-Rad). Void volume (20 mL) was determined with Blue Dextran. Elution
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profile was obtained testing the presence of carbohydrates. The presence of salts
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was tested by precipitation with AgNO3. Fractions containing carbohydrates
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(RTWI and RTWII) were pooled and lyophilized.
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For the structural analyses, RTWI (22 mg) was methylated according to Ciucanu and Kerek (1984) (powdered NaOH in dimethyl sulfoxide-
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iodomethane).The methylated polysaccharide was extracted with chloroform (4
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times), and the extracts dried off. The yield after the first methylation step
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(RTWI-m1) was 6.3 mg, and after the second (RTWI-m2), 3.1 mg.
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2.4. Monosaccharide composition
Sugar composition of the different fractions was determined by gas-liquid
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chromatography (GLC) after hydrolysis of the product with 2M trifluoroacetic
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acid (2 h at 121 ºC) during 90 min. The hydrolysates were then converted to their
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alditol acetates according to Albersheim et al. (1967).
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GLC of the alditol acetates and the partially methylated alditol acetates was
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carried out on a Hewlett–Packard 5890A gas-liquid chromatograph equipped with
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a flame ionization detector and fitted with a fused silica column (0.25 mm i.d.×30
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m) WCOT-coated with a 0.20 µm film of SP-2330. Chromatography was
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performed: (a) isothermally at 220 °C for the alditol acetates; and (b) from 160 °C
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to 210 °C at 2 °C min-1, then from 210 °C to 240 °C at 5 °C min-1 followed by a
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30-min hold for partially methylated alditol acetates (Shea and Carpita, 1988).
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Nitrogen was used as the carrier gas at a flow rate of 1 mL min-1with a split ratio
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80:1. The injector and detector temperatures were set at 240 °C. When necessary,
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GLC–MS analyses were carried out on a Shimadzu QP 5050 A (Kyoto, Japan)
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apparatus working at 70 eV using the same columns and conditions described
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above, but using helium as a carrier gas at a total flow rate of 1 mL min−1; the
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injector temperature was set at 240 °C.
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2.5. Sampling of microalgae Part of the frozen material (see 2.3) was separated for floristic determination. In the laboratory, at least four Eppendorf vials were thawed and subsamples were
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digested with 70% HNO3 and 98% H2SO4 during 24 h, followed by thorough
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rinsing with distilled water until neutral pH. Since significant amounts of
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carbonate residues contaminated the material, we proceeded to removal with 10%
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HCl for 24 h followed by rinsing in distilled water.
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For light microscopy, aliquots of clean valves suspension were dried on
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coverslips and mounted in Naphrax (Hasle and Fryxell, 1970). Slides were
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examined under a Polivar Reichert-Jung light microscope equipped with
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differential interference phase-contrast and digital photographic camera (Canon
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EOS 600D, Melville, New York, USA).
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Cell count proceeded by placing 500 µL of the digested subsamples on an
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object slide. Cell numbers were counted until a total of 400 cells under a known
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number of light fields in the microscope (at least two subsamples were analyzed
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for each sample).
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For scanning electron microscope (SEM) observations, clean valves were
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directly dried on a copper tape and platinum coated for observation in a Gemini
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Zeiss DSM 982 (Jena, Germany) equipped with a field emission gun (FEG) and a
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secondary electron detector (SE) working at a distance of 4 mm and accelerating
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voltage of 15 kV (CMA, FCEyN-UBA). For description of valve morphology we
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followed terminology suggested in Round et al. (1990).
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2.6. Grazing experiment
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Specimens of the Antarctic limpet N. concinna were collected by scuba diving
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at depth of 6-8 m from the subtidal zone of Potter Cove. Prior the experiment, the
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limpets were placed in aquaria with natural photoperiod and aerated seawater for
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an acclimatization period of 48 h (Ansaldo et al., 2007).
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Ten glass flasks (600 mL each) filled with filtered seawater were placed inside the containers. A continuous water flow from the cove inside the containers
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allowed maintaining water temperature inside the flasks identical to the one in the
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cove. Then, a colonized tile (see above) was included in each flask together with a
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previously weighed limpet specimen of Nacella concinna (average weight with
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shell 9±1.5g) during 24 h (n=10) in February 2013. For the purpose of the grazing
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experiment, each colonized tile was photographed at the beginning (time 0) and at
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the end (24 h after). Then, the tile´s photographs were analyzed using ImageJ
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1.47v free software (Rasband, 2016) in order to calculate the area grazed by the
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limpet.
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3. Results
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3.1. Monosaccharide composition and structural analysis of the extracts The composition of all the products obtained by aqueous extractions revealed
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the presence of both carbohydrates and proteins (Table 1). The product obtained at
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room temperature (RTW) showed a higher carbohydrate: protein ratio (2.3) than
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those extracted with hot water (HW1 and HW2) (0.90 and 1.3, respectively).
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Glucose predominated in RTW and HW1 (Table 1), but in the former there appeared smaller amounts of other monosaccharides. HW2 showed the greatest
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heterogeneity of sugars, with significant amounts of glucose, galactose, xylose
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and fucose. Gel permeation chromatography of RTW (Fig.2) allowed the separation of two
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peaks, RTWI and RTWII (yields %/w, 12 and 37% respectively). The higher yield
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of RTWII was due to heavy salt contamination, as revealed by the precipitation
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with AgNO3, a fact that had a negative incidence on its carbohydrate content
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(Table 1).
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Due to its higher carbohydrate: protein ratio, RTWI was selected for structural analysis. Methylation analysis of RTWI indicated the presence of a glucan,
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constituted by a 1,3-linked backbone with branching in C-6 and C-2 (Table 2).
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Undermethylation after the first methylation step was corroborated by the increase
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in trimethylated backbone glucose and the decrease in dimethylated residues after
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the second methylation attempt. Minor undermethylation in RTWI-m2 is
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suggested by small percentages of monomethylated glucose (1,3,4,5,6-tetra-O-
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acetyl-2-O-methylglucitol), and not methylated glucose (3.3% in total). Not taking
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into account these minor percentages in the following rationale, the ratio of
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tetramethylated terminal glucose residues (25.3%) to the rest of the glucose units
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(74.7%) indicates that the glucan backbone is substituted every five glucose
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residues in C-2 (11%), and every nine residues in C-6 (6 %).
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Noteworthy, after gel permeation more important amounts of galactose and
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mannose appeared in RTWI and RTWII in comparison to the original RTW
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product (Table 1). The presence of a galactomannan needs further confirmation by
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structural analysis.
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3.2. Diatom assemblages A total of 16 diatom genera were identified in the samples from the ceramic tiles (Table3) (Figs. 3, 4 and 5).Pseudogomphonema kamtschaticum was the
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dominant species in the analysed biofilm assemblages, followed by several
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Navicula species, Nitzschia pellucida and Synedra kerguelensis. Species such as
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Actinocyclus actinochilus, Entopyla ocellata, Minidiscus chilensis and
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Thalassiosira gracilis appeared very rarely on the tiles (Table 3).
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3.3. Grazing experiment
The area of removed biofilms from the tiles after N. concinna feeding ranged
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from 5 to 30% (Fig. 6). Grazed areas were denoted by a saw-like pattern on the
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tiles due to the action of the mollusks’ radula.
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4. Discussion
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In this work we reproduced diatom benthic diversity of Potter Cove on tiles as artificial substrate, which were employed in grazing experiments with the limpet
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N. concinna. Carbohydrate composition of the diatom biofilms mucilage and the
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identification of chrysolaminarin suggest their putative role as a carbohydrate
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source for the mollusk.
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4.1. EPS and chrysolaminarin as C-source for the limpet N. concinna
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The characterization of the different fractions of the EPS showed an increase in
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protein content with the progress of the aqueous extraction. Room temperature
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product represents the colloidal fraction (sensu Pierre et al., 2014) or the s-EPS
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(sensu Nielsen and Jahn, 1999). Instead, hot water extracted fractions correspond
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to b-EPS. But, as pointed out by Hanlon et al. (2006) it must be emphasized that
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these extraction procedures do not isolate specific polymer types but broadly
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separate different biofilm carbohydrates based on solubility and binding
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characteristics. Additionally, some contamination with intracellular material
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cannot be discarded in all the fractions, since the algae were frozen for
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preservation a fact that probably caused cell rupture and intracellular leaking
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(Passarelli et al., 2015). This could explain the abundance of glucose, originated
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from intracellular chrysolaminarin in all the products.
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Natural biofilms consist of consortia of different prokaryotic and eukaryotic
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organisms. Differing from unialgal laboratory cultures, the characterization and
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comparison of specific polysaccharides among different samples in natural
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biofilms relies on monosaccharide composition. Several similarities in
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monosaccharide composition of the different fractions obtained in the present
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work can be found with data in literature, such as predominance of glucose in
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RTW (equivalent to colloidal EPS) as reported by different authors (Taylor et al.,
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1999; Hanlon et al., 2010; Pierre et al., 2012, Passarelli et al., 2015) or more
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heterogeneous monosaccharide composition in HW2 (equivalent to b-EPS). In an
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attempt to establish a certain fingerprint pattern for biofilms based on
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monosaccharide composition, cluster analyses were carried out by Underwood
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and Paterson (2003) and Shniukova and Zolotareva (2015). The monosaccharide
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content of RTW, HW1 and RTWI resembles that of cluster E of these authors,
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described as “the group including attached, capsular or related to bottom
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sediments EPS and intracellular glucans and characterized by high levels of
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glucose”.
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Structural analyses of glucose-rich fractions confirmed the presence of chrysolaminarin, a β-(1→3)-D-glucan. In this case, the backbone chain exhibited
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branches in C-2 and C-6. A similar structure was reported for the chrysolaminarin
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of Skeletonema costatum (Paulsen and Myklestad, 1978), Stauroneis amphioxys
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(McConville et al., 1986) and Achnanthes longipes (Wustman et al., 1997). Yet it
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must be emphasized that structural analyses was not applied to unialgal samples,
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so the results represent an average structure for the storage polysaccharide.
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Benthic diatoms accumulate chrysolaminarin during daytime emersion period (Hanlon et al., 2006) which could be consumed by the limpet as a C-source. In
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gastropods, β-(1→3)-endoglucanase activity has been reported in Telescopium
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telescopium (Cutler and Yellowlees, 1979), Haliotis tuberculata (Lépagnol-
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Descamps et al., 1998) and Haliotis discus hannai (Kumagai and Ojima, 2009).
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Though identification of β-(1→3)-endoglucanase in N. concinna was not
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attempted in the present work, the fact that chrysolaminarin was abundant in the
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unique food (biofilms) offered and consumed by the limpets strongly supports the
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hypothesis of glucanase activity in the mollusk.
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In addition to being an energy-rich substrate, β-glucans (especially β-1,3/1,6-
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glucans) interact with receptors on immune cells and elicit specific biological
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responses, depending on the degree of substitution of the skeletal chain with
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lateral branches and on the length of these ramifications (Barsanti et al., 2011;
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Raa, 2015). Due to their properties they have been introduced as additives to
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improve the health and performance in marine ranching of fish and shrimp (Raa,
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2015 and references therein). Recently, antioxidant properties in the
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chrysolaminarin isolated from the diatom Odontella aurita have been informed by
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Xia et al. (2014), an interesting feature if we consider enhanced oxidative stress
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by UV exposure of Antarctic invertebrates in their natural environment.
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As feeding stock, diatoms are also a source of Polyunsaturated Fatty Acids
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(PUFAs) including docosahexaenoic acid (DHA), eicosapentaenoic acid (EPA),
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arachidonic acid (AA), γ-linolenic acid (GLA) (Barnech Bielsa et al., 2016),
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together with the antioxidant fucoxanthin (Xia et al., 2013).
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3.2. Biofilms development on artificial substrates
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In our experimental setting (tiles in the cabinets exposed to a continuous water flow from the Cove) microalgae settlement occurred with a similar diversity and
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abundance of species as the one reported in recent benthic diatoms inventories in
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Potter Cove (Al-Handal and Wulff, 2008a, 2008b). This indicates that both
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exposure time of artificial substrata and environmental factors in the containers
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were adequate to reproduce the conditions of the natural benthic microalgae
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community in the Cove. Yet, some differences were encountered. For example,
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Entopyla ocellata, Licmophora antarctica, Odontella litigiosa and Pinnularia
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quadratarea appeared rarely and very rarely on the tiles. Benthic diatoms are
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rapid colonizers in polar primary succession (Campana et al., 2009). Kang et al.
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(2002) reported that summer conditions (low salinity and high air temperature)
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were more suitable for the growth of Cocconeis sp and Pseudogomphonema
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kamtschaticum. The latter species was also abundant in our samples, but not the
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former one. It must be taken into account Cocconeis sp is more susceptible to cell
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rupture due to freezing of the tiles (Zacher et al., 2007), a fact that could lead to an
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underestimation of its abundance. On the other hand, mechanical robustness of the
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frustules of Synedra kerguelensis (Díaz et al., 2015) can result in its higher
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abundance in the processed samples.
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Colonization is subjected both to biotic and abiotic factors, which can modify the intrinsic growth rate of different diatom species, favoring those with high
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growth rates in efficient propagation on the substrate. The colonization process of
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hard-bottom polar benthic communities has been studied by clearing natural
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substrata and placement of artificial substrates, such as ceramic panels followed
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by direct observation, underwater photography or transference of colonized
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substrata to the laboratory (Barnes and Conlan, 2007; Beuchel and Gulliksen,
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2008; Campana et al., 2009 and references therein). Colonization on artificial
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substrata, as the ones employed in the present work, has been used to standardize
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sampling methods on sites of various natural bottom types or depths (Desrosiers et
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al., 2014 and literature within). Artificial substrates facilitate the testing of
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environmental conditions such as UV radiation and grazing (Wahl et al., 2004;
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Konar, 2007; Zacher et al., 2007; Fricke et al., 2008; Zacher and Campana, 2008)
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both in situ or after transfer to the laboratory.
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3.3. Grazing experiments
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Previous field observations indicated that the limpet is always found on rocky
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substrates or grazing upon macroalgae, but very rarely on soft bottoms, since
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these prevent firm attachment of the mollusk. Weak adhesion to the bottom turns
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the limpet more susceptible to the effect of waves or to predation by birds. Based
342
on the observations of the natural habitat of the limpet, we chose a hard artificial
343
substrate to be colonized by diatoms. The photographical record of the tiles along
344
the grazing experiments in the containers demonstrated that the limpet N.
345
concinna can consume diatom biofilms for nourishment. Taking into account that
346
the grazing marks on the tiles indicate a 5-30% level of activity of the limpet and
347
being N. concinna the most abundant mollusk in Antarctica, this would represent
348
an important transfer of organic matter from primary producers in the biofilms to
349
the limpet and to other secondary consumers (birds, urchins and others) in higher
350
levels of the food web. Massive growth of benthic diatoms in Marian Cove was
351
reported concomitant with the proliferation of a rich assemblage of benthic
352
megafauna comprising stalked ascidians, sponges, polychaetes, gastropods,
353
bivalves, bryozoans and gorgonians (Ahn et al., 2016), strongly suggesting that
354
the input of diatom biomass can support feeding requirements of at least part of
355
the associated fauna.
SC
M AN U
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The grazing on microphytobenthos by the limpet N. concinna has been
EP
356
RI PT
341
established in the subtidal of the Antarctic Peninsula (Dunton, 2001; Corbisier et
358
al., 2004) and in King George Island (Choy et al., 2011) with the δ13C isotope
359
analysis, that approximate the isotopic composition of their respective diets but to
360
the best of our knowledge this is the first report of the characterization of
361
chrysolaminarin in diatom biofilms grazed by the limpet as a possible carbon
362
source.
363
Acknowledgements
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The authors want to thanks especially to Dr. Nora I. Maidana (DBBE, FCEN,
365
UBA) for help in diatom identification and Dr. María Cristina Matulewicz (Depto.
366
Química Orgánica, CIHIDECAR, CONICET-UBA) for suggestions in structural
367
analysis. This work was funded by grants from UBA (GC 20020130100216BA)
368
and FONCyT- DNA (PICTO 0091). YD and HS are a Research Fellows of the
369
National Research Council of Argentina (CONICET).
371
SC
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571
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Tables Table 1: Monosaccharide composition (mol%) of the different mucilage fractions
575
extracted with water at room temperature (RTW), boiling water (HW1 and HW2)
576
and after RTW desalting in BioGel P2 (RTWI and RTWII)
RI PT
574
Constituent monosaccharides (mol%) Protein (%)
Carbohydrates (%)
Rha
Fuc
Ara
Xyl
Man
Glc
Gal
3.9
9.3
0.7
1.0
1.0
1.2
1.4
91.1
3.5
HW1
14.8
19.7
-
-
-
-
1.2
92.9
6.0
HW2
12.9
11.6
6.6
12.0
0.9
15.4
8.3
35.7 21.3
RTWI
n.d.
8.9
-
-
-
-
16.4 68.8 14.8
RTWII
n.d.
4.9
4.3
20.9
-
-
13.7 36.6 24.5
M AN U
SC
RTW
577
Abbreviations: Rha= rhamnose; Fuc= fucose; Ara= arabinose; Xyl= xylose; Man=
579
mannose; Gal= galactose; Glc= glucose.; n.d.= not determined
EP AC C
580
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578
27
ACCEPTED MANUSCRIPT
581
Table 2: Linkage analysis of constituent monosaccharides (mol %) produced by
582
methylation and hydrolysis of product RTWI after one (RTWI-m1) and two
583
(RTWI-m2) methylation steps.
Monosaccharidea
Deduced glycosidic linkage
RI PT
584
RTWI-m1 RTWI-m2
Terminal
20.1
25.3
2,4,6-Me3Glc
3-linked
44.1
55.1
4,6-Me2Glc
3-linked
16.6
10.2
M AN U
2-substituted
SC
2,3,4,6-Me4Glc
2,4-Me2Glc
3-linked
10.2
6.1
6.3
2.3
2.8
1.1
6-substituted 2- MeGlc
TE D
Glc 585
Abbreviations: 2-MeGlc=1,3,4,5,6-tetra-O-acetyl-2-O-methylglucitol; 2,4-
587
Me2Glc=1,3,5,6-tetra-O-acetyl- 2,4-di-O-methylglucitol; 4,6-Me2Glc= 1,2,3,5-
588
tetra-O-acetyl-4,6-di-O-methylglucitol; 2,4,6-Me3Glc = 1,3,5-tri-O-acetyl-2,4,6-
589
tri-O-methylglucitol; 2,3,4,6-Me4Glc=1,5-di-O-acetyl-2,3,4,6-tetra-O-
590
methylglucitol.
AC C
591
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586
592
28
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593
Table 3.Marine benthic diatom species recorded at Potter Cove, King George
594
Island, Antarctica, during the austral summer of 2013. A total of 400 valves were
595
counted.
Achnanthes bongrainii (Peragallo) Mann
r
Actinocyclus actinochilus (Ehrenberg) Simonsen
vr vr
Coccocneis costatavar. antárctica Manguin
M AN U
Coccocneis fasciolata (Ehrenberg) Brown
SC
Amphora sp.
RI PT
Relative abundance
r r r
Entopyla ocellata (Arnott) Grunow
vr
Licmophora antárctica G.W.F.Carlson
r
Minidiscus chilensis Rivera
vr
TE D
Coccocneis pinnata Gregory ex Greville
f
Navicula cf. perminuta Grunow
f
Navicula sp.
f
EP
Navicula directa (Smith) Ralfs
f
AC C
Nitzschia pellucida Grunow Odontella litigiosa (Van Heurck) Hoban
r
Parlibellus delognei (Van Heurck) Cox
r
Petroneis plagiostoma (Grunow)
r
Petroneis sp.
r
Pinnularia quadratarea (A. Schmidt)
r
Pseudogomphonema kamtschaticum (Grunow)
c
29
ACCEPTED MANUSCRIPT
Synedra kerguelensis Heiden
f
Thalassiosira gracilis (Karsten) Hustedt
vr
596
Relative abundance represents the occurrence of a species in all samples for a
598
given area; vr: very rare, found only once on a slide, r: rare <5%, f: frequent, 5–
599
25%, c: common >25%.
RI PT
597
AC C
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M AN U
SC
600
30
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Figure captions
602
Fig. 1: a Potter Cove map, b Containers, c a detail of flasks distribution inside the
603
containers, each with a colonized tile and a Nacella concinna specimen (black
604
arrowhead).
605
Fig. 2: Bio-Gel P-2 filtration elution profile of fraction RTW. Void volume (V0) is
606
indicated (small black arrow). Fractions indicated between the full black and
607
hatched arrows showed salt contamination.
608
Fig. 3: Marine benthic diatoms from 25 de Mayo (Potter Cove) Antarctica on
609
colonized ceramic tiles, viewed in LM. a Achnanthes bongrainii, b Entopyla
610
ocellata, c Parlibellus delognei, d Licmophora antarctica, e Odontella litigiosa, f
611
Amphora sp. g Actinocyclus actinochilus, h Cocconeis pinnata,I Cocconeis
612
costatavar. antarctica, j Cocconeis fasciolata. Scale bar= 10 µm.
613
Fig. 4: Marine benthic diatoms from 25 de Mayo (Potter Cove) Antarctica on
614
colonized ceramic tiles, viewed in LM. a Nitzschia pellucida, b Navicula directa,
615
c Petroneis sp., d Thalassiosira gracilis, e Navicula cf.perminuta, f Navicula sp, g
616
Petroneis plagiostoma, h Pinnularia quadratarea, I Synedra kerguelensis, j
617
Pseudogomphonema kamtschaticum, k Pleurosigma sp. Scale bar= d-f: 5 µm; for
618
the rest: 10 µm.
619
Fig. 5: Marine benthic diatoms from 25 de Mayo (Potter Cove) Antactica on
620
colonized ceramic tiles, viewed in SEM. a Odontella litigiosa, b Minidiscus
621
chilensis, c Navicula sp , d Navicula cf. perminuta, e-f Pseudogomphonema
622
kamtschaticum in external and internal valve view, g Synedra kerguelensis, h
623
Nitzschia pellucida. Scale bar= b: 1 µm; for the rest: 5 µm.
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31
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Fig. 6: Grazing experiment, a-bColonized tiles at time 0; c-dTiles after 24 h.
625
Scale= 1 cm.
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DOI:
10.1016/j.polar.2018.01.004
Reference:
POLAR 370
To appear in:
Polar Science
Received Date: 5 May 2017 Revised Date:
15 December 2017
Accepted Date: 26 January 2018
Please cite this article as: Daglio, Y., Sacristán, Herná., Ansaldo, Martí., Rodríguez, Marí.C., Benthic diatoms from Potter Cove, 25 de Mayo (King George) Island, Antarctica: Mucilage and glucan storage as a C-source for limpets, Polar Science (2018), doi: 10.1016/j.polar.2018.01.004. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Benthic diatoms from Potter Cove, 25 de Mayo (King George) Island,
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Antarctica: mucilage and glucan storage as a C-source for limpets.
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Yasmin Daglioa,Hernán Sacristánb, Martín Ansaldoc, María C. Rodrígueza,d*
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a
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Departamento de Química Orgánica, Consejo Nacional de Investigaciones
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Científicas y Técnicas- Centro de Investigación de Hidratos de Carbono
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(CIHIDECAR, CONICET-UBA), Ciudad Universitaria-Pabellón 2, Intendente
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Universidad de Buenos Aires, Facultad de Ciencias Exactas y Naturales,
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Güiraldes 2160, C1428EGA Buenos Aires, Argentina
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b
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V9410CAB Ushuaia, Argentina.
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c
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Martín, Buenos Aires, Argentina.
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d
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Departamento de Biodiversidad y Biología Experimental, Ciudad Universitaria-
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Pabellón 2, Intendente Güiraldes 2160, C1428EGA Buenos Aires, Argentina.
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Instituto Antártico Argentino, Avenida 25 de Mayo 1151, B1650HML San
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Universidad de Buenos Aires, Facultad de Ciencias Exactas y Naturales,
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Centro Austral de Investigaciones Científicas, CONICET, Houssay 200,
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* Corresponding author:
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E-mail address: [email protected]
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Abstract Biofilms were allowed to develop on ceramic tiles placed in closed containers on the shore of Potter Cove, 25 de Mayo (King George) Island. Water pumping
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from the cove inside the containers extended for 25 days. Diatoms were the
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dominant microalgae in these biofilms, which were removed from a set of tiles to
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a) characterize the extracellular mucilage, b) carry out floristic determination and
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c) perform grazing experiments with the limpet Nacella concinna. Biofilms
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mucilaginous matrix consisted of proteins and carbohydrates. Room temperature
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aqueous extraction of the freeze-dried material rendered a fraction enriched in the
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storage glucano chrysolaminarin, its identity confirmed by methylation structural
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analyses. Hot water extracted products showed greater heterogeneity in
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monosaccharide composition, including glucose, mannose, galactose, fucose,
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xylose and rhamnose. Diatom identification revealed that Pseudogomphonema
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kamtschaticum was the dominant species followed by several Navicula species,
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Nitzschia pellucida and Synedra kerguelensis. Photographical survey of colonized
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tiles placed in glass flasks together with a specimen of Nacella concinna exhibited
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between 5-30% removal of the biofilms coverage after 24 h of exposure to the
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limpet, suggesting that EPS and chrysolaminarin constitute a C-source for the
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gastropod.
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Keywords Benthic diatoms, Feeding, Gastropods, Nacella concinna, β-1,3-
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glucan, Antarctica, Potter Cove, 25 de Mayo (King George) Island.
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1. Introduction In the last 50 years, especially during austral winter, the Western Antarctic Peninsula (WAP) experienced rapid increases in temperature together with fast
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glacial retreat and strong sea ice decreases (Turner et al., 2009; Ducklow et al.,
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2013). In Potter Cove (25 de Mayo/King George Island) the retreat and melting of
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the Fourcade Glacier has left newly ice-free areas (Rückamp et al., 2011)
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available for benthic colonization (Quartino et al., 2013). These areas are exposed
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to high loads of sediment input from subglacial waters (Eraso and Domíınguez,
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2007), a fact particularly relevant for macroalgal and microalgal communities due
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to the reduction in light penetration within the water column (Schloss et al., 2012;
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Wiencke and Amsler, 2012; Quartino et al., 2013; Deregibus et al., 2016). On the
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other hand, nutrient income into the cove due to glacier underwater run-off has
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been reported to promote benthic microfilm blooms together with a proliferation
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of associated common benthic megafauna (Ahn et al., 2016).
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Diatoms frequently dominate in natural benthic biofilms which can be defined
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as consortia of autotrophic and heterotrophic organisms embedded in a matrix of
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polymers and particles (Decho, 2000).Within these biofilms, benthic diatoms are
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propelled in their gliding movement by mucilage secretion (Molino and
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Wetherbee, 2008). As the cells move on, they leave behind a trail of mucilage,
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which in time is hydrated and partially solubilized (Daglio et al., 2016).
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EPS (extracellular polymeric substances) are biopolymers of microbial origin
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in which biofilms microorganisms are embedded. The nature of EPS presents a
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complex biochemical profile comprising, among others, proteins, glycoproteins,
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sulfated and acid polysaccharides and glycolipids (for reviews see Underwood
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and Paterson, 2003; Delattre et al., 2016; Xiao and Zheng, 2016). This highly
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hydrated matrix attaches biofilms to the surfaces and allows the sequestering of
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dissolved and particulate substances on which biofilms organisms nourish
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(Fleming et al., 2007 and literature therein). Low molecular weight EPS are
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soluble (s-EPS), whereas high molecular weight polymers are associated with the
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cell surface or the substratum, constituting a jelly-like to a solid interface, termed
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bound EPS (b-EPS) (Nielsen and Jahn, 1999, Shniukova and Zolotareva, 2015).
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In addition, diatoms accumulate the storage polysaccharide chrysolaminarin.
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Its structure consists of a β-1,3-glucan backbone chain with different degree of
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substitution in C-2 and/or C-6 (Alekseeva et al., 2005; Storseth et al., 2006; Xia et
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al., 2014), resembling the laminaran of brown seaweeds, except for the lack of
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guluronic and mannuronic acid terminal groups.
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Regardless the chemical nature of the EPS, these polymers together with chrysolaminarin can constitute a carbon source for heterotrophic organisms, such
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as microscopic bacteria, viruses, eukaryotic protists (Pierre et al., 2014) and
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macroscopic grazers. Among the latter, the limpet Nacella concinna, a frequent
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invertebrate of the Antarctic and Sub-Antarctic nearshore fauna has been reported
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to feed on microbial films, on the microepiflora, calcareous rhodophytes, brown
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and red seaweed, bryozoans and sessile spirorbids (polychaetes) (Kim, 2001 and
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references therein; Ansaldo et al., 2007; Suda et al., 2015). N. concinna has a
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worldwide circumpolar distribution and lives in the intertidal up to depths ca. 100
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m. Its ubiquitous distribution in polar regions, together with its feeding habits
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throughout the year, turn this limpet a suitable model organism for histological,
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physiological, biochemical and environmental studies (Ansaldo et al., 2007 and
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references therein). In this work we analyzed the monosaccharide composition of the mucilage and
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intracellular fractions and the possible role of chrysolaminarin and EPS as a
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carbohydrate source in grazing experiments with the limpet Nacella concinna.We
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also present a floristic list of the epipelic and epilithic diatom taxa grown on an
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artificial substrate during the austral summer of 2012-2013 in the Potter Cove
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area, 25 de Mayo (King George) Island, Antarctica. We compare their diversity and relative abundance with previous reports on natural substrates.
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2. Materials y methods
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2.1. Study area
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Potter Cove is located on the southwest coast of 25 de Mayo (King George) Island (62°14'4.23" S, 58°39'53.90" W) in the South Shetland Archipelago of the
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Antarctic Peninsula (Fig. 1a).
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2.2. Diatom colonization on artificial substrate
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Two polypropylene containers (1 m3 capacity each) were located on the shore
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(Fig 1b). The containers received a continuous flow of seawater from the cove by pumping and they were sealed with a dark cover in order to prevent saturating
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natural light intensities. Inside, at the bottom of each of these containers, 30
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ceramic tiles with a rough face (5 x 5 cm) were used as artificial substrate (Fig
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1c). The rough side was placed up to be colonized by diatoms. Colonization was
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carried out during 25 days in January 2013.
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2.3. EPS fractionation and structural analysis Ten randomly selected tiles, colonized as described above, were gently scraped
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with a toothbrush to remove the biofilm and placed in Eppendorf vials for
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preservation at -20°C. Part of this material was used in floristic analysis (see
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below).
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In the laboratory, biofilms material was freeze-dried. The lyophilized material (980 mg) was resuspended and agitated in distilled water at room temperature.
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Cells were separated by centrifugation (10000 ×g for 30 min), and the supernatant
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lyophilized to obtain fraction RTW (room temperature extract, 617 mg). The
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cellular residue was freeze-dried and subjected to extraction in boiling water twice
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(100°C, 1 h), to obtain products HW1 (first hot water extract, 40.8 mg) and HW2
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(second hot water extract, 16.8 mg). Carbohydrate content was dosed by the
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phenol-sulfuric acid method (Dubois et al., 1956) using galactose as standard.
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Protein was determined according to Lowry et al. (1951) referring to bovine
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serum albumin.
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P-2 (Bio-Rad). Void volume (20 mL) was determined with Blue Dextran. Elution
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profile was obtained testing the presence of carbohydrates. The presence of salts
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was tested by precipitation with AgNO3. Fractions containing carbohydrates
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(RTWI and RTWII) were pooled and lyophilized.
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For the structural analyses, RTWI (22 mg) was methylated according to Ciucanu and Kerek (1984) (powdered NaOH in dimethyl sulfoxide-
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iodomethane).The methylated polysaccharide was extracted with chloroform (4
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times), and the extracts dried off. The yield after the first methylation step
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(RTWI-m1) was 6.3 mg, and after the second (RTWI-m2), 3.1 mg.
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2.4. Monosaccharide composition
Sugar composition of the different fractions was determined by gas-liquid
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chromatography (GLC) after hydrolysis of the product with 2M trifluoroacetic
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acid (2 h at 121 ºC) during 90 min. The hydrolysates were then converted to their
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alditol acetates according to Albersheim et al. (1967).
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GLC of the alditol acetates and the partially methylated alditol acetates was
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carried out on a Hewlett–Packard 5890A gas-liquid chromatograph equipped with
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a flame ionization detector and fitted with a fused silica column (0.25 mm i.d.×30
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m) WCOT-coated with a 0.20 µm film of SP-2330. Chromatography was
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performed: (a) isothermally at 220 °C for the alditol acetates; and (b) from 160 °C
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to 210 °C at 2 °C min-1, then from 210 °C to 240 °C at 5 °C min-1 followed by a
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30-min hold for partially methylated alditol acetates (Shea and Carpita, 1988).
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Nitrogen was used as the carrier gas at a flow rate of 1 mL min-1with a split ratio
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80:1. The injector and detector temperatures were set at 240 °C. When necessary,
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GLC–MS analyses were carried out on a Shimadzu QP 5050 A (Kyoto, Japan)
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apparatus working at 70 eV using the same columns and conditions described
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above, but using helium as a carrier gas at a total flow rate of 1 mL min−1; the
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injector temperature was set at 240 °C.
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2.5. Sampling of microalgae Part of the frozen material (see 2.3) was separated for floristic determination. In the laboratory, at least four Eppendorf vials were thawed and subsamples were
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digested with 70% HNO3 and 98% H2SO4 during 24 h, followed by thorough
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rinsing with distilled water until neutral pH. Since significant amounts of
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carbonate residues contaminated the material, we proceeded to removal with 10%
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HCl for 24 h followed by rinsing in distilled water.
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For light microscopy, aliquots of clean valves suspension were dried on
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coverslips and mounted in Naphrax (Hasle and Fryxell, 1970). Slides were
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examined under a Polivar Reichert-Jung light microscope equipped with
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differential interference phase-contrast and digital photographic camera (Canon
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EOS 600D, Melville, New York, USA).
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Cell count proceeded by placing 500 µL of the digested subsamples on an
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object slide. Cell numbers were counted until a total of 400 cells under a known
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number of light fields in the microscope (at least two subsamples were analyzed
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for each sample).
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For scanning electron microscope (SEM) observations, clean valves were
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directly dried on a copper tape and platinum coated for observation in a Gemini
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Zeiss DSM 982 (Jena, Germany) equipped with a field emission gun (FEG) and a
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secondary electron detector (SE) working at a distance of 4 mm and accelerating
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voltage of 15 kV (CMA, FCEyN-UBA). For description of valve morphology we
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followed terminology suggested in Round et al. (1990).
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2.6. Grazing experiment
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Specimens of the Antarctic limpet N. concinna were collected by scuba diving
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at depth of 6-8 m from the subtidal zone of Potter Cove. Prior the experiment, the
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limpets were placed in aquaria with natural photoperiod and aerated seawater for
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an acclimatization period of 48 h (Ansaldo et al., 2007).
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Ten glass flasks (600 mL each) filled with filtered seawater were placed inside the containers. A continuous water flow from the cove inside the containers
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allowed maintaining water temperature inside the flasks identical to the one in the
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cove. Then, a colonized tile (see above) was included in each flask together with a
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previously weighed limpet specimen of Nacella concinna (average weight with
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shell 9±1.5g) during 24 h (n=10) in February 2013. For the purpose of the grazing
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experiment, each colonized tile was photographed at the beginning (time 0) and at
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the end (24 h after). Then, the tile´s photographs were analyzed using ImageJ
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1.47v free software (Rasband, 2016) in order to calculate the area grazed by the
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limpet.
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3. Results
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3.1. Monosaccharide composition and structural analysis of the extracts The composition of all the products obtained by aqueous extractions revealed
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the presence of both carbohydrates and proteins (Table 1). The product obtained at
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room temperature (RTW) showed a higher carbohydrate: protein ratio (2.3) than
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those extracted with hot water (HW1 and HW2) (0.90 and 1.3, respectively).
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Glucose predominated in RTW and HW1 (Table 1), but in the former there appeared smaller amounts of other monosaccharides. HW2 showed the greatest
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heterogeneity of sugars, with significant amounts of glucose, galactose, xylose
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and fucose. Gel permeation chromatography of RTW (Fig.2) allowed the separation of two
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peaks, RTWI and RTWII (yields %/w, 12 and 37% respectively). The higher yield
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of RTWII was due to heavy salt contamination, as revealed by the precipitation
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with AgNO3, a fact that had a negative incidence on its carbohydrate content
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(Table 1).
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Due to its higher carbohydrate: protein ratio, RTWI was selected for structural analysis. Methylation analysis of RTWI indicated the presence of a glucan,
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constituted by a 1,3-linked backbone with branching in C-6 and C-2 (Table 2).
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Undermethylation after the first methylation step was corroborated by the increase
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in trimethylated backbone glucose and the decrease in dimethylated residues after
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the second methylation attempt. Minor undermethylation in RTWI-m2 is
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suggested by small percentages of monomethylated glucose (1,3,4,5,6-tetra-O-
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acetyl-2-O-methylglucitol), and not methylated glucose (3.3% in total). Not taking
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into account these minor percentages in the following rationale, the ratio of
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tetramethylated terminal glucose residues (25.3%) to the rest of the glucose units
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(74.7%) indicates that the glucan backbone is substituted every five glucose
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residues in C-2 (11%), and every nine residues in C-6 (6 %).
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Noteworthy, after gel permeation more important amounts of galactose and
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mannose appeared in RTWI and RTWII in comparison to the original RTW
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product (Table 1). The presence of a galactomannan needs further confirmation by
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structural analysis.
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3.2. Diatom assemblages A total of 16 diatom genera were identified in the samples from the ceramic tiles (Table3) (Figs. 3, 4 and 5).Pseudogomphonema kamtschaticum was the
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dominant species in the analysed biofilm assemblages, followed by several
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Navicula species, Nitzschia pellucida and Synedra kerguelensis. Species such as
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Actinocyclus actinochilus, Entopyla ocellata, Minidiscus chilensis and
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Thalassiosira gracilis appeared very rarely on the tiles (Table 3).
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3.3. Grazing experiment
The area of removed biofilms from the tiles after N. concinna feeding ranged
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from 5 to 30% (Fig. 6). Grazed areas were denoted by a saw-like pattern on the
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tiles due to the action of the mollusks’ radula.
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4. Discussion
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In this work we reproduced diatom benthic diversity of Potter Cove on tiles as artificial substrate, which were employed in grazing experiments with the limpet
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N. concinna. Carbohydrate composition of the diatom biofilms mucilage and the
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identification of chrysolaminarin suggest their putative role as a carbohydrate
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source for the mollusk.
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4.1. EPS and chrysolaminarin as C-source for the limpet N. concinna
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The characterization of the different fractions of the EPS showed an increase in
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protein content with the progress of the aqueous extraction. Room temperature
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product represents the colloidal fraction (sensu Pierre et al., 2014) or the s-EPS
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(sensu Nielsen and Jahn, 1999). Instead, hot water extracted fractions correspond
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to b-EPS. But, as pointed out by Hanlon et al. (2006) it must be emphasized that
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these extraction procedures do not isolate specific polymer types but broadly
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separate different biofilm carbohydrates based on solubility and binding
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characteristics. Additionally, some contamination with intracellular material
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cannot be discarded in all the fractions, since the algae were frozen for
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preservation a fact that probably caused cell rupture and intracellular leaking
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(Passarelli et al., 2015). This could explain the abundance of glucose, originated
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from intracellular chrysolaminarin in all the products.
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Natural biofilms consist of consortia of different prokaryotic and eukaryotic
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organisms. Differing from unialgal laboratory cultures, the characterization and
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comparison of specific polysaccharides among different samples in natural
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biofilms relies on monosaccharide composition. Several similarities in
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monosaccharide composition of the different fractions obtained in the present
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work can be found with data in literature, such as predominance of glucose in
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RTW (equivalent to colloidal EPS) as reported by different authors (Taylor et al.,
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1999; Hanlon et al., 2010; Pierre et al., 2012, Passarelli et al., 2015) or more
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heterogeneous monosaccharide composition in HW2 (equivalent to b-EPS). In an
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attempt to establish a certain fingerprint pattern for biofilms based on
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monosaccharide composition, cluster analyses were carried out by Underwood
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and Paterson (2003) and Shniukova and Zolotareva (2015). The monosaccharide
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content of RTW, HW1 and RTWI resembles that of cluster E of these authors,
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described as “the group including attached, capsular or related to bottom
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sediments EPS and intracellular glucans and characterized by high levels of
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glucose”.
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Structural analyses of glucose-rich fractions confirmed the presence of chrysolaminarin, a β-(1→3)-D-glucan. In this case, the backbone chain exhibited
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branches in C-2 and C-6. A similar structure was reported for the chrysolaminarin
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of Skeletonema costatum (Paulsen and Myklestad, 1978), Stauroneis amphioxys
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(McConville et al., 1986) and Achnanthes longipes (Wustman et al., 1997). Yet it
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must be emphasized that structural analyses was not applied to unialgal samples,
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so the results represent an average structure for the storage polysaccharide.
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Benthic diatoms accumulate chrysolaminarin during daytime emersion period (Hanlon et al., 2006) which could be consumed by the limpet as a C-source. In
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gastropods, β-(1→3)-endoglucanase activity has been reported in Telescopium
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telescopium (Cutler and Yellowlees, 1979), Haliotis tuberculata (Lépagnol-
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Descamps et al., 1998) and Haliotis discus hannai (Kumagai and Ojima, 2009).
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Though identification of β-(1→3)-endoglucanase in N. concinna was not
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attempted in the present work, the fact that chrysolaminarin was abundant in the
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unique food (biofilms) offered and consumed by the limpets strongly supports the
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hypothesis of glucanase activity in the mollusk.
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In addition to being an energy-rich substrate, β-glucans (especially β-1,3/1,6-
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glucans) interact with receptors on immune cells and elicit specific biological
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responses, depending on the degree of substitution of the skeletal chain with
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lateral branches and on the length of these ramifications (Barsanti et al., 2011;
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Raa, 2015). Due to their properties they have been introduced as additives to
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improve the health and performance in marine ranching of fish and shrimp (Raa,
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2015 and references therein). Recently, antioxidant properties in the
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chrysolaminarin isolated from the diatom Odontella aurita have been informed by
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Xia et al. (2014), an interesting feature if we consider enhanced oxidative stress
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by UV exposure of Antarctic invertebrates in their natural environment.
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As feeding stock, diatoms are also a source of Polyunsaturated Fatty Acids
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(PUFAs) including docosahexaenoic acid (DHA), eicosapentaenoic acid (EPA),
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arachidonic acid (AA), γ-linolenic acid (GLA) (Barnech Bielsa et al., 2016),
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together with the antioxidant fucoxanthin (Xia et al., 2013).
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3.2. Biofilms development on artificial substrates
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In our experimental setting (tiles in the cabinets exposed to a continuous water flow from the Cove) microalgae settlement occurred with a similar diversity and
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abundance of species as the one reported in recent benthic diatoms inventories in
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Potter Cove (Al-Handal and Wulff, 2008a, 2008b). This indicates that both
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exposure time of artificial substrata and environmental factors in the containers
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were adequate to reproduce the conditions of the natural benthic microalgae
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community in the Cove. Yet, some differences were encountered. For example,
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Entopyla ocellata, Licmophora antarctica, Odontella litigiosa and Pinnularia
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quadratarea appeared rarely and very rarely on the tiles. Benthic diatoms are
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rapid colonizers in polar primary succession (Campana et al., 2009). Kang et al.
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(2002) reported that summer conditions (low salinity and high air temperature)
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were more suitable for the growth of Cocconeis sp and Pseudogomphonema
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kamtschaticum. The latter species was also abundant in our samples, but not the
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former one. It must be taken into account Cocconeis sp is more susceptible to cell
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rupture due to freezing of the tiles (Zacher et al., 2007), a fact that could lead to an
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underestimation of its abundance. On the other hand, mechanical robustness of the
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frustules of Synedra kerguelensis (Díaz et al., 2015) can result in its higher
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abundance in the processed samples.
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Colonization is subjected both to biotic and abiotic factors, which can modify the intrinsic growth rate of different diatom species, favoring those with high
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growth rates in efficient propagation on the substrate. The colonization process of
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hard-bottom polar benthic communities has been studied by clearing natural
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substrata and placement of artificial substrates, such as ceramic panels followed
328
by direct observation, underwater photography or transference of colonized
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substrata to the laboratory (Barnes and Conlan, 2007; Beuchel and Gulliksen,
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2008; Campana et al., 2009 and references therein). Colonization on artificial
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substrata, as the ones employed in the present work, has been used to standardize
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sampling methods on sites of various natural bottom types or depths (Desrosiers et
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al., 2014 and literature within). Artificial substrates facilitate the testing of
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environmental conditions such as UV radiation and grazing (Wahl et al., 2004;
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Konar, 2007; Zacher et al., 2007; Fricke et al., 2008; Zacher and Campana, 2008)
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both in situ or after transfer to the laboratory.
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3.3. Grazing experiments
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Previous field observations indicated that the limpet is always found on rocky
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substrates or grazing upon macroalgae, but very rarely on soft bottoms, since
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these prevent firm attachment of the mollusk. Weak adhesion to the bottom turns
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the limpet more susceptible to the effect of waves or to predation by birds. Based
342
on the observations of the natural habitat of the limpet, we chose a hard artificial
343
substrate to be colonized by diatoms. The photographical record of the tiles along
344
the grazing experiments in the containers demonstrated that the limpet N.
345
concinna can consume diatom biofilms for nourishment. Taking into account that
346
the grazing marks on the tiles indicate a 5-30% level of activity of the limpet and
347
being N. concinna the most abundant mollusk in Antarctica, this would represent
348
an important transfer of organic matter from primary producers in the biofilms to
349
the limpet and to other secondary consumers (birds, urchins and others) in higher
350
levels of the food web. Massive growth of benthic diatoms in Marian Cove was
351
reported concomitant with the proliferation of a rich assemblage of benthic
352
megafauna comprising stalked ascidians, sponges, polychaetes, gastropods,
353
bivalves, bryozoans and gorgonians (Ahn et al., 2016), strongly suggesting that
354
the input of diatom biomass can support feeding requirements of at least part of
355
the associated fauna.
SC
M AN U
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The grazing on microphytobenthos by the limpet N. concinna has been
EP
356
RI PT
341
established in the subtidal of the Antarctic Peninsula (Dunton, 2001; Corbisier et
358
al., 2004) and in King George Island (Choy et al., 2011) with the δ13C isotope
359
analysis, that approximate the isotopic composition of their respective diets but to
360
the best of our knowledge this is the first report of the characterization of
361
chrysolaminarin in diatom biofilms grazed by the limpet as a possible carbon
362
source.
363
Acknowledgements
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The authors want to thanks especially to Dr. Nora I. Maidana (DBBE, FCEN,
365
UBA) for help in diatom identification and Dr. María Cristina Matulewicz (Depto.
366
Química Orgánica, CIHIDECAR, CONICET-UBA) for suggestions in structural
367
analysis. This work was funded by grants from UBA (GC 20020130100216BA)
368
and FONCyT- DNA (PICTO 0091). YD and HS are a Research Fellows of the
369
National Research Council of Argentina (CONICET).
371
SC
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571
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Tables Table 1: Monosaccharide composition (mol%) of the different mucilage fractions
575
extracted with water at room temperature (RTW), boiling water (HW1 and HW2)
576
and after RTW desalting in BioGel P2 (RTWI and RTWII)
RI PT
574
Constituent monosaccharides (mol%) Protein (%)
Carbohydrates (%)
Rha
Fuc
Ara
Xyl
Man
Glc
Gal
3.9
9.3
0.7
1.0
1.0
1.2
1.4
91.1
3.5
HW1
14.8
19.7
-
-
-
-
1.2
92.9
6.0
HW2
12.9
11.6
6.6
12.0
0.9
15.4
8.3
35.7 21.3
RTWI
n.d.
8.9
-
-
-
-
16.4 68.8 14.8
RTWII
n.d.
4.9
4.3
20.9
-
-
13.7 36.6 24.5
M AN U
SC
RTW
577
Abbreviations: Rha= rhamnose; Fuc= fucose; Ara= arabinose; Xyl= xylose; Man=
579
mannose; Gal= galactose; Glc= glucose.; n.d.= not determined
EP AC C
580
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578
27
ACCEPTED MANUSCRIPT
581
Table 2: Linkage analysis of constituent monosaccharides (mol %) produced by
582
methylation and hydrolysis of product RTWI after one (RTWI-m1) and two
583
(RTWI-m2) methylation steps.
Monosaccharidea
Deduced glycosidic linkage
RI PT
584
RTWI-m1 RTWI-m2
Terminal
20.1
25.3
2,4,6-Me3Glc
3-linked
44.1
55.1
4,6-Me2Glc
3-linked
16.6
10.2
M AN U
2-substituted
SC
2,3,4,6-Me4Glc
2,4-Me2Glc
3-linked
10.2
6.1
6.3
2.3
2.8
1.1
6-substituted 2- MeGlc
TE D
Glc 585
Abbreviations: 2-MeGlc=1,3,4,5,6-tetra-O-acetyl-2-O-methylglucitol; 2,4-
587
Me2Glc=1,3,5,6-tetra-O-acetyl- 2,4-di-O-methylglucitol; 4,6-Me2Glc= 1,2,3,5-
588
tetra-O-acetyl-4,6-di-O-methylglucitol; 2,4,6-Me3Glc = 1,3,5-tri-O-acetyl-2,4,6-
589
tri-O-methylglucitol; 2,3,4,6-Me4Glc=1,5-di-O-acetyl-2,3,4,6-tetra-O-
590
methylglucitol.
AC C
591
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586
592
28
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593
Table 3.Marine benthic diatom species recorded at Potter Cove, King George
594
Island, Antarctica, during the austral summer of 2013. A total of 400 valves were
595
counted.
Achnanthes bongrainii (Peragallo) Mann
r
Actinocyclus actinochilus (Ehrenberg) Simonsen
vr vr
Coccocneis costatavar. antárctica Manguin
M AN U
Coccocneis fasciolata (Ehrenberg) Brown
SC
Amphora sp.
RI PT
Relative abundance
r r r
Entopyla ocellata (Arnott) Grunow
vr
Licmophora antárctica G.W.F.Carlson
r
Minidiscus chilensis Rivera
vr
TE D
Coccocneis pinnata Gregory ex Greville
f
Navicula cf. perminuta Grunow
f
Navicula sp.
f
EP
Navicula directa (Smith) Ralfs
f
AC C
Nitzschia pellucida Grunow Odontella litigiosa (Van Heurck) Hoban
r
Parlibellus delognei (Van Heurck) Cox
r
Petroneis plagiostoma (Grunow)
r
Petroneis sp.
r
Pinnularia quadratarea (A. Schmidt)
r
Pseudogomphonema kamtschaticum (Grunow)
c
29
ACCEPTED MANUSCRIPT
Synedra kerguelensis Heiden
f
Thalassiosira gracilis (Karsten) Hustedt
vr
596
Relative abundance represents the occurrence of a species in all samples for a
598
given area; vr: very rare, found only once on a slide, r: rare <5%, f: frequent, 5–
599
25%, c: common >25%.
RI PT
597
AC C
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M AN U
SC
600
30
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Figure captions
602
Fig. 1: a Potter Cove map, b Containers, c a detail of flasks distribution inside the
603
containers, each with a colonized tile and a Nacella concinna specimen (black
604
arrowhead).
605
Fig. 2: Bio-Gel P-2 filtration elution profile of fraction RTW. Void volume (V0) is
606
indicated (small black arrow). Fractions indicated between the full black and
607
hatched arrows showed salt contamination.
608
Fig. 3: Marine benthic diatoms from 25 de Mayo (Potter Cove) Antarctica on
609
colonized ceramic tiles, viewed in LM. a Achnanthes bongrainii, b Entopyla
610
ocellata, c Parlibellus delognei, d Licmophora antarctica, e Odontella litigiosa, f
611
Amphora sp. g Actinocyclus actinochilus, h Cocconeis pinnata,I Cocconeis
612
costatavar. antarctica, j Cocconeis fasciolata. Scale bar= 10 µm.
613
Fig. 4: Marine benthic diatoms from 25 de Mayo (Potter Cove) Antarctica on
614
colonized ceramic tiles, viewed in LM. a Nitzschia pellucida, b Navicula directa,
615
c Petroneis sp., d Thalassiosira gracilis, e Navicula cf.perminuta, f Navicula sp, g
616
Petroneis plagiostoma, h Pinnularia quadratarea, I Synedra kerguelensis, j
617
Pseudogomphonema kamtschaticum, k Pleurosigma sp. Scale bar= d-f: 5 µm; for
618
the rest: 10 µm.
619
Fig. 5: Marine benthic diatoms from 25 de Mayo (Potter Cove) Antactica on
620
colonized ceramic tiles, viewed in SEM. a Odontella litigiosa, b Minidiscus
621
chilensis, c Navicula sp , d Navicula cf. perminuta, e-f Pseudogomphonema
622
kamtschaticum in external and internal valve view, g Synedra kerguelensis, h
623
Nitzschia pellucida. Scale bar= b: 1 µm; for the rest: 5 µm.
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31
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Fig. 6: Grazing experiment, a-bColonized tiles at time 0; c-dTiles after 24 h.
625
Scale= 1 cm.
AC C
EP
TE D
M AN U
SC
RI PT
626
32
AC C
EP
TE D M AN US C
RI P
ED
M AN
AC C
EP TE D
M AN US C
RI PT
CC EP TE D
M AN US C
RI P
AC C EP TE D
M AN US C
RI PT
EP TE D
M AN US C