A novel method for imaging internal growth patterns in marine mollusks: A fluorescence case study on the aragonitic shell of the marine bivalve Arctica islandica (Linnaeus)

In this article, we explore the use of fluorescence spectroscopy to image growth patterns in the marine bivalve Arctica islandica (L.). The method presented here does not require any chemical treatment of the polished shell section and yields results comparable to acetate peels of acid-etched shell sections and Mutveitreated shell sections. Further, our results indicate that the annual growth lines in A. islandica fluoresce in the blue light spectrum (450–490 nm), thus an ultraviolet source (mercury lamp) is not required. The reflected light entering the digital camera was filtered (510–540 nm) and later enhanced to emphasize the annual growth patterns. The fluorescence of annual growth lines was consistent among the four animals used in this study. Additionally, we measured growth increments in the umbo section of one A. islandica shell using both the traditional acetate method and fluorescence imaging. The two sets of measurements were highly correlated (r = 0.97; P < 0.0001). We suggest that the fluorescence imaging method presented here is a viable option for increment identification and measurement in this key marine archive. It is likely that the methods demonstrated here for A. islandica can easily be used/modified for other bivalve (mollusk) taxa. Fluorescence microscopy permits rapid analysis of shell growth patterns with minimal pretreatment and offers an objective method of determination of annual growth increments and lines. *Corresponding author: E-mail: [email protected]; Present Address: Department of Geological and Atmospheric Sciences, Iowa State University, Ames, IA 50011-3212, USA Acknowledgments This research is funded by EU Millennium (European Climate of the Last Millennium; Project no. 017008) and resulted from an Arctica islandica workshop held at the Gregynog Conference Centre, Wales in 2007. JDS acknowledges a Royal Society-Leverhulme Trust Senior Research Fellowship. We are grateful for helpful suggestions from three anonymous reviewers. Limnol. Oceanogr.: Methods 7, 2009, 673–681 © 2009, by the American Society of Limnology and Oceanography, Inc. LIMNOLOGY and OCEANOGRAPHY: METHODS sclerosponges, otoliths, statoliths) and the temporal context in which they formed (e.g., see 1st International Sclerochronology Conference, 2007 Jul 17–21, St. Petersburg, Florida, U.S.A., http://conference.ifas.ufl.edu/sclerochronology/; also see Oschmann 2009). Sclerochronology can be considered the aquatic counterpart of dendrochronology. Often, sclerochronological techniques are used to develop a master chronostratigraphy within biogenic carbonates, which is then used as a template for growth and geochemical analysis (Weidman et al. 1994; Hendy et al. 2003; Delong et al. 2007; Halfar et al. 2008). It has been demonstrated that master shell chronologies (Witbaard et al. 1997; Marchitto et al. 2000; Scourse et al. 2006; Helama et al. 2006; 2007; Butler et al. 2009a) can be developed on a level consistent with dendrochronological studies. In particular, shell-based records from the very long-lived bivalve Arctica islandica can be absolutely dated like tree rings (Thompson et al. 1980; Jones 1980), enabling the generation of ultra-highresolution and multicentennial paleoenvironmental records based on an absolute timescale (Butler et al. 2009a). These records have the potential to provide significant histories of marine environmental change, and to serve as a basis to document natural and anthropogenic change in key regions around the globe (Wanamaker et al. 2008a, 2008b). The ocean quahog (A. islandica L.), whose remarkable longevity has been found to exceed 400 years (Wanamaker et al. 2008a), is found along the continental shelves in the mid-to-high latitudes in the North Atlantic in water depths of <20 m to more than 200 m (Cargnelli et al. 1999). The usefulness of this proxy in ecosystem and ocean/climate studies is well established (e.g., Weidman and Jones 1993; Weidman et al. 1994; Marchitto et al. 2000; Witbaard et al. 2003; Schöne et al. 2003; 2005a; Scourse et al. 2006; Helama et al. 2007; Wanamaker et al. 2008a; 2008b; 2008c; Wanamaker et al. 2009; Butler et al. 2009a), and it is likely that A. islandica will be further established as a key marine archive from the North Atlantic (e.g., Schöne et al. 2005a; Wanamaker et al. 2008c) that will facilitate improved documentation and interpretation of recent and past environmental and climatic change. With the exception of shell thin sections, which are often difficult and time consuming to prepare, imaging of growth patterns in bivalves generally require that a polished section of the shell surface be treated or etched with an acid to enhance the microshell structure (see Ropes 1984; Schöne et al. 2005b). In many cases, an acetate peel replica is made of the etched shell surface (Ropes 1984), placed on a microscope slide and then photographed. Additionally, if geochemical sampling (isotopes, minor and trace elements) is necessary, a second shell section would be required. More recently, treatment of biogenic carbonates with Mutvei’s solution (a mixture of acetic acid, glutaraldehyde, and alcian blue; see Schöne et al. 2005b) has allowed the sectioned surface to be photographed without making an acetate peel. For many workers, the Mutvei method greatly improved their ability to visualize and interpret the growth record, but the acetate peel method and the Mutvei method both have some minor disadvantages. The acetate peel can introduce noise (blurring) unrelated to the growth record, or it may be unable to replicate adequately the physical structure of the increments. Treatment with Mutvei’s solution is slightly destructive to the shell over time (especially if re-etching is required), and the shell surface scratches easily after it is etched. Although the Mutvei method has been very useful in highlighting growth lines/structures, it requires the use of glutaraldehyde (a respiratory toxin) to fix the organics in the shell structure. An ideal method of imaging biogenic growth structures would dispense with the need to treat the polished shell section with chemicals and create replicates of the surface using acetate peels. Here, we image the annual growth increments in four specimens of the marine bivalve Arctica islandica (L.) using fluorescence microscopy. Fluorescence microscopy is a widely used technique in both geological and biological sciences (Isdale 1984; Scoffin et al. 1989; Baker et al. 1998, 1999; Proctor et al. 2000; Charman et al. 2001; Hendy et al. 2003). Typical components of a fluorescence microscope are the light source (typically xenon or mercury lamps), an excitation filter, a beam splitter, and an emission filter. The filters and beam splitter are chosen to match the spectral excitation and emission characteristics of the fluorescent material being analyzed. This fluorescent material may be intrinsic to the sample under analysis (e.g., natural organic matter preserved in geological materials such as stalagmites (Baker et al. 1993) and particulate organic matter in sedimentary deposits (Hart 1986),or it may be an artificially added fluorophore (e.g., fluorescent dyes or probes used to tag DNA; for example in fluorescent in-situ hybridization; Wilkinson 1999). In this article, we investigate the use of fluorescence microscopy as a suitable alternative to chemical-etching techniques as a method of imaging the internal growth patterns of polished shell sections of A. islandica. Methods and procedures The four A. islandica shells used in this study (WG060329, WG061254, WG061271, WG061290) were live-collected along the north Icelandic shelf in 2006 in 80 m water depth (see Wanamaker et al. 2008a for details), and prepared using methods outlined by Scourse et al. (2006), which are summarized below. The left-shell valves were embedded in resin (MetPrep; Kleer Set type FF) and the shells sectioned radially from the umbo to the shell margin (Fig. 1). The embedded left valve was sectioned through the center of the hinge tooth using a diamond saw. The cut surfaces were ground on progressively finer grades of silicon carbide paper (MetPrep; P120, P400, P1200) and polished with dilute diamond paste (Presi; 3 μm) on rotary magnetic pads (Presi; 200 mm, 3106 and 0307). Polished sections of the four A. islandica shells were analyzed episcopically using a Zeiss Axiotech fluorescence microscope fitted with a Q imaging Micropublisher 3.3RTV camera (CCD color; 3.3 million pixels; 2048 × 1536) and three filter sets; Zeiss Wanamaker et al. Imaging mollusk shells via fluorescence