2012 NORTHEASTERN NATURALIST 19(2):201–216 Spatial and Temporal Variation in Otolith Chemistry for Tautog (Tautoga onitis) along the US Northeast Coast Ivan Mateo1,*, Edward G. Durbin2, David A. Bengtson1, Richard Kingsley2, Peter K. Swart3, and Daisy Durant4 Abstract - Elemental concentrations and stable (δ 18O, δ 13C) isotopic ratios in otoliths of young-of-the year (YOY) Tautoga onitis (Tautog) captured in nurseries in Rhode Island, Connecticut, New Jersey, and Virginia were determined using otolith micro-chemistry. Multi-chemical signatures differed significantly among the distinct nurseries among regions (MANOVA: P < 0.001) and between years (MANOVA: P < 0.001). Classification accuracy for Tautog nurseries among regions ranged from 92% to 96% for each of the two years. Since accurate classification of juvenile Tautog to their nursery sites was achieved, otolith chemistry can potentially be used as a natural habitat tag in assigning adult Tautog to their respective estuarine nurseries, but it is important to consider that the chemical signals may change annually. Introduction Connectivity is an important property in marine conservation, and determines colonization patterns of new habitats, the resiliency of populations to harvest, and the design of marine protected areas (MPAs). However, quantifying connectivity in marine organisms is extremely difficult because the natal or nursery origins of most adult fish remain unknown. This lack of knowledge is primarily due to the difficulty of conducting mark-recapture studies in species that are characterized by the production of large numbers of small planktonic offspring and suffer high juvenile mortality (Eldson et al. 2008, Thorrold and Hare 2002). Recently, innovative techniques using natural isotopic and elemental markers in calcareous structures such as otoliths/statoliths have been developed for species that are not able to be tagged or recaptured using conventional approaches (Eldson et al. 2008, Thorrold and Hare 2002). Otoliths are acellular mineral structures, associated with orientation and motion, located in the inner ear of fish. Otoliths are made primarily of aragonitic calcium carbonate (>90%) embedded in an organic matrix (less than 8%), with the remainder (ca. 2%) consisting of trace and minor elements (Campana 1999). Otoliths are not susceptible to resorption or metabolic activity once the material is laid down (Campana and Thorrold 2001). Elements are incorporated from the endolymph onto the otoliths surface (Campana 1999, Campana and Neilson 1985, 1University of Rhode Island, Department of Fisheries, Animal and Veterinary Science, Kingston, RI 02881. 2University of Rhode Island, Graduate School of Oceanography, Narragansett, RI 02882. 3Division of Marine Geology and Physics, Rosenstiel School of Marine and Atmospheric Sciences, University of Miami, Miami, fl33149. 4Narragansett Bay National Estuarine Research Reserve, PO Box 151, Prudence Island, RI 02872. *Corresponding author - imateo32@cox.net. 202 Northeastern Naturalist Vol. 19, No. 2 Markwitz et al. 2000) throughout the fish’s lifetime. Elements that are considered as the best environmental recorders are those that are not under strict physiological control and whose incorporation into the otolith consistently reflects the water chemistry the fish inhabit (Campana 1999). Strontium (Sr) is the most widely used element in otolith chemistry studies. Sr and calcium (Ca) are alkaline earth metals with similar chemical composition, which allows Sr to be readily substituted for Ca in the aragonitic crystal lattice of otoliths due to its similar valence, ionic radius, and crystal structure (Farrell and Campana 1996, Radtke and Shafer 1992). Though the Sr:Ca ratio has been the most widely used elemental ratio in otolith microchemistry studies, other alkaline earth and alkaline metals, such as magnesium (Mg) and barium (Ba), have been successfully used in otolith microchemistry studies (Arslan and Paulson 2003). There are many factors influencing the otolith chemistry composition, including geology, water chemistry, temperature, physiology, and diet (Arslan and Paulson 2003). However, several studies have shown that the incorporation of many alkaline earth metals into the otoliths is directly related to the ambient water, thus demonstrating their potential utility (Bath et al. 2000; Elsdon and Gillanders 2003; Wells et al. 2000, 2003). Analysis of stable isotope ratios has also been shown to be useful in otolith microchemistry studies. Stable isotopic variation in biogenic carbonates (e.g., corals, foraminiferans, and bivalves) has been used extensively in paleoclimate studies. They are also useful proxies of temperature, salinity, and trophic level (Campana 1999). The oxygen (O) and carbon (C) isotopes are the most extensively studied (Campana 1999). Oxygen (O18/O16, expressed as δ18O) isotopes are incorporated into otoliths in ratios in close relationship to those found in the water from their environment (Kalish 1991a, Kennedy et al. 1997, Thorrold et al. 1997a). Differences in δ18O in the water are related to sea surface temperatures, evaporation rates, and freshwater inputs (Edmonds et al. 1999). Otolith carbonate (13C/12C, expressed as δ13C) is incorporated in isotopic disequilibrium with ambient seawater. It is thought that variations in δ13C are related to metabolic changes, dietary changes, or the δ13C of the dissolved inorganic carbon (Gannes et al. 1997, Thorrold and Hare 2002). Applications of chemical analysis in otoliths The application of otolith microchemistry to fish ecology studies is based on the idea that the chemistry of the water a fish inhabits provides a natural tag in the otolith, that can then be used to discriminate among groups of fish from different spawning and nursery grounds (Campana 1999). This application has been widely used to identify unique stocks for marine fisheries management and conservation goals (e.g., Campana and Gagné 1995; Campana et al. 1994, 2000; Edmonds et al. 1991; Gillanders et al. 2001; Patterson et al. 1999). Variation of otolith chemical composition to changes in the temperature, salinity, and water chemistry the fish experience throughout their lives (e.g., Gauldie et al. 1986, Radtke and Kinzie 1996) is another major application of otolith microchemistry. Stable oxygen isotope ratios (δ18O; Kalish 1991b, Thorrold et al. 1997a), and Sr:Ca ratios (Radtke et al. 1990; Townsend et al. 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 203 1989, 1992, 1995) have both been used as proxies for temperature. Salinity histories have also been determined using Sr:Ca ratios (e.g., Limburg 1995, Radtke et al. 1998, Tzeng et al. 1997) and stable oxygen isotopes (e.g., Meyer- Rochow et al. 1992, Northcote et al. 1992) as salinity proxies. Most salinity histories studies using Sr:Ca ratios have documented evidence of anadromous and catadromous migration (Campana 1999). Furthermore, changes in otolith elemental composition resulting from movement from one environment to another (e.g., freshwater to marine) have been used to tell apart anadromous from non-anadromous (resident) fish (e.g., Kalish 1990, Rieman et al. 1994, Zimmerman and Nielsen 2003) as well as hatchery from wild fish (e.g., Campana 1999). In this application, measurements are made with probe-like instruments along the otolith growth axis or the entire otolith in order to determine migration history (e.g., Arai and Miyazaki 2001, Limburg 1995, Limburg et al. 2001, Secor 1992, Thorrold et al. 1997b). Chemical habitat tags in the otoliths of juvenile fish have been used (with a high degree of accuracy) to differentiate individuals from different estuarine/ riverine systems (Gillanders 2002b; Gillanders and Kingsford 2000; Thorrold et al. 1998a, 1998b) and alternative types of nearshore habitats, including estuary versus rocky reef (Gillanders and Kingsford 1996) and estuary versus exposed coastal habitats (Forrester and Swearer 2002, Yamashita et al. 2000). In addition, through chemical analysis of the juvenile portion of adult otoliths, natural habitat tags have been used to determine the proportion of the adult population that resided in different juvenile habitats (Gillanders 2002a, Thorrold et al. 2001, Yamashita et al. 2000). Application of otolith chemistry approach in Tautog Tautoga onitis L. (Tautog) is an economically and ecologically important species found in the waters of eastern North America from the Gulf of Maine to North Carolina. While adults may live off the coast in waters as deep as 25 m (Arendt 1999, Dorf and Powell 1997), they move inshore for reproduction purposes (Arendt 1999). Juvenile Tautog are known to depend upon the coastal zone for nursery habitats where they are safe from high levels of predation and can find necessary food resources (Arendt 1999, Dorf and Powell 1997). However, the relative importance of open coastline versus enclosed bays and lagoons as nursery habitat for Tautog is still poorly understood (Sogard et al. 1992). In Narragansett Bay, RI, Zostera marina L. (Common Eelgrass) has declined since the early 1950s because of water pollution, coastal development, harbor dredging, and other factors (Meng et al. 2001). In 1996, scientists determined that of the 1000 acres of eelgrass that were in Narragansett Bay at the turn of the century, fewer than 100 acres remained. Furthermore, Eelgrass has decreased 41% in coastal ponds due to increased nitrogen loads (Cottrell et al. 1999). In light of the fact that the northeastern coast of the United States has experienced a major loss of its estuarine habitats due to human alteration of the coastal zone (Cottrell et al 1999, Meng et al 2001), data are needed to quantify the importance of specific coastal habitat types in sustaining Tautog populations. 204 Northeastern Naturalist Vol. 19, No. 2 The specific purpose of this study was to determine if juvenile nurseries can be distinguished for Tautog populations among regions within the US Northeast coast using otolith chemistry. This is a crucial step in order to quantify the relative contribution and connectivity of estuarine habitats for adult Tautog populations. Materials and Methods Sampling of juvenile fish In Rhode Island, young-of-the-year (YOY) Tautog of 45–64 mm fork length (FL) were sampled in Narragansett Bay: Rose Island (RS; 41°29'48.19"N, 71°20'35.69"W) (Fig. 1). The samples were obtained by seine net in cooperation with Rhode Island Department of Environmental Management, Division of Marine Fisheries (RIDEM) during August and September of 2005 and 2006, respectively. In 2006, data from the closest point to RS (Fort Wetherill, Jamestown), which is about 1.5 km east , showed average surface temperatures of 17.4 °C and salinities of 30.8‰. We also collected Tautog from Connecticut and New Jersey over the same two years in which those from Rhode Island were collected. In Connecticut, YOY Tautog were sampled from the New Haven Harbor/Quinnipiac River estuary (41°15'26.64"N, 72°53'39.13"W) by staff from the National Marine Fisheries Service at Milford, CT using seine nets. In New Jersey, fish were sampled around Great Bay-Little Egg Harbor (39°29'30.21"N, 74°18'58.40"W) by staff from the Rutgers University, Marine Field Station using minnow traps. In Virginia, YOY Tautog were collected from one area of the Chesapeake Bay estuarine system (South Bay Eastern Shore) (37°26'1.26"N, 75°59'33.86"W) by staff from the Virginia Institute of Marine Science (VIMS) as part of their normal seine-net sampling program; however, samples were collected in 2005 but not in 2006 because the Tautog experienced poor recruitment (Jacques Vont Monfrans, VIMS, Gloucester Point, VA, pers. comm.). All these stations are located in lower areas of their respective estuaries with low fresh water input and with salinities around 30‰. A total of twenty juveniles per site per year were captured for analysis. Sampled fish were kept frozen until otolith removal. Water sampling was not obtained nor conducted due to lack of funding. Laboratory processing of samples Prior to otolith removal, each fish was weighed (wet weight to the nearest 0.1 g) and measured (fland standard length [SL] to the nearest 0.1 mm). Both sagittal otoliths were removed from each fish, cleaned of adhering tissue, rinsed 3 times with Milli-Q water, and allowed to dry in a Class 100 laminar-flow hood. The left sagittal otolith was used for trace-metal analysis, while the right otolith was used for stable-isotope analysis. Of the original number of otoliths, about 12% were lost during the decontamination process. Therefore, they were removed from subsequent analyses. A total of 113 otoliths were prepared for trace metal analysis. Each otolith was weighed on a Thermo Cahn microbalance (± 0.01 mg). Samples were then placed in acid-washed 2.5-ml polypropylene containers. The otolith weights 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 205 ranged from 0.08 to 0.34 mg and averaged 0.18 mg. Otoliths used for trace-metal analysis were transferred to clean 5-ml polypropylene tubes and 0.2 ml of tripledistilled 17% HNO3 was added to insure complete dissolution. An internal thulium single-element standard spike was added to the solution to correct for variable matrix effects during the inductively coupled plasma mass spectrometry (ICPMS) analyses, and subsequently it was diluted to 1.8 Figure 1. Map of Tautog sampling stations in high-salinity areas in the US northeast coast for the years 2005 and 2006. 206 Northeastern Naturalist Vol. 19, No. 2 ml with triple-distilled water. This dilution resulted in a Ca concentration of approximately 40 ppm in the analyzed otolith solution. Otolith chemistry analysis Elemental concentrations of YOY otoliths were determined through solutionbased ICP-MS at the University of Rhode Island (URI) Graduate School of Oceanography (GSO). All measurements were carried out on a Finnigan Element High-Resolution ICP-MS (Thermo Fisher Scientific). A procedural blank was prepared in the same manner as the samples, but without an otolith. The procedural blank was then compared to the system blank to determine if contamination occurred. System blanks using the same acid concentration as sample dissolution were run every four samples. A driftcorrection standard was prepared by gravimetrically spiking a CaCO3 standard solution with appropriate concentrations of Na, K, Rb, Mg, Ca, Mn, Ni, Cu, Zn, Sr, Ba, Co, and Pb, to match the typical elemental composition of the otoliths. This drift-correction standard was analyzed every four samples as well to track for instrument drift. Thirteen elements (Na, K, Rb, Mg, Ca, Mn, Ni, Cu, Zn, Sr, Ba, Co, and Pb) were selected based on previous studies using trace elements as natural tags (Gillanders 2002b; Gillanders and Kingsford 2000; Thorrold et al. 1998a, 1998b). Results were expressed as absolute concentrations of molar ratios with respect to calcium—element:Ca ratios, expressed as units of mmol mol-1 or μmol mol-1—as the elements are considered to substitute for calcium within the crystalline lattice of the otolith. The elements used for subsequent analysis (Rb, Mg, Ca, Sr, and Ba) were always found in concentrations above their limit of detection (LOD). The LODs were as follows (values in ppm): Rb (0.007), Mg (0.02), Sr (0.077), and Ba (0.014). These values were much less than the 3 ppm to 2000 ppm range for these elements found in the sample otoliths. The LODs were calculated to be 3 times the standard deviation of the isotope’s sensitivity (in counts per second or cps) found in acid blanks divided by the corresponding sensitivity in cps/ppm of the CRM22 carbonate standard. The precision of the analyses can be estimated from the average relative standard deviations (RSD) measured in multiple runs of the CRM22 standard and were as follows: Rb (3%), Mg (10%), Ca (1%), Sr (1%), and Ba (5%). Stable carbon and oxygen isotopes of these otolith samples were determined at the Rosenstiel School of Marine and Atmospheric Sciences, University of Miami, using an automated carbonate device (Kiel III) attached to a Thermo- Finnigan Delta Plus stable isotope mass spectrometer. Data are expressed using the conventional δ notation relative to V-PDB (Vienna Peedee Belemnite). Data have been corrected for the usual isobaric interferences. The external precision (calculated from replicate analyses of an internal laboratory calcite standard) was 0.04% for δ13C and 0.08% for δ18O. Statistical analysis A two-way ANOVA was used to test for differences in fish body length among stations and years. We also examined relationships between otolith weight and 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 207 otolith elemental composition and isotopic signatures with analysis of covariance (ANCOVA). If a significant relationship was observed, we removed the effect of size (otolith weight used as a proxy for fish size) to insure that differences in fish size among samples did not confound any site-specific differences in otolith chemistry. Element concentrations were weight-detrended by subtraction of the product of the common within-group linear slope times the otolith weight from the observed concentration (Campana et al. 2000). We used analysis of variance (ANOVA) to test for differences in mean individual elemental concentration ratios of (Sr, Ba, Mg, Rb) and otolith isotopic signatures (δ13C, δ 18O) among sites (RI, CT, NJ, and VA) within each year (2005, 2006). Multivariate analyses of variance (MANOVA) were used to test for differences in the multi-chemical signatures combined of mean elemental concentration ratios (Sr, Ba, Mg, Rb) and otolith isotopic signatures (δ13C, δ18O) among sites (RI, CT, NJ) and between years (2005 and 2006). Pillai’s trace was chosen as the multivariate test statistic because it is more robust to small sample sizes, unequal cell sizes, and situations in which covariances are not homogeneous. Tukey’s HSD test was used to detect a posteriori differences among means (α = 0.05). Before statistical testing, residuals were examined for normality and homogeneity among stations. To meet model assumptions, all analyses of trace element concentrations were performed using natural log transformed data. Linear discriminant function analyses (DFAs) were used on Tautog juvenile data to visualize spatial differences within sites and to examine classification success for juveniles from different sites or regions. The DFA develops an algorithm to classify fish according to their nursery habitats based on the elemental composition of their otoliths. This multivariate approach was used to determine whether the relationship among several independent variables could be used to predict nursery origin of an individual. Classification success is translated as the percentage of fish that were correctly assigned to their capture location based upon the chemical signature of each fish. Cross validations were performed using jackknifing procedures in SYSTAT to test the model accuracy and robustness. Results Size distribution Mean flof juvenile Tautog at all stations ranged from 45–64 mm (Table 1). There were significant differences in mean length among stations within regions within each year (2005: F3,68 = 8.65, P < 0.001; 2006: F2,54=10.45, P < 0.001 ). In both 2005 and 2006, the mean length of NJ fish was significantly greater than those from CT, VA, and RI (Tukey test: P < 0.05). Individual element concentrations Elemental concentrations of Ba and δ 18O varied significantly in 2005 and 2006 among regions (ANOVA: P < 0.001; Table 2, Fig. 2). For those two years, δ 18O was highest in otoliths of fish from the RI station (Rose Island), whereas it was most depleted in those from the CT station. 208 Northeastern Naturalist Vol. 19, No. 2 Table 1. Mean fork length (FL) in mm and number of fish (n) captured at each station by year. Numbers in parenthesis are SE. Abbreviations provided after each station will be used in subsequent tables. State Station n fl(mm) 2005 Rhode Island Rose Island (RI) 20 52.1 (1.2) Connecticut Morris Cove New Haven Harbor (CT) 16 50.1 (2.2) New Jersey Tuckerton Rutgers Field Station (NJ) 17 63.5 (2.1) Virginia Virginia Eastern Shore Peninsula (VA) 19 56.1 (2.3) 2006 Rhode Island Rose Island (RI) 17 45.5 (3.5) Connecticut Morris Cove New Haven Harbor (CT) 20 57.2 (1.8) New Jersey Tuckerton Rutgers Field Station (NJ) 20 63.4 (2.9) Virginia Virginia Eastern Shore Peninsula (VA) * * *Tautogs were not captured in this station in 2006. Table 2. Results of ANOVA of trace element concentration and stable isotope composition measured in otoliths of YOY Tautog collected at high salinity stations on the US northeast coast in 2005 and 2006. Tukey honestly significant different (HSD) test was based on Ln (x + 1) transformed data. Stations are labeled as 1 = RI, 2 = CT, 3 = NJ, 4 = VA (see Table 1 for abbreviations of station names). The HSD column shows the pairs of stations that were significantly different. 2005 2006 Source d.f. MS F P HSD Source d.f MS F P HSD [Sr/Ca] [Sr/Ca] Station 3 0.44 35.46 0.001 1,2 Station 2 0.01 1.03 0.363 Error 62 0.26 1.27 1,4 Error 53 0.01 2,3 2,4 [Ba/Ca] [Ba/Ca] Station 3 2.89 13.67 0.001 2,4 Station 2 1.05 5.11 0.009 1,2 Error 63 0.21 3,4 Error 52 0.21 1,3 [Mg/Ca] [Mg/Ca] Station 3 0.28 0.50 0.682 Station 2 0.70 0.64 0.533 Error 62 0.56 Error 52 1.09 [Mn/Ca] [Mn/Ca] Station 3 2.33 5.20 0.003 1,2 Station 2 4.35 7.64 0.001 1,3 Error 63 0.45 1,3 Error 52 0.57 2,3 1,4 [Rb/Ca] [Rb/Ca] Station 3 0.04 2.79 0.047 Station 2 0.08 9.09 0.001 1,2 Error 63 0.01 Error 49 0.01 1,3 [13C] [13C] Station 3 2.04 7.71 0.001 1,2 Station 2 0.64 2.60 0.084 Error 64 0.15 1,4 Error 53 0.24 [18O] [18O] Station 3 3.18 28.16 0.001 1,2 Station 2 21.45 174.62 0.001 1,2 Error 64 0.32 1,3 Error 50 0.12 1,3 2,3 2,3 3,4 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 209 Multi-elemental fingerprint results There were significant differences for Tautog chemical signatures between regions (MANOVA: P < 0.05; Table 3) and years (MANOVA: P < 0.001; Table 3). Significant interaction was found among regions and year (MANOVA: P < 0.001; Table 3). Classification success among regions within each year varied from 92–96% (Table 4). Figure 2. Mean elemental concentrations molar ratios (with standard errors) with respect to Ca and stable isotopes measured in otoliths of YOY Tautog collected from highersalinity stations along the US Northeast coast in 2005 and 2006. All trace element data (Element/Ca x 103) are Ln (x + 1) transformed. The y-axes for the two stable isotopes are inverted. State station codes are RI = Rhode Island (Rose Island), CT = Connecticut, NJ = New Jersey, and VA = Virginia. 210 Northeastern Naturalist Vol. 19, No. 2 Discussion The elemental concentrations of juvenile Tautog otoliths varied considerably among estuaries and between years. We found very strong differences in the concentrations of Sr, Ba, and Rb as well as in the stable isotopic signatures of δ 13C and δ 18O among the regions. High classification success rates (generally >90%) of the discriminant functions derived from trace element and stable isotope signatures together demonstrated that these otolith elemental fingerprints may be potentially used as a natural tag of the estuarine nursery area of juvenile Tautog. However, in order to consider otolith elemental signatures as an effective natural marker to study connectivity in estuarine fish nurseries, certain requirements must be met: (1) ensure that all the potential nursery sites with unique distinct signatures have been sampled and are representative for the species in question, and (2) show there is direct relationship of the otolith elemental signatures with the environment (Campana et al. 2000, Elsdon et al. 2008). Differences among geographic sites in this study were presumably related to surface-water mass characteristics and associated geo-chemical differences; however, the source of the elemental fingerprint differences remains unknown. Table 3. Results of MANOVA for trace element concentration and stable isotope composition in otoliths of YOY Tautog collected at high-salinity stations along the US northeast coast (RI, CT, NJ). All statistical analyses were done on Ln (x + 1) transformed data and on Pillai’s statistic. VA was removed from analysis as YOY Tautog were not collected in 2006. Source Value F Numerator Denominator P Year 0.163 13.775 7 86 0.027 Station 1.051 2.399 14 174 0.001 Year*Station 0.778 7.917 14 174 0.001 Table 4. Jackknifed classification success using linear discriminant function analysis for Tautog otoliths collected from high-salinity stations along the US northeast coast during 2005 and 2006. Based on solution-based ICPMS using combined trace metals (Sr, Ba, Mg, and Rb) and stable isotopes (δ18O, δ13C) (see Table 1 for abbreviations of station names). CT NJ RI VA Cross validation accuracy 2005 CT 15 0 0 0 100 NJ 0 13 1 2 81 RI 0 0 19 0 100 VA 0 2 0 14 88 Total 92 2006 CT 13 0 0 100 NJ 0 15 2 88 RI 0 0 17 100 VA * * * * * Total 96 *Tautog were not captured in this station in 2006. 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 211 Thus, the observed elemental compositions represent only those sites and years that were sampled. Because the elemental composition of the otolith is metabolically stable, the YOY juvenile portion of the otolith serves as a natural tag of the nursery area independent of cause (Campana et al. 2000). The study did not sample all the potential Tautog nursery and habitat (e.g., salinity gradient) within the respective states since the main objective of the study was to determine if differences exist among the nurseries areas sampled. Given that we found signifi cant differences, future work should be focusing on resolving the spatial scale across which Tautog nurseries can be distinguished. It is noteworthy to say that our results would be more robust if we had performed chemical analysis of the ambient water to support our data, as there could be potential interactions of salinity and temperature influencing elemental concentrations in otoliths. Despite not analyzing the water chemistry surrounding the Tautog nurseries in this study, there is evidence that water chemistry can be used to predict otolith chemistry, providing there is a strong correlation between the two variables in natural environments. Experimental work has demonstrated strong effects of ambient Sr and Ba concentrations on otolith composition (e.g., Bath et al. 2000, Elsdon and Gillanders 2003, Kraus and Secor 2004). Recently, Walther and Thorrold (2009) also found that Sr:Ca and Ba:Ca in juvenile Alosa sapidissima (Wilson) (American Shad) otoliths were correlated with ambient levels in river waters. Our approach using solution-based ICPMS to distinguish Tautog nursery areas with high rate of classification success was suitable for this species due to its early life-history characteristics. Some of the factors contributing to this high classification success were: (1) Tautog has a larval period of 17 to 20 days, so the larval signature is very small compared to the juvenile signal (Dorrf and Powell 1997); and (2) once settled, Tautog juveniles only move about 20 m during the first year of life (Able et al. 2005). Thus, with Tautog, there is small chance to have ontogenetic movement to different habitats within a year of life. Therefore, we thought that solution-based ICPMS was appropriate to describe the chemical signatures for their entire first year of life compared to other techniques such as laser ablation ICPMS (LA ICPMS) or electron ion microprobe. Elemental fingerprints, however, should not be regarded as temporally and spatially stable markers of actual estuarine habitat or environment (Forrester and Swearer 2002, Swearer et al. 2003). Estuarine habitats are very dynamic, with seawater properties and composition at a particular location varying over tidal to annual time scales (Church 1986, Eldson and Gillanders 2003, Peters 1999). As a result, it might be expected that the variations in elemental fingerprints in otoliths among estuaries will not remain constant over time. The significant inter-annual differences we report among year classes in age-0 Tautog otolith elemental signatures, are similar to inter-annual differences in otolith chemistry reported for other marine fishes (Gillanders 2005, Gillanders and Kingsford 2000, Milton et al. 1997). Despite inter-annual differences, there were distinct spatial trends in otolith isotopic signatures’ concentrations among nursery regions that were 212 Northeastern Naturalist Vol. 19, No. 2 similar among the cohorts. For example, δ18O concentrations remained similar in the high-salinity stations of New Jersey and Connecticut for the two years. At the RI (Rose Island) station, Tautog individuals had the most enriched 18O for the two years, whereas CT had the most depleted. Successful discrimination of some Tautog estuarine nurseries along the US east coast in the present study was accomplished through the combined multielemental fingerprints of otolith elemental concentrations and stable isotopic signatures, fulfilling one of the requirements for their possible use as natural tags (Campana et al. 2000). We hope to use this tool to determine the magnitude and timing of the movement of Tautog and other estuarine-dependent fish populations between estuaries, and to identify which estuaries are responsible for recruitment to these populations in offshore habitats. Trace element and isotopic signatures in otoliths have shown promise in the present study as natural tags of nursery area in Tautog. With more comprehensive sampling of putative nursery grounds, it should be possible to track movement patterns of adult Tautog by using the YOY juvenile portion of the adult otoliths. The present results are an important advancement towards establishing evidence of estuarine fish juvenile movement to adult offshore habitats, which must be examined in fish nursery evaluation studies (Beck et al. 2001). Understanding habitat linkages between estuarine nurseries and adult populations is important in fisheries management and marine conservation because it can aid in prioritizing conservation efforts towards important nursery habitats contributing to the overall fisheries production. Acknowledgments For helping in the collection of fish, we would like to thank Chris Powell, Michelle Burnett, and Brian Murphy from RIDEM, as well as Prentice Stout from Camp Fuller, and Robert Dickau from Pond Shore Association. Special thanks go to Bryan Taplin, Richard Pruell, and the late Dr. Lesa Meng from EPA, and to Dr. Kathleen Castro from URI Sea Grant Fisheries Extension for support and inspiration for this project. We also like to thank Robin Weber from Narragansett Bay National Estuarine Research Reserve for her help with the map figures. This study was funded through University of Rhode Island Sea Grant Program and the Nature Conservancy Global Marine Initiative. Literature Cited Able, K.W., L.S. Hales, and S.M. Hagan. 2005. Movement and growth of juvenile (age 0 and 1+) Tautog (Tautoga onitis L.) and Cunner (Tautogolabrus adspersus Walbaum) in a southern New Jersey estuary. Journal of Experimental Marine Biology and Ecology 327:22–35. Arai, T., and N. Miyazaki. 2001. Use of otolith microchemistry to estimate the migratory history of the Russian Sturgeon, Acipenser guldenstadti. Journal of the Marine Biological Association of the United Kingdom 81:709–710. Arendt, M.D. 1999. Seasonal residence, movement, and activity of adult Tautog (Tautoga onitis) in lower Chesapeake Bay. M.Sc. Thesis. School of Marine Science, College of William and Mary, Gloucester Point, VA. 104 pp. 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 213 Arslan, Z., and A.J. Paulson. 2003. Solid-phase extraction for analysis of biogenic carbonates by electrothermal vaporization inductively coupled plasma mass spectrometry (ETV-ICP-MS): An investigation of rare-earth element signatures in otolith microchemistry. Analytica Chimica Acta 476:1–13. Bath, G.E., S.R. Thorrold, C.M. Jones, S.E. Campana, J.W. McLaren, and H. Lam. 2000. Strontium and barium uptake in aragonitic otoliths of marine fish. Geochimica et Cosmochimica Acta 64:1705–1714. Beck, M.W., K.L. Heck, Jr., K.W. Able, D.L. Childers, D.B. Eggleston, B.M. Gillanders, B. Halpern, C.G. Hays, K. Hoshino, T.J. Minello, R.J. Orth, P.F. Sheridan, and M.P. Weinstein. 2001. The identification, conservation, and management of estuarine and marine nurseries for fish and invertebrates. BioScience 51:633–641. Campana, S.E. 1999. Chemistry and composition of fish otoliths: Pathways, mechanisms, and applications. Marine Ecology Progress Series 188:263–297. Campana, S.E., and J.A. Gagné. 1995. Cod stock discrimination using ICPMS elemental assays of otoliths. Pp. 671–691, In D.H. Secor, J.M. Dean, and S.E. Campana (Eds.). Recent Developments in Fish Otolith Research. University of South Carolina Press, Columbia, SC. Campana, S.E., and J.D. Neilson. 1985. Microstructure of fish otoliths. Canadian Journal of Fisheries and Aquatic Sciences 42:1014–1032. Campana, S.E., and S.R. Thorrold. 2001. Otoliths, increments, and elements: Keys to a comprehensive understanding of fish populations? Canadian Journal of Fisheries and Aquatic Sciences 58:1–9. Campana, S.E., A.J. Fowler, and C.M. Jones. 1994. Otolith elemental fingerprinting for stock identification of Atlantic Cod (Gadus morhua) using laser ablation ICPMS. Canadian Journal of Fisheries and Aquatic Sciences 51:1942–1950. Campana, S.E., G.A. Chouinard, J.M. Hanson, A. Frechet, and J. Brattey. 2000. Otolith elemental fingerprints as biological tracers of fish stocks. Fisheries Research 46:343–357. Church, T.M. 1986. Biogeochemical factors influencing the residence time of microconstituents in a large tidal estuary, Delaware Bay. Marine. Chemistry. 18:393–406. Cottrell, H., I. Huber, L. Carlson, and A. Lipsky. 1999. Coastal wetland habitat mapping in Narragansett Bay, Rhode Island/Massachusetts. Proceedings of Estuarine Research Federation 1st Biennial International Conference, New Orleans, LA. Dorf, B.A., and J.C. Powell. 1997. Distribution, abundance, and habitat characteristics of juvenile Tautog (Tautoga onitis, Family Labridae) in Narragansett Bay, Rhode Island, 1988–1992. Estuaries 20:589–600. Edmonds, J.S., N. Caputi, and M. Morita. 1991. Stock discrimination by trace-element analysis of otoliths of Orange Roughy (Hoplostethus atlanticus), a deep-water marine teleost. Australian Journal of Marine and Freshwater Research 42:383–389. Edmonds, J.S., R.A. Steckis, M.J. Moran, N. Caputi, and M. Morita. 1999. Stock delineation of Pink Snapper and Tailor from Western Australia by analysis of stable isotope and strontium/calcium ratios in otolith carbonate. Journal of Fish Biology 55:243–259. Elsdon, T.S., and B.M. Gillanders. 2003. Relationship between water and otolith elemental concentrations in juvenile Black Bream Acanthopagrus butcheri. Marine Ecology Progress Series 260:263–272. Elsdon, T.S., B.K. Wells, S.E. Campana, B.M. Gillanders, C.M. Jones, K.E. Limburg, D.H. Secor, S.R. Thorrold, and B.D. Walther. 2008. Otolith chemistry to describe movements and life-history parameters of fishes: Hypotheses, assumptions, limitations, and inferences. Oceanography and Marine Biology: An Annual Review 46:297–338. 214 Northeastern Naturalist Vol. 19, No. 2 Farrell, J., and S.E. Campana. 1996. Regulation of calcium and strontium deposition on the otoliths of juvenile Tilapia, Oreochromis niloticus. Comparative Biochemistry and Physiology 115A:103–109. Forrester, G.E., and S.E. Swearer. 2002. Trace elements in otoliths indicate the use of open-coast versus bay nursery habitats by juvenile California Halibut. Marine Ecology Progress Series 241:201–213. Gannes, L.Z., D.M. O’Brien, and C.M. del Rio. 1997. Stable isotopes in animal ecology: Assumptions, caveats, and a call for more laboratory experiments. Ecology 78:1271–1276. Gauldie, R.W., D.A. Fournier, D.E. Dunlop, and G. Coote. 1986. Atomic emission and proton microprobe studies of the ion content of otoliths of Chinook Salmon aimed at recovering the temperature life history of individuals. Comparative Biochemistry and Physiology 84A:607–615. Gillanders, B.M. 2002a. Connectivity between juvenile and adult fish populations: Do adults remain near their recruitment estuaries? Marine Ecology Progress Series 240:215–223. Gillanders, B.M. 2002b. Temporal and spatial variability in elemental composition of otoliths: Implications for determining stock identity and connectivity of populations. Canadian Journal Fish Aquatic Sciences 59:669–679. Gillanders, B.M. 2005. Using elemental chemistry of fish otoliths to determine connectivity between estuarine and coastal habitats. Estuarine Coastal Shelf Science 64:47–57. Gillanders, B.M., and M.J. Kingsford. 1996. Elements in otoliths may elucidate the contribution of estuarine recruitment to sustaining coastal reef populations of a temperate reef fish. Marine Ecology Progress Series 141:13–20. Gillanders, B.M., and M.J. Kingsford. 2000. Elemental fingerprints of otoliths of fish may distinguish estuarine “nursery” habitats. Marine Ecology Progress Series 201:273–286. Gillanders, B.M., P. Sanchez-Jerez, J. Bayle-Sempere, and A. Ramos-Espla. 2001. Trace elements in otoliths of the two-banded bream from a coastal region in the southwest Mediterranean: Are there differences among locations? Journal of Fish Biology 59:350–363. Kalish, J.M. 1990. Use of otolith microchemistry to distinguish the progeny of sympatric anadromous and non-anadromous salmonids. Fishery Bulletin 88:657–666. Kalish, J.M. 1991a. 13C and 18O isotopic disequilibria in fish otoliths: Metabolic and kinetic effects. Marine Ecology Progress Series 75:191–203. Kalish, J.M. 1991b. Oxygen and carbon stable isotopes in the otoliths of wild and laboratory- reared Australian Salmon (Arripis trutta). Marine Biology 110:37–47. Kennedy, B.P., C.L. Folt, J.D. Blum, and C.P. Chamberlain. 1997. Natural isotope markers in Salmon. Nature 387:766–767. Kraus, R.T., and D.H. Secor. 2004. Incorporation of strontium into otoliths of an estuarine fish. Journal of Experimental Marine Biology and Ecology 302(1):85–106. Limburg, K.E. 1995. Otolith strontium traces environmental history of subyearling American Shad, Alosa sapidissima. Marine Ecology Progress Series 119:25–35. Limburg, K.E., P. Landergren, L. Westin, M. Elfman, and P. Kristiansson. 2001. Flexible modes of anadromy in Baltic Sea Trout: Making the most of marginal spawning streams. Journal of Fish Biology 59:682–695. 2012 I. Mateo, E.G. Durbin, D.A. Bengtson, R. Kingsley, P.K. Swart, and D. Durant 215 Markwitz, A., D. Grambole, F. Herrmann, W.J. Trompetter, T. Dioses, and R.W. Gauldie. 2000. Reliable micro-measurement of strontium is the key to cracking the life-history code in the fish otolith. Nuclear Instruments and Methods in Physics Research B 168:109–116. Meng, L., J.C. Powell, and B. Taplin. 2001. Using Winter Flounder growth rates to assess habitat quality across an anthropogenic gradient in Narragansett Bay, Rhode Island. Estuaries 24:576–584. Meyer-Rochow, V.B., I. Cook, and C.H. Hendy. 1992. How to obtain clues from the otoliths of an adult fish about the aquatic environment it has been in as a larva. Comparative Biochemistry and Physiology 103A:333–335. Milton, D.A., S.R.Chenery, M.J. Farmer, and S.J.M. Blaber. 1997. Identifying the spawning estuaries of the tropical shad, Terubok tenualosa Toli, using otolith microchemistry. Marine Ecology Progress Series 153:1–13 Northcote, T.G., C.H. Hendy, C.S. Nelson, and J.A.T. Boubee. 1992. Tests for migratory history of the New Zealand Common Smelt (Retropinna retropinna (Richardson)) using otolith isotopic composition. Ecology of Freshwater Fish 1:61–72. Patterson, H.M., S.R. Thorrold, and J.M. Shenker. 1999. Analysis of otolith chemistry in Nassau Grouper (Epinephelus striatus) from the Bahamas and Belize using solutionbased ICP-MS. Coral Reefs 18:171–178. Peters, H. 1999. Spatial and temporal variability of turbulent mixing in an estuary. Journal Marine Research 57:805–845. Radtke, R.L., and R.A. Kinzie. 1996. Evidence of a marine larval stage in endemic Hawaiian Stream Gobies from isolated high-elevation locations. Transactions of the American Fisheries Society 125:613–621. Radtke, R.L., and D.J. Shafer. 1992. Environmental sensitivity of fish otolith microchemistry. Australian Journal of Marine and Freshwater Research 43:935–951. Radtke, R.L., D.W. Townsend, S.D. Folsom, and M.A. Morrison. 1990. Strontium:calcium concentration ratios in otoliths of Herring larvae as indicators of environmental histories. Environmental Biology of Fishes 27:51–61. Radtke, R.L., J.B. Dempson, and J. Ruzicka. 1998. Microprobe analyses of anadromous Arctic Charr, Salvelinus alpinus, otoliths to infer the life-history migration events. Polar Biology 19:1–8. Rieman, B.E., D.L. Myers, and R.L. Nielsen. 1994. Use of otolith microchemistry to discriminate Oncorhynchus nerka of resident and anadromous origin. Canadian Journal of Fisheries and Aquatic Sciences 51:68–77. Secor, D.H. 1992. Application of otolith microchemistry analysis to investigate anadromy in Chesapeake Bay Striped Bass Morone saxatilis. Fishery Bulletin 90:798–806. Sogard, S.M., K.W. Able, and M.P. Fahay. 1992. Early life history of the Tautog, Tautoga onitis, in the Mid-Atlantic Bight. Fishery Bulletin 90:529–539. Swearer, S.E., G.E. Forrester, M.A. Steele, A.J. Brooks, and D.W. Lea. 2003. Spatio-temporal and interspecific variation in otolith trace-elemental fingerprints in a temperate estuarine fish assemblage. Estuarine Coastal Shelf Science 56:1111–1123. Thorrold, S.R., and J.A. Hare. 2002. Otolith applications in reef fish ecology. Pp. 243–264, In P.F. Sale (Ed.). Coral Reef Fishes: Dynamics and Diversity in a Complex Ecosystem. Academic Press, San Diego, CA. Thorrold, S.R., S.E. Campana, C.M. Jones, and P.K. Swart. 1997a. Factors determining δ13C and δ18O fractionation in aragonitic otoliths of marine fish. Geochimica et Cosmochimica Acta 61:2909–2919. 216 Northeastern Naturalist Vol. 19, No. 2 Thorrold, S.R., C.M. Jones, and S.E. Campana. 1997b. Response of otolith microchemistry to environmental variations experienced by larval and juvenile Atlantic Croaker (Micropogonias undulatus). Limnology and Oceanography 42:102–111. Thorrold, S.R., C.M. Jones, S.E. Campana, J.W. McLaren, and H. Lam. 1998a. Trace element signatures in otoliths record natal river of juvenile American Shad (Alosa sapidissima). Limnology and Oceanography 43:1826–1835. Thorrold, S.R., C.M. Jones, P.K. Swart, and T.E. Targett. 1998b. Accurate classification of juvenile Weakfish, Cynoscion regalis, to estuarine nursery areas based on chemical signatures in otoliths. Marine Ecology Progress Series 173:253–265. Thorrold, S.R., C. Latkoczy, P.K. Swart, and C.M. Jones. 2001. Natal homing in a marine fish metapopulation. Science 291:297–299. Townsend, D.W., R.L. Radtke, S. Corwin, and D.A. Libby. 1992. Strontium:calcium ratios in juvenile Atlantic Herring, Clupea harengus L., otoliths as a function of water temperature. Journal of Experimental Marine Biology and Ecology 160:131–140. Townsend, D.W., R.L. Radtke, D.P. Malone, and J.P. Wallinga. 1995. Use of otolith strontium:calcium ratios for hindcasting larval Cod, Gadus morhua, distributions relative to water masses on Georges Bank. Marine Ecology Progress Series 119:37–44. Townsend, D.W., R.L. Radtke, M. Morrison, and S. Folsom. 1989. Recruitment implications of larval Herring overwintering distributions in the Gulf of Maine, inferred using a new otolith technique. Marine Ecology Progress Series 55:1–13. Tzeng, W.N., K.P. Severin, and H. Wickström. 1997. Use of otolith microchemistry to investigate the environmental history of European Eel, Anguilla anguilla. Marine Ecology Progress Series 149:73–81. Walther, B.D., and S.R. Thorrold. 2009. Inter-annual variability in isotope and elemental ratios recorded in otoliths of an anadromous fish. Journal of Geochemical Exploration 102:181–186. Wells, B.K., G.E. Bath, S.R. Thorrold, and C.M. Jones. 2000. Incorporation of strontium, cadmium, and barium in juvenile Spot (Leiostomus xanthurus) scales reflects water chemistry. Canadian Journal of Fisheries and Aquatic Sciences 57:2122–2129. Wells, B.K., B.E. Rieman, J.L. Clayton, D.L. Horan, and C.M. Jones. 2003. Relationships between water, otolith, and scale chemistries of West Slope Cutthroat Trout from the Coeur d'Alene River, Idaho: The potential application of hard-part chemistry to describe movements in freshwater. Transactions of the American Fisheries Society 132:409–424. Yamashita, Y., T. Otake, and H. Yamada. 2000. Relative contributions from exposed inshore and estuarine nursery grounds to the recruitment of Stone Flounder, Platichthys bicoloratus, estimated using otolith Sr:Ca ratios. Fisheries Oceanography 9:316–327. Zimmerman, C.E., and R.L. Nielsen. 2003. Effect of analytical conditions in wavelength dispersive electron microprobe analysis on the measurement of strontium-to-calcium (Sr/Ca) ratios in otoliths of anadromous salmonids. Fishery Bulletin 101:712–718.