Identification of the optimal fluorochrome for marking larvae of the pulmonate limpet Siphonaria australis.
KEY WORDS: connectivity, fluorochrome, limpet, Siphonaria australis
Determining levels of population connectivity is vital if the resilience of marine species is to be ensured in the face of increasing anthropogenic threats (i.e., climate change, overfishing, and habitat degradation) (Hoffmann & Gaines 2008). In itself, population connectivity consists of larval dispersal and postlarval survival, with the latter being more easily quantified than the former (Pineda et al. 2007). Efforts to determine larval dispersal necessitate some form of population marker be used to identify individuals of interest (e.g., genetic differences, trace elements, or chemical marks in calcified structures) (Levin 2006). Moreover, to be successful, a population marker must satisfy four key criteria: low cost, benign in terms of mortality, benign in terms of behavior of tagged individuals, and able to mass mark large numbers of individuals that can be unambiguously identified from among nonmarked individuals (Thorrold et al. 2002). To this end, fluorochrome compounds make ideal candidates for tracking the dispersal of shelled invertebrate larvae (Thorrold et al. 2002).
Fluorochrome stains act by chelating the calcium of the larval shell thereby causing a permanent mark that fluoresces when viewed under ultraviolet (UV) wavelengths (Day et al. 1995). Because of these properties, fluorochromes have been widely used in studying vertebrate and invertebrate taxa, especially for the validation of growth marks in otoliths and shells (Levin 1990). However, some fluorochromes can elevate mortality levels depending on the species immersed and the compound used (Purcell & Blockmans 2009). Fortunately, a number of fluorochromes are available to researchers, of which alizarin red S (ARS), calcein, and oxytetracycline (OTC) are often used in experimental studies (Thorrold et al, 2002). Thus, it is essential that the optimal fluorochrome for larval tagging be identified experimentally for a species.
An issue with comparisons of ftuorochrome efficacy is that subjective methods (i.e., qualitative scales) have been used to compare mark brightness among fluorochromes. To reduce such bias, the use of complex mathematical algorithms can be used, but they require a familiarity with software programming beyond nonsubject specialists (Purcell et al. 2006). With the advent of image analysis software freely available on the Web (e.g., ImageJ, http://rsbweb.nih.gov/ij), quantitative measures can be achieved without such specialist knowledge, and they may be of use to nonacademic end users (e.g., community groups) looking to use fluorochrome staining in community-based conservation projects.
We sought to identify the optimal fluorochrome for staining the coastal gastropod Siphonaria australis (Quoy and Gairmard, 1883), a very common species on rocky shores throughout New Zealand. This species is highly fecund, laying gelatinous egg masses holding up to 18,000 capsule-bound larvae, at every new moon (deSilva 2001). Larvae develop for an additional 6-10 d within capsules before hatching into the water column to complete their planktonic development and dispersal phase (Russell & Phillips 2009). Such reproductive characteristics make this species ideal for use in studies of larval dispersal, because the young remain in their natal conditions throughout the staining process. Furthermore, monthly immersions of fresh egg masses are possible, and their allocation within a reciprocal transplant regime among sites allows the exposure of spatiotemporal patterns in larval dispersal. However, such methods are contingent upon a successful fluorochrome staining process that can penetrate the gelatinous egg masses and capsules within which larval S. australis develop.
The aim of this study, therefore, was to provide quantitative measures of mark brightness using Image J software to identify which of three commonly used fluorochromes (ARS, calcein, OTC) was optimal for marking developing S. australis larvae still contained within capsules and egg masses. Qualities sought in a fluorochrome were that 100% of the larvae immersed were marked with the brightest mark possible, and that mortality was negligible compared with other treatments and a control with no chemical treatment.
Larval Collection and Maintenance
Live S. australis larvae were obtained by collecting egg masses in September 2009 and January 2010 for the mortality and brightness experiments, respectively. Newly laid egg masses were gently scraped from the rocky shore at Matheson Bay, northern New Zealand (36[degrees]18'02" S, 174[degrees]48'07" E). Back at the laboratory, individual egg masses were maintained for 1 wk before experimentation in separate 20-mL glass vials containing 1 [micro]m filtered and UV-treated seawater. Vials were held at ambient seawater temperatures (15 [+ or -] 1 [degrees]C, mortality experiment; 20 [+ or -] 1 [degrees]C, brightness experiments) via a flow-through water bath. To minimize stress, the seawater in the vials was exchanged every 48 h.
Larval Immersion in Fluorochrome Solutions
Stock solutions (1 g/L) of the fluorochromes ARS, OTC, and calcein (see Table 1 for more detailed chemical information) were prepared by mixing each fluorochrome in distilled water. Sodium bicarbonate (NaHC[O.sub.3]) was added to maintain solutions at a pH of 7 and to enhance the solubility of the compounds (Wilson et al. 1987, Moran 2000, Moran & Marko 2005). Addition of the stock fluorochrome solution to each vial holding S. australis egg masses was performed to achieve the desired concentrations of 200 mg/L.
To assess brightness of the fluorochrome mark, 10 vials of each fluorochrome were set up and a randomly selected egg mass was allocated to each vial. Controls consisted of 10 vials of 1 [micro]m filtered seawater with an egg mass held within each. After 24 h of immersion, all egg masses were removed and washed 3 times with filtered seawater, then maintained in 1 [micro]m filtered seawater for 24 h. Vials containing the egg masses were then agitated on a vortex mixer for 2 min, thereby separating the egg capsules from the egg mass as described in Przeslawski et al. (2004). Individual egg capsules were randomly selected and removed from the vials using a glass pipette and placed in concave microscope slides for inspection under a fluorescence microscope (Leitz Diaplan, Sydney, Australia). To visualize larvae, a series of filter cubes were used for excitation of the fluorochromes (Table 2).
Digital images of the egg capsules were captured using a Leica DFC 490 camera and AnalySiS LifeScience software from Olympus NZ. Images were then analyzed for brightness using ImageJ software's freehand selection tool and histogram analysis function. This function quantifies the brightness of each pixel within the selected area, as a value between 0 (black) and 255 (white). The number of pixels at each value within the selected area is plotted on a histogram and the overall mean brightness of the selected area is established. The mean brightness was recorded as a measure of total chemical fluorescence from each larva, thus providing a nonsubjective measure of brightness not found in previous studies on fluorochrome marking.
To assess larval mortality resulting from fluorochrome immersion, a similar experiment was monitored. A total of 40 egg masses were randomly selected and allocated to separate vials, 10 ten for each of the 3 candidate fluorochromes and 10 for the control treatments. Prior to immersion in the fluorochrome solutions, an initial assessment of mortality (quantities of dead vs. live larvae) was conducted to obtain mortality levels at t = 0. To ensure a consistent sampling regime, egg masses were visually divided into 4 quarters, and the first 25 larvae sighted within each quarter were assessed for movement for 10 sec under a compound microscope at 100 x magnification. If no movement was observed within this time period, larvae were recorded as dead. This was repeated after immersion and a subsequent 24-h recovery period to allow pre- and postimmersion comparisons.
A Levenes test revealed nonhomogeneity of variances, thus differences in mean brightness between fluorochrome treatments were tested using Welch's analysis of variance (ANOVA), which is robust to nonequal variance among treatments (Zar 2010). Statistically significant differences in fluorochrome brightness was then identified via Tukey's HSD tests, with [alpha] = 0.05 for all statistical comparisons. To determine any significant change in mortality resulting from fluorochrome immersion, the before-and-after means were analyzed using nested ANOVA, fitted to a Poisson distribution because of a lack of normality in the distribution of the experimental data. The statistical program SAS (SAS Institute, Cary, NC) was used for all data analyses.
For all three fluorochromes there was 100% marking success, with calcein- and OTC-immersed larvae found to be fluorescing brighter than control larvae 48 h after immersion (Fig. 1). No significant difference in brightness could be found between ARS-immersed larvae and controls. Among fluorochrome treatments, mean brightness of the larvae differed significantly (Welch's ANOVA; df = 3, F = 73.75, P < 0.001), with larvae immersed in a 200-mg/L solution of calcein (mean brightness, 30.97 [+ or -] 1.05 (SEM)) significantly brighter than larvae immersed in 200 mg/L OTC (mean brightness, 20.55 [+ or -] 0.38) or ARS (mean brightness, 15.45 [+ or -] 0.57; Fig. 1). Finally, the brightness exhibited by control shells indicates that S. australis autofluoresces when viewed under UV light.
When comparing with preimmersion values, larvae immersed in ARS experienced significantly higher levels of mortality postimmersion than larvae immersed in either calcein or OTC (Table 3). Although there was a trend for the number of dead larvae to increase in the control after immersion, this was not statistically significant (Table 3). This finding resulted from high levels of mortality (4 dead larvae preimmersion, 97 dead larvae postimmersion) within 1 of the 10 egg masses allocated to the control treatment, thereby inflating levels of variance in mean mortality.
Using three common fluorochromes (ARS, calcein, and OTC), we sought to identify an effective protocol for staining larval S. australis still contained within gelatinous egg masses. Efficacy of staining protocols was measured in terms of mark brightness and larval mortality postimmersion.
[FIGURE 1 OMITTED]
Consistent marking of all stained larvae is a critical component of successful chemical marking methodologies (Thorrold et al. 2002). In this study, all S. australis larvae immersed in the three fluorochrome treatments were found to fluoresce 48 h postimmersion. This result agrees with previous studies in which immersion of abalone (Haliotis rubra) in OTC and calcein, scallop (Argopecten irradians) and mussel larvae (Mytilus trossulus) in calcein, and the saucer scallop (Amusium balloti) in calcein, OTC, and ARS all resulted in 100% marking success (Day et al. 1995, Moran & Marko 2005, Lucas et al. 2009). Furthermore, we have shown that fluorochromes can effectively penetrate the gelatinous matrix of gastropod egg masses and offer a reliable method for staining the developing prodissoconchs within. This is of value to studies of larval dispersal dynamics in coastal species where transplants of stained gastropod larvae among sites are to be undertaken, because movement of larvae brooded within egg masses is logistically simpler than free-living broadcast individuals. In addition, this method does not involve intervention in other larval characteristics, such as feeding condition, in hatchery cultured larvae.
Quantifying fluorochrome mark brightness in molluscan shells using the freely available software ImageJ has provided a simpler and more accessible method for nonspecialist end users of fluorochrome marking techniques than the mathematical algorithms outlined in a previous study (i.e., Purcell & Blockmans (2009)). Using a quantitative, rather than a subjective, approach allows unequivocal comparisons of the brightness of fluorochrome marks to be made.
In our work, the mark calcein produced in stained S. australis larvae was 25% brighter than that achieved for OTC and 50% brighter than that of ARS. Using similar quantitative methods, calcein consistently produced brighter marks than OTC in juvenile sea cucumbers (Holothuria scabra) when immersed for 24 h (Purcell & Blockmans 2009). Moreover, in studies in which concentrations of calcein solutions were 90% lower than OTC, a brighter mark was still recorded for calcein (e.g., in the Australian abalone, H. rubra stained for 24 h (Day et al. 1995)). However, such a trend is not clear-cut across all species, with OTC rather than calcein producing the brightest mark in saucer scallop (A. balloti) spat (Lucas et al. 2009). Thus, it is advisable that the optimal fluorochrome be identified from a range of compounds before attempts to mark marine species are undertaken.
In our study, immersion of larval S. australis in either calcein or OTC for 24 h did not significantly increase larval mortality above preimmersion values. As a compound for marking marine mollusc species, calcein appears comparatively benign, with negligible mortality reported in stained mussels (Perna perna and M. trossulus), whelks (Nucella ostrina), and scallops (A. irradians) (Kaehler & McQuaid 1999, Moran 2000, Bashey 2004, Moran & Marko 2005).
However, OTC immersion has, in certain cases, elevated mortality levels of the study species (i.e., red drum (Sciaenops ocellatus)) larvae and juveniles (Thomas et al. 1995). The exact pathology of OTC staining is unknown, but does appear species specific with abalone (H. iris) and mussel (Lampsilis cardium) species unaffected by OTC staining (Pirker & Schiel 1993, Eads & Layzer 2002). The mortality of S. australis larvae was elevated by 24-h immersion in ARS; accordingly, we do not recommend its use for staining S. australis larvae in future studies. As a compound, ARS has had variable success in marking, with high mortality and reduced marking success noted in white sucker fish larvae (Catostomus commersoni) immersed in ARS (Beckman & Schulz 1996). In addition, some species exhibit increased mortality depending on life stage when immersion occurred, with mortality of 5-d-old whitefish (Coregonus lavaretus) larvae twice that of 14-d-old larvae (Eckmann 2003). Such results again emphasize the need to identify optimal fluorochromes for staining experimental animals, particularly when sensitive life stages such as larvae are to be tested.
A synthesis of our results reveals that although more expensive (Table
1), calcein is the best fluorochrome candidate for the staining of S. australis larvae contained within egg masses. Although budgetary restraints are a significant consideration in the design of future experiments, the superior brightness and lower mortality levels produced by calcein staining offsets its higher purchase price.
Given that a reliable and noninvasive marking protocol has been designed for S. australis, future work is now needed on assessing mark durability under ambient conditions and whether increased predation of marked versus nonmarked individuals occurs. Once established, there is then the opportunity to track the dispersal of a highly fecund and ecologically important species, and to stain thousands of individuals without the need for the laborious efforts and expensive shellfish hatchery facilities often required in these studies.
This study was made possible by a University of Auckland Faculty Research Development Fund to B. D. and A. J. Statistical advice was provided by Brian McArdle and Nick Shears.
Bashey, F. 2004. A comparison of the suitability of alizarin red S and calcein for inducing a nonlethally detectable mark in juvenile guppies. Trans. Am. Fish. Soc. 133:1516-1523.
Beckman, D. W. and R. G. Schulz. 1996. A simple method for marking fish otoliths with alizarin compounds. Trans. Am. Fish. Soc. 125:146-149.
Day, R. W., M. C. Williams & G. W. Hawkes. 1995. A comparison of fluorochromes for marking abalone shells. Mar. Freshw. Res. 46:599-605.
deSilva, D. 2001. Reproductive biology and population ecology of pulmonate limpet Siphonaria zelandica. MS thesis, Leigh Marine Laboratory, University of Auckland. 186 pp.
Eads, C. B. & J. B. Layzer. 2002. How to pick your mussels out of a crowd: using fluorescence to mark juvenile freshwater mussels. J. North Am. Benthol. Soc. 21:476-486.
Eckmann, R. 2003. Alizarin marking of whitefish, Coregonus lavaretus, otoliths during egg incubation. Fish. Manag. Ecol. 10:233-239.
Hoffmann, G. E. & S. D. Gaines. 2008. New tools to meet new challenges: emerging technologies for managing marine ecosystems for resilience. Bioscience 58:43-52.
Kaehler, S. & C. D. McQuaid. 1999. Use of the fluorochrome calcein as an in situ growth marker in the brown mussel Perna perna. Mar. Biol. 133:455-460.
Levin, L. A. 1990. A review of methods for labeling and tracking marine invertebrate larvae. Ophelia 32:115-144.
Levin, L. A. 2006. Recent progress in understanding larval dispersal: new directions and digressions. Integr. Comp. Biol. 46:282-297.
Lucas, T., P. J. Palmer, S. Wang, R. Scoones & E. O'Brian. 2009. Marking the shell of the saucer scallop Amusium balloti for sea ranching using oxytetracycline, calcein, and alizarin red S. J. Shellfish Res. 27:1183-1188.
Moran, A. L. 2000. Calcein as a marker in experimental studies of newly-hatched gastropods. Mar. Biol. 137:893-898.
Moran, A. L. & P. B. Marko. 2005. A simple technique for physical marking of larvae of marine bivalves. J. Shellfish Res. 24:567-571.
Pineda, J., J. A. Hare & S. U. Sponaugle. 2007. Larval transport and dispersal in the coastal ocean and consequences for population connectivity. Oceanography (Wash. DC) 20:22-39.
Pirker, J. G. & D. R. Schiel. 1993. Tetracycline as a fluorescent shell-marker in the abalone Haliotis iris. Mar. Biol. 116:81-86.
Przeslawski, R., A. R. Davis, et al. 2004. Effects of ultraviolet radiation and visible light on the development of encapsulated molluscan embryos. Mar. Ecol. Prog. Ser. 268:151-160.
Purcell, S. W. & B. F. Blockmans. 2009. Effective fluorochrome marking of juvenile sea cucumbers for sea ranching and restocking. Aquaculture 296:263-270.
Purcell, S. W., B. F. Blockmans & W. J. Nash. 2006. Efficacy of chemical markers and physical tags for large-scale release of an exploited holothurian. J. Exp. Mar. Biol. Ecol. 334:283-293.
Russell, J. & N. Phillips. 2009. Synergistic effects of ultraviolet radiation and conditions at low tide on egg masses of limpets (Benhamina obliquata; and Siphonaria australis) in New Zealand. Mar. Biol. 156:579-587.
Thomas, L. M., S. A. Holt & C. R. Arnold. 1995. Chemical marking techniques for larval and juvenile red drum (Sciaenops ocellatus) M. E. R. otoliths using different fluorescent markers. In: D. H. Secor, J. M. Dean & S. E. Campana, editors. Recent developments in fish otolith research. Columbia: University of South Carolina Press. pp. 703-717.
Thorrold, S. R., G. P. Jones, M. E. Hellburg, R. S. Burton, S. E. Swearer, J. E. Neigel, S. G. Morgan & R. R. Warner. 2002.
Quantifying larval retention and connectivity in marine populations with artificial and natural markers. Bull. Mar. Sci. 70:291-308.
Wilson, C. A., D. W. Beckman & J. M. Dean. 1987. Calcein as a fluorescent marker of otoliths of larval and juvenile fish. Trans. Am. Fish. Soc. 116:668-670.
Zar, J. H. 2010. Biostatistical analysis. Upper Saddle River, N J: Pearson Education. 944 pp.
M. P. FITZPATRICK, (1) A. G. JEFFS (1) AND B. J. DUNPHY (2), *
(1) Leigh Marine Laboratory, University of Auckland, Private Bag 92019, Auckland 1143, New Zealand; (2) School of Biological Sciences, University of Auckland, Private Bag 92019, Auckland 1143, New Zealand
* Corresponding author. E-mail: email@example.com
TABLE 1. Sigma-Aldrich CAS information and price for fluorochromes used in this study. Fluorochrome Chemical Synonym Alizarin red S 3,4-Dihydroxy-9,10-dioxo-2-anthracenesulfonic acid sodium salt Calcein Bis[N,Nbis(carboxymethyl)aminomethyl] fluorescein Oxytetracycline 2-Naphthacenecarboxamide Sigma Fluorochrome Product No. CAS No. Cost $US/g Alizarin red S 33010 130-22-3 3.47 Calcein C0875 1461-15-0 17.3 Oxytetracycline 2756 152-51-6 4.81 TABLE 2. Filter blocks used for mark visualization of the S. australis larvae immersed in fluorochrome solutions. Filter Excitation Split Emission Fluorochrome Cube Type (nm) (nm) (nm) Oxytetracycline A Longpass 340-380 400 >425 Calcein I3 Longpass 450-590 510 >515 Alizarin red S N2.1 Longpass 515-560 580 >590 TABLE 3. Mean mortality of larvae within S. australis egg masses before and after immersion in either alizarin red S, calcein, or oxytetracycline. Pre-immersion Post-immersion ([+ or -] SEM) ([+ or -] SEM) Alizarin red S 0.80 ([+ or -] 0.29) 11.0 ([+ or -] 4.70) Calcein 0.90 ([+ or -] 0.35) 1.0 ([+ or -] 0.63) Oxytetracycline 3.90 ([+ or -] 2.08) 3.10 ([+ or -] 1.68) Control 3.20 ([+ or -] 1.58) 15.1 ([+ or -] 9.46) ANOVA, df (1,18) Alizarin red S F = 14.84, P < 0.05 Calcein F = 0.09, P > 0.05 Oxytetracycline F = 4.84, P > 0.05 Control F = 1.9, P > 0.05 Significant results are indicated in bold type (P < 0.05).
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|Author:||Fitzpatrick, M.P.; Jeffs, A.G.; Dunphy, B.J.|
|Publication:||Journal of Shellfish Research|
|Date:||Dec 1, 2010|
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