ROBERT C. SIEGFRIED, II. Louis Berger & Associates, Inc. 1001 E. Broad Street. Richmond, Virginia 23219. MICHAEL P. WEINSTEIN. Envirosphere Company.
Estuaries Vol. 12, No. 3, p. 180-185 September 1989
Validation of Daily Increment Deposition in the Otoliths of Spot (Leiostomusxanthurus) ROBERT C. SIEGFRIED, I I
Louis Berger & Associates, Inc. 1001 E. Broad Street Richmond, Virginia 23219 MICHAEL P. WEINSTEIN
Envirosphere Company 160 Chubb Avenue Lyndhurst, New Jersey 07010 ABSTRACT: The otoliths o f 200 postlarval spot (Leiostomus xanthurus) were marked by immersing the fish for 24 hr in a solution o f 0.4 g tetracycline HCI 1-1 o f salt water (15 %~). The resulting time-mark could later be identified under ultraviolet (UV) light with an epifluorescence microscope. Spot were held in the laboratory or in the field for up to 30 d. Two validation methods were employed to document daily increment formation. The total number o f increments per otolith, or the number of increments formed after the time-mark, were regressed against days held in an experiment. The hypothesis o f one increment per day would result in a regression coefficient not significantly different from 1.0. In the field both the total count method (b = 1.05) and the time-mark method (b = 0.90) supported a one-increment-per-day hypothesis and were not significantly different from each other (p > 0.05). In the laboratory the total count method (b = 0.56) and the time-mark method (b = 0.36) were significantly different from 1.0 (p < 0.05), and thus did not support the one-increment-per-day hypothesis. Failure to resolve a oneincrement-per-day deposition rate in the laboratory was primarily attributed to food limitation in the experiments.
foundation for developing more complex daily aging research, yet it is often ignored. The preferred method for documenting daily increments involves the use of known-age fish (Taubert and Coble 1977). This method is restricted to species that can be spawned in the laboratory; for those species validation is a relatively simple process. The use of fish of unknown-age for documenting daily increment deposition presents more problems and often leads investigators into making broad assumptions so as to avoid the need for validation experiments. Few investigators have actually examined alternative validation methods to establish efficient, yet accurate, methods for aging fish. Validation experiments can also be conducted by total counts of increments in an otolith, or partial counts with the use of a time-marker. Campana and Neilson (1982) successfully attempted a field and laboratory comparison of tetracycline-marked fish. This paper examines the validation of daily increments by employing both total and partial counts of increments from groups of spot (Leiostomus xanthurus) held in the laboratory and in the field.
Introduction
The use of daily increment counts in otoliths of young fishes has become commonplace for evaluating growth, mortality, age-frequency distributions, and other population parameters (Brothers et al. 1976; Struhsaker and Uchiyama 1976; Barkman 1978; Brothers 1978; Methot 1983; Campana and Neilson 1985). A prerequisite to the use of these methods in new applications, however, is the validation of daily increment counts. As Jones (1986) noted, validation of daily increment deposition under the natural range of experience of young fishes is fundamental to accurate estimation of age-size relationships in wild fish. Previously, daily increments have been assumed from lengthfrequency data (Oxenford and Hunte 1983), and have been extrapolated from one species to another within families (Warlan 1982) and even entire ecosystems (Brothers et al. 1983). Beamish and McFarlane (1983) noted a similar disregard for age validation in annual aging research. As early as 1941 Van Oosten stated that the scale aging method was taken "too much for granted in the belief that it is a simple one which can be applied without technical knowledge." This statement holds true for daily aging techniques. Documenting the periodicity of increment formation provides the © 1989 Estuaries Research Federation
Methods
Postlarval spot used in these experiments were collected in April 1984 in the York River, Virginia, 180
0160-8347/89/030180-06501.50/0
Spot, Otolith Daily Increments, Validation
by fishing from a 4.9-m otter trawl with a 1.0-mm mesh liner. An attempt was made to capture the earliest recruiting spot (mean total length -- 16.0 mm) in order to reduce age and size variation in the samples. Prior to the experiments, fish were acclimated to laboratory conditions for 2 to 3 d to allow for mortality due to capture and handling. They were then immersed in a 15-1 tank containing a solution of 0.4 g tetracycline HC11-1 of salt water (15 %), a concentration that produces a fluorescent mark or increment that can be identified later. The 200 marked fish used in the experiments were randomly divided between laboratory and field holding containers. Twenty fish were collected from each experiment every tenth day for 30 d. The actual number of fish comprising each test sample (Table 1) varied according to the success of processing readable otoliths. Laboratory experiments were conducted in 220-1 aquaria under ambient lighting (natural photoperiod). Temperature was maintained at 20 + I°C by submersible heaters; salinity was maintained over the range 12 to 15%. Ad libitum feeding was provided once a day with a commercial flaked food. The field experiment was conducted in submersible cages (constructed of 2-in. PVC pipe and 2-mm fiberglass window screening) placed along the bank of a tidal marsh creek. T h e screens, which were periodically cleaned, allowed food and water to flow through the cage, but excluded other postlarvae and predators. During the 30 d of the field experiment, salinity and temperature increased from 5 to 8.5% and from 15°C to 20°C, respectively. All fish were preserved in 95% E T O H (replenished after 24 hr); sagittae were removed and stored in individual wells of a tissue culture tray. Details of the otolith preparation procedure are described in Haake et al. (1982). Briefly, otoliths mounted in Spurr blocks (Polysciences, Inc.) were rough cut with a fine-toothed saw to produce a transverse grinding plane, and affixed to a square of glass with a thermoplastic cement (Crystalbond, Aremco). After grinding on 600 grit sandpaper and polishing with 0.3-mm alumina, the section was reversed and similarly ground to reveal the otolith's primordium. Two methods were used to validate the periodicity of increment formation in postlarval spot. In both cases the null hypothesis that one increment per day was deposited was tested by regressing the number of increments counted against days held in an experiment and testing for a significant departure from a regression coefficient of 1.0. In the total count method the total number of increments in an otolith was determined. One otolith from each fish was read up to five times along as many
181
TABLE 1. Sample size, mean number of increments, and standard deviation arranged according to method of counting, type of experiment, and days held in an experiment for juvenile spot, Leiostomus xanthurus. DaysHeld
n
Field Mean
SD
n
Total Increment Count Method 0 18 71.3 7.2 18 10 12 81.0 7.5 16 20 14 92.8 5.8 18 30 17 102.4 7.4 16 Total sample n 61 68 10 20 30 Total sample n
Time-mark Method 14 9.7 1.7 13 16 19.6 1.1 16 16 27.7 1.8 17 46 46
Laboratory Mean SD 71.3 81.8 86.4 87.9
7.2 6.9 5.3 8.8
9.5 13.6 16.7
2.4 2.2 2.9
different transects as possible under 400-1,000 x magnification. Higher magnifications and multiple counts were occasionally required to resolve differences in closely spaced increments and to help stabilize the variability in counts. The median of these readings was used for analysis. In the second validation experiment, the number of increments between the tetracycline time-mark and the otolith's outer edge was determined. T h e time-mark was located under UV light with an epifluorescence microscope (400 x) and the increments were then counted under normal transmitted light. Two to five readings were made and the median was used for the regression analysis. Results Table 1 contains the sample size, mean number of increments, and standard deviation for each sampling date arranged according to type of experiment and method of verification. T h e regression of total number of increments against days held in the field or laboratory is plotted in Figs. 1 and 2, respectively. Figures 3 and 4 are the regressions for the number of increments counted after the time-mark for the field and laboratory experiments, respectively. Within the same verification method and experiment, the variances of the number of increments for each separate sampling date were not significantly different (p < 0.05), except for the time-mark method from the laboratory experiment. T h e regression coefficients from the total count method (b = 1.05) and the time-mark method (b = 0.90) for fish held in the field were not significantly different from 1.0 (p > 0.05) or from one another (p > 0.05). In the laboratory experiment, the regression coefficients from the total count method (b = 0.56) and the time-mark method (b = 0.36) were significantly different from a slope of 1.0 (p > 0.05). T h e regression coeffi-
182
R.C. Siegfried, il and M. P. Weinstein
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Fig. 1. Regression of otolith daily increment counts versus days held in the field (total count method) for spot, Leiostomus
Fig. 3. Regression of otolith daily increment counts versus days held in the field (time-mark method) for spot, Leiostomus
xanthurus.
xanthurus.
cients for the field experiments were significantly different from those of the laboratory experiments
ments in a small percentage of otoliths and the marks not being intense enough to be seen at magnifications above 400 x, The second problem can be overcome by increasing the dosage to 0.6-0.8 g tetracycline HC11 -] salt water, which would increase the intensity of the marks. Similarly, decreasing the exposure time from 24 to 12-18 hr should reduce the number of double-marked increments. The time-mark method has been widely used in daily aging research. Most previous attempts, however, involved injection of large juvenile or adult fishes (Wild and Foreman 1980; Campana and Neilson 1982; Campana 1983a). Injection would obviously be difficult for larval or small juvenile fishes. Hettler (1984) reported a partially successful method of marking otoliths of postlarval spot. He immersed spot (13-16 mm SL) for 60 to 120
(p < o.o5). T h e R 2 values for the total count method were 0.77 and 0.42 in the field and laboratory, respectively. T h e R 2 values for the time-mark method were 0.96 and 0.58 in the field and laboratory, respectively. Discussion T h e tetracycline time-marking method was very successful; over 99% of the spot had clearly discernible fluorescent marks visible at 4 0 0 x magnification. This method appears to subject the fish to a minimum of stress, and is inexpensive and adaptable to a variety of situations. Two slight drawbacks to this marking procedure, which could be corrected easily, are the marking of two incre-
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Fig. 2. Regression of otolith daily increment counts versus days held in the laboratory (total count method) for spot, Leios-
Fig. 4. Regression of otolith daily increment counts versus days held in the laboratory (time-mark method) for spot, Leios-
tomus xantb+tr~ts.
tomus xanthurus.
Spot, Otolith Daily Increments, Validation
min in a 0.5 g oxytetracycline 1-1 of 1% saline solution. Our dosage was comparable to Hettler's (1984), but the exposure times were much longer, 24 hr compared to 1-2 hr. Hettler reported using a 1% saline solution to avoid tetracycline chelating with magnesium and calcium in salt water. He also reported that the unbuffered tetracycline solution (pH = 2.5-3.5) was lethal. Neither problem was encountered here, perhaps partially due to the form of tetracycline used. The results of the field experiment clearly support a one-increment-per-day hypothesis. The regression coefficients from both the total count method and the time-mark method were not significantly different from 1.0; thus, increment formation occurred daily under field conditions. The results of the laboratory experiment were quite different. T h e regression coefficients for the total count method and the time-mark method were significantly different from 1.0, thus not supporting daily increment formation. A closer examination of the laboratory results revealed that the increase in mean number of increments between sampling dates decreased over time. In the total count method the increase was 10.5 increments between day 0 and day 10, which decreased to 1.5 increments between day 20 and day 30. When counted by the time-mark method, however, 9.5 increments formed between day 0 and day 10, but only 3.1 between day 20 and day 30. The fish collected on the tenth day of the laboratory experiment supported a one-incrementper-day hypothesis, but those collected thereafter did not. Some factor must have differed between the field and the laboratory to account for the difference in increment formation. Since photoperiod and temperature were similar, food availability was probably the major difference between the experiments. T h e fish held in the field grew from a mean SL of 16.6 mm to 26.4 mm in 30 d. This growth (0.32 mm d -1) was comparable to field estimates of average growth rates for spot (Weinstein and Walters 1981; Beckman and Dean 1984). T h e average growth rate for the laboratory experiment was 0.08 mm d -1, much lower than the field experiment. In the field, spot could feed in unrestricted fashion on ostracods, amphipods, and copepods. Feeding in the laboratory was limited to once daily, a typical feeding regime, with a commercial flaked food. T h e laboratory food rations may have been near or below maintenance levels, as indicated by the slow growth rates. Thus, f o o d limitation a n d / o r other factors contributed to a failure to resolve increment formation in laboratory-held fish. This seems supported by the higher variances seen in the time-mark method from the
183
laboratory experiment, a possible reflection of the difficulty in counting increments in these slower growing individuals. Campana (1983b) starved steelhead trout, Salmo gairdneri, and starry flounder; Platichthys steUatus (>70 mm SL), for 32 d, but increment formation did not break down. These larger fish had much greater energy reserves than the postlarval spot; thus, longer periods of starvation may be necessary before increment resolution deteriorates. Methot and Kramer (1979) found that the larvae of northern anchovy, Engraulis mordax, may not produce increments during starvation. T h e effects of starvation on increment deposition may be dependent on the size of a fish and its energy reserves. Marshall and Parker (1982) showed that under starved conditions sockeye salmon, Oncorhynchusnerka, grew in length but not in weight. They hypothesized the utilization of stored lipid reserves to continue linear growth despite starvation. After three weeks of starvation under cold conditions, the sockeye salmon exhibited reduced increment deposition (b = 0.57). Short periods of food limitation do not appear to affect increment deposition in larger juvenile and adult fishes, but may alter increment formation during the larval and early juvenile stages, thus producing unreliable increment counts at later dates. Another explanation for the apparent breakdown of increment formation in the laboratory experiment would be decreased microscopic resolution due to reduction of increment width a n d / or increment definition. Campana (1983a) reported that starved fish produced narrower increments with a "decrease in visual contrast." Stress lowers calcium uptake, and stable warm temperatures decrease the protein/Ca ratio, which results in narrower increments with reduced visual definition (Campana 1983a). The laboratory experiments, with more stress and higher and more stable temperatures, could produce the lower regression coefficient as a result of underestimation, instead of absence, of the increments. This was recently verified in experiments conducted by Jones and Brothers (1987). Under optimal laboratory feeding conditions, striped bass otolith i n c r e m e n t s were deposited daily through the first two months of life and were discernible with the light microscope. Under restricted feeding conditions, counts with the light microscope did not reveal true age; however, the same otolith counted under scanning electron microscopy (SEM) revealed daily increments that could not be resolved with light microscopy. Similarly, Campana et al. (1987) reported that resolution limitations appear to explain all of the reported instances of apparent non-daily increment formation.
184
R.C. Siegfried, II and M. P. Weinstein
The time-mark method produced results similar to those in the total count method. The field experiments supported a one-increment-per-day hypothesis, while the laboratory experiments did not. The time-mark method produced slightly lower, but not significantly different, regression coefficients than the total count method. The tetracycline treatment produced marks visible at 400 x but not at 1,000x, which was the magnification used in the total count method. Thus, the lower resolution at 400x may be responsible for the slightly lower regression coefficients. Both validation methods produced similar results, yet each method had its advantages and disadvantages. More time was spent in grinding and reading with the total count method than with the time-mark method. The latter method required additional labor and expense in the marking process. Some concerns have been expressed over tetracycline being photolytic (Choate 1964), but the otoliths in this study, which were stored in the dark, showed no reduction in intensity after 6 to 10 months. Because the otoliths of some species of fish require etching for reliable readings, the timemark method may not be appropriate, as tetracycline marks may be destroyed by etching. A major advantage of the time-mark method is its ability to reduce the effects of the initial age variation of fish captured from the field as evidenced by the higher R ~ values obtained for this method. As seen in this research, laboratory validation experiments may not reliably represent the rate of increment formation. Yet field validation has been used only rarely in daily aging research. Wild and Foreman (1980) employed a mark-recapture program to verify daily increment counts in adult yellowfin tuna, Thunnus albacares, but this method was very labor-intensive and not applicable to larval and juvenile fish. Struhsaker and Uchiyama (1976) followed a cohort of nehu, Stolephorus purpures, over time, but this method precluded examination of species with protracted or multiple spawning periods. Thus, if field validation proves impossible, certain measures should be taken to ensure that reliable laboratory experiments can be conducted. The use of the larger fish for a short period of time or a feeding regime with adequate high growth efficiency may be one such approach to ensuring the experiment against unreliable results. Confirmation of results with SEM would further strengthen the relationship when increments are closely spaced. Documentation of increment formation is a necessary part of any daily age and growth research. T h e conflict between field and laboratory results emphasizes the importance of conducting validation exoeriments under conditions as close to field
conditions as possible. The use of submersible cages and the tetracycline time-marks can provide quick, simple, and inexpensive field validation for suitable species. At a minimum, validation experiments in the laboratory should be Conducted with an appropriate 24-hr light : dark cycle and a successful feeding regime. ACKNOWLEDGMENTS The authors thank L. Cadman and S. Szedlymayer for help in the field. This study was supported by EPA Grant No. R810334 to MPW. LITERATURE CITED BARKMAN, R. C. 1978. The use of otolith growth rings to age young Atlantic silversides, Menidia menidia. Trans. Am. Fish. Soc. 107:790-792. BEAMISH, R. J., AND G. A. MCFARLANE. 1983. The forgotten requirement for age validation in fisheries biology. Trans. Am. Fish. Soc. 112:735-744. BECKMAN,D. W., ANDJ. M. DEAN. 1984. The age and growth of young-of-the-year spot, Leiostomus xanthurus Lacepede, in South Carolina. Estuaries 7:487-496. BROTHERS, E. B. 1978. Exogenous factors and the formation of daily and subdaily growth increments in fish otoliths. Am. Zool. 18:631. BROTHERS,E. B., C. P. MATHEWS,AND R. LASKER. 1976. Daily growth increments in otoliths from larval and adult fishes. Fish. Bull., U.S. 74:1-8. BROTHERS, E. B., D. McB. WILLIAMS, AND P. F. SALE. 1983. Length of larval life in twelve families of fishes at "One Tree Lagoon," Great Barrier Reef, Australia. Mar. Biol. 76:319324. CAMPANA, S. E. 1983a. Calcium deposition and otolith check formation during periods of stress in coho salmon, Oncorhynchus kisutch. Comp. Biochem. Physiol. 75A:215-220. CAMPANA,S. E. 1983b. Feeding periodicity and the production of daily growth increments in otoliths of steelhead trout (Salmo gairdneri) and starry flounder (Platichthys stellatus). Can. J. Zool. 61:1591-1597. CAMPANA, S. E., A. J. GAGNE, AND J. MUNRO. 1987. Otolith microstructure of larval herring (Clupea harengus): Image or reality? Can. J. Fish. Aquat. Sci. 44:1922-1929. CAMPANA, S. E., ANDJ. D. NEILSON. 1982. Daily growth increments in otoliths of starry flounder (Platichthys stellatus) and the influence of some environmental variables in their production. Can. J. Fish. Aquat. Sci. 39:937-942. CAMPANA, S. E., AND J. D. NEILSON. 1985. Microstructure of fish otoliths. Can. J. Fish. Aquat. Sci. 42:1014-1032. CHOATE, J. 1964. Use of tetracycline drugs to mark advanced fry and fingerling brook trout (Salvelinus fontinalis). Trans. Am. Fish. Soc. 93:309-311. HAAKE, P. W., C. A. WILSON, ANDJ. M. DEAN. 1982. A technique for the examination of otoliths by SEM with application to larval fish, p. 12-15. In C. F. Bryan, J. V. Conner, and F. M. Truesdale (eds.), Proc. Fifth Annual Larval Fish Conference. LSU Press, Baton Rouge, Louisiana. HETTLER, W. F. 1984. Marking otoliths by immersion of marine fish larvae in tetracycline. Trans. Am. Fish. Soc. 112:370373. JONES, C. 1986. Determining age of larval fish with the otolith increment technique. Fish. Bull., U.S. 84:91-104. JONES, C., AND E. B. BROTHERS. 1987. Validation of otolith increment aging technique for striped bass, Morone saxatilis, larvae reared under sub-optimal feeding conditions. Fish. Bull., U.S. 85:171-178.
Spot, Otolith Daily Increments, Validation MARSHALL, S. L., AND S. S. PARKER. 1982. Pattern identification in the microstructure of sockeye salmon (Oncorhynchus
nerka) otoliths. Can. J. Fish. Aquat. 8ei. 39:542-547. METHOT, R. D., JR. 1983. Seasonal variation in survival of larval northern anchovy, Engraulis mordax, estimated from age distribution of juveniles. Fish. Bull., U.8. 81:741-750. METHOT, R. D., JR., AND D. KRAMER. 1979. Growth of the northern anchovy, Engraulis mordax, larvae in the sea. Fish. Bull., U.S. 77:413-423. OXENFORD, H. A., AND W. HUNTE. 1983. Age and growth of dolphin Coryphaena hippurus, as determined by growth rings in otoliths. Fish. Bull., U.S. 81:906-909. STRUHSAKER,P., ANDJ. H. UCHIYAMA. 1976. Age and growth of the nehu, Stolephorus purpures (Pisces: Engraulidae), from the Hawaiian Islands as indicated by daily growth increments of sagittae. Fish. Bull., U.S. 74:9-17. TAUBERT,B. C., AND O. W. COBLE. 1977. Daily rings in otoliths of three species of Lepomis and Tilapia mossambica. J. Fish. Res. Board Can. 34:332-340.
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VAN OOSTEN,J. 1941. The age and growth of freshwater fishes, p. 196-205. In A Symposium on Hydrobiology. Univ. Wisconsin Press, Madison, Wisconsin. WARLAN, S. M. 1982. Age and growth of larvae and spawning time of Atlantic croaker in North Carolina, Proc. Annu. Conf. Southeast. Assoc. Fish Wildl. Agen. 34:204-214. WEINSTEIN,M. P., ANDM. P. W'ALTERS. 1981. Growth, survival, and production of Leiostomus xanthurus Lacepede residing in tidal creeks. Estuaries 4:185-197. WILD, A., AND T.J. FOREMAN. 1980. The relationship between otolith increments and time for the yellowfin and skipjack tuna marked with tetracycline. Inter-Am. Trop. Tuna Comm. Bull. 17:509-600.
Received for consideration, January 3, 1989 Accepted for publication, March 27, 1989
