Brown & Wooden—Golden perch daily growth-increments New Zealand Journal of Marine and Freshwater Research, 2007, Vol. 41: 157–161 0028–8330/07/4102–0157 © The Royal Society of New Zealand 2007
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Short communication
Age at first increment formation and validation of daily growth increments in golden perch (Macquaria ambigua: Percichthyidae) otoliths Paul Brown* Ian Wooden Narrandera Fisheries Centre NSW Department of Primary Industries P.O. Box 182, Narrandera NSW 2700, Australia * Present
address: Primary Industries Research Victoria (PIRVic), Department of Primary Industries, Private Bag 20, Alexandra, Victoria, 3714 Australia. email:
[email protected]. Abstract The micro-increments in otoliths from a series of golden perch (Macquaria ambigua) larvae of up to 15 days known age (n = 124, total length (TL) 4.14 to 10.0 mm) were examined and counted to establish the age at formation of the first increment, and periodicity of formation of subsequent increments. The regression of true age on increment count (true age = 3.54 +1.36 × increment count) was significant (r2 = 0.78, P < 0.001) and the slope of the overall regression (1.36 day–1) was not significantly different to 1 (t = 0.09, P = 0.05). First increment formation occurred from day 3 in some larvae, and was completed by day 13 in all larvae. A median estimate for age at first-increment deposition was 6.0 days, when 50% of the daily sample (n = 10 fish) had at least one increment. However, solving the regression for age when increment number equaled 1.0 provided an estimate of mean age at first-increment deposition of 4.9 days. Our study validated daily growth increment deposition for golden perch larvae over the first 15 days of development and cautions that age at first-increment formation varied within a single cohort of larvae by 10 days (i.e., youngest larvae 3 days, oldest larvae 13 days old).
M07002; Online publication date 27 April 2007 Received 9 January 2007; accepted 8 March 2007
Keywords growth; first increment; otolith; daily growth increment; validation; hatch-date Introduction By understanding the pattern and process of otolith creation we can infer details of the daily lives of fish from birth until death (Secor et al. 1991). Information about an individual fish’s growth-rates, the chemical composition of its ambient environment, and the precise timing of its age and development are all stored within a series of micro-incremental depositions (Campana & Neilson 1985; Kalish 1990). Studies of larval dynamics, spawning cues or environmental factors correlated with recruitment often use backcalculations of spawning, or hatching-date frequency distributions to “position” such events in time (Prince et al. 1991; Jordan 1994; Vilizzi 1998). To determine the timing of spawning events, the rate of formation of otolith micro-increments and the age at which the larvae deposit their first increment must both be known. The former is most accurately established with known-age fish (Campana 2001). The latter is usually reported as a single age, in days, with few estimates of its variability (Campana & Neilson 1985). The variability of first-increment formation can only be determined by examining larvae resulting from spawning events where the timing is known. Pragmatically, the potential uncertainty over timing of spawning in wild stocks often restricts observations to those made under experimental conditions. Studies to date have shown that under natural photoperiod, temperature and growth conditions juveniles of most fish species produce otolith microincrements at a rate of one per day (Secor & Dean 1989; Wright 1991; Jordan 1994). First increment formation is highly variable between species and can occur at or before hatching as in eastern rainbowfish (Humphrey et al. 2003); or within a day of hatching as in white crappies (Pomoxis annularis) (Sweatman & Kohler 1991), sticklebacks (Gasterosteus aculeatus) (Wright & Huntingford 1993), and
158 New Zealand Journal of Marine and Freshwater Research, 2007, Vol. 41 sandeels (Ammodytes marinus) (Wright 1993); or corresponding to an ontogenetic “landmark” such as the transition from endogenous to exogenous feeding as for jack mackerel (Trachurus declivis) (Jordan 1994) or herring (Clupea harengus) (Moksness 1992). Although golden perch (Macquaria ambigua) (Richardson, 1845) larvae have been collected in the wild (Brown & Neira 1998), quantitative relationships between environmental stimuli (e.g., stream flows, water temperature) and spawning events for wild stocks of golden perch have not yet been established. To do so is important in the context of habitat and flow-related rehabilitation strategies for management of the species. The lack of accurate information about environmental spawning “cues” is partly owing to uncertainty about the relationship between age and otolith micro-increment counts. Validation of daily deposition of otolith growth increments will enable accurate estimates of the age of wild-caught golden perch larvae, to be made with known precision. Determining accurate larval age estimates will permit a better understanding of the environmental conditions that may be correlated to spawning events.
solution (0.1 g litre–1) and measured (TL ± 0.01 mm) using the graduated reticule of a dissecting micro scope. Initially otoliths were identified and extracted from freshly killed larvae under a compound microscope (40× magnification) using transmitted light and a polarising filter. Sagittae were identifiable, and discernible from lapidae and asteriscae by their slightly larger size from day 0. Each pair of sagittae were mounted whole in a drop of DePX mounting medium and protected with a coverslip. No further preparation was required. The entire set of otoliths (n = 124) was read by the first reader then a sub-sample of 45 otoliths were read a second time by an independent reader as an evaluation of precision. Neither reader knew the true age or larval identity of otoliths being read. Analysis of variance (PROC GLM, SAS Cor poration) was used to compare the precision of ageestimates by each reader. Linear regression best described the relationship between increment-count and true age and the 95% prediction confidence estimates. A 2-tailed Student’s t test was used to test whether the slope estimate from the regression was significantly different to 1 (Zar 1974). A statistical significance level of a = 5% was assumed throughout.
Materials and Methods Artificial propagation techniques (after Rowland 1983) produced a batch of eggs from a single female golden perch, which were incubated and hatched in indoor tanks under reduced natural light conditions (i.e., room contained large windows and no blinds) (30 October – 5 November 1991). A single batch of eggs was used to reduce variability in timing of fertilisation and subsequent development. Water temperatures ranged naturally 24 to 26°C. Hatching took place over c. 24 h. Nominally, samples of 10 larvae were taken daily from the day when hatching started (day = 1) until day 15 (excluding days 10 and 11 when no samples were taken). On six sampling days the final sample size was 9 owing to a failure in one of the preparation stages, resulting in a total sample size of n = 124. Swim bladder inflation started on day 5. Food was offered at this point and some larvae were observed feeding on Artemia nauplii by day 6. The cohort of larvae was stocked into an outdoor rearing pond on day 6 and experienced abundant plankton densities (P. Brown pers. obs.), and natural fluctuations in ambient light and temperature from this day. Larvae were killed by immersion in benzocaine
Results and Discussion First-increment formation began on day 3, 3 days before any feeding on Artemia or natural pondplankton. From day 3, the proportion of each daily sample with at least one increment increased until 50% of the sample had at least one increment by day 6 (Fig. 1). Thus, day 6 was the median age of first increment formation and coincided with transfer of the larvae to an outdoor pond and a gradual increase in increment width. Apart from a single 13-day-old larva with no discernable increments, after day 9 all fish sampled had at least one increment. Comparison of the precision of age-estimates by each reader showed no significant difference (F = 221, d.f. = 43, P > 0.001) between readers. Therefore data from the full sample of 124 otolith age estimates by the first reader were used in regression analysis. Linear regression of the increment-count on true age (Fig. 2) (n = 124) provided a slope estimate of 1.36 increments per day and a predicted intercept of 3.54 days (age = 3.54+1.36 × increment count) (r2 = 0.78, P < 0.001). Biologically, the intercept is the mean age of larvae with no increments. The
Brown & Wooden—Golden perch daily growth-increments
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Fig. 1 Proportion of daily sample of Macquaria ambigua larvae showing a first sagittal micro-increment. Arrow shows median age at first-increment formation. No increments observed on days 1 and 2, and no samples collected on day 10 and 11. Sample sizes in parentheses.
slope estimate was not significantly different to 1 (t = 0.09, d.f. = 122, P = 0.05). The mean age at firstincrement formation was obtained by solving the regression for age when increment count equals 1.0, which provided an estimated age at first-increment formation of 4.9 days (c. 5 whole days). The 95% prediction confidence interval on the regression (Fig. 2) shows that where a single increment was observed, the estimated true age in whole days was 5 days, although we can be 95% confident that the true age lies only between 1 and 8 days. This study has shown that in golden perch larvae under natural environmental conditions of daylength and temperature, otolith micro-increments are deposited with daily periodicity. As the progeny of a single female was examined, no attempt was made to determine how increment formation may vary across the population. Most daily-increment age validation studies either use wild caught larvae that are the offspring of many parents (Moksness 1992; Jordan 1994), or use captive populations potentially with multiple parents (Humphrey et al. 2003). In doing so, these authors’ assumptions about the applicability of their results to the population may be more robust. However, in the present study, time to first increment formation varied considerably for individuals within a larval cohort. It seems likely that this variation would increase if offspring of multiple parents were examined; however, further work is needed to determine parental influence on timing of larval first increment formation. The observed 10-day variation in age of first-increment formation among sibling individuals undergoing identical environmental conditions has not been reported previously in the literature. Many studies report the age at first-increment formation with no indication of its variability (Moksness 1992; Jordan 1994; Vilizzi 1998). Studies relying on daily growth increment
Fig. 2 Linear regression between true age and otolith micro-increment count (r2 = 0.78, P < 0.001) for golden perch, Macquaria ambigua larvae (n = 124, age = 3.54 + 1.36 × count). Bars indicate 1 SE, dashed lines 95% prediction CI. The slope of the regression (1.36 increments per day) was not significantly different to 1.
counts for back-calculation need to account for this variation and its effect on the precision of spawning or hatching-date estimates. Golden perch larvae are typical of a pelagic spawning species (Brown & Neira 1998). They have a brief yolk-sac stage, and the estimated median
160 New Zealand Journal of Marine and Freshwater Research, 2007, Vol. 41 age at first-increment deposition coincided with gas bladder inflation and first-feed as reported for Norwegian herring (C. harengus) and jack mackerel (T. declivis) (Moksness 1992; Jordan 1994). Similarly, in striped bass (Morone saxatilis) (Secor & Dean 1989) the first increment is deposited on day 4–5 which also corresponds to the onset of exogenous feeding and the start of gas bladder inflation (Hadley et al. 1987). The temperature range in the present study was close to that noted by Lake (1967), as necessary to induce spawning (>24°C). It is unlikely that the minor temperature fluctuations (24–26°C) observed in the early part of the present study played a significant part in the variation in timing of firstincrement formation observed. Houde & Morin (1990) showed that for striped bass (M. saxatalis) and white perch (Morone americanus), the age when the first increment formed varied linearly with water temperature. For these species, firstincrements formed from 2 to 5 days after hatching over a range of 15–21°C. The temperatures which yolk-sac M. ambigua larvae were exposed to during this study are typical of those which wild populations encounter (Mackay et al. 1988). However, given the broad geographical range of the species (22° to 37° latitude), and that spawning may occur over a period of several months, e.g., December–March (Lake 1967; Mackay 1973), considerable variation in temperature may be encountered by golden perch larvae sampled from the wild. We cannot be certain whether the variability in increment counts for each known age reflects some reader-inaccuracy (i.e., measurement error) alone, or is a combination of measurement-error and true variation in periodicity of increment formation. To establish a useable age estimate for back-calculated spawning-date estimates of wildcaught golden perch larvae, either the median age (6 days) or mean age (5 days) of first increment formation are reasonable values. However, owing to variation of first-increment formation within a larval cohort, a precision estimate of ±4 days is realistic for back-calculated estimates of hatch-date for golden perch. To optimise back-calculations of hatch-date for particular golden perch stocks, further studies should investigate the relationship between known-age and increment count in older juveniles. Juveniles older than 15 days are more easily and often sampled in the field (Brown & Neira 1998), therefore this age-validation study could be extended to include older juveniles. Evaluation of how first increment formation varies with the broad range of
ambient water temperatures that are anticipated over the species’ geographic range, would further extend the applicability of this study. References Brown P, Neira F 1998. Percichthyidae: basses, perches and cods. In: Neira F, Misciewicz A, Trnski T ed. Larvae of temperate Australian fishes. Nedlands, Western Australia, University of Western Australia Press. Pp. 259–265. Campana SE, Neilson JD 1985. Microstructure of fish otoliths. Canadian Journal of Fisheries and Aquatic Sciences 42: 1014–1032. Campana SE 2001. Accuracy, precision and quality control in age determination, including a review of the use and abuse of age validation methods. Journal of Fish Biology 59: 197–242. Hadley CG, Rust MB, Van Eenennaam JP, Doroshov SI 1987. Factors influencing initial swim bladder inflation by striped bass. American Fisheries Society Symposium 2: 164–169. Houde ED, Morin LG 1990. Temperature effects on otolith daily increment deposition in Striped Bass and white perch larvae. Collected papers of the council meeting of the International Council for the Exploration of the Sea; 4–12 October 1990; Copenhagen, Denmark. ICES. 19 p. Humphrey C, Klumpp D, Pearson R 2003. Early development and growth of the eastern rainbowfish, Melanotaenia splendida splendida (Peters) II. Otolith development, increment validation and larval growth. Marine and Freshwater Research 54: 105–111. Jordan AR 1994. Age, growth and back-calculated birthdate distributions of larval jack mackerel, Trachurus declivis (Pisces: Carangidae), from Eastern Tasmanian coastal waters. Australian Journal of Marine and Freshwater Research 45: 19–33. Kalish JM 1990. Use of otolith microchemistry to distinguish the progeny of sympatric anadromous and non-anadromous salmonids. Fishery Bulletin 88(4): 657–666. Lake JS 1967. Rearing experiments with five species of Australian freshwater fishes I: Inducements to spawning. Australian Journal of Marine and Freshwater Research 18: 137–153. Mackay N, Hillman T, Rolls J 1988. Water quality of the River Murray. Canberra: Murray Darling Basin Commission. 62 p. Mackay NJ 1973. Histological changes in the ovaries of the golden perch, Plectroplites ambiguus, associated with the reproductive cycle. Australian Journal of Marine and Freshwater Research 24: 95–101.
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Sweatman JJ, Kohler CC 1991. Validation of daily otolith increments for young-of-the-year white crappies. North American Journal of Fisheries Management 11(4): 499–503. Vilizzi L 1998. Age, growth and cohort composition of 0+ carp in the River Murray, Australia. Journal of Fish Biology 52: 997–1013. Wright PJ 1991. The influence of metabolic rate on otolith increment width in Atlantic Salmon parr, Salmo salar L. Journal of Fish Biology 38: 929–933. Wright PJ 1993. Otolith microstructure of the lesser sandeel, Ammodytes marinus. Journal of the Marine Biological Association of the United Kingdom 73(1): 245–248. Wright PJ, Huntingford FA 1993. Daily growth increments in the otoliths of the three spined stickleback, Gasterosteus aculeatus L. Journal of Fish Biology 42: 65–77. Zar JH 1974. Biostatistical analysis. New Jersey, PrenticeHall. 620 p.
