Running Title: Tripletail Reproductive Status

Evaluation of reproductive status for tripletail, Lobotes surinamensis, nearshore Jekyll Island,

GA, USA1

R.T. Parr2

University of Georgia Warnell School of Forestry and Natural ResourcesAthens, GA 30602

C.A. JenningsUS Geological SurveyGeorgia Cooperative Fish and Wildlife Research UnitWarnell School of Forestry and Natural ResourcesUniversity of GeorgiaAthens, GA 30302

N. D. DenslowUniversity of FloridaDepartment of Physiological Sciences and Center for Environmental and Human ToxicologyGainesville, FL 32610

1 This draft manuscript is distributed solely for purposes of scientific peer review. Its content is deliberative and predecisional, so it must not be disclosed or released by reviewers. Because the manuscript has not yet been approved for publication by the U.S. Geological Survey (USGS), it does not represent any official USGS finding or policy.2 Current Address: 41 Park of Commerce Way Suite 303, Savannah, Georgia 31405

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K. J. KrollUniversity of FloridaDepartment of Physiological Sciences and Center for Environmental and Human ToxicologyGainesville, FL 32610

R.B. Bringolf3

University of Georgia Warnell School of Forestry and Natural ResourcesAthens, GA 30602

3 Author to whom correspondence should be addressed: Tel.: 1-706-542-1477; email: bringo@uga.edu

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Evaluation of reproductive status for tripletail Lobotes surinamensis nearshore Jekyll Island, GA,

USA

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Abstract

Tripletail (Lobotes surinamensis) aggregate nearshore off the Coast of Jekyll Island, Georgia,

United States from March through July. Why tripletail aggregate to these nearshore waters is

unknown, however anecdotal evidence has suggested the possibility of a spawning area. The

objective of the study was to evaluate the reproductive status of tripletail with a non-lethal

sampling method, plasma vitellogenin (VTG) analysis, concurrently with traditional lethal

sampling methodologies, gonadosomatic index (GSI) and histology. A total of 224 (122 males

and 102 females) tripletail were sampled for reproductive evaluation. Most of the male tripletail

(115 of 122) were in the spawning-capable reproductive phase. Female tripletail were found in

all reproductive phases, with few (10 of 102) in the spawning capable phase. The estimated

length at which 50% (L50) of females reached maturity was 459 mm at an approximate age of

1.17 years. The L50 for male tripletail could not be determined because of the lack of immature

fish within the study sample; however, the age at 50% maturity was estimated at 0.55 years.

Gonadosomatic index ranks for females were significantly higher than males (P <0.0001), as

well as among reproductive phases within females (P <0.0001). Females had significantly

higher VTG levels than males (P = 0.0006). Vitellogenin concentrations in spawning capable

females were higher than all phases except the developing phase. Linear regression demonstrated

a strong relationship (r2 = 0.873, n= 77) between VTG and GSI in females. The study indicates

that VTG levels in developing and spawning capable females may be distinguished from other

female reproductive phases and male tripletail; however, a greater sample size of spawning

capable females will be required to further evaluate the use of VTG as a non-lethal reproductive

status indicator.

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Keywords: fish, reproductive assessment, non-lethal monitoring, GSI

INTRODUCTION

Tripletail Lobotes surinamensis (Bloch 1790) are medium-sized, deep-bodied fish that

occur in tropical and subtropical waters worldwide (Baughman 1941; Hardy 1978). The species

is the only member from the monophyletic family Lobotidae, which is found in the western

Atlantic Ocean (Hardy 1978). Tripletail are known to occur from Massachusetts, USA to

Argentina, including the Gulf of Mexico and the Caribbean Sea, with higher abundances south of

North Carolina, USA (Hardy 1978). Tripletail are migratory; however, detailed information

about their exact movement patterns is scarce or lacking (Merriner and Foster 1974; Franks et

al.1998; Streich et al. 2013).

Tripletail growth appears to be rapid during early life stages; wild-caught and captive fish

have demonstrated the ability to grow greater than 500 mm total length at age-1 (Armstrong et

al. 1996; Franks et al. 1998; Strelcheck et al. 2004; Parr 2011). Scale-based total length (TL) at

age of tripletail from North Carolina was: age-0 (n=1) 190 mm, age-1 (n=6) 445- 591 mm, age-2

(n=5) 562-706 mm, and age-3 (n=2) 568-706 mm (Merriner and Foster 1974). Spine-derived

ages of tripletail also show similar results (Franks et al. 1998; Strelcheck et al. 2004).

Peer-reviewed literature with tripletail reproductive data is scarce, and much of the

available information remains in unpublished agency reports. The size at which 50% of males

reach maturity has not been determined because of a lack of data for age-0 and immature males;

however, Brown-Peterson and Franks (2001) estimated that 50% of males reach maturity by at

least 290 mm. Estimates for female age-at-sexual maturity range from one to two years and

lengths from 350 – 500 mm TL depending upon criteria used for determination of maturity status

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(Brown-Peterson and Franks 2001; Strelcheck et al. 2004). Brown-Peterson and Franks (2001)

and Strelcheck et al. (2004) estimate that 50% of females reach maturity at approximately one

year and approximately 490 mm TL. Strelcheck et al. (2004) note that size at 50% maturity for

females could change significantly if the presence of vitellogenic oocytes is used as the gauge for

maturity rather than presence of oocytes in the cortical alveolar stage.

Tripletail appear to be protracted multiple-batch spawners, with the ability to spawn once

every three to five days during the spawning season (Brown-Peterson and Franks 2001; Cooper

2002). Baughman (1941) documented gravid females during the months of July and August on

the Atlantic Coast, but other anecdotal reports of gravid females have not been

confirmed(Gudger, 1931, Baughman, 1941). In the Gulf of Mexico, running-ripe males have

been captured from May through September, and females in late ovarian maturation phases have

been found from June through August (Brown Peterson and Franks 2001; Strelcheck et al. 2004).

Ditty and Shaw (1994) captured larval tripletail in >100 m of water in plankton surface tows and

suggested that tripletail may use offshore spawning. Juvenile tripletail also have been captured in

offshore waters in the Gulf of Mexico in association with sargassum patches (Franks et al. 2001).

Histological analysis can yield accurate information on oocyte development and

reproductive phase; however, this analysis requires more time and expense compared to visual

staging based on macroscopic appearance of the gonads or calculation of gonadal somatic index

(GSI) values (West 1990). However, all of the aforementioned techniques require sacrifice of

the fish, and non-lethal alternatives for determining reproductive status are desired for species

such as tripletail, for which definitive population dynamics data are scarce or lacking completely.

Non-lethal methods have been used to determine reproductive status of other fish species

and often include evaluation of blood plasma for the egg yolk precursor, vitellogenin (VTG;

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Heppell and Sullivan 2000; Ceapa et al. 2002; Wildhaber et al. 2007; Heise et al. 2009).

Vitellogenin is found at highest concentrations during the final oocyte maturation phase in most

fishes. Therefore, VTG can be an effective tool for determining sex, and in some cases,

reproductive phase -- although increased VTG levels are not always indicative of spawning

location (Heppell and Sullivan 2000; Ceapa et al. 2002; Wildhaber et al. 2007; Heise et al. 2009).

In Georgia (GA, USA), tripletail are found associated with structure in estuaries and

nearshore of some of the barrier islands. Tripletail are also found free-floating in the nearshore

Atlantic Ocean waters immediately east of Jekyll Island from March to July (Figure 1). Some

investigators (Brown-Peterson and Franks 2001; Cooper 2002; Strelcheck et al. 2004) have

suggested that this period coincides with the spawning season for tripletail. Atypically, many of

the fish near Jekyll Island are not associated with any structure, but rather are free swimming

near the surface of waters ranging from 2 to 4 meters deep. As tripletail angling pressure

increases in GA and elsewhere, additional information about their life history such as age,

growth, and reproduction data, is needed to inform fisheries managers to adequately protect this

species.

The lack of published tripletail data, particularly for the western Atlantic Ocean,

underscores the general scarcity of information on the life history of the species. The goal of the

current study was to describe reproductive characteristics of a nearshore tripletail aggregation in

the western Atlantic Ocean, specifically fish found near Jekyll Island, GA, USA. Specific

objectives were to compare results of non-lethal techniques with traditional lethal methods to

determine the gender and reproductive status of tripletail and to determine size at maturity to

inform fishery management actions. The current study represents the first attempt to evaluate

tripletail plasma VTG levels as a non-lethal method to determine gender and reproductive phase.

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MATERIALS AND METHODS

Study Site

Tripletail sampling was conducted from March 30 to August 10, 2009 and March 14 -

August 6, 2010 in the Atlantic Ocean nearshore Jekyll Island, GA, USA. Tripletail were sampled

primarily around the northeast to central part of the island and on channel markers, range

markers, buoys, and other structures within St. Simons Sound, north and west of Jekyll Island

(Figure 1).

Field Sampling

This project was conducted under the auspices of the University of Georgia Animal Use

Protocol #A2009 02-030-R2. Sampling for tripletail and associated water quality variables was

performed on multiple dates during April to August during 2009 and 2010. On each sampling

trip, surface water temperature (ºC), salinity (ppt), and dissolved oxygen (mg/L) were measured

with an YSI 85 dissolved oxygen meter and recorded upon arrival at the sample site (Figure 1).

Hook-and-line methods from an open-cockpit fiberglass boat were used to sample tripletail. For

near shore sampling, the study area was searched visually for tripletail near the surface; when a

tripletail was spotted, they were directly targeted fish by casting a live white shrimp Litopenaeus

setiferus (Linneaus, 1758) or brown shrimp Farfantepenaeus aztecus (Ives, 1891)) or striped

mullet Mugil cephalus (Linneaus, 1758) with a spinning rod equipped with braided line (14 kg

test) attached to a popping cork rig with a Kahle™ live bait hook (size 1). Total numbers of

tripletail observed and captured were recorded and duration of the sampling event was recorded

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to the nearest minute. The capture location for each fish was recorded with a global positioning

system (GPS).

Tripletail sampling also occurred around structures in the St. Simons Sound and adjacent

shipping channel. We used hook-and-line sampling methods to sample structures but with

different tactics than described for near shore (open water) sampling. We sampled approximately

two hours prior to, and post, slack tide. Heavier tackle was necessary to prevent line breakage

around the structure; we used a heavy-action spinning rod equipped with braided line (36.2 kg

test) with a slip float rig, a monofilament leader (36.2 kg), and a 7-g jig head hook baited with

live penaeid shrimp or striped mullet. We approached the structure by boat and fished the bait

throughout the water column next to the structure. Upon capture of a fish, data gathering

procedures were identical to the aforementioned sampling methodology. In 2009, we sampled

opportunistically on inshore structures when weather did not permit sampling nearshore. In 2010,

we began sampling the nearshore area and structures (in St. Simons sound) at a 50:50 ratio by

sampling day and allowed weekly catch success to determine where our effort was best allocated

the following week.

In addition to sampling with hook-and-line gear nearshore Jekyll Island, tripletail from

the aggregation were also sampled with a 12.2-meter fish trawl with 76.2-mm mesh (bar)

deployed from the GA-DNR Research Vessel (R/V) Anna. Vessel location (latitude and

longitude was determined with GPS), and heading of the R/V Anna was recorded at the

beginning of each tow, with trawl net--in, and at the end of each tow, with trawl net--out. Speed

was determined based on the amount of time required to get from the trawl net in GPS point to

the trawl net out GPS point. A maximum tow time of 10 minutes was used to minimize the risk

of lethal interactions with tripletail and other non-target species. The number of tripletail caught

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was recorded, and procedures for handling the fish were identical to nearshore sampling

methodology by hook and line. Tripletail were also sampled opportunistically from local

tournaments and recreational anglers. As a result of the differing condition of the donations (i.e.,

carcass, fresh dead fish, live fish), not all fish sampled were included in all statistical analyses.

The sampling target was a maximum of 10 fish ≤ 610 mm per week for histological

evaluation: five individuals <457 mm total length (TL) and five individuals ≥ 457 mm (TL). The

457-mm (18 inch) designation represents the minimum size limit for possession of tripletail in

Georgia, established by the Georgia Department of Natural Resources (GADNR). Based on the

literature, all tripletail ≥ 610 mm were assumed to be mature fish and therefore were sacrificed.

Any captured fish that exceeded the target number were measured to the nearest millimeter for

TL and standard length (SL), tagged with a uniquely numbered Hallprint™ PDS plastic dart tag

inserted behind the base of the third dorsal spine, and released.

Upon capture of tripletail within the sampling target size range, blood samples were

immediately obtained from the caudal vein with a heparinized 3-mL syringe equipped with a

Luer-lok 18-guage hypodermic needle. The blood samples were carefully expelled into

numbered 1.0-mL centrifuge tubes and immediately placed on ice. To minimize degradation of

VTG, blood samples collected in 2009 contained the protease inhibitor aprotinin, and 2010

samples were treated with a protease inhibitor cocktail (P2714, Sigma-Aldrich, St. Louis, MO,

USA). Specimens were then placed on ice and returned to the lab within six hours of collection.

Once blood samples were collected, fish were marked with a unique tag affixed to the fish

through their mouth and out of their operculum via a zip tie and placed on ice until processing

when researchers returned to the lab at the GADNR Coastal Regional Division (CRD)

headquarters in Brunswick, GA.

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At the CRD lab, plasma was separated from blood cells by centrifugation (1500 x g) for

10 min and was then extracted with a micropipette, placed into a cryogenic vial and frozen in

liquid nitrogen. Samples were later transferred from liquid nitrogen to an ultracold (-80ºC)

freezer until VTG analysis. Sacrificed tripletail were measured for TL and SL (nearest mm) and

weighed (nearest 1.0 g) with an electronic platform scale (max 20 kg; Northern Industrial R-

2553). Gonads were dissected from the fish, weighed (nearest 0.1g), and preserved in 10%

buffered formalin until further processing for histological analysis.

Total length and total weight (TW) data were examined for normality with the Shapiro-

Wilk test. These data were not normally distributed; therefore, they were log10 transformed to

achieve normality. Levene’s test was used to evaluate homogeneity of variances. A 2 X 2

factorial analysis of variance (ANOVA) was used to determine if there were significant

differences among TL and TW of samples collected in 2009 and 2010, sexes, or the interaction

between capture year and sex. Data were pooled if the relationships were not significantly

different. A chi-square analysis was used to determine if monthly sex ratios varied from the

expected 50:50 ratio (α = 0.05). March samples were excluded from the chi-square analysis

based on insufficient data (< 5 samples) during this month.

Gonad Histology

A subsample of 15 males and 15 females was used to determine if development within

the gonads was uniform throughout the length of the gonad. Gonads were removed from 10%

buffered formalin and three sections (i.e., one each from the posterior, central, and anterior

portions of the gonads) were placed in tissue cassettes. Standard histological procedures

including dehydration, embedding in paraffin, thin sectioning (4 um), staining with hematoxylin,

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and counterstaining with eosin, were performed at the University of Georgia Veterinary

Medicine Diagnostic Lab. Development of gametes was confirmed to be uniform throughout the

gonad and all subsequent gonad samples were collected only from the central portion of the

gonad. Reproductive phases for both males and females were evaluated based on criteria

described by Brown-Peterson et al. (2011) by examination of the mounted and stained gonad

sections with a compound microscope. Females were classified as immature, developing (sub-

phase: early developing), spawning capable (sub-phase: actively spawning), regressing or

regenerating. Immature females contained only oogonia and primary growth (PG) oocytes.

Females entered the developing phase with the appearance of cortical alveoli (CA) oocytes.

Primary vitellogenic and secondary vitellogenic oocytes are also present during the developing

reproductive phase. Primary, secondary, and tertiary vitellogenic oocytes are determined based

on the relative increase in yolk deposition (Lowerre-Barbieri et al. 2011). Males were classified

as immature, developing (sub-phase: early developing), spawning capable (sub-phase: early

germinal epithelium (GE), mid-GE, and late-GE), regressing or regenerating (Brown-Peterson et

al. 2011). A second reader evaluated the reproductive status of a randomly selected subsample (n

= 25) for both males and females for quality assurance and control.

Length at 50% Maturity

Length at which 50% of individuals reached maturity for male and female tripletail was

evaluated for TL and age, based on the equation described by Goncalves and Erzini (2000):

Pi=1

1+e−b¿¿¿

where Pi represents the proportion of mature adults in each 50-mm length bin, b represents the

slope of the maturity curve, L50 is the length at which 50% of the fish are mature, and length or

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age class at 50% maturity (Li ). Ages for fish in this study were based on a concurrent tripletail

aging project (Parr 2011). All male and female tripletail that were not in the immature

reproductive phase were classified as mature for determination of length at 50% maturity. Non-

linear regression (PROC NLIN) was used to model the parameters for the length at 50% maturity

(SAS 9.1; Freund and Littell 1991).

Gonadosomatic Index

Gonadosomatic index (GSI) values were calculated based on the equation originally

described by Nikolsky (1963):

I g=(W g

W t)× 100

where, Ig represents the percentage of gonad weight to total body weight, Wg represents gonad

weight, and Wt represents total weight of the fish. Gonadosomatic index values were evaluated

for normality with the Shapiro-Wilk test. Data were not normally distributed and could not be

transformed to fit a normal distribution; therefore, the GSI values were ranked for further

(nonparametric) statistical analyses. As a result of the relatively small, annual sample sizes, data

were combined from the two years of the study. Sex, reproductive phase, and capture month

variables were analyzed individually with one-way ANOVA to evaluate their relationships with

the GSI values (α = 0.05). Small sample sizes in some months and reproductive phases precluded

the use of a factorial ANOVA design to evaluate the interaction and effects of month and

reproductive phase on GSI (Appendix A). Trends were examined for GSI values across the

reproductive phases and capture months.

Plasma Vitellogenin

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Tripletail plasma samples were shipped on ice via overnight express courier to the

University of Florida for VTG analysis. Vitellogenin was purified from plasma of estrogen

treated males by anion exchange chromatography with the BIOCAD Sprint Perfusion system

(Perseptive Biosystems) as described by Denslow et al. (1999). Briefly, plasma was diluted in

running buffer (10mM bis-tris propane, 50 mM NaCl, pH 9.0) and loaded onto a strong anion

exchange resin (POROS 20 HQ). Vitellogenin was eluted from the column with a linear gradient

of NaCl (50→1000 mM). The VTG fractions were pooled, pH adjusted to 7.0, and were then

concentrated with a 30,000 MWCO Centricon (Amicon) filter. Protease inhibitor, (Aprotinin, 10

KIU/ml), bacteriocide (azide, 0.02%), and cryprotectant (glycerol, 1:2) were added to the

purified VTG. The standard stored in this form is stable for 1-2 years (-20°C) and circumvents

freeze/thaw protein fracture.

Concentrations of plasma VTG were determined by direct enzyme-linked immunosorbent

assay (ELISA) with a monoclonal antibody, 3G2 (HL1393), that was developed for striped bass

VTG. Plasma samples were diluted 1:100, 1:10,000, and 1:100,000 with 10 mM phosphate, 150

mM NaCl, 0.02% azide, 10 KIU/mL Aprotinin, pH 7.6 (PBSZ-AP). Tripletail VTG standards (0,

0.005, 0.01, 0.02, 0.04, 0.06, 0.08, 0.1, 0.2, 0.4, 0.6, 0.8, 1.0 µg/mL) containing 1:100, 1:10,000

and 1:100,000 male plasma (in PBSZ-AP) were used to make a species-specific standard curve.

Male plasma was added to the standards to account for matrix effect found in direct ELISAs.

Samples and standards were loaded onto a 96-well ELISA plate in triplicate and stored overnight

at 4 ºC in a humidified Tupperware® container. The following day the plates were washed four

times with phosphate buffered saline solution and were then blocked with 1% bovine serum

albumin in 10 mM Tris, 150 mM NaCl, 0.05% Tween, 0.02% azide, 10 KIU/mL Aprotinin, pH

7.6 (1% BSA/TBSTZ-AP) for 2 hr at room temperature. The plates were rewashed with PBSZ (4

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times) and the 1° monoclonal antibody, 3G2, was loaded into wells on each plate. The lowest

dilution (1:100) was probed with 1µg/mL of the monoclonal antibody and dilution > 1:10,000

with 0.1 µg/mL. After the addition of the monoclonal antibody, the plates were stored at 4 ºC

overnight in the humidified container. The following day the plates were washed and the

biotinylated secondary antibody, goat anti-mouse IgG-biotin (Pierce) was added to each well at

1:1,000 dilution in 1% BSA/TBSTZ-AP and incubated at room temperature for 2 hr. The plates

were washed and a strepavidin-alkaline phosphatase conjugate (Pierce) was added at 1:1,000

dilution in 1% BSA/TBSTZ-AP and incubated for 2 hr at room temperature. After a final wash

of the plates, the color was developed by adding 1 mg/ml p-nitro-phenyl phosphate in carbonate

buffer (30 mM carbonate, 2 mM MgCl2, pH 9.6) and the color was measured with an ELISA

plate reader (SpectraMax Plus384, Applied Biosystems) at 405 nm. Concentrations of the

unknowns were determined from the standard curves by using SoftMax® Pro analysis program.

The limit of detection for tripletail VTG was 1ug/mL. All assays were performed in triplicate

and reported as the mean of the three measurements. The coefficient of variation was <10% for

all samples analyzed. Inter- and intra-assay variability were measured by analyzing positive

controls on several plates and were <10%, and <5%, respectively.

Plasma VTG data was evaluated for normality with the Shapiro-wilk test and evaluated

for equality of variances with Levene’s test. Results of these tests indicated the data were not

normally distributed and their variances were unequal; therefore, VTG concentration levels were

ranked to facilitate further (nonparametric) statistical evaluation with a Kruskal-Wallis test. A t-

test was used to evaluate differences between male and female VTG ranks. Similar to GSI data,

VTG data from the two years were combined because of the small sample size. A one-way

ANOVA was performed for each sex to evaluate differences between the VTG ranks between

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reproductive phases and capture month. As with GSI evaluations, some capture months and

reproductive phases did not contain a sufficient minimum sample size and restricted the use of a

factorial design. Linear regression was used to evaluate relationships between VTG and GSI for

males and females.

All statistical analyses were conducted with program SAS 9.1(SAS Institute, Inc; Cary,

NC). Duncan's multiple range test was used to evaluate significant differences among means for

all one-way ANOVAs (α = 0.05).

RESULTS

Field Sampling

The project included 177 sampling events with a total of 385 hours of ‘near shore’

sampling, 95 hours of ‘structure’ sampling, and 9 hours of ‘trawl’ sampling. Average water

temperature was 25.6 ºC, with a range of 13.7 – 33.0 ºC. Average salinity was 30.5 ppt, with a

range of 14.5 to 38.7 ppt. Dissolved oxygen averaged of 6.18 mg/L and ranged from 3.34 – 9.02

mg/L. A total of 432 tripletail were captured during the study, and 224 (122 males and 102

females) were sampled for reproductive evaluation. The remaining 208 fish were tagged and

released. Tripletail TL ranged from 241 to 822 mm and TW ranged from 265 to 14,152 g.

Differences in mean TL (F1,216 = 0.87, P = 0.3517) and TW (F1,216 = 0.89, P = 0.3476) were not

significant between the years; therefore, the data for both years were pooled. Mean female TL

(489 mm) was not significantly different from mean male TL (455 mm) (ANOVA: F1,216 = 3.85,

P = 0.0512). Mean female TW (3,116 g) was significantly heavier than mean male TW (2,314 g)

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(ANOVA: F1,216 = 4.96, P = 0.00270). Ratio of males and females was not significantly different

than the expected 1:1 ratio (X2 = 2.3937, P = 0.0828).

Gonad Histology

Nearly all (115 of 122, 94%) of the male tripletail captured were in the spawning-capable

phase (Table I). Only one male (0.8 %) was in an immature reproductive phase. Female tripletail

were captured in all reproductive phases (Table II), but only two (2.0 %) of the 102 females were

classified in the ‘actively spawning’ sub-phase of the spawning-capable category. One actively

spawning female captured in June 2009 was 615 mm and contained oocytes that were

undergoing lipid coalescence, which is indicative of final oocyte maturation. The other actively

spawning female was 355 mm and was captured in April 2010; the ovaries contained hydrated

oocytes and post-ovulatory follicles, which indicated ovulation had occurred within 48 hours.

Length at Maturity

The estimated length at which 50% of females reached maturity (L50) was 459 mm with a

slope (b) of 0.02, and the age at which 50% of females reached maturity was estimated at 1.17

years with a slope of 4.21 (Figures 2 and 3). Length at which 50% of males reached maturity

could not be estimated by regression techniques because of a lack of convergence caused by the

lack of immature fish in the sample (Figure 2); however, the age at which 50% of males became

mature was 0.55 years with a slope of 9.74 (Figure 3).

Gonadosomatic Index

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Gonadosomatic index values for female tripletail (n = 101) ranged from 0.07 to 10.9%

with a mean of 1.1% (Figure 4). Male (n = 116) GSI values ranged from 0.04 to 0.22% with a

mean of 0.11% (Figure 4). Females had significantly higher GSI ranks than males (T-test, DF =

210, t-value = -3.518, P <0.0001), and female GSI values in March were significantly higher

than May and June values (ANOVA: F5, 95 = 2.98, P = 0.0153). Significant differences were not

detected for male GSI ranks among sample months (ANOVA: F4,111 = 1.51, P = 0.2040)

Female GSI values were significantly different among reproductive phases (ANOVA:

F4,96 = 25.21, P = <0.0001). Gonadosomatic Index values for females in the immature

reproductive phase were lower than all other phases, and values for the regenerating phase were

higher than the immature phase and lower than all other reproductive phases. Differences were

not detected among the developing, spawning-capable, and regressing female reproductive

phases but were higher than all other reproductive phases. Significant differences were not

detected among male reproductive phases for GSI values (ANOVA: F3,111 = 1.89, P = 0.1357).

Vitellogenin

Plasma VTG concentrations for female tripletail (n = 77) ranged from below detection

(1.0 µg/ml) to 4000 µg/ml with a mean of 209 µg/ml and a standard deviation of 696 µg/ml

(Figures 5 and 6). Male tripletail (n = 98) VTG levels ranged from below detection to a

maximum of 170 μg/ml with a mean of 36 μg/ml and a standard deviation of 28 μg/ml (Figures 5

and 6). A t-test indicated that females had significantly higher VTG concentrations than males

(DF = 173, t-value = -3.49, P = 0.0006). Vitellogenin did not differ among months for males

(ANOVA: F4,92 = 0.64, P = 0.6351). Significant differences were detected among months for

female VTG concentrations (ANOVA: F4,73 = 3.33, P = 0.0146). May female VTG concentration

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was lower than April and August but was not different from June and July. Mean VTG in

immature females was lower than all other phases except the regenerating phase (ANOVA: F4,73

= 9.09, P < 0.0001). Differences were not detected among females in developing, regressing, and

regenerating phases. Mean VTG concentration in spawning-capable females was higher than all

phases except the developing phase. Vitellogenin concentrations did not differ among male

reproductive phases (ANOVA: F3,93 = 0.24, P = 0.8684).

Plasma VTG and GSI values showed similar trends across the reproductive phases for

males and females (Figure 7). Male tripletail did not exhibit any peaks in GSI or VTG during our

sampling period, and linear regression analysis of VTG and GSI for males showed a very weak

relationship (r2=0.186; n = 94). Female tripletail in the spawning capable phase increased in

VTG and GSI values, but VTG and GSI across all other reproductive phases were similar. Linear

regression demonstrated a strong relationship (r2 = 0.873, n= 77) between VTG and GSI in

females.

DISCUSSION

Male tripletail captured near Jekyll Island, GA from March to August were in a

spawning-capable reproductive phase; however, most female tripletail in this aggregation were

not in spawning condition. Plasma VTG analysis was unable to decisively distinguish males and

females, although spawning-capable females were distinguishable from all other reproductive

phases. A larger sample size than was available in the present study, particularly of females >525

mm, is needed to provide more conclusive evidence for the utility of VTG to differentiate sexes.

Increased sampling of females >500 mm is needed to determine if the variability in the GSI and

VTG data occurring during the year is natural or is an artifact of the small data set. The

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limitations of this data set required us to evaluate relationships between GSI and VTG with one-

way ANOVA based on ranks instead of actual data, which increased uncertainty in interpreting

our results.

Based on histology in the present study, 50% of male tripletail reached maturity at

approximately 0.5 years old, although our estimate is limited by the lack of age-0 fish (Figure 3).

The lack of immature males precluded calculation of L50 for males; however, the smallest male

tripletail found in a spawning-capable phase was 261 mm. The only immature male was 336 mm

and our data suggests that the L50 for males is < 261 mm (Figure 2). Brown-Peterson and Franks

(2001) and Strelcheck et al. (2004) reported that all males in their studies were sexually mature

and suggested that length at which 50% of the males were mature would likely be ≤290 mm.

Male GSI data remained relatively constant throughout the sampling period, mean value of

0.11% and a range of 0.04 to 0.22%, as would be expected from the primarily spawning capable

male population. Overall, male tripletail captured in the present study near Jekyll Island, GA

were spawning capable through the size range (261- 714 mm) and through the entire sampling

period.

Through histological analysis it was determined that 50% of female tripletail in GA

reached maturity at 449 mm or 1.17 years (Figure 2 and 3). Previous studies have reported that

50% of female tripletail in the Gulf of Mexico reached maturity at approximately 490 mm

(Brown-Peterson and Franks 2001; Strelcheck et al. 2004). Similarly, Brown-Peterson and

Franks (2001) reported that all females >525 were mm sexually mature. Our study indicates that

the L50 for females (449 mm) was slightly below the current minimum size limit (457 mm) for

tripletail caught in Georgia waters.

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Female tripletail had a wider range of GSI values than males, 0.07 - 10.9%, with a mean

of 1.1%; female GSI generally increased from April to August. The high GSI value in March is a

result of a two females captured in 2009 and could be indicative of a protracted spawning period

of tripletail or could be an artifact of small sample size. The asynchronous oocyte maturation and

relatively low mean GSI values throughout the sampling period is consistent with previous

tripletail studies (Brown-Peterson and Franks 2001; Cooper 2002), as well as other multiple-

batch spawning species such as wahoo Acanthocybium solandri (Brown-Peterson et al. 2000).

Plasma VTG concentrations were significantly greater in female tripletail than in males

as expected; however, the difference appears to be heavily influenced by high VTG

concentrations in a few spawning-capable females. Statistical interpretations of the data were

limited by the small sample size, but the trend of higher GSI and VTG for females in the

spawning-capable phase suggests that a more robust dataset than the one used in this study could

provide the ability to differentiate spawning females from other female reproductive phases and

from males (Figures 5 and 6). The relationship between VTG and GSI was strong (r2= 0.873) but

should be interpreted with caution because of the small sample size of high VTG and high GSI,

which may have disproportionately affected the coefficient of determination. The VTG

concentration in females also appears to increase into August, which suggests that peak

spawning may occur in August or later into the year (Figure 6). It should be noted that increased

VTG levels are not always clear indicators of temporal and spatial distribution of spawning fish,

but can be used as a line of evidence for elucidating reproductive activities (Ceapa et al. 2002).

Plasma VTG in spawning capable females was similar to those reported in other fish species

including spawning stellate sturgeon Acipenser stellatus (Ceapa et al. 2002), gag grouper

Mycteroperca microlepis (~3000 µg/ml;Heppell and Sullivan 2000), and presumed spawning

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female Gulf sturgeon Acipenser oxyrinchus desotoi (> 1000 µg/ml;Heise et al. 2009). Tripletail

VTG levels were greater (~ 400 µg/ml) than concentrations found in spawning shovelnose

sturgeon Scaphirhynchus platorhynchus (Wildhaber et al. 2007).

Previous estimates of length and age at which 50% of the population is mature and those

of the present study are based on the premise that females in a developing stage during the

predicted spawning season are mature fish. Other investigators (Strelcheck et al. 2004) have

warned that if fish in the developing phase do not spawn in that season, length- and age-at-sexual

maturity may be underestimated. Strelcheck et al. (2004) further suggested that if early

vitellogenic oocytes were used as the minimum qualification for maturity status, length at which

50% of the females are mature would increase from 494 to 594 mm, which corresponds to an age

of 1 or 2 years. In the present study, classification of immature and developing females as

immature and all other female reproductive phases as mature resulted in an increase in length at

50% maturity from 449 to 491 mm, which corresponds to an age of 1.17 to 1.47 years of age.

This change in maturity classification indicates that a more conservative approach of classifying

maturity status yields only slightly higher length and age estimates; however, the increase to

491mm does raise the length at 50% maturity above Georgia’s current minimum size restriction,

of 457 mm, meaning less than 50% of female fish are mature when they become vulnerable to

harvest.

Preliminary tagging data (Streich et al. 2013) indicates that tripletail remain near the

Georgia coast as late as October; therefore, spawning in the Atlantic could occur later in the year

than previously reported for fish captured in the Gulf of Mexico and represents a possible bias in

the study design. Cooper (2002) suggested that spawning could occur year-round in tropical

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waters, although spawning seems to occur primarily between April and September off the eastern

coast of Florida.

Currently the state of Georgia’s recreational fishing harvest regulations require a

minimum size limit of 457 mm and a creel limit of two fish per person per day. This level of

harvest seems to allow 100% of the males and approximately 50% of the females to reach sexual

maturity prior to vulnerability of harvest. An increase in the minimum size would allow a

greater percentage of females to reach sexual maturity. In the present study, only one female

tripletail greater than 500 mm (1 of 39) was in an immature reproductive phase; therefore, an

increase in the Georgia minimum size limit to 500-525 mm may allow nearly all females to reach

maturity prior to susceptibility to exploitation by anglers. Currently, fisheries managers do not

have sufficient data to understand the effects of fishing mortality on tripletail populations.

The inability to use entanglement gear to capture tripletail in their primary habitat

presents researchers with a challenge when attempting to obtain representative samples across all

size ranges. The use of hook-and-line sampling presents the most cost effective method for

sampling tripletail; however, the use of these sampling methodologies can bias sampling, often

leading to the disproportionate capture of fish in the lower end of the size range, a trend evident

in this and other tripletail studies (Brown-Peterson and Franks 2001; Cooper 2002; Strelcheck et

al. 2004). The lack of imminently spawning females in the present study could be attributed to

the possibility that imminently spawning females may not actively feed; therefore, other methods

to capture tripletail must continue to be explored. Future research should expand the sampling to

a larger temporal and spatial scale and use tagging information to better understand migratory

patterns for more cost-effective sampling. A larger spatial scale should include more inshore and

nearshore locations on the Atlantic coast but should also include an offshore component. A large

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offshore effort may be cost prohibitive because of the random distribution of tripletail and

association with non-stationary floating debris.

Although the role of near shore aggregations in the life history of tripletail in the western

Atlantic remains unanswered, the current study indicates that plasma VTG concentrations could

provide an effective non-lethal sampling technique for determining reproductive status of female

tripletail. The primary need for future research is an increased sample size of sexually mature

female tripletail (> ~525 mm) throughout the year. Increasing the sample size of large females

throughout the year could elucidate trends in GSI and plasma VTG concentrations. Future

laboratory studies of VTG and sex steroid hormone (e.g., testosterone, estradiol, progesterone)

concentrations in maturing tripletail may be a cost effective means for better understanding

tripletail reproductive biology.

Acknowledgements

This project was funded by the Georgia Department of Natural Resources, Coastal

Resources Division (CRD) and the Federal Sportfish Restoration Fund. D. Haymans and S.

Woodward helped all along the way to the project’s completion. C. Belcher provided critical

guidance and technical assistance. B. Warren provided insightful editorial comments to an

earlier draft of this manuscript. J. Franks provided invaluable technical advice on many aspects

of tripletail life history and biology. Staff at GADNR CRD including P. Geer, C. Kinstle,

C.Kalinowsky, G. Gaddis, S. Jordan, E. Robillard, D. McDowell, K. Herrin, L. Willis, K. Wolfe,

G. Meeks, J. Mericle, D. Guadagnoli, P. Medders, W. Hughes, J. Page, B. Readdick, R.

Flournoy, E. Butler and D.Varnadoe provided field and technical assistance with the project.

Anglers including those from the Coastal Conservation Association of Georgia and especially G.

Hildreth provided assistance with field sampling. The GA Cooperative Research Unit is jointly

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sponsored by the GA Department of Natural Resources, the University of Georgia, the U.S. Fish

and Wildlife Service, the U.S. Geological Survey and the Wildlife Management Insitute. Any

use of trade, product, or firm names is for descriptive purposes only and does not imply

endorsement by the U.S. Government.

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Brown-Peterson, N. J., Wyanski D. M., Saborido-Rey, F., Macewicz, B. J., &

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vitellogenin levels in stellate sturgeon (Acipenser Stellatus Pallas) during spawning

migration in the Danube River. Journal of Applied Ichthyology 18, 391-396.

Cooper, D. C. (2002). Spawning Patterns of Tripletail, Lobotes surinamensis, on the Atlantic

Coast of Florida. Master’s Thesis. Florida Institute of Technology, Melbourne, FL.

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aggregation nearshore Jekyll Island, GA, USA. MS Thesis. University of Georgia.

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Table I. Reproductive phases of individual male tripletail (n = 122) captured in 2009 and 2010 near Jekyll Island, GA, USA. All males

in this study in the regressing phase were spawning capable but were not undergoing active spermatogenesis. GE = germinal

epithelium.

Developing Spawning Capable

Month Immature Early Developing Early GE Mid GE Late GE Regressing Regenerating

March 0 0 0 0 0 0 0 0

April 0 1 0 0 2 1 1 0

May 0 1 4 6 9 6 8 0

June 1 0 0 5 20 21 3 0

July 0 0 0 3 12 12 1 0

August 0 0 0 0 2 2 1 0

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Table II. Reproductive phases of individual female tripletail (n = 102) captured in 2009 and 2010 near Jekyll Island, GA, USA.

Developing Spawning Capable

Month Immature Early DevelopingSpawning Capable

Actively Spawning Regressing

Regenerating

March 0 1 0 1 0 0 0

April 7 6 0 0 1 0 2

May 14 2 0 0 0 2 5

June 18 0 0 0 1 2 15

July 4 0 3 5 0 0 9

August 0 0  0 2  0 1 1

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Figure 1. Tripletail sampling location near Jekyll Island, GA, USA. Inset shows a map of the

southeast United States with the Jekyll Island area indicated by the arrow. Each point on the map

indicates that one or more tripletail were captured at that location from March to August 2009

and 2010.

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Figure 2. Non-linear regression of sexually mature female tripletail in 50-mm length bins

captured near Jekyll Island, GA, USA from March to August 2009 and 2010. Closed data points

represent actual cumulative percent maturity of females, whereas males are represented by the

open data points. The dashed line denotes the length (459 mm) at which 50% of female tripletail

are mature. The dotted line represents the minimum size limit (457 mm) currently enforced by

the Georgia Department of Natural Resources. Non-linear regression models for males were

unable to converge and therefore were unable to be modeled; however, only one male captured

in this study was classified as immature.

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Figure 3. Non-linear regression of sexually mature female and male tripletail by age class

captured near Jekyll Island, GA, USA from March to August 2009 and 2010. Closed data points

represent actual cumulative percent maturity of females; males are represented by the open data

points. The dotted line denotes the age at which 50% of males (0.55 years) and females (1.17

years) are mature.

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Figure 4. Mean gonadosomatic index values for (A) female (n = 101) and (B) male (n = 116)

tripletail captured near Jekyll Island, GA, USA from March to August 2009 and 2010. Error bars

represent 95% confidence intervals.

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Figure 5. Blood plasma vitellogenin concentrations of tripletail captured near Jekyll Island, GA,

USA from March to August 2009 and 2010. Females are represented by the triangles and males

are represented by open circles. Reproductive phases are represented by Imm- immature, Dev-

developing, SC- spawning capable, Regress- regressing, and Regen- regenerating.

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Figure 6. Mean plasma vitellogenin concentrations (mug/ml) for (A) female (n = 77) and (B)

male (n = 98) tripletail captured near Jekyll Island, GA, USA from March to August 2009 and

2010. Error bars represent 95% confidence intervals.

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Figure 7. Comparison of plasma vitellogenin and gonadosomatic index values across

reproductive phases for (A) female (n=77) and (B) male (n=98) tripletail captured near Jekyll

Island, GA, USA during March to August 2009 and 2010. Vitellogenin means are represented

by triangles and GSI mean values are represented by circles. Error Bars represent 95%

confidence intervals. Reproductive phases are represented by Imm- immature, Dev- developing,

SC- spawning capable, Regress- regressing, and Regen- regenerating

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