An essential component for assessing and managing populations of elasmobranchs (sharks, skates, and rays) is determining each species' specific reproductive biology, e.g., when sexual maturity occurs, the timing of seasonal cycles, or gestation length (Walker 2004, 2005). Despite the importance of such data to fisheries management (Walker 2004), they are still unavailable for many species of elasmobranchs (Castro et al. 1999; Walker 2004; Pinhal et al. 2008), which has resulted in approximately 40% of the elasmobranch population being classified as data deficient (IUCN 2011). The lack of life history data, combined with direct and indirect fishing pressures, make elasmobranchs susceptible to overexploitation worldwide, which can lead to population declines and localized extinction (Dulvy et al. 2003).

Historically, studying the reproductive biology of elasmobranchs entailed sacrificing a relatively large number of specimens and observing the changes in gross reproductive morphology over the reproductive cycle (e.g., Hisaw and Albert 1947; Zeiner andWolf 1993;Walmsley-Hart et al. 1999; Francis et al. 2001; Ebert 2005; Ruocco et al. 2006).While this approach is still currently appropriate for elasmobranchs with sustainable population sizes (Heupel and Simpfendorfer 2010), it is not suitable for many because of their relatively lowor unknown species abundance (IUCN 2011; Hammerschlag and Sulikowski 2011). Because of this limitation, alternative methods for determining size at maturity and/or reproductive cycles in elasmobranchs are being developed and validated (Sulikowski et al. 2007b).

One particular approach that has been utilized to nonlethally study the reproductive biology of elasmobranchs is the analysis of blood steroid hormone concentrations (e.g., Perez and Callard 1993; Sulikowski et al. 2006, 2007a, 2007b; Kneebone et al. 2007; Awruch et al. 2008; Henningsen et al. 2008). Reproductive hormones directly correlate with morphological changes within the reproductive tract, specifically at maturation (e.g., Gelsleichter et al. 2002; Sulikowski et al. 2006) and during reproductive cycles (e.g., Kneebone et al. 2007; Sulikowski et al. 2007a, 2007b). However, obtaining blood from large species is impractical and can be problematic because of the difficulties involved with specimen handling. These restrictions also serve to protect the animals from stresses induced by handling procedures on large species (Skomal 2007). This situation is further compounded by the restrictions associated with sample collection from threatened and endangered species. Although plasma steroid hormone analysis is particularly useful for studying the reproductive biology of elasmobranchs, the aforementioned constraints make it necessary for new, less invasive approaches to be developed, especially for studying vulnerable species.

While steroid hormones are easily isolated from blood plasma, they are present in other body depots as well (Hoffmann 1978). For example, steroid hormones have been extracted from milk and/or muscle tissues from cows (Noppe et al. 2008), sheep, and chickens (Sawaya et al. 1998); the urine and feces of free-ranging wildlife (Lasley and Kirkpatrick 1991; Holt and Pickard 1999); blubber from marine mammals (Mansour et al. 2002; Kellar et al. 2006); and the skeletalmuscle tissue of a chimaera, the Spotted Ratfish Hydrolagus colliei (Barnett et al. 2009), for various reproductive analyses. Despite the successful extraction of steroid hormones from body depots other than plasma, this approach has never been taken with an elasmobranch. Given the lack of knowledge pertaining to the reproductive biology of elasmobranch fishes, the objectives of the present study were to determine (1) whether reproductive steroid hormones can be quantified from elasmobranch skeletal muscle tissue and (2) whether there is a relationship between the muscle and plasma concentrations of these reproductive hormones.

METHODS

Specimen collection.—Based on their relatively high abundance and ease of collection, the aplacental viviparous Spiny Dogfish Squalus acanthias and yolk sac placental viviparous Atlantic Sharpnose Shark Rhizoprionodon terraenovae were selected as study organisms. Female Spiny Dogfishwere primarily captured by otter trawl off the coast of Rhode Island aboard the 13.7-m bottom trawler FV Proud Mary between December 2010 and February 2012 in an area centered around 71.600°W and 41.365°N. Additional Spiny Dogfish samples were captured by gill net in the Gulf of Maine aboard the 14-m bottom trawler FV Lady Victoria in November 2010 in an area centered around 70.257°W and 42.917°N. Female Atlantic Sharpnose Sharks were captured by bottom longline in the Gulf of Mexico aboard the 51.8-m RV Oregon II during the 2011 NOAA Bottom Longline Survey in an area centered around 82.628°Wand 25.822°N (Figure 1).

Sampling.—Immediately after the capture of an individual, an 8-mL aliquot of blood was collected by caudal venipuncture using a heparinized needle and Vacutainer. Blood samples were placed in a cooler (4°C) for up to 24 h before hematocrit analysis and centrifugation at 1,242 × g for 5 min. The separated plasma was stored at -20°C in the laboratory until extraction. Gross morphological parameters were recorded, including mass (kg), FL, and natural TL, whichweremeasured to the nearest centimeter over a straight line along the axis of the body from the tip of the snout to the posterior notch of the caudal fin and the posterior tip of the upper lobe of the caudal fin while in its natural position, respectively. Afterwards, all sharks were euthanized by severing the cervical vertebrae. Sharks captured in the Gulf ofMaine and off the coast of Rhode Island were kept in a shipboard cooler at 4°C before transport on ice to the University of New England's Marine Science Center and dissected approximately 24 h after capture. All sharks captured in the Gulf of Mexico were dissected approximately 1 h after capture. During dissections, the following were recorded: ovary and oviducal gland mass (to the nearest g); oviducal gland and the five maximum follicle diameters (to the nearest mm); and, if present, pup sex and stretch total length (STL), which was measured to the nearest millimeter over a straight line along the axis of the body from the tip of the snout to the posterior tip of the upper lobe of the caudal fin while fully extended along the axis of the body. In addition, a 5-g white skeletal muscle tissue sample from behind the second dorsal fin was collected and immediately stored at −20°C until extraction.

Plasma steroid hormone extraction.—Each plasma sample was extracted for 17β-estradiol ([E2]), testosterone ([T]), and progesterone ([P4]) following the protocols from Tsang and Callard (1987) and Sulikowski et al. (2004). A 500-µL aliquot of each plasma sample was extracted twice for each hormone with 10 volumes of ethyl ether (ACS grade), and the liquid phase was evaporated at 37°C in a heat block under a stream of nitrogen. To account for procedural loss, each sample was spiked with 1,000 counts/min of tritiated [E2], [T], or [P4] (Perkin-Elmer, Waltham, Massachusetts) prior to extraction. Extracts were reconstituted in phosphate-buffered saline with 0.1% gelatin (PBSG).

FIGURE 1.

Centers of sampling areas for Spiny Dogfish (1 and 2) and Atlantic Sharpnose Sharks (3).

Skeletal muscle tissue steroid hormone extraction.—Two grams of white skeletal muscle tissue from each shark were homogenized with 8 mL of cold phosphate-buffered saline (PBS) for 30 s using a Kinematica PT 10–35 polytron (Bohemia, New York). Then, 500-µL aliquots of homogenate, in quadruplicate, were extracted for each hormone. Nonradiolabeled [E2], [T], and [P4] (Steraloids, Inc., Newport, Rhode Island) were used to make stocks (6.4 µg/mL for [E2], and 80 µg/mL for [T] and [P4]: all in ethanol, ACS grade) to “cold-spike” samples to calculate recoveries following extraction and generate the standard curves for radioimmunoassay. A 100-µL aliquot of each stock was evaporated under a stream of nitrogen, reconstituted in PBSG, and serially diluted to make working stocks with final concentrations of 200 pg/100 µL for [T] and [P4] and 320 pg/100 µL for [E2]. Working stocks were utilized to cold-spike muscle homogenates for calculating sample recovery. Of the four homogenate replicates, two were cold-spiked with a 50-µL aliquot of the corresponding working stock (100 pg of [T] or [P4], 160 pg of [E2]) to track sample recovery. All samples were heated in a 50°C water bath for 15 min, briefly vortexed, and then cooled to room temperature. Samples were extracted with 10 volumes (5 mL) of 2:1 chloroform : methanol (ACS grade; histology grade), and vortexed for 1 min before snap-freezing in a dry ice acetone bath. The liquid phasewas removed by piercing the frozen homogenate layer with a pasture pipette and transferring it to a newtest tube. The liquid phasewas then evaporated at 37°C in a heat block under a stream of nitrogen. OnemL of 70% methanol (histology grade) was added to the dry extract, and the samples were stored at -20°C for 24 h. Samples were centrifuged at 962 × g for 10 min at 4°C, and the liquid phase was decanted and evaporated at 37°C in a heat block under a stream of nitrogen. Extracts were reconstituted in 200 µL of PBSG.

Individual muscle sample recoveries were calculated by subtracting the mean concentration of hormone found in the two nonspiked replicates from the mean concentration of hormone found in the two cold-spiked replicates. The resulting concentration of hormone was then divided by the concentration of hormone that was initially added to the two cold-spiked replicates (100 pg of [T] or [P4], 160 pg of [E2]). To calculate the final concentration of hormone in each skeletal muscle sample (in pg/g), the mean concentration of hormone found in the two nonspiked replicates was calculated, multiplied by a dilution factor of 8 (to convert the units to pg/g), and finally divided by the individual sample recovery.

Radioimmunoassay.—Steroid hormone concentrations from both plasma and muscle extracts were determined following a modified radioimmunoassay procedure from Tsang and Callard (1987). Previously prepared nonradiolabeled [E2], [T], and [P4] stocks (6.4 µg/mL for [E2] and 80 µg/mL for [T] and [P4]) were utilized. Antibodies of [E2], [T], and [P4] (Gordon Niswender, Colorado State, Fort Collins, Colorado), were diluted to final concentrations of 1:18,000, 1:10,000, and 1:2,500, respectively. Tritiated hormones and antibodies were added to the reconstituted plasma samples (100 and 50 µL in duplicate) and to muscle samples (200 µL) using PBSG to bring the total volume to 400 µL. Assays were incubated at 4°C for 24 h before separating free from bound hormone by the addition of a carbon (0.2%; Acros Organics, Fairlawn, New Jersey) and dextran 70 (0.02%; Tokyo Chemical Industry Co., Ltd., Tokyo, Japan) suspension followed by centrifugation at 1,242 × g for 10 min at 4°C. The resulting supernatant was then decanted into scintillation vials and 3.5 mL of Ecolume (MPO Biomedicals, Solon, Ohio) was added. The radioactivity level was then measured by a Perkin-Elmer Tri-Carb 2900TR liquid scintillation analyzer (Waltham,Massachusetts). The mean intra-assay coefficients of variation were 8, 10, and 9% for [E2], [T], and [P4], respectively, in the Spiny Dogfish plasma assays and 7, 7, and 6% in the Atlantic Sharpnose Shark plasma assays. The mean interassay coefficients of variation were 10, 11, and 12% for [E2], [T], and [P4], respectively, in the Spiny Dogfish plasma assays and 8, 8, and 9% in the Atlantic Sharpnose Shark plasma assays. The mean intra-assay coefficients of variation were 8, 10, and 8% for [E2], [T], and [P4], respectively, in the Spiny Dogfish muscle assays and 10, 8, and 9% in the Atlantic Sharpnose Shark muscle assays. The mean interassay coefficients of variation were 10, 11, and 12% for [E2], [T] and [P4], respectively, in the Spiny Dogfish muscle assays. In calculating the means ± SEs for the concentrations of each steroid hormone per stage, any value that was nondetectable was assigned the lowest possible concentration that the assay would have been able to detect in the aliquots utilized.

Statistical analysis.—Linear regressions were performed for [E2], [T], and [P4] quantified from plasma and skeletal muscle. Data were transformed if the variables failed tests of normality or homogeneity of variance. If transformed variables still violated the assumptions the nonparametric Kendall's tau rank correlation was conducted. All data were analyzed using R 2.13.1 (R-Core Development, 2011), and means and SEs were graphed in Sigmaplot 12 (Systat Software, San Jose, California).All tests were considered significant at α = 0.05.

RESULTS

A total of 31 female Spiny Dogfish (78–90 cm FL, 2.7– 4.9 kg) were collected and divided into discrete reproductive stages (Table 1) based on previous literature (Hisaw and Albert 1947; Tsang and Callard 1987); the group included six preovulatory females, six females containing candles, five early-gestation females containing 62–88-mm embryos, eight mid-gestation females containing 190–240-mm embryos, and six late-gestation females containing 250–275-mm embryos. Ten female Atlantic Sharpnose Sharks (68–85 cm FL, 2.4–4.6 kg) were collected and divided into arbitrary discrete reproductive stages (Table 2), including one preovulatory female, two early-gestation females containing 28–55-mm embryos, four early—mid-gestation females containing 56–83-mm embryos, and three mid-gestation females containing 85–139-mm embryos. The overall mean recoveries of plasma [E2], [T], and [P4] extractions were 80, 87, and 70% for Spiny Dogfish and 71, 85, and 68% for Atlantic Sharpnose Sharks, respectively, while the overall mean recoveries of muscle [E2], [T], and [P4] extractions were 21, 23, and 20% for Spiny Dogfish and 59, 39, and 55% for Atlantic Sharpnose Sharks, respectively.

Spiny Dogfish

In preovulatory female Spiny Dogfish, [P4] concentrations in both plasma (658 ± 186 pg/mL) and muscle (543 ± 285 pg/g) were similar. However, plasma [P4] (60 ± 22 pg/mL) decreased by 10-fold during the candle stage, while muscle [P4] (2,412 ± 420 pg/g) increased by about fivefold. Females in early gestation displayed relatively elevated plasma (1,004 ± 225 pg/mL) and muscle (1,150 ± 209 pg/g) [P4] concentrations before they declined (plasma [159 ± 38 pg/mL] and muscle [386 ± 89 pg/g]) through the remainder of gestation (Figure 2A). Overall, muscle [P4] concentrations were not significantly related to plasma [P4] concentrations (Kendall's tau = 0.16; P = 0.23).

TABLE 1.

Mean ± SE maximum follicle diameter (MFD) and pup STL for Spiny Dogfish, by gestational stage (preovulatory [P/O], candle, early, mid, and late); N = sample size.

TABLE 2.

Mean ± SE maximum follicle diameter (MFD) and pup STL for Atlantic Sharpnose Sharks, by gestational stage. See Table 1 for additional information.

During the candle stage, [T] concentrations in Spiny Dogfish plasma (23 ± 8 pg/mL) and muscle (74 ± 24 pg/g) were low, and they remained that way through mid gestation (plasma [47 ± 11 pg/mL] and muscle [134 ± 34 pg/g]). However, during late gestation, plasma [T] increased (423 ± 127 pg/mL), while muscle [T] remained unchanged (78 ± 18 pg/g). The highest muscle (447 ± 54 pg/g) and plasma (668 ± 255 pg/mL) [T] concentrations were found in the preovulatory stage (Figure 2B). Similar to [P4], muscle [T] was not significantly related to plasma [T] in Spiny Dogfish (Kendall's tau = 0.16; P = 0.30).

From preovulation to mid gestation, Spiny Dogfish had low plasma (65 ± 12 pg/mL) and muscle (113 ± 39 pg/g) [E2] concentrations. But concentrations of [E2] increased in plasma (864 ± 116 pg/mL) and muscle (364 ± 73 pg/g) through the second half of gestation (Figure 2C). Unlike [P4] and [T], Spiny Dogfish muscle [E2] concentrations were significantly related to plasma [E2] (Kendall's tau = 0.49; P = 0.0017).

Atlantic Sharpnose Sharks

From preovulation to early gestation, concentrations of [P4] and [T] in Atlantic Sharpnose Sharks were relatively low in plasma ([P4]: 162 ± 76 pg/mL; [T]: 151 ± 119 pg/mL) and muscle ([P4]: 182 ± 76 pg/g; [T]: 84 ± 34 pg/g). However, a marked elevation of these two hormones occurred in plasma ([P4]: 3,582 ± 1,767 pg/mL; [T]: 4,840 ± 884 pg/mL) and muscle ([P4]: 3,325 ± 1,253 pg/g; [T]: 693 ± 84 pg/g) samples in early—mid-gestation females. Concentrations of [P4] and [T] then decreased in the plasma ([P4]: 132 ± 9 pg/mL; [T]: 83 ± 23 pg/mL) and muscle ([P4]: 391 ± 63 pg/g; [T]: 141 ± 91 pg/g) through mid gestation (Figure 3). The muscle [P4] and [T] of Atlantic Sharpnose Sharks were found to be significant predictors of plasma [P4] and [T] (linear regression; [P4]: R2 = 0.65, P = 0.0048; [T]: R2 = 0.84, P = 0.00022).

FIGURE 2.

Mean concentrations of (A) progesterone, (B) testosterone, and (C) estradiol in plasma and muscle of Spiny Dogfish, by gestational stage (preovulatory [P/O], candle, early, mid, and late). The numbers above the xaxes are sample sizes; the error bars represent SEs. Note the difference in scale between the plasma and muscle.

The preovulatoryAtlantic Sharpnose Shark female contained relatively elevated concentrations of [E2] in plasma (4,885 pg/mL) and muscle (232 pg/g). Concentrations of [E2] progressively decreased in both plasma (762 ± 167 pg/mL) and muscle (72 ± 12 pg/g) through early to mid gestation (Figure 3C). Similar to those of [P4] and [T], the concentrations of Atlantic Sharpnose Shark muscle [E2] were found to be significant predictors of plasma [E2] (linear regression; R2 = 0.79, P = 0.00059).

FIGURE 3.

Mean concentrations of (A) progesterone, (B) testosterone, and (C) estradiol in plasma and muscle of Atlantic Sharpnose Sharks, by gestational stage. See Figure 2 for additional information.

DISCUSSION

Concentrations of reproductive hormones were successfully detected in the muscle and plasma of female Spiny Dogfish and Atlantic Sharpnose Sharks during specific chronological events in the reproductive cycle. In addition, significant relationships were found between plasma and skeletal muscle concentrations of [E2] in Spiny Dogfish and [E2], [P4], and [T] in Atlantic Sharpnose Sharks, suggesting that the concentrations of these hormones in the skeletal muscle of elasmobranchs have the potential to be an indicator of gestational stage.

CONCLUSIONS

In the present study, we developed a method of quantifying steroid hormones in skeletal muscle and tested their efficacy, relative to plasma hormone concentrations, in assessing the reproductive status of sharks. An advantage of using skeletal muscle rather than plasma is that muscle tissue can be obtained nonlethally and with minimal invasion. The results suggest that white skeletal muscle tissue of Spiny Dogfish and Atlantic Sharpnose Sharks is a depot for the reproductive hormones [P4], [T], and [E2], which can be reliably measured by radioimmunoassay. Additionally, the patterns of muscle reproductive hormones, which are coincident with gestational stage, paralleled those in plasma despite the different reproductive modes exhibited by these two sharks. Our results suggest that skeletal muscle steroid hormone content correlates well with gestational stage in yolk sac placental species. As such, future Atlantic Sharpnose Shark sampling efforts will focus on collecting individuals in the later stages of gestation to determine whether the parallel patterns observed in plasma and muscle steroid hormone concentrations are consistent throughout the reproductive cycle. While there were no significant relationships between plasma and muscle [P4] and [T] concentrations in Spiny Dogfish, the similar patterns exhibited by these two hormones in plasma and skeletal muscle suggest that they could still be used to broadly assess gestational stage in that species. Certainly, because of the stronger relationship between plasma and skeletal muscle [E2] concentrations, this hormone may be a better indicator of gestational stage in Spiny Dogfish.