Sex Reversal in Yellow Perch (Perca Flavescens) to Produce Functional Neomale Sperm Donors

Sex Reversal in Yellow Perch (Perca flavescens) to Produce Functional Neomale Sperm Donors

THESIS

Presented in Partial Fulfillment of the Requirements for the Degree of Master of Science in the Graduate School of The Ohio State University

Kristen Towne

Graduate Program in Environment and Natural Resources

The Ohio State University

2016

Master’s Examination Committee:

Dr. Konrad Dabrowski, Advisor

Dr. Robert Gates

Dr. Suzanne Gray

Dr. Roman Lanno

Copyrighted by Kristen Marie Towne 2016

Abstract

Yellow perch (Perca flavescens) are a popular game and food fish in the Great Lakes region. However, intensive culture of this species in the North Central Region is still in its infancy. In this species, females grow faster and to a larger size than males, making them more valuable in aquaculture. In fact, a population of 1,000 female yellow perch can gross approximately $555 more than the same number of fish exhibiting a 1:1 male:female ratio at the current fillet value of $14 per pound. Attempts have been made to produce monosex populations of yellow perch through sex reversal, but have resulted in the formation of nonfunctional, intersex fish due to the initiation of treatment after the beginning of gonadal differentiation. Yellow perch require more elaborate techniques to conduct sex reversal than many other species due to the beginning of sexual differentiation occurring before the transition to formulated feeds. This thesis consists of two sex reversal experiments carried out in 2015 and 2016. The 2015 studies evaluated the success of hormonal masculinization using the synthetic steroid hormone 17α- methyltestosterone (MT) by two treatment methods, diet enrichment and immersion, as well as two initiation times with respect to fish size, 12 mm and 14 mm total length. This study also evaluated the feminizing capacity of another steroid hormone, estradiol-17β, in yellow perch exposed via immersion treatments beginning at the same two size classes.

All fish were checked for spermiation by manual stripping in late January/early February

ii of 2016. Although the groups that had been exposed to the masculinizing hormone beginning at 12 mm total length displayed significantly more spermiating males than those exposed beginning at 14 mm (P < 0.01), the control groups of both size classes also displayed a sex ratio significantly deviated toward males (P < 0.01). In addition, the 12 mm feminizing hormone-treated group displayed a sex ratio significantly deviated toward males (P < 0.05). During the hormone exposure phase, fish were maintained at water temperatures that were much higher than those found in Lake Erie during this time of ontogenetic development to optimize growth. Therefore, it was hypothesized that these high water temperatures inadvertently induced masculinization. In the spring of 2016, two additional sex reversal experiments were conducted, this time directly evaluating the effect of temperature on gonadal differentiation. Three egg ribbons were obtained from

Mill Creek Perch Farm, LLC, and divided into two groups each: one exposed to water temperatures of 15.6 ± 1.9o C, and the other to temperatures of 24.0 ± 1.3o C for a duration of 43 days. The sex ratio of each group was checked by dissection in September of 2016, with no group showing a sex ratio that significantly differed from the expected

1:1 sex ratio (P < 0.05). The other temperature sex reversal experiment was conducted on the progeny of a sample of the control males from the 2015 hormonal sex reversal experiment. Similar to the other temperature experiment, 10 day old larvae were separated into warm (23.1 ± 0.2o C) and cool (16.4 ± 1.0o C) water systems for a duration of 23 days. The sex ratio was checked by dissection in October of 2016. Of the five fathers evaluated, two produced progeny populations that significantly deviated toward females

(P < 0.05), indicating that these two fish were likely masculinized females/neomales. The

iii remaining three fathers produced progeny populations that did not significantly differ from the expected sex ratio. Additionally, the sex ratios of individual males of the warm groups did not significantly differ from the cool groups (P < 0.05). The results from these two temperatures used in these experiments indicate that high water temperatures do not affect gonadal differentiation in juvenile yellow perch. Therefore, the skewed sex ratios in the control groups of the 2015 hormonal sex reversal experiment may have been caused by some environmental or food-containing factor(s) yet to be determined.

However, to our knowledge, this is the first work to produce all-female populations of yellow perch from functional neomale sperm donors.

Acknowledgements

First and foremost, I would like to thank my advisor, Dr. Konrad Dabrowski, for his endless hours of support and guidance. I would also like to thank the other members of my examination committee and the kind and helpful staff of the School of Environment and

Natural Resources. I would like to thank my dear friends and family for their support and understanding during stressful spawning seasons. Lastly, this research would not have been possible without the patience, guidance, and assistance of Mohammad Alam,

Thomas Delomas, Kevin Fisher, John Grayson, Megan Kemski, Mackenzie Miller, and Shib

Nath Saha.

Vita

June 2009……………….Brecksville-Broadview Heights High School, Broadview Heights, Ohio

May 2013……………………………………………………..……….B.S. Animal Science, Cornell University

January 2015 to present…...Graduate Teaching Associate, SENR, The Ohio State University

Field of Study

Major Field: Environment and Natural Resources Specialization: Fisheries and Wildlife Science

Table of Contents

Abstract………………………………………………………………………………….………………………………...…..ii

Acknowledgements………………………………………………………………..……………………………..……….v

Vita…………………………………………………………………………….……………………………………….………..vi

Field of Study………………………………………………………………………..…………………………………….…vi

Table of Contents……………………………………………………………….………………………………………..vii

List of Tables…………………………………..………………………………………………………………………………x

List of Figures………………………………………………….…………………………………………………………….xi

Chapter 1: Introduction to Yellow Perch and Sex Reversal……………………………………..………1

Taxonomy and Physical Characteristics……………………………………………………………….1

Market Demand for Yellow Perch………………………………………………….……………………1

Sex Reversal………………………………………………………………………………………………………..3

Hormonal Sex Reversal……………………………………………………………………………4

Environmental Sex Reversal………………………………………………………………….11

Goals and Objectives……………………………………………..……………………………..………….13

Hypotheses……………………………………………………………………………………………………….14

Chapter 2: Determination of the Optimum Timing and Exposure Method for Hormonal Sex Reversal in Yellow Perch……………………………………..…………………………….……….16

Introduction…………………………………………………………………..…………………………………16

vii

Methods…………………………………………………………….…………………………………………….20

Gamete Collection and Incubation……………………………………………..…………21

Larval Rearing…………………………………………….…………………………………………22

Enrichment of Artemia nauplii……………………………………………..……………….23

Sex Reversal…………………………………………….……………………………………………24

Growout………………………………………………………………………………..………………28

Analysis of F2 Generation……………………………………………………………..………29

Statistical Analysis……………………………………………………………..………………….31

Results…………………………………………………..………………………………………………………….32

Method of Exposure………………………………………………………..……………………32

Time of Initiation……………………………………………………..……………………………34

Evaluation of F2 Generation……………………………………..…………………………..35

Discussion…………………………..…………………………………………………………………………….35

Chapter 3: Exploration of the Potential for Temperature-Induced Sex Reversal in Yellow Perch…………………………………………..……………………………………………………………………42

Introduction………………………………………..……………………………………………………………42

Methods…………………………………………….…………………………………………………………….44

Experiment 1……………………………………………………………..………………………….44

Experiment 2………………………………………………………..……………………………….47

Statistical Analysis……………………………………..………………………………………….50

Results……………………………..……………………………………………………………………………….52

viii

Experiment 1…………………………………………………..…………………………………….52

Experiment 2………………………………………………..……………………………………….53

Discussion………………………..……………………………………………………………………………….55

Future Research……………………………………….………………………………………………………………….58

Literature Cited (AFS)……………………………………………….………………………………………………….60

Appendix A: T Test and ANOVA Tables……………………………………………..………………………….74

List of Tables

Table 1. Gonadosomatic indices of yellow perch fed diets containing different levels of methyltestosterone (MT) or cholesterol (C, control) for 84 days. Numbers are expressed as mean ± SE. Different letters next to the numbers within a column indicate a significant difference between groups (P < 0.01). Adapted from Malison et al. (1986)………………………………………………….18

Table 2. Summary of the hormone exposures for both Group 14 mm and Group 12 mm…………....27

Table 3. Water quality parameters during the hormone exposure phase of Yellow Perch larvae in Group 14 mm. Values are given as mean ± standard deviation. No values within a column differed significantly (ANOVA, P < 0.05)…………………………………………………………………………………………………..27

Table 4. Water quality parameters during the hormone exposure phase of Yellow Perch larvae in Group 12 mm. Values are given as mean ± standard deviation. No values within a column differed significantly (ANOVA, P < 0.05)…………………………………………………………………………………………………..28

Table 5. Total weight (TW), gonadosomatic index (GSI), and percent females of progenies (n = 5) produced by two control males from Group 14 mm (Chapter 2) that had been raised at 23.1 ± 0.2o C (Warm) or 16.4 ± 1.0o C (Cool). Values are given as mean ± standard deviation. Single values represent those from a single individual. Both sires were in the “large” group of the two that were PIT tagged…………………………………………………………………………………………………………………………………55

Table 6. Total weight (TW), gonadosomatic index (GSI), and percent females of progenies (n = 5) produced by three control males from Group 12 mm (Chapter 2) that had been raised at 23.1 ± 0.2o C (Warm) or 16.4 ± 1.0o C (Cool). Values are given as mean ± standard deviation. Single values represent those from a single individual. All sires were in the “small” group of the two that were PIT tagged…………………………………………………………………………………………………………………………………55

List of Figures

Figure 1. Examples of genital duct abnormalities resulting in nonfunctional masculinized Rainbow Trout Oncorhynchus mykiss. (A) Normal morphology; (B, C, D) absence of the deferent duct (DD), with a conjunctive rudiment (GR) displaying periodic (B) or continuous (C, D) oedematous vesicles (OE); (E, F) the deferent duct does not connect to the papilla (Cousin-Gerber et al. 1989)…………….7

Figure 2. Flow chart with the three phases of the experiment: fertilization and larval rearing, hormone exposure, and growout. Larvae were separated into the four treatment groups, each in triplicate, for the hormone exposure phase. Fish remained separated by treatment group and, for the 12 mm Group, also by replicate following hormone exposure. Eventually, all fish were fin clipped by treatment group and for the 12 mm Group, also tagged with a visible implant elastomer by replicate, and combined into one common growout tank. This design was followed by each size group separately, so the two were never combined…………………………………………………………………..20

Figure 3. Recirculating aquaria system used to raise larval and juvenile perch. System consisted of 12 tanks with individual aeration, biofilter system, 100 μm mesh outlets, feeding rings, heater in a lower reservoir, and 1,800 gph pump………………………………………………………………………………………22

Figure 4. Incubation setup for Artemia nauplii enrichment. Enrichments followed the methods of Grayson (2014), with slight modification to substitute 17α-methyltestosterone for the polyunsaturated fatty acid. Both enrichments were set 24 hours after the Artemia cysts began the hydration process……………………………………………………………………………………………………………………..24

Figure 5. Important points in the timelines of 2015 hormonal sex reversal experiments. Time bars are in relation to each other (i.e., Group 12 mm fertilization occurred at 25 days post-fertilization (DPF) for Group 14 mm………………………………………………………………………………………………………………25

Figure 6. (A) Closed, 6-L tanks with individual aeration used to house juvenile fish during hormone treatment. Treatments were randomly distributed across the 12 tanks; (B) Tanks were supplied with algae to reduce aggression between the fish and salt (3 ppt) to extend the lifespan of the Artemia diet………………………………………………………………………………………………………………………………26

Figure 7. Survival rates during the hormone exposure phase of fish exposed to one of two hormones (MT or E2) by one of two methods (diet or immersion), beginning at either 12 mm or 14 mm TL. Survival rates between treatments within one initiation size did not significantly differ for either Group 12 mm or Group 14 mm. Error bars represent standard deviation. There were no significant differences in survival rate (P < 0.05)…………………………………………………………….…………..32

Figure 8. Percent spermiating males in Yellow Perch exposed to one of two hormones (17α- methyltestosterone (MT) or estradiol-17β (E2)) by one of two treatment methods (Diet or Immersion (Imm.)), compared to control. Error bars represent standard deviation. Error bars are not present of Group 14 mm because the replicates were combined prior to checking for spermiation to compensate for differing stocking densities. Letters above columns indicate a significant difference between fish size groups (one-way ANOVA, Tukey HSD). Stars above columns indicate a significant deviation from the expected 1:1 sex ratio (chi-squared test). Significance was accepted at P < 0.05…………………………………………………………………………………………34

Figure 9. Water bath housing warm group tanks to more accurately maintain temperature (24.4 ± 0.4o C). System consists of four tanks holding 6 liters of water each, aeration in each tank, water heater, and aeration in water bath to ensure water movement………………………………………………….46

Figure 10. Important points in the timeline of Experiment 2. From 56 days post-fertilization (DPF) onward, weighings and samplings were based on degree days instead of age to reflect the size differences of the fish caused by increased growth rates in warm groups. Values are presented as mean ± standard deviation………………………………………………………………………………………………………..51

Figure 11. Percent males in four groups of fish from three unrelated dams following exposure to high temperatures throughout the period of gonadal differentiation. Results showed no significant differences between groups (ANOVA, α = 0.05), as well as no significant deviations from the expected 1:1 male to female sex ratio (chi-squared test, α = 0.05). Numbers above columns indicate sample sizes…………………………………………………………………………………………………………………53

Figure 12. Percent females produced by each of the five sires from the 2015 control Yellow Perch. Five fish were sampled from each group for sexing. Stars within columns indicate a significant deviation from the expected sex ratio (chi-squared test). Significance was accepted at P < 0.05…54

xii

Chapter 1: Introduction to Yellow Perch and Sex Reversal

Taxonomy and Biological Characteristics

Yellow perch belong to the family Percidae, which includes Walleye Sander vitreus, Sauger

S. canadensis, and darters (Ammocrypta spp., Crystallaria spp., Etheostoma spp., Percina spp., and others) (Hart et al. 2006). The species is characterized by two separated dorsal fins, the anterior fin having 13-15 spines and the posterior fin having 1-2 spines, the remainder of which consists of 12-15 soft rays (Thorpe 1977). Pelvic and anal fins also have one and two spines, respectively, and each opercula carry a spine as well (Thorpe

1977). Their coloration is characterized as an olive green color dorsally, fading to cream ventrally, with 6-8 dark vertical bars (Hart et al. 2006). Adults typically range from 15-30 cm (6-12 in.) total length (TL) (Hart et al. 2006). Yellow Perch are similar in morphology to the Eurasian Perch P. fluviatilis, and the two are recognized as sister species and, in some cases, biologically equivalent (Thorpe 1977). The third member of the genus, the Balkhash

Perch P. schrenkii, which is native to China and Kazakhstan, is also similar in meristic characters to Yellow Perch (Mamilov 2015). Yellow Perch is the only species in the Perca genus that is native to North America (Stepien & Haponski 2015).

Market Demand for Yellow Perch

Yellow Perch have proven to be an attractive game fish in the Great Lakes region of the

United States over the last several decades. Their low muscle fat content (< 2%), firm flesh, and high protein level (19.5%) (Hinshaw 2006) make them a desirable food fish, resulting in a peak harvest of more than 15,000 metric tons (33 million pounds) per year in the 1960s (Malison 2003). This species has several attributes of a quality food fish, including lower percent fat than many other popular cultured food fishes (e.g., catfishes, salmonids) and a comparable amount of omega-3 fatty acids (0.3%) (Hinshaw 2006).

Yellow Perch has also risen in popularity due to its long shelf life, ability to resist freezer burn, and simplicity to cook, all as a result of its low crude fat and high phospholipid content (Malison 2003), which comprise 70-80% of total lipids (Dabrowski et al. 2015).

Approximately 70% of the Yellow Perch sales in the United States occur within 50 miles of the Great Lakes region, with another large market in major Canadian cities along the

St. Lawrence River and Lake Ontario. The annual US harvest from wild caught Yellow Perch has dropped steadily in the years since its peak, reaching 5-8,000 metric tons (11-18 million pounds) per year in the 1980s and 1990s, and less than 5,000 metric tons (10 million pounds) per year in the early 2000s (Malison 2003). The Canadian yield, although larger, also follows the same downward trend (Hudson & Ziegler 2014). The precipitous decline led to the species’ listing as “high priority” for production-oriented research by the North Central Regional Aquaculture Center (NCRAC) Industry Advisory Council (IAC).

In the years since Yellow Perch were listed, the species has been identified as a good

2 candidate for aquaculture because of its acceptance of formulated feeds, their lack of aggression or cannibalism as adults, and their high tolerance for intensive culture conditions (Malison et al. 1993).

Yellow Perch are the third most valuable food fish in the North Central Region, with an annual profit of approximately $432,000 (Census of Aquaculture 2013). As of 2005, it was estimated that 30 fingerling producers supplied 6-12 million fingerlings per year, which translates to less than 226 metric tons per year after grow-out (Hart et al. 2006). This grow-out involves raising fingerling Yellow Perch to market size, which took place at only

12 farms (Census of Aquaculture 2013). However, Malison (2003) estimated the aquaculture facilities in the Midwestern United States produced a mere 90.8 metric tons every year. Wallet et al. (2005) also stated that Yellow Perch aquaculture has not expanded rapidly. Undoubtedly, the yield of farmed Yellow Perch will have to increase substantially if the market demand is to be met.

Sex Reversal

Like many fish species, Yellow Perch exhibit sexually dimorphic growth, with females growing faster, maturing later, and reaching a larger final size than males (Diana & Salz

1990; Marsden & Robillard 2004). Understandably, this has major implications for the aquaculture of this species. In fact, an all-female population of Eurasian Perch weighed

3 approximately 30% more than a 1:1 male:female population after their first year (Rougeot

2015). At this time, the mean body mass of the all-female population was 140 g and that of the mixed-sex population was 100 g (Rougeot 2015). Consequently, a tank of 1,000 female fish at a price of $13.99/lb for fillets (Frank’s Fish and Seafood Market, Columbus,

Ohio, June 2016) and an estimated 45% fillet yield (Malison 2003) will gross approximately

$555 more than mixed-sex populations. Therefore, it is highly beneficial to produce the maximum number of females possible.

Hormonal sex reversal

Sex control has long been used in aquaculture for a number of reasons: to produce greater numbers of a more desirable sex (either by differential growth rate, egg production, or for the aquarium trade), to limit growth depression caused by high rates of reproduction in aquaculture ponds, or to reduce the reproductive potential of fishes released to the environment (Singh 2013; Strüssmann et al. 2005). Several sex reversal studies have been conducted on a variety of species to create phenotypically male organisms from genetic females that can act as sperm donors to obtain a new generation of all female fish (Lin et al. 2012; Malison et al. 1986; Rougeot et al. 2002; Singh 2013). Several fish species have been shown to undergo sex inversion when exposed to exogenous steroids, such as 17α- methyltestosterone (MT), inducing masculinization, and estradiol-17β (E2), inducing feminization (Singh 2013). According to Budd et al. (2015), MT in particular has been used to successfully masculinize approximately 35 fish species. 17α-methyltestosterone is

4 most notably used to masculinize Nile Tilapia Oreochromis niloticus to prevent high rates of reproduction and subsequent growth stunting in overpopulated ponds (Mateen &

Ahmed 2015). Additionally, males of this species grow larger than females, making them more profitable (Gale et al. 1999). However, for many species, production of 100% monosex populations have previously not been reliable due to the influence of environmental factors on phenotypic sex in fishes (Strüssmann et al. 2005).

There are four methods of introducing hormones into the fish body: diet treatment, immersion treatment, injection, and silastic implantation (Pandian & Sheela 1995). Only two of these, diet treatment and immersion treatment, are feasible in Yellow Perch because of the size at which treatment must begin (< 16 mm total length; Malison et al.

1986). Although hormonal injection may produce a quicker sex reversal while using less hormone, it is more laborious, expensive, difficult, and there is greater potential for injury and infection. Silastic implantation of the hormone can be highly beneficial because of the ability to release the hormone in a slow, uniform manner, but it is also more difficult and expensive than diet or immersion treatments. Both injection and implantation also require the fish be fairly large, so they must undergo sex differentiation at a relatively late period (Pandian & Sheela 1995).

Therefore, Yellow Perch undergoing sex reversal must be exposed to the hormone either via dietary treatment or immersion treatment. Both methods have the advantages of being inexpensive and requiring relatively simple protocols. Diet treatments are

5 disadvantageous in that the hormone is metabolized in the digestive tract, and the purity, solubility, and distribution in the feed may vary (Pandian & Sheela 1995). According to

Ong et al. (2012), only 5-10% of all MT ingested enters the blood stream because most of it is destroyed by the liver. Varying sizes of the fish may also lead to a variation in feed

(and therefore hormone) uptake (Pandian & Sheela 1995). However, immersion treatments are limited in that they have practically been restricted to use in embryos and larvae. Both treatment methods also carry the risk of sterility or paradoxical feminization should the incorrect dosage or duration be used.

Regardless of the treatment method, three major factors contribute to success of hormonal sex reversal in fishes: dose, time of initiation, and duration of treatment, all of which vary with species (Budd et al. 2015; Rougeot et al. 2002). Treatment must be initiated before the onset of sexual differentiation in gonochoristic fishes (Rougeot et al.

2002) and extend throughout differentiation for both masculinization and feminization

(Budd et al. 2015; Nakamura 2013) to avoid the production of nonfunctional fish (Figure

1). Sex determination is defined as “the mechanisms directing sex differentiation whereas sex differentiation is development of testis or ovaries from the undeveloped

(undifferentiated) gonad” (Hayes 1998). The time of hormone initiation is likely the most vital factor influencing the success of sex reversal (Rougeot 2015). For instance, gynogenetic Northern Pike Esox lucius fed 30 mg MT/kg diet beginning at 26-30 mm TL resulted in 83% male and 17% intersex fish (Luczynski et al. 2003). The same treatment initiated at 30-41 mm TL resulted in 76% male and 24% intersex fish. The authors

6 concluded that larger fish had already begun sexual differentiation at the time that hormone treatment began, causing higher proportions of intersex fish. Additionally, there is evidence that germ cells begin physiological differentiation before any discernible histological differentiation of the gonads. Hormone treatments must be extended throughout both these physiological and morphological differentiations in order to be successful (Nakamura 2013).

Although the time of initiation of hormone treatment may perhaps be the most important factor influencing the success of sex reversal, the dosage of hormone used plays an important role as well. Orally administered MT was shown to induce masculinization at a dosage of 50 mg/kg diet, initiated at 7 days after hatching for a duration of 18 days in

Mozambique Tilapia Oreochromis mossambicus (Nakamura 1975). On the contrary, the same hormone at a concentration of 1,000 mg/kg diet, initiated at the same time and for the same duration, had no effect on females and formation of an ovarian cavity in males

(Nakamura 1975). This phenomenon that results from the transformation of androgens to estrogens, known as “paradoxical feminization”, has also been observed in Channel

Catfish Ictalurus punctatus (Goudie et al. 1983), Blackchin Tilapia Sarotherodon melanotheron, and Jack Dempsey Cichlid Cichlasoma biocellatum (Nakamura 2013), among others. The paradoxical feminization proved to be functional and permanent in all species studied (Nakamura 2013). The potential for intersex traits and subsequent sterilization is also present in the cases that do not result in complete feminization

(Luczynski et al. 2003). Excessive doses of hormone may also result in increased mortality, as was observed in Rainbow Trout Oncorhynchus mykiss alevins (21-67%) exposed to 30-

300 mg MT/L (Van den Hurk & Slof 1981).

Optimizing the time of initial hormone application for certain species is made more difficult by sexual differentiation beginning at different times in males and females.

Ovarian development has been shown to occur 8 days before hatching in Coho Salmon

Oncorhynchus kisutch, but testicular development does not begin until 6 days post-hatch

(DPH) (Piferrer & Donaldson 1989). Similarly, Channel Catfish begin to sexually differentiate at 19 and 90 DPH for females and males, respectively (Patiño et al. 1996).

These results stress the necessity to perform thorough histological analyses in embryonic, larval, and juvenile stages of fish to determine the appropriate time for initial hormone exposure in order to increase the likelihood for successful sex reversal and minimizing hermaphroditism.

Lastly, the duration of treatment is an important factor contributing to the success of sex reversal. In Chinook Salmon Oncorhynchus tshawytscha exposed to a single immersion treatment of 20 μg MT/L water, groups that were exposed to longer treatment durations produced a higher proportion of males (Baker et al. 1988). Additionally, groups exposed to two immersion treatments displayed a greater proportion of males than those only receiving one treatment (Baker et al. 1988). Species that show more successful sex reversal when exposed to multiple hormone immersion treatments likely have an extended labile period (Budd et al. 2015). The same holds true for diet treatments.

Chevassus & Krieg (1992) fed all-female Brown Trout Salmo trutta a diet consisting of either 0.5 or 3 mg MT/kg diet for a duration of either 60 or 80 days. At the 3 mg/kg dose, an 80 day treatment duration resulted in 84.7% males, while the 60 day duration led to only 60.4% males. The same pattern was followed at the 0.5 mg/kg dose: 80 days of exposure led to 28.3% male, and the 60 day duration period led to 14.8% male. All of the doses and durations resulted in similar percentages of functional males (i.e., those with open sperm ducts). However, Cousin-Gerber et al. (1989) produced 82% functional male

Rainbow Trout when fish were fed 0.5 mg MT/kg diet for a duration of 60 days. That percentage dropped to 72.6% functional males when the duration was extended to 90 days. An even larger decrease in functional males occurred when the dose was increased to 3 mg MT/kg diet for 60 or 90 days, producing 35.6% and 19.2% functional males, respectively. The duration of treatment should be as short as possible to limit the potential for hormone toxicity, assuming the dosage is optimal (Nakamura 2013).

Although there are several benefits to hormonal sex reversal, there also are a number of challenges. Overdoses and long durations can lead to increased mortality, growth depression, and deformities or paradoxical feminization. In addition, the potential for adverse human health effects for those that handle hormones require researchers and farmers take precautions when administering them to the fish (Budd et al. 2015).

Synthetic hormones such as the ones used in sex reversal studies also have the potential to influence the sex ratio of wild populations should the hormone-treated water be discarded improperly (Budd et al. 2015; Mlalila et al. 2015). Although studies show that hormones such as MT do not persist in the fish tissues 24-72 h following uptake (Rinchard et al. 1999), hormone application to commercial food fish is still prohibited in India,

Ecuador, Costa Rica, European Union countries, and some sub-Saharan African countries

(Mlalila et al. 2015). However, the United States allows the use of sex reversal hormones in commercial food fish with an Investigational New Animal Drug (INAD) permit (Mlalila et al. 2015), and no countries have regulations regarding hormonal sex reversal of broodfish (Budd et al. 2015).

Environmental sex reversal

Environmental manipulation has been used in addition to hormonal treatment to induce sex reversal in fishes. Environmental variables have been shown to control sex determination in more than 60 fishes across diverse families (Baroiller et al. 2009; Budd et al. 2015). Although extreme salinity, pH, density, social status, and hypoxia have all been shown to influence sexual differentiation of various fish species, temperature extremes have been the most prevalent environmental factor governing sexual differentiation. According to Baroiller et al. (2009), there are three possible types of environmental sensitivity to sex differentiation:

1. Environmental factors have little influence on sex differentiation (e.g., Rainbow

Trout, Common Carp Cyprinus carpio, Medaka Oryzias latipes).

2. Fish are highly sensitive to one or two specific environmental factors, such as

temperature or pH (e.g., tilapias, Pejerrey Odontesthes bonariensis).

3. Fish are highly sensitive to a multitude of environmental (biotic and abiotic)

factors (e.g., Zebrafish Danio rerio)

The first evidence of environmental influence on sex differentiation in a gonochoristic fish was presented by Conover & Kynard (1981) in the Atlantic Silverside Menidia menidia.

Since this time, several other species have been shown to be affected by high

11 temperatures that resulted in a disproportionately large numbers of males, such as

Zebrafish (Abozaid et al. 2012), tilapias (Oreochromis spp.) (Baroiller & Toguyeni 1996),

Pejerrey (Strüssmann et al. 1997), and Japanese Flounder Paralichthyes olivaceus (Kitano et al. 1999). Of the thermosensitive species, the response to temperature extremes may fall into one of three categories (Baroiller & D’Cotta 2001):

1. High temperatures increase the percentage of males and low temperatures either

decrease or have no effect on the percentage of males.

2. High temperatures increase the percentage of females and low temperatures

either decrease or have no effect on the percentage of females.

3. High and low temperatures increase the percentage of males, with intermediate

temperatures producing a 1:1 male to female ratio.

Most thermosensitive species fall into group 1, including species in the Atherinidae,

Poeciliidae, Cichlidae, Cyprinidae, and Pleuronectidae families, among others. Far fewer species belong to groups 2 (e.g., Channel Catfish, Patiño et al. 1996) and 3 (e.g., Japanese

Flounder, Yamamoto 1999). To our knowledge, there is currently no research addressing the influence of temperature on sexual differentiation in Yellow Perch.

Challenges for environmental sex reversal exist as well. While extreme temperatures may skew the population toward one sex, most fishes do not produce 100% monosex populations under these conditions, indicating the genotype is inhibiting complete sex

12 reversal in a manner not yet understood. There is also immense diversity in the types of stimuli and the effectiveness of the manipulations, making the development of specific procedures for even a single species complex (Budd et al. 2015).

Goals and Objectives

The ultimate goal of this project was to determine an effective procedure to produce an all-female generation of Yellow Perch through sex reversal. My research was conducted in the Aquaculture Lab at The Ohio State University, under the supervision of Dr. Konrad

Dabrowski. Percid aquaculture is the major research focus in his laboratory. My specific research objectives were as follows:

1. Determine the most effective exposure method for steroid hormone

treatment: immersion in a hormone bath, or incorporation of the hormone

into live Artemia nauplii tissues. To accomplish this, I:

 Fertilized eggs from two broodstock females using the sperm from two

broodstock males (one male per female).

 Incubated the eggs and raised the larvae through swim bladder

inflation.

 Separated the swim bladder-inflated fish into stagnant water tanks and

assigned them to Control, E2-Immersion, MT-Immersion, and MT-Diet

groups, in triplicate.

 Fed the Control, E2-Immersion, and MT-Immersion groups live Artemia

nauplii that had been fed an emulsion with no hormone, and fed the

MT-Diet treatment group nauplii that had been fed an emulsion

containing MT for a duration of three weeks.

 Subjected the E2-Immersion and MT-Immersion groups to a bath of the

appropriate hormone for four hours, after which they were returned

to fresh water.

 After a 20 day hormone phase, returned the fish to flow-through or

recirculating system for growout.

 Checked for spermiation via stripping during the following spawning

season (spring of 2016).

2. Determine the optimum time of initiation for hormone treatment: 12 mm or

14 mm total length. To accomplish this, I:

 Began hormone treatment at approximately 12 mm total length,

labeled Group 12 mm, for one parental pair and at approximately 14

mm total length, labeled Group 14 mm, for the second parental pair.

Hypotheses

For the present work, I hypothesized that:

1. Masculinization in Yellow Perch is more successful when fish are exposed via

dietary treatment as opposed to immersion treatments.

2. Sex reversal in Yellow Perch is more successful when initiated at 12 mm total

length compared to 14 mm.

3. Fish populations exposed to water temperature in excess of 22o C beginning

before and extending throughout sexual differentiation would be skewed toward

males.

Although other researchers have conducted sex reversal studies on this species, they were unsuccessful because they started too late. Yellow Perch are a more complicated species on which to conduct sex reversal because sexual differentiation begins before the transition to formulated feeds (Best 1981). Prior to my work, there have been no studies that have attempted to masculinize Yellow Perch through incorporation of the hormone into live Artemia tissues, a process that was demonstrated by Stewart et al. (2001). This would allow for the sex reversal process via diet treatments to begin before sexual differentiation. If aquaculturists are able to produce all-female populations and subsequently increase their overall yield, there will be a greater supply, and the wild stocks will not be as pressured by harvest as they are currently. An increase in the profitability of the business may also cause more people to be interested in owning their own facility, leading to an even greater supply of farm-raised Yellow Perch.

Chapter 2: Determination of the Optimum Timing and Exposure Method for Hormonal

Sex Reversal in Yellow Perch

Introduction

To increase the profitability and popularity of farm-raised Yellow Perch, methods must be developed to increase the proportion of female fish. Yellow Perch exhibit sexually dimorphic growth, with females growing faster and larger than males, making them more valuable in aquaculture. An all-female population of Eurasian Perch reared in a constant temperature of 23o C weighs approximately 30% more than a 1:1 male to female population after their first year (Rougeot 2015). This translates to a substantial profit difference for monosex female populations. Sex reversal is one promising area of research to achieve all-female populations. The Perca genus is characterized by a genetic sex determination system with female homogamety (Rougeot et al. 2002), similar to humans.

So, by obtaining fish that are genetically female and phenotypically male (known as

“neomales” or “masculinized females”), sperm donors can be produced that can only contribute an X chromosome to progeny (Singh 2013). When combined with the X eggs from the mothers under normal conditions, they will generate an entire generation of all- female Yellow Perch.

Three major factors contribute to the success of sex reversal in fish: dose, time of application, and duration of hormonal treatment (Rougeot et al. 2002), all of which vary depending on species (Singh 2013). Initiation of hormone exposure must be before the onset of morphological features of sexual differentiation to produce fully functional sex- reversed fish. The dose and duration of treatment must also not be too high, causing sterilization, or too low, resulting in incomplete or no sex reversal (Rougeot et al. 2002).

Malison et al. (1986) conducted an experiment attempting to reverse sex of Yellow Perch juveniles ranging from 20 to 35 mm TL by using two diet-administered hormones: estradiol-17β (E2) and 17α-methyltestosterone (MT). The authors also conducted histological analyses of fish that were not exposed to hormones (control) to determine the relationship between age, body length, and sexual differentiation in this species.

Malison et al. (1986) found that hormonal treatment must be initiated before the fish reach 16 mm total length (TL) for the development of fully-functional gonads to occur.

Although all MT-treated groups displayed approximately 50% fish with paired testes (i.e., normal male morphology) and 50% sex reversed or intersex fish, no sex-reversed fish displayed functional sperm ducts. Additionally, the gonadosomatic index (GSI) of the masculinized females, which displayed a single ovotestis, were significantly lower than the ovary of controls (Table 1; Malison et al. 1986). Interestingly, groups exposed to the highest doses of MT (30 and 60 μg MT/g feed) displayed a significantly lower GSI of the ovotestis than those fed the lower doses (1.5 and 15 μg MT/g feed). Control fish GSIs were also significantly greater than those of both sexes for every E2-treated group. The authors concluded that “sex-reversed” fish were too large at the onset of hormone exposure, and

17 therefore displayed abnormal gonad morphology. Although masculinized females produced sperm, it was not possible to collect the sperm without sacrificing the fish, and were therefore deemed “non-functional males”. Sperm collected from the sacrificed fish did, however, successfully produced a monosex female population (Malison et al. 1986).

Table 1. Gonadosomatic indices of Yellow Perch fed diets containing different levels of methyltestosterone (MT) or cholesterol (C, control) for 84 days.

Steroid and dose Single Paired Testes Single Ovary (μg/g diet) Ovotestes C 0 0.118 ± 0.009ab ----- 0.446 ± 0.079a C 120 0.117 ± 0.009ab ----- 0.362 ± 0.043a MT 1.5 0.182 ± 0.043a 0.120 ± 0.024 0.060 ± 0.101b MT 15 0.120 ± 0.036a 0.080 ± 0.022 0.014b MT 30 0.084 ± 0.020b 0.028 ± 0.007 0.009 ± 0.002c MT 60 0.065 ± 0.015b 0.016 ± 0.005 0.026 ±0.008c Numbers are expressed as mean ± SE. Different letters next to the numbers within a column indicate a significant difference between groups (P < 0.01). Adapted from Malison et al. (1986).

In a later study on Eurasian Perch, females were treated with MT dissolved in 95% ethyl alcohol and mixed into the formulated diet (Rougeot et al. 2002). The purpose of this study was to determine an effective concentration, treatment duration, and fish size at initiation for hormonal sex reversal in this species. Rougeot et al. (2002) found that of the dosages, durations, and initiation times tested, 40 mg MT/kg diet at 70 mg initial body weight of fish for a duration of 30 days was the only effective method to produce masculinized females. It was later shown that sperm density and motility from Eurasian

Perch neomales did not differ from that of the normal males. Therefore, it is likely that

18 they would produce normal fertilization rates while still resulting in an all-female generation (Rougeot et al. 2004). However, these neomales also did not have functional sperm ducts, and the sperm had to be collected by dissection and maceration of the testes, as in the neomales produced by Malison et al. (1986). Further research must be conducted for the ability to routinely produce monosex female Yellow Perch populations.

Although there are several methods of introducing sex reversal hormones, two are practiced commonly due to their relative ease: incorporation of the hormone into the diet and immersion of the fish in a hormone bath. Sex reversal in Yellow Perch is particularly difficult compared to other commonly sex reversed fish (e.g., Nile Tilapia) because the onset of sexual differentiation in Eurasian and Yellow Perch occurs before juveniles begin accepting formulated feeds (Best 1981; Kestemont et al. 2015; Ljunggren et al. 2003).

However, Stewart et al. (2001) demonstrated the ability of Artemia nauplii to be enriched with steroid hormones that could be ingested.

The goal of my research was to establish the optimum method for introduction of the masculinizing hormone (diet or immersion treatment) and to determine the effectiveness of hormonal sex reversal in Yellow Perch at initial sizes of 12 mm versus 14 mm TL. My hypotheses were:

1. Groups of Yellow Perch juveniles exposed to MT through the diet will have a

higher percentage of functional males compared to those exposed to MT by

immersion due to more homogenous exposure to the hormone of all fish

despite variation in feed intake/acceptance.

2. Groups of Yellow Perch juveniles that underwent hormone exposure

beginning at 12 mm TL will have a higher percentage of functionally sex

reversed fish compared to those exposed to the hormone beginning at 14 mm

TL.

Methods

Fish in this experiment underwent three phases: fertilization and larval rearing, hormone exposure, and growout (Figure 2 ).

Figure 2. Flow chart with the three pha ses of the experiment: fertilization and larval rearing, hormone exposure, and growout. Larvae were separated into the four treatment groups, each in triplicate, for the hormone exposure phase. Fish remained separated by treatment group and, for the 12 mm Group, also by replicate following hormone exposure. Eventually, all fish were fin clipped by treatment group and for the 12 mm Group, also tagged with a visible implant elastomer by replicate, and combined into one common growout tank. This design was fol lowed by each size group separately, so the two were never combined.

Gamete collection and incubation

Broodstock Yellow Perch (4 years old) from the Aquaculture Lab at The Ohio State

University (Columbus, Ohio) were placed on an accelerated photothermal regime and injected with luteinizing hormone-releasing hormone analog (LHRHa; Sigma-Aldrich, St.

Louis, MO), first with a priming dose (10 μg/kg) on 11 March 2015, and again with a higher dose (100 μg/kg) on 13 March 2015 to induce ovulation. The gametes from one female

(229 g post-strip, GSI 23.6%) in this group were used (“Group 14 mm”). The female was anesthetized in tricaine methanesulfonate (MS-222) at a concentration of 75 mg/L water and stripped on 17 March 2015. The female used to generate the second group (“Group

12 mm”) was an age-3 female (228 g post-strip, GSI 27.6%) that was not related to the fish that produced Group 14 mm. The Group 12 mm female was obtained from Mill Creek

Perch Farm, LLC, where it was kept in a pond throughout its life. This female was anesthetized and stripped without hormone injection on 11 April 2015. The eggs were kept in a dry, air-tight plastic container and floated in 8o C water for no more than one hour before in vitro fertilization. Egg quality/ribbon firmness and color were qualitatively evaluated immediately after stripping and were similar for both groups.

Sperm used for the 14 mm Group was collected and pooled from three age-1 males and cryopreserved in 100-µL pellets on 12 February 2015 according to methods described by

Ciereszko & Dabrowski (1993). The appropriate number of pellets were thawed immediately prior to fertilization. Sperm used for Group 12 mm was collected into an

Eppendorf tube from three age-4 males and immediately used to fertilize the eggs. For both groups, 12.5 µL sperm was added per 1 g of eggs, followed by 0.8 mL of activating solution (4.4 g NaCl/L water) per 1 g eggs.

Embryos of the 14 mm Group were incubated in a vertical California tray incubation system at approximately 13o C. Those of the 12 mm Group were incubated in rectangular flow-through tanks at approximately 15o C. Both groups were force-hatched by siphoning the embryos into a bucket before they were transferred to a recirculating aquaria system

(Figure 3).

Larval rearing

Larvae of the 14 mm Group were transferred to a recirculating aquaria system 10 days post fertilization (DPF). The transferred larvae were fed rotifers (Brachionus spp.) and

Artemia nauplii (Premium Grade Brine Shrimp Eggs; Brine Shrimp Direct, Ogden, Utah) when they reached an appropriate size. Tanks were cleaned and mortalities were counted and recorded twice daily at 07:30 and 17:00. Light was maintained at 15 hours light to 9 hours dark. Nannochloropsis spp. algae paste (Nanno 3600TM Instant Algae®; Reed

Mariculture, Campbell, CA) was also added three times daily to increase turbidity to 10-

15 NTU (Grayson 2014). Lastly, salt was added twice daily to maintain the system around

3 ppt to increase the survival of live feeds. The larvae were fed decapsulated Artemia cysts as feed along with live Artemia nauplii between 11 and 23 DPF. Larvae of the 12 mm

Group were also transferred to a recirculating aquaria system at 10 DPF and received the same care as those of the 14 mm Group. However, these larvae were only fed live Artemia nauplii.

Enrichment of Artemia nauplii

Live Artemia nauplii were exposed to an enrichment approximately 24 hours after the beginning of the hydration process (Figure 4). The enrichment procedure was slightly modified from the methods of Grayson (2014) to include 1 mL MT dissolved in 100% ethanol (or 1 mL of 100% ethanol without hormone, in the case of the control diet) instead of the polyunsaturated fatty acid mixture. The ethanol or ethanol/MT mixture, 0.4 mL

23 chicken egg yolk, 0.5 mL Nannochloropsis spp. algae paste, and 1 mL ethyl ester fish oil

(AlaskOmega EE600200M®; Organic Technologies, Coshocton, Ohio) were homogenized in 35 mL fresh water for two minutes. This preparation was then added to the Artemia nauplii growth medium. The amounts of the ingredients listed were based on the requirement of Artemia necessary for a single treatment group to be fed at a rate of 20% wet body weight/day (Stewart et al. 2001). The nauplii were allowed to incubate in the enrichment 24 hours after the cyst hydration process began and for a duration of at least eight hours before being fed to fish. Uneaten Artemia was properly discarded 48 hours after the Artemia cyst hydration process began.

Sex reversal

At 26 DPF (Figure 5), larvae from the 14 mm Group were randomly divided and transferred to twelve 6- L stagnant water containers at a density of 100 fish per container

(Figure 6A) . Larvae were selected for swim bladder inflation at the time of stocking. Each tank had its own aeration and was maintained at 21.2 ± 1.1 o C. The tanks were also kept at 3 ppt salinity to extend the lifespan of live feeds . A small amount of Nannochloropsis sp p. algae paste was added to the tanks to increase turbidity and potentially reduce aggression amongst the larval fish at the time of stocking and shortly thereafter (Figure

6B) . The mean total fish length was 14.3 ± 1.8 mm (mean ± standard deviation) at the time of stocking.

Larvae from Group 12 mm were also selected for swim bladder inflation, randomly divided, and transferred to the same 6-L containers at a density of 60 fish per container at 23 DPF. These fish were kept in the same salinity (3 ppt), temperature (21.7 ± 0.7 o C) ,

25 and algae-induced turbidity as described above. The mean total fish length was 12.1 ±

0.44 mm (mean ± standard deviation) at the time of stocking.

A B

The 12 tanks of each group were randomly assigned one of four treatments, each with three replicates. The treatments were control, MT-Diet, MT-Immersion, and E2-

Immersion (Table 2). The control, MT-Immersion, and E2-Immersion groups were fed live

Artemia nauplii that were exposed to the control emulsion, while the MT-Diet group was fed Artemia that were fed an emulsion enriched with MT. All fish were fed these diets for a total of 20 days. MT-Immersion and E2-Immersion were exposed to MT and E2, respectively, through a 4 h hormone bath at a concentration of 400 µg/L water during the

20 days of enriched feeding. A stock solution of 24 mg hormone/10 mL 100% ethanol was covered in aluminum foil and kept in a refrigerator. One milliliter of the appropriate stock solution was added to each tank to achieve the desired hormone concentration at the time of each immersion treatment. The hormone treated water was removed, and the

26 fish were returned to clean, untreated water at the conclusion of the immersion treatment. The 14 mm Group underwent four immersion treatments, at 25, 29, 40, and

46 DPF. The 12 mm Group underwent the same number of immersion treatments at 24,

28, 32, and 38 DPF. Survival during this time period was tracked, and water quality parameters (temperature, dissolved oxygen, pH, ammonium concentration, and ammonia concentration) were monitored daily (Table 3 and Table 4).

Table 2. Summary of the hormone exposures for both Group 14 mm and Group 12 mm. Treatment Hormone Dose Treatment Duration Control N/A N/A MT-Diet 40 mg MT/kg diet 20 days MT-Immersion 0.4 mg MT/L 4 treatments last 4 hours each over 20 days

E2-Immersion 0.4 mg E2/L 4 treatments last 4 hours each over 20 days

Table 3. Water quality parameters during the hormone exposure phase of Yellow Perch larvae in Group 14 mm.

o + Treatment Temperature ( C) O2 (mg/L) pH NH4 (mg/L) NH3 (mg/L) Control 21.5 ± 1.3 8.76 ± 0.53 8.13 ± 0.08 3.30 ± 0.96 0.20 ± 0.07 MT-Diet 21.4 ± 1.4 8.76 ± 0.41 8.14 ± 0.07 3.36 ± 0.90 0.21 ± 0.08 MT-Immersion 21.4 ± 1.3 8.42 ± 0.99 8.11 ± 0.12 2.76 ± 0.78 0.16 ± 0.06

E2-Immersion 21.4 ± 1.3 8.63 ± 0.74 8.11 ± 0.10 2.84 ± 0.84 0.17 ± 0.06

Total 21.4 ± 1.3 8.63 ± 0.74 8.12 ± 0.09 3.04 ± 0.91 0.18 ± 0.07 Values are given as mean ± standard deviation. No values within a column differed significantly (ANOVA, P < 0.05).

Table 4. Water quality parameters during the hormone exposure phase of Yellow Perch larvae in Group 12 mm.

o + Treatment Temperature ( C) O2 (mg/L) pH NH4 (mg/L) NH3 (mg/L) Control 21.8 ± 0.7 8.45 ± 0.56 8.02 ± 0.09 3.02 ± 0.83 0.14 ± 0.04 MT-Diet 21.8 ± 0.6 8.52 ± 0.3 8.19 ± 0.13 2.98 ± 0.85 0.12 ± 0.03 MT-Immersion 21.8 ± 0.6 8.61 ± 0.43 7.96 ± 0.13 2.58 ± 1.76 0.10 ± 0.03 E2-Immersion 21.58 ± 0.7 8.53 ± 0.40 8.53 ± 0.40 2.53 ± 1.02 0.11 ± 0.04

Total 21.8 ± 0.4 8.53 ± 0.43 7.98 ± 0.12 2.78 ± 1.19 0.12 ± 0.04 Values are given as mean ± standard deviation. No values within a column differed significantly (ANOVA, P < 0.05).

Growout

The three 14 mm Group replicates of each treatment group were combined and then evenly and randomly distributed to three aquaria in a 12 aquaria recirculation system

(Figure 2) after the 20 day enrichment period. Replicates of this group were combined in order to eliminate growth differences based on the wide variety in density, caused by a mass mortality event at 34 DPF. Fish were treated with a chloramine-T solution (5 mg/L) to combat a presumed outbreak of a bacterial disease. The replicates of the 12 mm Group juveniles had similar densities at completion of the hormone exposure phase, and so remained separated when they were transferred to the recirculating aquarium system.

The salinity of the recirculation system was decreased from 3 to 0 ppt over three days.

Fish of both groups were trained to accept a formulated diet (Otohime B1; Aquasonic Ltd.,

Wauchope, Australia). At 134 DPF (3.4 ± 1.5 g wet weight), Group 14 mm fish were anesthetized in 50 mg/L MS-222, fin clipped by treatment and combined into one tank

28 for common growout. Group 12 mm fish were anesthetized and fin clipped by treatment following the same methods as Group 14 mm at 116 DPF (3.8 ± 0.6 g wet weight). Fish in this group were also anesthetized (50 mg/L MS-222) and tagged by replicate with a visible implant elastomer (Visible Implant Elastomer Tags; Northwest Marine Technology, Inc.,

Shaw Island, WA) and transferred to a round 400 L fiberglass tank common growout at

164 DPF. Fish in the 14 mm Group were not tagged with the elastomer because the three replicates were combined following the hormone exposure phase as a result of the differences in stocking density. All fish in each treatment were checked for spermiation by attempting to manually strip sperm on 25 January 2016 (314 DPF) and 1 February 2016

(296 DPF) for Group 14 mm and Group 12 mm, respectively. Spermiating fish were counted as “male” and non-spermiating fish were counted as “female/unknown” due to the potential presence of intersex fish or immature males.

Analysis of the F2 generation

The reproductive capability of the masculinized females/neomales in the F1 generation

(Group 12 mm and Group 14 mm) was evaluated by determining fertilization rate and embryo survival to 48 HPF and 10 DPF of their progeny (F2 generation). On 7 March 2016, seven large and seven small fish from both Group 12 mm (23.0 ± 1.4 g and 8.8 ± 1.0 g, respectively) and Group 14 mm (38.6 ± 8.7 g and 11.7 ± 1.3 g, respectively) were PIT tagged. The tags and the needle were soaked in 100% EtOH before and between uses, and the injection site was rubbed with the EtOH following injection.

Eggs were collected from an age-3 female (266 g post-strip, GSI 22.0%) that had been raised in the Aquaculture Lab on 10 May 2016. The female was anesthetized in 50 mg/L of MS-222 before eggs were collected by stripping. Eggs were placed into a dry, sealed plastic container and floated in 15o C water until fertilization.

Sperm was collected from large and small control males that had been previously PIT- tagged in both Group 12 mm (seven large, three small males) and Group 14 mm (three large, three small males). Sperm samples were held in 1-mL Eppendorf tubes and kept on ice until use (45 to 60 min following collection). An outside control male (age-3) was also used to fertilize a portion of eggs to verify their quality. This male was part of the broodstock held at the Aquaculture Lab.

Fertilization began one hour after eggs were stripped from the female. Each male was used to fertilize 4.5 g of eggs, with the exception of the outside control, which was used to fertilize 1.8 g. Although the original plan was to use 12.5 μL of sperm and 0.8 mL of activator (4.4 g NaCl/L water) per 1 g of eggs, five of the 16 males did not produce enough sperm to achieve this level. Activator was added directly into the Eppendorf tube with the milt, and the entire volume was transferred to the eggs to minimize the waste of sperm for these males. All other sperm samples were added directly to the eggs, and activator was added last.

Embryos were incubated in a vertical tray incubation system at 10o C. Each group was housed in a basket made of PVC pipe with a 600 μm mesh bottom, which was mixed several times daily to ensure water movement over the eggs. Photographs were taken using a dissecting microscope across several fields of a sample from each group at three stages of development: two cell stage (fertilization rate), 48 HPF, and 10 DPF.

Fertilization/survival in each group was evaluated at a later date from these photographs.

Statistical analyses

Differences in the variances between groups were analyzed with a Levene’s test.

Differences in the sex ratio and survival rates between groups with equal variance were analyzed using a one-way ANOVA and Tukey HSD test. Survival rates between two initiation sizes of one treatment with unequal variance were analyzed with a t test.

Survival rates of treatments at one initiation size with differences in variance were analyzed using a Kruskal-Wallis analysis. Survival differences between the two size groups of one treatment that displayed different variances were analyzed with a Wilcoxon test.

Deviations from the expected 1:1 male to female sex ratio were analyzed using a chi- squared test. Statistical analysis of the fertilization (2-cell) and further embryo survival rates could not be completed for the control sample due to the presence of only one replicate. All tests were carried out using JMP (Version 12), and significance was accepted at P < 0.05.

Results

Method of Exposure

Group 14 mm

Survival rates at the end of the hormone exposure phase were highly variable in this group due to a disease outbreak in some tanks but not others. The variance between treatment groups did not differ (Levene’s test, P = 0.0692). No groups differed in survival significantly from others or the control (ANOVA, F ratio = 0.81, P = 0.523) (Figure 7).

Figure 7. Survival rates during the hormone exposure phase of Yellow Perch exposed to one of two hormones (MT or E2) by one of two methods (diet or immersion), beginning at either 12 mm or 14 mm TL. Survival rates between treatments within one initiation size did not significantly differ for either Group 12 mm or Group 14 mm. Error bars represent standard deviation. There were no significant differences in survival rate (P < 0.05).

Variances between groups in percentage of spermiating males were not different (P =

0.7552). There was no significant deviation from the expected sex ratio in the MT-Diet (χ2

2 2 = 1.88; P = 0.168), MT-Immersion (χ = 1.32; P = 0.248), and E2-Immersion (χ = 0.07; P =

0.786) treatment groups (Figure 8). The sex ratio of these treatments also did not differ from each other. However, the control treatment group (89.29%) displayed a significantly higher percentage of spermiating males than expected (χ2 = 34.57; P < 0.0001) (Figure 8).

The percent spermiating males in the control group was not higher than the MT-Diet (P =

0.050) group, but it was significantly higher than the MT-Immersion (P = 0.001) and E2-

Immersion (P = 0.002) groups.

Group 12 mm

Survival rates at the 12 mm initiation size were much less variable compared to Group

14mm, but the variance between the four treatments was different (Levene’s test, P =

0.0046). Survival rates did not differ between any groups (Kruskal-Wallis, P < 0.05) (Figure

7). No disease was observed in this experiment during the hormone exposure phase.

The control group displayed a higher percentage of spermiating males than the E2-

Immersion group (P = 0.002), but did not differ from either the MT-Diet (p = 0.977) or the

MT-Immersion (p = 0.999) groups (Figure 8). However, all four groups displayed a significant deviation from the expected 1:1 sex ratio, with control, MT-Diet, and MT-

Immersion groups containing 88.3 ± 6.8% (χ2 = 63.04; P < 0.0001), 92.1 ± 3.4% (χ2 = 97.96;

P < 0.0001), and 86.2 ± 4.6% (χ2 = 70.74; P < 0.0001) spermiating males, respectively. The

E2-Immersion group (59.6 ± 5.3% spermiating males), which displayed fewer spermiating males than the Control, MT-Diet (P = 0.001), and MT-Immersion (P = 0.003) groups, still deviated from the expected sex ratio (χ2 = 5.74; P = 0.017).

a * a * ab * a *

c * bc c

Time of Initiation

The MT-Diet (P = 0.010) and MT-Immersion (P = 0.001) treatment groups of the 12 mm

Group displayed a significantly higher percentage of spermiating males than those of the

14 mm Group (Figure 8). The percent spermiating males in the control and E2-Immersion

34 treatment groups did not significantly differ between the two size classes of fish (P =

0.876).

The variance in survival differed between the two initiation sizes of the MT-Diet (P =

0.0214) and MT-Immersion (P = 0.0225), but not control (P = 0.0955) or E2-Immersion (P

= 0.1055). The survival of fish in both MT-Diet (P = 0.0809) and MT-Immersion (P = 0.0765) treatments did not differ between initiation sizes (Wilcoxon test). Survival of the two size classes in each treatment group did not differ between the control (t(2.64) = -0.56, P =

0.6166) and E2-Immersion (t(2.67) = -2.22, P = 0.1237) groups either. (Figure 7).

Evaluation of F2 Generation

Fertilization rates ranged from 21.0% to 56.3% (n = 16) among embryos of PIT-tagged fish

(31.8 ± 10.4%, mean ± standard deviation), while the control group had a fertilization rate of 52.0%. Mean survival to 48 HPF in the treatment group (38.3 ± 12.7%) was similarly low compared to the control group (58.6%). However, survival to 10 DPF was more similar between the two groups (treatment: 42.6 ± 14.2%; control: 48.6%).

Discussion

Our results indicate that hormonal masculinization in Yellow Perch is effective when initiated at 12 mm TL in both hormone delivery methods (Figure 8). Both size groups

35 began hormone treatment before the reported onset of sexual differentiation in this species at 16 mm TL (Malison et al. 1986). Therefore, Group 12 mm may have had a greater proportion of functional males due to germ cells that began physiological differentiation before any histological indication could be observed, as suggested in tilapia by Nakamura (2013). The initiation of hormone treatment in the present study was earlier than those presented by Malison et al. (1986) and Rougeot et al. (2002), and was likely the reason for the increase in functional males. Similarly, hormonal masculinization of gynogenetic Northern Pike beginning 26-30 mm TL and 30-41 mm TL resulted in 83% male/17% intersex fish and 76% male/24% intersex fish, respectively (Luczynski et al.

2003).

The immersion dose chosen for the present study was lower than the highest successful dose for Nile Tilapia masculinization (Gale et al. 1999), but higher than that in Chinook

Salmon masculinization (Baker et al. 1988). Baker et al. (1988) also found that longer treatment durations resulted in a higher proportion of males, which lead to the decision to expose our immersion groups to four separate hormone baths of four hour duration each. In gonochoristic fishes, sex reversal techniques must extend throughout sexual differentiation (Nakamura 2013). Therefore, the immersion groups received additional hormone baths to cover the extended labile period of Yellow Perch, instead of only two as were proven successful in Chinook Salmon (Baker et al. 1988) and Nile Tilapia (Gale et al. 1999). The hormonal dose for masculinization by dietary treatment (40 mg MT/kg dry weight Artemia nauplii) was chosen based on the findings of Rougeot et al. (2002), which

36 stated that hormonal masculinization was successful in Eurasian Perch fed 40 mg MT/kg formulated feed. The failure to produce functional Eurasian Perch neomales at this dose was likely a result of the initiation size and not the hormone dose, which our findings support.

The lack of a difference in survival between the four treatment groups within each initiation size follows the findings of Vera Cruz & Mair (1993) in masculinization of Nile

Tilapia, those of Baker et al. (1988) in masculinization of Chinook Salmon, and those of

Strüssmann et al. (1996) in the feminization of Pejerrey. Initiation size also did not have an effect on survival during the hormone exposure phase (Figure 7). However, the disease outbreak in Group 14 mm caused an unusually high amount of variability in survival rates, so these results are not yet conclusive. Initiation size also had no effect on survival in masculinized Eurasian Perch (Rougeot et al. 2002).

Additionally, there was no difference in masculinization success between the diet treatment and the immersion treatment. However, these conclusions also can only be made with some reservation due to the significant bias toward males in the control groups of both size classes. This deviation suggests a variable other than exogenous hormones may have influenced sexual differentiation.

The highly male-biased sex ratios in control groups at both 12 and 14 mm initiation sizes is completely counterintuitive. Rearing techniques (e.g., turbidity increase, salt addition,

Artemia and rotifer sources, incubation temperatures) used in this study were not different than those used in previous years in this laboratory, which had not resulted in skewed sex ratios. There are two possible scenarios that might have led to the observed male-biased sex ratios (Figure 8). First, there is potential for some environmental variable to have inadvertently influenced sexual differentiation. Some fish, such as Atlantic

Silverside, display environmental sex determination and rely solely on environmental variables for sexual differentiation (e.g., temperature, in Atlantic Silverside) (Baroiller et al. 2009). Several other species typically display genetic sex determination, but that system may be overridden when fish are exposed to extreme environmental conditions.

Environmental variables have been shown to influence sex differentiation in more than

60 fishes across diverse families. Although variables such as pH, salinity, and stocking density have been shown to control sexual differentiation, the most dominant variable is temperature (Baroiller et al. 2009). No species in the Percidae family is documented to be thermosensitive in sexual differentiation, but high temperatures have been found to masculinize at least one species in three families belonging to the same order as Yellow

Perch (Baroiller & D’Cotta 2001). Additionally, temperatures of containers housing the fish in this study during sexual differentiation (21.2 ± 1.1o C and 21.7 ± 0.7o C for Group

14 mm and Group 12 mm, respectively) were considerably higher than Lake Erie temperatures during this period (12 – 16o C, as reported by Reichert et al. (2010)).

Therefore, it is possible that the high temperatures used in the present study, intended to maximize growth, inadvertently induced masculinization.

It is also possible that the skewed sex ratios seen in the control groups at both initiation sizes were caused by other environmental variables that influence sexual differentiation.

For instance, Green Swordtail Xiphophorus helleri populations favor males under acidic conditions (pH 6.2) and females under basic conditions (pH 7.8) (Rubin 1985). Water acidity has also been shown to influence sexual differentiation in another poeciliid, the

Blackbelly Limia Limia melanogaster (Römer & Belsenherz 1996), as well as the Kribensis

Cichlid Pelvicachromis pulcher (Oldfield 2005) and five Apistogramma species (Römer &

Belsenherz 1996; Rubin 1985). Although pH during the hormone exposure phase was basic for both the 12 mm Group (7.98 ± 0.12) and the 14 mm Group (8.12 ± 0.09) (Table

3 and Table 4), these values are within the believed pH range of another percid, the

Walleye (6.0-9.0) (Schnieder et al. 2002). However, little research has been conducted on the upper pH limit in Yellow Perch.

There is also evidence to suggest that hypoxia may influence sexual differentiation in

Zebrafish. According to Shang et al. (2006), hypoxic conditions (0.8 mg O2/L) result in male-skewed populations (74.4 ± 1.7% males) of Zebrafish compared to normoxic conditions (5.8 mg O2/L) (61.9 ± 1.6% males). It is unlikely that dissolved oxygen content alone influenced sex ratios in the present study, considering the oxygen concentrations for both the 12 mm Group (8.53 ± 0.43 mg/L) and the 14 mm Group (8.63 ± 0.74 mg/L)

(Table 3 and Table 4) during the hormone exposure phase were well above the lethal limit for Yellow Perch (~2 mg/L) (Feiner & Höök 2015).

One of the more difficult environmental variables to evaluate for sex control is growth rate because it can be influenced by other factors, such as temperature, hypoxia, and stocking density, all of which have been shown to affect sexual differentiation in some species (Baroiller et al. 2009). Paull et al. (2009) found a significant deviation towards males in slow growing Roach (Rutilus rutilus) populations, so it is possible that the growth rate of the fish in this experiment resulted in the male-skewed populations, and not some other environmental factor(s).

Another possible cause for the male-skewed control populations is the potential presence of some endocrine active substance in the control Artemia nauplii enrichment. A previous publication that had used this enrichment recipe (Grayson 2014) did not raise the fish to a large enough size to determine the sex ratio, so the masculinizing capacity of this particular enrichment has not been evaluated. Lastly, the potential for additive effects of genetic, environmental, and/or exogenous substances (i.e., steroid hormones or endocrine active substances in the diet) create a substantial number of causes for the skewed sex ratios observed in these experiments. Penman & Piferrer (2008) present the example of European Seabass Dicentrarchus labrax as having additive effects of temperature and genetics influencing sexual differentiation. In the present study, environmental factors such as temperature and pH may have together resulted in skewed sex ratios.

There are some limitations in design of the present study. First, only one dam was used for each size group of fish. Future studies investigating optimum initiation sizes for hormonal treatment of Yellow Perch would benefit from having the fertilized egg ribbon from each female split into multiple sections, each of which to begin hormone treatment at different larval sizes. Each section would also be separated into treatment groups (e.g., control and multiple hormone concentrations). This design would then be repeated for several male/female pairs.

Regardless of the driving cause of the masculinization, to our knowledge, this is the first demonstration of production of monosex female populations of Yellow Perch from functional neomale sperm donors. Malison et al. (1986) and Rougeot et al. (2002) succeeded in producing monosex female progenies, but males had to be sacrificed in order to collect sperm. This process destroys any hope to collect sperm and produce additional generations in the future. The ability to maintain confirmed neomale sperm donors for future spawning seasons allows for greater economic feasibility due to limited needs for repeated masculinization and use of hormone treatments. Aquaculturists may elect to sex reverse a small portion of the fish in each spawning season to replace the aging broodstock. With the help of identification tags or isolation of the neomales (XX genotype), producers would no longer be required to test the sex ratio of the progeny of the hormone-exposed fish beyond the F2 generation. Additionally, cryopreserved sperm may allow for the production of all-female progenies at several facilities for many future generations.

Chapter 3: Exploration of the Potential for Temperature-Induced Sex Reversal in Yellow

Perch

Introduction

Despite the potentially profound benefits of understanding the influence of biotic and abiotic factors on sex determination and its application for explaining sex reversal in wild stocks of Yellow Perch (Marsden & Robillard 2004), little research has been conducted in this area for representatives of the Percidae family. Application of extreme temperatures before and during sexual differentiation has been proven to reverse sex of individuals of several species in diverse families, including Atherinidae, Poeciliidae, Cichlidae,

Cyprinidae, and Pleuronectidae (Abozaid et al. 2011; Baroiller and D’Cotta 2001; Ito et al.

2005). Fish species that exhibit temperature-induced sex control under extreme temperature conditions fall into one of three categories: (1) high temperatures increase the percentage of males and low temperatures either decrease or have no effect on the percentage of males, (2) high temperatures increase the percentage of females and low temperatures either decrease or have no effect on the percentage of females, and (3) high and low temperatures increase the percentage of males, with intermediate temperatures producing a 1:1 male to female ratio (Baroiller & D’Cotta 2001). Evidence

42 in some species is incomplete. For instance, according to Patiño et al. (1996), only one temperature (34o C) has a significant effect on the sex ratio of channel catfish, resulting in feminization. All other temperatures produce the expected 1:1 sex ratio.

Most thermosensitive species fall into group one, including all of the families previously listed. Far fewer species fall into group 2 (e.g., Channel Catfish (Patiño et al. 1996)) and group 3 (e.g., Japanese Flounder (Yamamoto 1999)). To our knowledge, the effects of temperature on sexual differentiation have not been evaluated for any species in the

Percidae family.

The previous study investigating optimum exposure methods and times of initiation for hormonal treatment to induce sex reversal in Yellow Perch, conducted in spring 2015, produced approximately 90% spermiating males in the control groups of both size classes

(12 and 14 mm total length), a large deviation from the expected 1:1 male to female sex ratio. In order to optimize growth during the time of hormone exposure, fish were subjected to temperatures of 21.2 ± 1.1o C or 21.7 ± 0.7o C for Group 14 mm and Group

12 mm, respectively. These temperatures are considerably higher than the temperature of Lake Erie when Yellow Perch undergo sexual differentiation (12-16o C, as reported by

Reichert et al. (2010)). High temperature during the larval phase might have altered sex ratio during the laboratory experiments. However, the impact on sex ratio may have been the result of the genetic makeup of the parental fish, or some contamination of the

Artemia source by environmental endocrine disruptors (McMaster 2001).

The purpose of this study was to determine the potential for high water temperatures during the period of sexual differentiation to cause masculinization in Yellow Perch. We hypothesized that temperatures in excess of 22o C, when initiated at the onset of sexual differentiation and extended throughout that period, would lead to a male-skewed sex ratio (i.e., production of “neomales”, XX-genotype males)

Methods

Experiment 1

Yellow Perch embryos were collected from three females (designated as A, B, and C) at

Mill Creek Perch Farms, LLC (Marysville, OH) when they were at 9 DPF. Fertilization took place in a single pond, so the male source cannot be determined. The embryos were transported in separate coolers with water, and temperature was maintained at 15o C during this time and throughout the rest of incubation. The fertilized egg ribbons were given a Povidone treatment (100 μL/L water) to reduce the transfer of diseases from the farm to the indoor facility after arrival at the Aquaculture Lab. Incubation took place in

40-L rectangular flow-through tanks for the remainder of the embryonic stage.

Hatching began at 11 DPF, and larvae were transferred to two recirculating aquarium systems on 14 DPF. Larvae of each female were randomly distributed to tanks in two

44 identical recirculating aquarium systems kept at different temperatures. Group A exhibited a much higher fertilization rate than the other two groups, so the larvae were stocked into two tanks in each system of 1,000 fish each. These groups were designated as A(1) and A(2). Larvae of groups B and C were each stocked into one tank in each system of 1,000 fish per tank, making a total of eight tanks. The cool system was maintained at

15.6 ± 1.9o C from 15-58 DPF, while the warm system was kept at 24.0 ± 1.3o C over the same time period. However, at 23 DPF, the warm groups were transferred to closed 6-L containers housed in a heated water bath to more consistently maintain temperature

(Figure 9). Nannochloropsis spp. algae paste (Nanno 3600TM Instant Algae®; Reed

Mariculture, Campbell, CA) and salt (3 ppt) were added to all containers to reduce aggressive interactions and cannibalism, and to extend the lifespans of live feeds, respectively. The warm groups were fed rotifers (Brachionus spp.) from 16-19 DPF and then Artemia nauplii (Great Salt Lake Artemia Cysts; Artemia International, Fairview, TX) from 20-61 DPF. Fish were then transitioned to a formulated feed (Otohime B2; Reed

Mariculture, Campbell, CA). The cool groups were fed rotifers at 17 DPF and transitioned to Artemia nauplii at 27 DPF, and then weaned to the same formulated feed as the warm groups at 127 DPF. All tanks were cleaned and water quality was tested daily.

The cool groups had experienced significant mortalities by 55 DPF, and were eliminated from the experiment. Warm groups were kept in flow-through tanks from 59 DPF through the remainder of the experiment.

Due to differential stocking densities, two of the warm groups were large enough to be sampled to conduct morphological sex determination earlier than the others. Groups B and C were sampled at 175 DPF, while both groups A(1) and A(2) were sampled at 201

DPF. Fish were anesthetized in an ice slurry and euthanized by pithing. Ten fish were sampled from each group except for Group B, which only had 5 fish due to high mortality caused by disease. All five of the fish in this group were sampled. Total weight (TW), TL, and gonad weight (GW) were recorded, and gonads were placed in 10% neutral buffered formalin for histological preparation. GSI was calculated from these measurements.

Mean growth differences, measured in mm/day, were calculated and analyzed using measurements taken throughout the experiment. Mean specific growth rate (% weight/day) was also calculated using the equation provided by Saillant et al. (2001). It

46 was not possible to determine individual growth rates by either metric due to the small size of the fishes and our inability to tag them. Therefore, the mean weights and lengths for each group were used to calculate both metrics, and consequently lack standard deviations.

Experiment 2

The progeny of control males from the 2015 hormonal sex reversal experiment were also subjected to two experimental temperatures. Eggs were collected from an age-3 female that were raised in the Aquaculture lab. The female was anesthetized in 50 mg/L of MS-

222 before eggs were collected by stripping. Eggs were placed into a dry, sealed plastic container and floated in 15o C water until fertilization one hour later.

Sperm was collected from large and small control males that had previously been PIT- tagged in both Group 12 mm (seven large, three small males) and Group 14 mm (three large, three small males). Sperm samples were held in 1-mL Eppendorf tubes and kept on ice until use. An outside control male (age-4) was also used to fertilize a portion of eggs to verify their quality. This male was part of the broodstock held at the Aquaculture Lab.

Fertilization began one hour after eggs were stripped from the female. Each male was used to fertilize ~4.5 g of eggs, with the exception of the outside control male, which was used to fertilize 1.8 g. Although the original plan was to use 12.5 μL of sperm/gram of eggs and 0.8 mL of activator (4.4 g NaCl/L water)/gram of eggs, five of the 16 males did

47 not produce enough sperm to achieve this goal. For these males, activator was added directly into the Eppendorf tube with the milt, and the entire volume was transferred to the eggs to minimize the waste of sperm. All other sperm samples were added directly to the eggs, and activator was added last.

Embryos were incubated in a vertical tray incubation system at 10o C. Each sample was housed in a basket made of PVC pipe with a 600 μm mesh bottom, which was mixed several times daily to ensure water movement over the eggs.

Embryos were force-hatched at 16 DPF (Figure 10) and stocked to the same recirculating aquarium system as the fish of experiment 1. The six baskets from the 12 mm Group

(three large males, three small males) with the highest hatch rate and all baskets from the 14 mm Group were chosen to stock. Embryos from the outside control male were discarded after the quality of eggs was evaluated. Aquaria were cleaned and mortalities counted and logged daily. Algae and salt were added to the system in similar fashion as experiment 1. Temperature was maintained at 15.6 ± 1.1o C. Rotifers (Brachionus spp.) were introduced at 20 DPF, and although food was observed in the gut immediately in some groups, others were not observed to eat after 27 DPF. These groups had nearly total mortality and were discarded. The remaining five groups were transitioned to Artemia nauplii at 29 DPF. For convenience, these groups will be referred to as male 1, 2, 3, 4, and

Approximately half of the population of each group of larvae sired by each male was transferred to an identical recirculation system containing warm water (23.1 ± 0.2o C) at

32 DPF to determine the potential for high temperature-induced masculinization. The cool system was maintained at 16.4 ± 1.0o C during this time. Fish were kept in the two systems until 56 DPF, when the warm water and cool water fish were transferred to a new recirculating aquarium system with the optimum temperature for growth (22.9o C).

Acclimation of the “cool” groups to the warmer temperature occurred at a rate of 1o C per day. The “warm” group of fish were transitioned to a formulated feed (Otohime B2) at 65 DPF, while the “cool” groups were transitioned to the same feed at 78 DPF. All fish remained in this system until they reached an appropriate size (>5 g wet weight) for morphological sex determination. Fish from each group were randomly selected, anesthetized in an ice slurry, and euthanized by pithing. Five fish were sampled from each group to save the majority of fish for further analyses (e.g., spermiation). TW and TL were recorded before the gonad was dissected, weighed, and placed in 10% neutral buffered formalin to be eventually prepared for histological analysis. Gonadosomatic index was calculated from these measurements.

Histological processing for both experiments was conducted by the Ohio State University

Comprehensive Cancer Center Pathology and Mouse Phenotyping Shared Resource.

Sections were cut at 5 μm thickness along the length of the gonad for samples weighing

0.15 g or less, while the remainder were cut at the same thickness, but transversely. All samples were stained with haematoxylin and eosin and were analyzed by light microscopy

49 for male and female germ cells. The stage of germ cell development was classified based on the descriptions for yellow perch provided by Blazer (2002).

Mean daily growth rates, measured in mm/day, were calculated (Graeb et al. 2004) and analyzed using measurements taken throughout the experiment. The growth rates for the five groups were averaged to obtain the mean and standard deviation in warm and cool water raised groups.

Statistical Analysis

Differences in variances were analyzed with a Levene’s test. Deviations from the expected

1:1 male to female sex ratio were analyzed using a chi-squared test. Differences between sex ratios and growth rates in warm and cool groups were evaluated using a two sample t-test. All statistics were conducted using JMP (Version 12), and significance was accepted at P < 0.05.

days - days -

gonads gonads lues are presented presented are lues Cool groups sampled for for sampled groups Cool Warm groups sampled for for sampled groups Warm degree ~3200 DPF, 169 158 DPF, ~3200 degree ~3200 DPF, 158

days - fertilization (DPF) onward, weighings and samplings were based based were samplings and weighings onward, (DPF) fertilization

days - days -

degree

Cool groups weighed and and weighed groups Cool 78 DPF, ~ 1100 degree 1100 ~ DPF, 78 transferred to formulated feed formulated to transferred

group fish group nt 2. From 56 days post days 56 From 2. nt

days - C)

o 58 DPF, 685 degree 685 DPF, 58 Warm groups weighed and and weighed groups Warm

65 DPF, ~1100 DPF, 65 Weighed and measured cool cool measured and Weighed transferred to formulated feed formulated to transferred 56 DPF 56 warm group fish group warm Fish combined to to combined Fish one system (23 one system Weighed and measured measured and Weighed 57 DPF, 820 degree 820 DPF, 57 C vs. C

o ion. ion.

DPF 32 29 DPF 29 16.4 ± 0.4 16.4 ±

Began feeding feeding Began transferred to warm warm to transferred o Artemia nauplii Artemia Sample of each group each group of Sample system (23.1 ± 0.2 ± (23.1 system standard deviat standard

± DPF 20 DPF 16

Hatching and and Hatching (15.6 ±1.1 (15.6 stocking to aquaria aquaria to stocking Began feeding rotifers feeding Began Figure 10. Important points in the timeline of Experime of timeline the in points Important 10. Figure Va groups. warm in rates growth increased by caused fish the of differences size the reflect to age of instead days degree on as mean

Fertilization 51

Results

Experiment 1

Juveniles in groups A(1), A(2), B, and C had sex ratios that did not differ from the expected

1:1 sex ratio (Figure 11). Microscopic examination of the gonads revealed normal male and female histology in 100% of the samples (i.e., no evidence intersex characteristics).

Of all the gonad samples examined histologically, 100% of the females were in the previtellogenesis stage and exhibited perinucleolar oocytes. Of the males sampled for histology, 66.7% were in early spermatogenesis with primarily spermatocytes and spermatids, with some spermatozoa present. The remainder were in mid- spermatogenesis, with roughly equal numbers of spermatocytes, spermatids, and spermatozoa. No samples displayed any indication of intersex tissues.

The specific growth rates of groups A(1), A(2), B, and C were 5.54%, 5.30%, 5.57%, and

5.80%, respectively, during 23-127 DPF. All groups increased in length at a rate of 0.5 mm/day over the same time period except for group C, which grew at a rate of 0.6 mm/day.

10 10 10

Figure 11. Percent males in four groups of Yellow Perch from three unrelated dams following exposure to high temperatures throughout the period of gonadal differentiation. Results showed no significant differences between groups (ANOVA, P = 0.05), as well as no significant deviations from the expected 1:1 male to female sex ratio (chi-squared test, P = 0.05). Numbers above columns indicate sample sizes.

Experiment 2

Variances in growth rates were not different between warm and cool groups (Levene’s, P

= 0.1391). Analysis of growth rates during gonadal differentiation of the warm (0.59 ±

0.09 mm/day) and cool (0.35 ± 0.05 mm/day) groups revealed a difference between the two (t(6.5) = 5.27, P < 0.01). Growth rates also differed between the warm (0.77 ± 0.07 mm/day) and cool (0.65 ± 0.06 mm/day) groups after gonadal differentiation (t(8.0) =

2.78, P = 0.02).

The variances of sex ratios did not differ (Levene’s, P < 0.05). Male 1 (χ2 = 1.80; P = 0.180), male 2 (χ2 = 0.20; P = 0.655), and male 3 (χ2 = 0.2000; P = 0.6547) did not produce progeny in the warm groups that differed from the expected sex ratio. However, males 4 and 5 both produced progeny populations biased toward females (χ2 = 5.0000; P = 00253). This pattern was followed in the sex ratios of the cool groups (Figure 12). The sex ratio of the warm and cool groups did not differ (t(8) = 0.19, P = 0.8548). The wet weights and gonadosomatic indexes of the sampled progenies also did not differ (Table 5 and Table

6).

* * * *

Table 5. Total weight (TW), gonadosomatic index (GSI), and percent females of Yellow Perch progenies (n = 5) produced by two control males from Group 14 mm (Chapter 2) that had been raised at 23.1 ± 0.2o C (Warm) or 16.4 ± 1.0o C (Cool). Both sires were in the “large” group of the two that were PIT tagged.

Male 1 Male 3 Warm Cool Warm Cool Male TW (g) 12.60 ± 0.01 16.77 ± 1.04 9.02 ± 3.84 9.17 ± 4.16 Male GSI (%) 0.08 ± 0.02 0.09 ± 0.00 0.12 ± 0.03 0.16 ± 0.05 Female TW (g) 18.46 ± 0.58 9.01 ± 3.11 11.87 6.74 Female GSI (%) 0.2 ± 0.02 0.19 ± 0.02 0.175 0.224 Values are given as mean ± standard deviation. Single values represent those from a single individual.

Table 6. Total weight (TW), gonadosomatic index (GSI), and percent females of Yellow Perch progenies (n = 5) produced by three control males from Group 12 mm (Chapter 2) that had been raised at 23.1 ± 0.2o C (Warm) or 16.4 ± 1.0o C (Cool). All sires were in the “small” group of the two that were PIT tagged.

Male 2 Male 4 Male 5 Warm Cool Warm Cool Warm Cool Male TW (g) 12.7 ± 8.17 2.68 ------Male GSI (%) 0.10 ± 0.03 0.153 ------Female TW (g) 15.77 ± 3.95 6.77 ± 1.86 9.87 ± 3.57 6.98 ± 3.52 7.08 ± 2.19 5.55 ± 2.35 Female GSI (%) 0.21 ± 0.02 0.27 ± 0.07 0.20 ± 0.02 0.26 ± 0.06 0.21 ± 0.03 0.28 ± 0.05 Values are given as mean ± standard deviation. Single values represent those from a single individual.

Discussion

The results indicated that the extremely high temperatures used in the present experiment (23-24o C) had no influence on gonadal differentiation of Yellow Perch.

However, it is possible that temperature-induced sex reversal does still exist in Yellow

Perch. The temperatures used in my study may have not been high or low enough to

55 influence sexual differentiation. Therefore, the skewed sex ratios in the control groups of the 2015 hormonal sex reversal experiments were likely overridden by other abiotic factors (e.g., hypoxia or pH), the presence of endocrine-active substances in the fish diet, or the additive effects of these factors and/or genetics. One smaller possibility is the influence of nitric oxide on sexual differentiation. Nitric oxide (NO) is formed as part of the denitrification process and has been identified as an aromatase (i.e., the enzyme responsible for converting testosterone to estrogen) inhibitor in human gonadal tissues

(Snyder et al. 1996). However, to our knowledge, there is no research concerning its influence on sexual differentiation in fish.

Garrett (1989) conducted a similar study on Largemouth Bass Micropterus salmoides to determine the success of hormonal sex reversal by diet treatments utilizing formulated feed versus live Artemia nauplii. The control Artemia-fed group in this experiment of consisted of 89% males. Although the author attributed the heavily skewed sex ratio to contamination of Artemia due to the close proximity of the hormone-treated incubations, it was not recognized at that time that female Largemouth Bass are heterogametic for sex

(i.e., this species displays a ZZ/ZW genetic sex determination system) (Glennon et al.

2012). This fact would significantly impact the sex ratio following hormone treatment. In a follow up study, the control Largemouth Bass had a normal sex ratio while the hormone treated groups displayed 100% monosex populations (Garrett 1989).

While several environmental factors have been shown to influence sexual differentiation in other fishes, more research is needed on Yellow Perch to explore, for instance, temperatures higher than 23o C, time of initiation of exposure, and/or duration of the high temperature treatment. Differential growth rates should also be explored as a means of controlling sex in Yellow Perch. Populations of Roach that grow rapidly during sexual differentiation display a greater proportion of females compared to their slower growing counterparts (Paull et al. 2009). Presumably, there are more males present in slower growing populations because a small body size limits fecundity in females, but small males are still able to reproduce fairly well (Paull et al. 2009). Our results suggest that this pattern is not followed in Yellow Perch, given the significant difference in growth rates between the warm and cool groups in experiment 2 and the lack of a difference in the resulting sex ratios among individual progenies. However, the temperature and stocking density of the two groups also differed and resulted in growth differences (Table 3; Table

4). A definitive conclusion cannot be made regarding the influence of growth rate on sexual differentiation in Yellow Perch from this experiment. Although it is well documented that female Yellow Perch grow faster and larger than males, that dimorphism is not significant until the fish are 15-20 g (Schott et al. 1978), well after sexual differentiation is complete.

Future Research

Given the large amount of unknowns in this research, several directions in follow up studies should be pursued. First, since the control and MT-treated Artemia incubations in the 2015 hormonal sex reversal experiments were not analyzed for hormone concentration, a follow up study should include MT accumulation and metabolization in

Artemia nauplii, and subsequently validate the dosage for inducing masculinization in larval Yellow Perch. Time of initiation perhaps no longer requires testing because the results presented here provide evidence that masculinization by either diet or immersion treatments is more effective when initiated at 12 mm TL compared to 14 mm, assuming that no selective mortality occurred during the hormone treatment phase. Additionally, this follow up study can focus on solely the immersion treatment since the 2015 results do not indicate a significant difference between diet and immersion methods. Immersion treatments are far less time and labor consuming than preparation of hormone-enriched

Artemia nauplii. The alternative of lower hormone dosage is particularly important to examine. We have used MT and E2 concentrations similar to those used in salmonids

(Baker et al. 1988). In warm water fishes, steroid concentrations used are much lower and of longer duration (e.g., zebrafish; Maack & Segner 2004).

Another interesting path a future study can take is to explore the potential for other environmental factors to influence sexual differentiation in this species, such as extreme salinity, density, and pH. The effect of environmental factors on gonadal differentiation has not been studied in any species within the Percidae family. The possibility of utilizing environmental manipulations to control sex in this group is exciting, given the high number of commercially relevant species in the family. Controlling sex using environmental factors would allow for a cheaper and safer alternative to sex reversal by hormone exposure methods, and should therefore be further explored.

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Appendix A: T Tests and ANOVA Tables

One-Way ANOVA and Tukey HSD test results for significant effects of hormone treatment on percentage of spermiating males in Yellow Perch from the 2015 experiments.

Analysis of Variance Source DF Sum of Squares Mean Square F Ratio Prob > F Treatment 7 4881.9966 697.428 26.2532 <0.0001* Error 8 212.5232 26.565 C. Total 15 5094.5198

Ordered Differences Report Level ―Level Difference Std Err Dif Lower CL Upper CL p-Value 14 mm Control 14 mm MT-I 52.45000 7.289088 23.6063 81.29368 0.0014* 14 mm Control 14 mm E2-I 37.44000 7.289088 8.5963 66.28368 0.0119* 14 mm Control 14 mm MT-D 28.82000 7.289088 -0.0237 57.66368 0.0502 14 mm MT-D 14 mm MT-I 23.63000 7.289088 -5.2137 52.47368 0.1247 14 mm MT-D 14 mm E2-I 8.62000 7.289088 -20.2237 37.46368 0.9166 14 mm E2-I 14 mm MT-I 15.01000 7.289088 -13.8337 43.85368 0.5025 12 mm Control 12 mm MT-I 2.14000 4.208357 -14.5129 18.79291 0.9992 12 mm Control 12 mm E2-I 28.70667 4.208357 12.0538 45.35957 0.0019* 12 mm MT-D 12 mm Control 3.79667 4.208357 -12.8562 20.44957 0.9773 12 mm MT-D 12 mm MT-I 5.93667 4.208357 -10.7162 22.58957 0.8310 12 mm MT-D 12 mm E2-I 32.50333 4.208357 15.8504 49.15624 0.0008* 12 mm MT-I 12 mm E2-I 26.56667 4.208357 9.9138 43.21957 0.0033* 14 mm Control 12 mm Control 0.99667 5.951515 -22.5541 24.54743 1.0000 12 mm MT-I 14 mm MT-I 49.31333 5.951515 25.7626 72.8641 0.0005* 12 mm MT-D 14 mm MT-D 31.62000 5.951515 8.0692 55.17077 0.0097* 12 mm E2-I 14 mm E2-I 7.73667 5.951515 -15.8141 31.28743 0.8763

One-Way ANOVA test results for significant effects of hormone treatment on growth rate of Yellow Perch during treatment from the 2015 experiments.

Analysis of Variance Source DF Sum of Squares Mean Square F Ratio Prob > F Treatment 3 0.6675000 0.222500 3.3242 0.0774 Error 8 0.5354667 0.066933 C. Total 11 1.2029667

T Test results for significant effects of high temperature on percentage of males in Yellow Perch from the 2016 Experiment 2.

T Test Difference 0.40000 t Ratio 0.188982 Std Err Dif 0.21166 DF 8 Upper CL Dif 0.52809 Prob > │t│ 0.8548 Lower CL Dif -0.44809 Prob > t 0.4274 Confidence 0.95 Prob < t 0.5726

T Test results for significant effects of high temperature on growth rate in Yellow Perch from the 2016 Experiment 2.

T Test Difference 0.236000 t Ratio 5.266598 Std Err Dif 0.044811 DF 6.46104 Upper CL Dif 0.343779 Prob > │t│ 0.0015* Lower CL Dif 0.128221 Prob > t 0.0008* Confidence 0.95 Prob < t 0.9992