GROOVER E.Pdf

Home , Ballan wrasse, Deep sand bed, Hawaiian cleaner wrasse, Ornate wrasse, Tropical fish

ASSESSMENT OF CULTURE TECHNIQUES FOR TWO Halichoeres WRASSES, H. melanurus AND H. chrysus

ELIZABETH MARIE GROOVER

A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE

UNIVERSITY OF FLORIDA

2018

To all who have supported me in pursuit of my passion

ACKNOWLEDGMENTS

I would like to thank my friends and family, especially my parents, Michael and

Donna Groover, whose unconditional love and support have made me the woman I am today. I would like to acknowledge my major professor, Dr. Matthew DiMaggio, for his experience, guidance, encouragement, and confidence in my abilities as a scientist and fish culturist. I would also like to acknowledge my committee members, Dr. Cortney Ohs and Dr. Joshua Patterson, for their assistance and expertise in the field. Thank you to my fellow colleagues Dr. Katharine Starzel, Michael Sipos, Tim Lyons, Taylor Lipscomb,

Dr. Marion Hauville, and Shane Ramee for their advice, assistance, knowledge, and friendship. Additionally, I would like to thank all staff and students at University of

Florida’s Tropical Aquaculture Lab for their support, knowledge, guidance, and willingness to help whenever it was needed. I would like to thank Segrest Farms and

Kevin Kohen of Doctors Foster and Smith Live Aquaria.com for their generous donations of wrasse broodstock, as well as Instant Ocean Spectrum Brands, Larry’s

Reef Services, Fritz Industries, Inc., Piscine Energetics, and Petco for providing additional elements needed for these trials. Lastly, I would like to thank Dr. Judy St.

Leger, Rising Tide Conservation, and the Sea World and Busch Gardens Conservation

Fund for providing the funding for my research.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 8

LIST OF FIGURES ...... 9

ABSTRACT ...... 11

CHAPTER

1 GENERAL INTRODUCTION ...... 13

Marine Aquarium Trade ...... 13 Marine Ornamental Aquaculture ...... 14 Labridae ...... 17 Halichoeres ...... 19

2 THE MELANURUS WRASSE Halichoeres melanurus: A DESCRIPTION OF SPAWNING, CULTURE PROTOCOLS, LARVAL DEVELOPMENT, AND COMPLETION OF LIFE CYCLE ...... 21

Introduction ...... 21 Methods ...... 24 Acquisition of Broodstock ...... 24 Quarantine Protocol ...... 24 Broodstock Husbandry ...... 26 Egg Collection and Analysis ...... 27 Larval Growth and Development Trial ...... 27 Results ...... 31 Quarantine Protocol ...... 31 Spawning Frequency and Quantity ...... 31 Larval Growth and Development Trial ...... 31 Completion of Life Cycle...... 35 Discussion ...... 36

3 DEVELOPMENT OF EARLY LARVICULTURE PROTOCOLS FOR Halichoeres melanurus ...... 50

Introduction ...... 50 Materials and Methods...... 56 Acquisition and Quarantine of Broodstock...... 56 Broodstock Husbandry ...... 58 Egg Collection and Analysis ...... 59 Experimental Design of Embryo Incubation Experiments ...... 60

Experiment 3-1 Temperature - Embryo Incubation Time and Larval Survival at Hatch ...... 61 Experiment 3-2 Temperature - Larval Size at Hatch ...... 61 Experimental Design of Larviculture Experiments ...... 62 Experiment 3-3 Algal Density - Larval Growth and Survival ...... 64 Experiment 3-4 Shading Method - Larval Growth and Survival ...... 64 Experiment 3-5 Algal Density and Prey Availability Period - Larval Feeding Percentage ...... 65 Experiment 3-6 First Feed Type and Prey Availability Period - Larval Feeding Percentage ...... 66 Experiment 3-7 Prey Density and Prey Availability Period - Larval Feeding Percentage ...... 66 Statistical Analysis ...... 67 Results ...... 68 Experiment 3-1 and 3-2 Temperature - Incubation Time, Larval Size and Survival ...... 68 Experiment 3-3 Algal Density - Larval Growth and Survival ...... 69 Experiment 3-4 Shading Method - Larval Growth and Survival ...... 69 Experiment 3-5 Algal Density and Feed Availability Period - Larval Feeding Percentage ...... 70 Experiment 3-6 First Feed Type and Feed Availability Period - Larval Feeding Percentage ...... 70 Experiment 3-7 Prey Density and Feed Availability Period - Larval Feeding Percentage ...... 70 Discussion ...... 71

4 EXPLORATION OF EARLY LARVICULTURE PROTOCOLS FOR Halichoeres chrysus ...... 83

Introduction ...... 83 Materials and Methods...... 89 Acquisition and Quarantine of Broodstock...... 89 Broodstock Husbandry ...... 91 Egg Collection and Analysis ...... 92 Experimental Design of Embryo Incubation Experiments ...... 93 Experiment 4-1 Temperature - Embryo Incubation Time and Larval Survival at Hatch ...... 94 Experiment 4-2 Temperature - Larval Size at Hatch ...... 94 Experimental Design of Larviculture Experiments ...... 95 Experiment 4-3 Algal Density - Larval Growth and Survival ...... 97 Experiment 4-4 Shading Method - Larval Growth and Survival ...... 97 Experiment 4-5 Algal Density and Prey Availability Period – Larval Feeding Percentage ...... 98 Experiment 4-6 Prey Density and Prey Availability Period - Larval Feeding Percentage ...... 99 Experiment 4-7 Prey Type and Prey Availability Period - Larval Feeding Percentage ...... 99

Statistical Analysis ...... 99 Results ...... 100 Experiment 4-1 and 4-2 Temperature - Incubation Time, Larval Size and Survival ...... 100 Experiment 4-3 Algal Density - Larval Growth and Survival ...... 101 Experiment 4-4 Shading Method - Larval Growth and Survival ...... 101 Experiment 4-5 Algal Density and Feed Availability Period - Larval Feeding Percentage ...... 102 Experiment 4-6 Prey Density and Feed Availability Period - Larval Feeding Percentage ...... 102 Experiment 4-7 Prey Type on and Feed Availability Period - Larval Feeding Percentage ...... 103 Discussion ...... 103

5 CONCLUSIONS ...... 117

LIST OF REFERENCES ...... 121

BIOGRAPHICAL SKETCH ...... 130

LIST OF TABLES

Table page

2-1 Water parameters throughout 45-day H. melanurus larval rearing trial...... 44

3-1 Water quality parameters for H. melanurus larviculture experiments ...... 78

4-1 Water quality parameters for H. chrysus larviculture experiments ...... 110

LIST OF FIGURES

Figure page

2-1 H. melanurus sexual dimorphism ...... 44

2-2 Feeding regime throught 45-day H. melanurus larval rearing trial...... 45

2-3 F0 melanurus wrasse spawning data ...... 46

2-4 Larval development of Halichoeres melanurus ...... 47

2-5 Larval growth of H. melanurus over 45-day culture period ...... 48

2-6 F1 melanurus wrasse spawning data ...... 49

3-1 Hatch rate of H. melanurus embryos in response to temperature ...... 78

3-2 Survival at hatch of H. melanurus embryos in response to temperature ...... 79

3-3 Notochord length at hatch of H. melanurus larvae in response to temperature .. 79

3-4 Survival of H. melanurus larvae in response to algal density ...... 80

3-5 Notochord length of H. melanurus larvae in response to shading method ...... 80

3-6 Survival of H. melanurus larvae in response to shading method ...... 81

3-7 Feeding percentage of H. melanurus larvae in response to algal density ...... 81

3-8 Feeding percentage of H. melanurus larvae in response to prey density...... 82

4-1 H. chrysus sexual dimorphism ...... 111

4-2 F0 H. chrysus broodstock spawning data ...... 112

4-3 Hatch rate of H. chrysus embryos in response to temperature ...... 113

4-4 Survival at hatch of H. chrysus embryos in response to temperature ...... 113

4-5 Notochord length at hatch of H. chrysus larvae in response to temperature ..... 114

4-6 Survival of H. chrysus larvae in response to algal density ...... 114

4-7 Notochord length of H. chrysus larvae in response to shading method ...... 115

4-8 Survival of H. chrysus larvae in response to shading method ...... 115

4-9 Feeding percentage of H. chrysus larvae in response to algal density ...... 116

4-10 Feeding percentage of H. chrysus larvae in response to prey density ...... 116

Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science

ASSESSMENT OF CULTURE TECHNIQUES FOR TWO Halichoeres WRASSES, H. melanurus AND H. chrysus

Elizabeth Marie Groover

August 2018

Chair: Matthew DiMaggio Major: Fisheries and Aquatic Sciences

To date, only five ornamental wrasse species have been successfully cultured and no commercial production exists. The melanurus and yellow wrasses (Halichoeres melanurus and H. chrysus, respectively) were chosen as candidates for development of culture methodologies due to their popularity in the aquarium trade and short larval duration. Melanurus wrasse larvae initiated feeding, swimbladder inflation, flexion, and metamorphosis at 3, 10, 15, and 37 days post hatch (DPH), respectively, with 0.5% survival. First generation wrasses began spawning at ~8 months of age, signifying closure of the life cycle. For both wrasse species, an inverse relationship was found between water temperature and embryo incubation period. Incubation temperatures of

25 and 28 degrees Celsius produced highest survival and smaller larvae at hatch.

Examination of melanurus and yellow wrasse larvae prior to first feeding established that algal densities greater than or equal to 50,000 cells mL-1 or artificial shading resulting in ~300 lux, enhanced growth and survival. Melanurus wrasse first feeding trials revealed that algal densities of 300,000 and 500,000 cells mL-1, provision of

Parvocalanus crassirostris nauplii (<75 microns), and prey densities greater than or

equal to 5.0 items mL-1 produced elevated larval feeding success. Yellow wrasse first feeding culture trials examining similar parameters did not elucidate any advantageous culture protocols, indicating the necessity for further exploration of these parameters and others affecting larval first feeding success. Continued investigation into larviculture techniques influencing growth, survival, and feeding success should establish methodologies leading to commercial protocols for these and other marine ornamental species.

CHAPTER 1 GENERAL INTRODUCTION

Marine Aquarium Trade

The global aquarium trade is a multi-billion-dollar industry that supports one of humanities’ most popular hobbies, aquarium keeping (Livengood and Chapman, 2007).

Although freshwater fish comprise the largest portion of ornamental fish in the aquarium industry by volume (Dawes, 1998), marine ornamental fish are also popular with a higher number of species present in the trade (Dawes, 1998; Livengood and Chapman,

2007). An estimated 27 million marine ornamental fishes are globally traded (Townsend,

2011) and between 7-11 million fishes from over 1,800 species and 125 families are imported yearly into the United States (U.S.) alone (Rhyne et al., 2012, 2017). These marine fish will ultimately find their way into public aquariums, research institutions, businesses, and approximately 2.5 million households (APPA, 2018), making the U.S. the largest importer of ornamental fish (Livengood and Chapman, 2007). Other major importers of marine ornamental fish include Europe, Japan, and China (Calado et al.,

2017; Livengood and Chapman, 2007; Rhyne et al., 2017).

The majority of tropical marine ornamental fish are harvested sustainably from richly biodiverse coral reef ecosystems in Indonesia and the Philippines using nets and other non-destructive methods (Olivotto et al., 2017; Rhyne et al., 2012). In coastal developing nations, ornamental fish collection accounts for a large percentage of the local economy and provides employment for many families, promoting conservation of local reef habitats (Tlusty, 2002). However, unsustainable and potentially destructive methods, such as the use of cyanide (Barber and Pratt, 1998; Dee et al., 2014; Rubec et al., 2001; Thornhill, 2012; Wabnitz et al., 2003), are sometimes employed, which can

negatively impact native fish populations and the coral reefs they reside on (Calado et al., 2017; Wabnitz et al., 2003).

One possible option to mitigate environmental and ecological impacts that may be associated with irresponsible collection practices is the captive culture of certain marine ornamental fish species (Tlusty, 2002). The refinement of practices for species currently in production and elucidation of protocols and techniques for the culture of new commodities, will help to support the growth and development of the marine ornamental aquaculture industry. This progression will contribute to the continued availability and expansion of appropriate captive bred alternatives to certain wild caught specimens.

Marine Ornamental Aquaculture

Captive production of marine fish for the aquarium trade is a relatively new and less recognized segment of the aquaculture industry compared to foodfish aquaculture

(Calado et al., 2017). However, the marine ornamental aquaculture industry has made much progress in the last 10 to 20 years, in part, due to recent advances in technology and culture techniques (Calado et al., 2017). In 1998, the only marine ornamental fish species available as captive bred were from the families Pomacentridae (clownfish and damselfish), Pseudochromidae (dottybacks), and Gobiidae (gobies) (Tlusty 2002). Just two decades later, the number of marine aquarium ﬁsh species successfully bred in captivity has been brought to 358 across 47 families (Sweet and Pedersen, 2018), representing approximately 20% of the total number of species available in the marine aquarium trade (Rhyne et al., 2012). Although this is a significant achievement, it is also deceiving, as only about 37 species are considered commonly available as captive bred, 33 species as low to moderate, and 51 as scarce, a combined total of 121 species

(Sweet and Pedersen, 2018). These species only comprise about 7% of the total

number of species traded globally for the marine aquarium industry, a small percentage when compared to the freshwater ornamental fish industry, where over 90% of species are available as captive bred (Calado et al., 2017; Livengood and Chapman, 2007;

Olivotto et al., 2011; Tlusty, 2002; Wabnitz et al., 2003).

Demersal spawners comprise ~74% of marine ornamental species that have been cultured commercially or as part of research (Moorhead and Zeng, 2010). These species deposit large adhesive eggs on a substrate, which often receive some level of parental care, and result in precocial newly hatched larvae that can consume easily cultured live food items such as rotifers (Brachionus spp.) and brine shrimp (Artemia spp.) (Calado et al., 2017; Olivotto et al., 2011, 2017). Examples of demersal spawning families of marine fish commonly produced for the aquarium trade are clownfish

(Pomacentridae), gobies (Gobiidae), dottybacks (Pseudochromidae), and blennies

(Blennidae). Conversely, culture methods for pelagic spawning species are relatively unexplored and few are commercially produced (Calado et al., 2017; Moorhead and

Zeng, 2010). These fish release gametes into the water column and provide no parental care, resulting in altricial (Gopakumar et al., 2009) newly hatched larvae with small mouths that can often only consume smaller live feed items such as copepod nauplii and ciliates that are more difficult and expensive to culture (Calado et al., 2017; Olivotto et al., 2011, 2017). Examples of pelagic spawning families of marine ornamental fish include angelfish (Pomacanthidae), tangs (Acanthuridae), anthias (Serranidae), and wrasses (Labridae).

There are many bottlenecks associated with the culture of pelagic spawning species that hinder captive propagation. Development of reliable and productive

broodstock is often a challenge due to limited knowledge concerning required nutritional composition of diets, social structure, and environmental cues needed to stimulate spawning behavior (Calado et al., 2017; Olivotto et al., 2011, 2017; Thresher, 1984).

Embryos resulting from pelagic spawning species are often small and develop rapidly, normally hatching within 24 hours, and are heavily impacted by environmental parameters such as temperature (Beacham and Murray, 1990; Gray, 1928; Pauly and

Pullin, 1988; Pepin, 1991; Pepin et al., 1997). Decrease in incubation temperature has been shown to increase larval size at hatch in several species, which is often correlated to increased larval survival (Barden et al., 2013; Gray, 1928; Pepin et al., 1997).

However, it is critical to indentify appropriate incubation temperature because deviation outside of this range can disrupt proper development of embryos and increase mortality

(Gray, 1928; Pepin, 1991).

Newly hatched yolk sac larvae of pelagic spawning marine ornamental fish are typically fragile and require specific environmental parameters such as water flow, turbidity, light, and oxygen for survival. Maximizing survival during this period of endogenous feeding is important since the transition to exogenous feeding often results in high larval mortality (May, 1974; Thresher, 1984; Turingan et al., 2005; Yúfera and

Darias, 2007). Successful transition from yolk sac and oil globule absorption to external feed items requires factors such as algal density, prey type, and prey density to be appropriate for the species and culture environment. The physical size of prey items is often the most limiting factor for altricial larvae due to their reduced mouth gape

(Baensch and Tamaru, 2009; Holt, 2003; Turingan et al., 2005; Wittenrich et al., 2009;

Wittenrich and Turingan, 2011). Common first feed items such as rotifers Brachionus

sp. and Instar I Artemia sp. nauplii are usually too large, lack proper nutritional elements, or do not elicit a feeding response for these larvae, thus alternative live prey is required such as copepod nauplii (Calado et al., 2017; Davis et al., 2018; Moorhead and Zeng, 2010; Olivotto et al., 2011). Identifying these first feed requirements is paramount for successful first feeding and subsequent larval survival.

Labridae

Wrasses are one of the largest families of fish with 60 genera and 548 species

(Kuiter, 2010; Parenti and Randall, 2018). These fish are diverse in size ranging from the humphead wrasse Cheilinus undulates, with length and weight exceeding two meters and 190 kg, respectively (Hirai et al., 2013), to much smaller ornamental wrasses under 10 cm in length. Smaller wrasses are very popular in the aquarium trade due to their beautiful and diverse coloration, active behavior, adaptability to captivity, and general hardiness. They are the second most imported family of fish into the United

States by volume and the most diverse in terms of number of species (228) (Rhyne et al., 2012). The most recent marine aquarium trade data (www.aquariumtradedata.org) estimated that over 1.7 million individual wrasses were imported into the United States in 2008, 2009, and 2011 collectively. Most of these wrasses were sourced from

Southeast Asia (primarily Indonesia and Philippines) then transshipped to U.S. ports such as Miami, FL and Los Angeles, CA. The high market demand for Labrids and lack of production further substantiates the need for applied research that supports the development of commercial aquaculture protocols (Murray and Watson, 2014).

Despite their immense popularity in the aquarium trade, wrasses are inherently difficult to culture due to early life characteristics common to pelagic spawning species.

In fact, only ten wrasse species have been cultured past metamorphosis, with most

successes being a one-time success (Sweet and Pedersen, 2018). Wrasse species cultured to date include: the rainbow wrasse Parajulis poecilopterus (Kimura and

Kiriyama, 1993), hogfish Lachnolaimus maximus (Colin, 1982), humphead wrasse (Hirai et al., 2013), tautog Tautoga onitis (Perry et al., 1998, 2001), ballan wrasse Labrus bergylta (Hamre et al., 2013a; Oie et al., 2017; Ottesen et al., 2012; Skiftesvik et al.,

2013), Cuban hogfish Bodianus pulchellus (Ohs et al., 2018), Hawaiian cleaner wrasse

Labroides phthirophagus (Montalvo, personal communication, 2017), bluestreak cleaner wrasse Labroides dimidiatus (Sweet, 2013), ornate wrasse Halichoeres ornatissimus

(Baensch, personal communication, 2017), and the melanurus wrasse Halichoeres melanurus (Barden et al., 2016; Groover et al., 2018). Ballan wrasses, as well as several other cold-water wrasse species, have been used since the late 1980s for removal of salmon louse Lepeophtheirus salmonis parasites from cultured salmon

Salmo salar and rainbow trout Oncorhynchus mykiss (D’Arcy et al., 2012; Hamre et al.,

2013a; Skiftesvik et al., 2013). Ballan wrasse aquaculture became important when drug resistant salmon lice proliferated and wild caught ballan wrasses could not meet the demands of the salmon industry (Hamre et al., 2013a; Skiftesvik et al., 2013). The ballan wrasse is the sole species of the Labridae family to be commercialized, although several bottlenecks still need to be overcome to achieve large scale production (Hamre et al., 2013a; Ottesen et al., 2012). The Cuban hogfish, Hawaiian cleaner wrasse, bluestreak cleaner wrasse, ornate wrasse, and melanurus wrasse are considered ornamental species, none of which are cultured commercially due to many impediments associated with broodstock management and larviculture.

Halichoeres

Wrasses in the genus Halichoeres are very popular in the aquarium trade with non-specific Halichoeres sp. ranking 11th for wrasses imported into the U.S.

(www.aquariumtradedata.org). Currently 80 species comprise this genus (Parenti and

Randall, 2018) with most species found in tropical to sub-tropical waters of the Pacific and Western Atlantic (Kuiter, 2010). A study conducted by Victor (1986) used the daily otolith-increment ageing-technique and determined that Halichoeres wrasses have relatively short larval durations ranging from 20.8 to 41.1 days (average 26.5 days). This short larval duration is advantageous and suggests Halichoeres wrasses may be more suitable for commercial production than other species with more prolonged larval durations.

The melanurus wrasse Halichoeres melanurus, first described by Bleeker (1851), is one of the top 20 most imported warsse species into the U.S. for the aquarium industry (www.aquariumtradedata.org) and is found widespread throughout the Western

Pacific including Indonesia, Micronesia, and Samoa (Kuiter, 2010). Like most wrasse species in the genus Halichoeres, the melanurus wrasse has been shown to have a relatively short larval duration (mean 22.1 days) likely increasing the economic potential for commercial production of this species (Victor, 1986a). The first successful culture of this species through metamorphosis was achieved in 2015 at the University of Florida

Tropical Aquaculture Laboratory and was supported by Rising Tide Conservation, laying the groundwork for the following research (Barden et al., 2016).

The yellow wrasse Halichoeres chrysus, first described by Randall (1981), is one of the most popular wrasses in the marine aquarium trade, being the third most imported wrasse species into the U.S. (www.aquariumtradedata.org). Found throughout

the Western Pacific from Bali to Christmas Island in the Indian Ocean, these small

(maximum length of 12 cm) bright yellow wrasses are found in small harems on shallow coastal reef habitats (Kuiter, 2010). This wrasse species also has a short larval duration averaging 26.1 days, as is characteristic of most Halichoeres wrasses (Victor, 1986a).

This species has not been cultured to date.

The objective of this study was to elucidate crucial elements of broodstock husbandry and management as well as larviculture protocols for the melanurus and yellow wrasse that will contribute to the development of empirically based culture protocols and ultimate commercial production of these valuable marine ornamental species. Quarantine procedures and broodstock husbandry techniques will be explored.

Spawning characteristics and larval developments will be evaluated. Through replicated experimentation, larval culture conditions were assessed. The effect of temperature on embryo incubation period, as well as on larval survival and size at hatch will be assessed for both species. Pre-exogenous feeding larval rearing experiments will be conducted to evaluate the effect of algal density and shading method on growth and survival from hatch to 3 DPH for both species. First feeding trials will be conducted for melanurus and yellow wrasse larvae which examined the effect of algal density, prey type, and prey density on first feeding success at 3 DPH. Elucidation of these husbandry and culture protocols will significantly contribute to the limited information available about captive culture of the melanurus and yellow wrasse, acting as a precursor to the formation of commercial production protocols.

CHAPTER 2 THE MELANURUS WRASSE Halichoeres melanurus: A DESCRIPTION OF SPAWNING, CULTURE PROTOCOLS, LARVAL DEVELOPMENT, AND COMPLETION OF LIFE CYCLE

Introduction

The global aquarium hobby is a multi-billion dollar industry with an estimated 27 million marine ornamental fish traded annually (Townsend, 2011). Primary consumers of marine ornamentals are the United States (U.S.), Europe, Japan, and China (Calado et al., 2017; Rhyne et al., 2017) with the U.S. importing between 7-11 million individual fish from over 1,800 species and 125 families every year (Rhyne et al., 2012, 2017).

These fish will ultimately find their way to public aquariums, research institutions, private businesses, and approximately 2.5 million households (2% of the U.S. population)

(APPA, 2018).

Currently, less than 10% of all marine ornamental fish species in the aquarium trade are cultured, meaning the vast majority of species are harvested directly from richly biodiverse coral reef ecosystems (Calado et al., 2017; Wabnitz et al., 2003). Most commercially cultured species exhibit demersal spawning behavior, depositing large adhesive eggs on a substrate. These embryos often receive some level of parental care and result in precocial newly hatched larvae that can consume readily cultured live food organisms such as rotifers (Brachionus spp.) and brine shrimp (Artemia spp.)(Calado et al., 2017; Olivotto et al., 2011, 2017). Conversely, very few pelagic spawning species are currently produced in commercial quantities due to the early life history traits they share. Generally, these fish broadcast gametes into the water column and provide no parental care following a spawning event. Newly hatched larvae are often altricial with a small mouth gape that limits their diet to smaller live feed items such as copepod nauplii

and ciliates that are more difficult and costly to culture (Calado et al., 2017; Olivotto et al., 2011, 2017).

Wrasses (Labridae) are one such family of pelagic spawning fish which exhibit many of these early life history characteristics, resulting in limited culture success to date. To the authors knowledge, only ten wrasse species have been cultured past metamorphosis, with most successes being a singular event with no further investigation (Barden et al., 2016; Sweet and Peterson, 2018). Historically, much of the information regarding the culture of ornamental wrasses has appeared in “grey” literature or has been referenced anecdotally in magazine articles and online media.

Currently, there is no commercial production of any ornamental wrasse species in the marine aquarium industry. However, wrasses are very popular in the aquarium trade, being the second most imported family of fish into the U.S. by volume and first by number of species (228) (Rhyne et al., 2012). According to the most recent marine aquarium trade data (www.aquariumtradedata.org), it was estimated that over 1.7 million wrasses were imported into the U.S. in 2008, 2009, and 2011 collectively. The high market demand for Labrids and absence of captive production highlights the need for research to develop commercial aquaculture protocols (Murray and Watson, 2014).

An important consideration when choosing a marine ornamental fish species for commercial production is length of larval duration. This ontogenetic period is generally characterized by numerous physiological and morphological developments, which often times can result in considerable larval mortality. A shorter larval duration may allow for increased economic potential for commercial production by decreasing the investment of time, labor, and resources needed to raise the fish through this challenging phase.

Halichoeres wrasses have relatively short larval durations, averaging 26.5 days, compared to most other wrasses which can be as long as 121 days in the blacktail wrasse Thalassoma ballieui (Victor, 1986a).

Consistent with other Halichoeres wrasses, the melanurus wrasse Halichoeres melanurus has been shown to have an abbreviated larval duration (mean 22.1 days) in the wild (Victor, 1986a). Found widespread throughout the Western Pacific, mainly on shallow, inshore coral reefs (Kuiter, 2010), the melanurus wrasse is one of the most popular wrasse species in the aquarium trade. With a maximum reported length of approximately 12 cm (Kuiter, 2010) this wrasse is a desirable size for many home aquaria. Melanurus wrasses can also assist with pest control in the aquarium. The species is carnivorous, with a natural diet mainly consisting of small invertebrates such as polychaetes and crustaceans that can become pests in aquaria. The sex of a melanurus wrasse is relatively easy to identify. In addition to being protogynous hermaphrodites, where males are normally larger and more colorful than females, melanurus wrasses are sexually dimorphic according to the number of dorsal ocelli and presence of facial barring (Kuwamura et al., 2000). Juveniles possess two dorsal ocelli

(one anterior and one slightly enlarged and centered) and one caudal ocellus (Figure 2-

1A). Females also display two dorsal ocelli and one dorsal ocellus, with all three being of similar size. Larger melanurus wrasses with one dorsal ocellus and one caudal ocellus can be female or male (Figure 2-1B). Terminal males either completely lack ocelli or display one caudal ocellus, normally accompanied by pink facial barring and yellow patches around the pectoral fins (Figure 2-1C). The first successful culture of this species through metamorphosis was achieved in 2015 at the University of Florida

Tropical Aquaculture Laboratory (UF-TAL), laying the groundwork for the following research (Barden et al., 2016).

The objective of this trial aimed to further elucidate and improve captive propagation techniques for the melanurus wrasse by describing reproductive behavior and larval growth and development in an aquaculture setting for the first time.

Furthermore, description of an effective and reliable larval feeding regime was of great interest as this information is essential to the formulation of commercial culture protocols.

Methods

Acquisition of Broodstock

Melanurus wrasse broodstock were originally collected by commercial fishers from coastal waters surrounding the Philippines and Fiji. Fish were shipped via air freight to a tropical fish wholesale company in 2016 (Segrest Farms, in Gibsonton, FL,

USA) and then subsequently transported to the UF-TAL in Ruskin, FL.

Quarantine Protocol

Wrasses were first prophylactically treated with a hyposaline bath (0 mg L-1 for 5 min) to dislodge any external parasites. The fish were then acclimated to the salinity and temperature of the quarantine system, which was maintained at 30 ± 1 g L-1 and 27

± 1 °C respectively. Salinity was maintained below full strength seawater to match that of transport water and decrease osmoregulatory stress. The recirculating system

(~1,500 L total volume) was composed of three 378 L, round, fiberglass tanks, with a dark interior, a fluidized sand bed, degassing column, and nylon sock filters (200 and

100 m). Tanks were covered with 0.635 cm mesh to prevent fish from jumping. A sand bed with an approximate depth of 5 cm covering about ¼ of the tank bottom was added

to accommodate the wrasses’ natural burying behavior. An assortment of polyvinyl chloride (PVC) pipes were provided for shelter. Fish were quarantined for 30 days.

During this time, the following antibiotic treatment protocol was adhered to because of previous incidents of bacterial infections identified in wrasses throughout previous quarantine periods. An Oxylinic acid (Fishman Chemical LLC, Vero Beach, FL,

USA) immersion treatment (1 mg L-1 for 24 hours every day for ten days) was administered as a precautionary measure for bacterial pathogens. Simultaneously, an oral dose of Oxylinic acid (0.333 mg g-1 of food) was delivered by incorporation into

Shrimp Souffle Scavenger Gel Premix (Repashy Ventures, Inc., Oceanside, CA, USA) and fed three times daily for ten days.

Following the completion of the antibiotic treatment, the diet consisted of two feedings per day until apparent satiation and was composed of a 0.840–1.410 mm sinking crumble (Reed Mariculture, Campbell CA, USA: APBreed Top Dressed

Otohime-C2, 49% protein, 14% fat, 3.5% fiber, 6.5% moisture, 15% ash, 1.5%

Phosphorous, and min 250 mg/L Astaxanthin) and PE Frozen Mysis Shrimp (Mysis relicta) (Piscine Energetics, Vernon, British Columbia, Canada, 69.5% protein, 8.35% fat, 2.75% fiber, and 5.5% ash). Salinity was gradually increased to that of full strength seawater (35 g L-1) by the end of the quarantine period in preparation for movement into the broodstock system. The following water quality parameters were maintained throughout the quarantine period: 0 mg L-1 total ammonia nitrogen (TAN), 0 mg L-1 nitrite, < 40 mg L-1 nitrate, pH 8.0 ± 0.1, alkalinity 171  17.1 g L-1 calcium carbonate, and dissolved oxygen (DO) >6.0 mg L-1. Once quarantine was complete, 14 female wrasses were transferred to the broodstock system.

Broodstock Husbandry

The broodstock were maintained in a 2,650 L round, fiberglass, tank with black walls and a white bottom. The tank was connected to a recirculating system (~34,000 L total volume) that included a bead filter, fluidized sand bed, protein skimmer, moving bed bioreactor with Kaldnes biomedia, and two 150-watt UV sterilizers. A circular laminar flow was achieved to aid in passive collection of buoyant embryos.

Vigorous aeration was provided from one airstone positioned at the tank bottom.

A 40 x 32 x 15 cm “sandbox” with at least 8 cm of thoroughly rinsed Quikrete® Pool

Filter Sand (particle size 0.85-0.425 mm) was also placed on the tank bottom to accommodate natural burrowing behavior. Additional structure in the form of PVC pipes and artificial coral provided shelter. Wrasse broodstock were fed 4-5 times per day until apparent satiation. The diet consisted of a 0.840–1.410 mm sinking crumble (Reed

Mariculture, Campbell CA, USA: APBreed Top Dressed Otohime-C2), PE Frozen Mysis

Shrimp (M. relicta, Piscine Energetics, Vernon, BC, Canada), LRS Fertility Frenzy

(Larry’s Reef Services, Advance, NC, USA, 15% protein, 3% fat, 1% fiber, 82% moisture, and 1 million CFU/gram microorangisms/probiotics), and LRS shad roe

(American shad Alosa sapidissima, Larry’s Reef Services, Advance, NC, USA, 23.24% protein, 0.91% fat, 1.12% fiber, 74.58% moisture).

Salinity was maintained between 33-36 g L-1 and measured every other day using a digital refractometer (Milwaukee Instruments, Inc., Rocky Mount, NC). Water temperature was maintained using external heat pumps set to 26.5 ± 1°C. Water quality parameters including TAN, nitrite, nitrate, alkalinity, DO, and pH were tested at least once every other week. TAN, nitrite, and alkalinity were evaluated using standardized colorimetric protocols (Hach, Loveland, CO, USA). Nitrate was tested using the API

Nitrate Test Kit (Mars Fishcare North America, Inc., Chalfont, PA, USA). DO was measured using a YSI ProODO meter (YSI Incorporated, Yellow Springs, OH, USA). A

YSI EcoSense pH100A meter was used to measure pH. Tanks were siphoned once per week to remove algal growth, uneaten feed, and fecal matter.

Egg Collection and Analysis

Buoyant pelagic wrasse embryos were collected daily using directed overflow from the broodstock system into a 20 L bucket lined with 200 µm mesh (DiMaggio et al.,

2017). The collector was set before dusk and retrieved the following morning.

Embryos were concentrated in a 250 µm mesh sieve for gross inspection, then transferred into an appropriately sized graduated container (1 L or 100 mL dependent on spawn size) for volumetric enumeration. Five 3 mL samples were taken from the graduated container of homogenized embryos and the number of fertilized eggs were counted in each sample. The total number of fertilized eggs was calculated by finding the mean number of fertilized eggs mL-1 and multiplying by the total volume (mL) of liquid in the container.

Embryos were positioned flush on a Sedgewick Rafter Counting Chamber to provide scale and photographed under a trinocular dissecting microscope capable of image capture (Jenoptik, 40 ProgRes Capture Pro v2.8.8, Jupiter, FL, USA). Embryo diameter was measured from digital images using ImageJ (U. S. National Institutes of

Health, Bethesda, Maryland, USA) software.

Larval Growth and Development Trial

The larval culture system was comprised of a series of bag filters (25 µm, 100

µm, and 200 µm), degassing column, protein skimmer, moving bed bioreactor with

Kaldnes biomedia, and 150-watt UV sterilizer. A round, 125 L, fiberglass tank with black

interior and a single white stripe at the bottom (to assist with visualizing larvae) was used for culturing larvae. The tanks’ internal central drain was fitted with a vertical PVC pipe covered with nylon mesh that excluded larvae from exiting the tank while allowing excess live feeds to be flushed. An initial mesh size of 250 µm was used for the central drain and then changed to 750 µm when larvae began consuming Instar 1 and 2

Artemia sp. An external standpipe was used to dictate water level.

A spawn of 3,150 fertilized eggs, originating from a harem of one dominant terminal male and 13 females, was stocked into a single larval tank at a density of 25 fertilized eggs L-1. Light airflow was provided by a single airstone secured next to the central drain throughout the duration of the trial. Fluorescent lights (32-watt) were positioned above the tank yielding a light intensity at the surface and tank bottom

(without algae) of 1,473 ± 29 and 371 ± 12 lux, respectively. Photoperiod was kept consistent at 16 hours of light and eight hours of dark (16L:8D) throughout the duration of the trial. The culture tank was kept static with no water flow from 0-5 days post hatch

(DPH). Flow rate was increased to 1 tank turnover per day (TOD) from 6-11 DPH and 3

TOD from 12 DPH until completion of metamorphosis. Water quality parameters were measured every other day during the static phase and every three days after water flow was introduced. At 25 DPH, a layer of Quikrete Filter Sand (approximately 1.5 cm deep) was slowly added to the tank through a 2.54 cm PVC pipe that extended from above the water surface down to the tank bottom. This sand bed covered ~25% of the tank bottom and served as a settlement cue for the larvae.

Live Tetraselmis chuii microaglae was used in larval rearing trials to provide countershading for developing larvae and aid in prey capture. T. chuii was harvested

from carboy cultures and cell densities (cells mL-1) were determined daily using a hemocytometer. Algae was temperature acclimated to match conditions in the larval tank prior to introduction. Algae was gradually added to the culture tank every morning prior to the lights turning on from 0-35 DPH, until a density of 300,000 cells mL-1 was achieved (Figure 2-2).

Parvocalanus crassirostris copepods were used in larval rearing trials as the initial live feed item. P. crassirostris was batch cultured at UF-TAL and fed a diet of live mircroalgae (Tetraselmis chuii, Tisochrysis lutea, and Chaetoceros muelleri). Nauplii

(<75 µm) were harvested daily from stock cultures and culture densities (nauplii mL-1) were determined. Live feeds were added to the larval tank through a drip bucket that slowly released its contents for minimal disturbance. P. crassirostris <75 µm (~Instar 1-

3) nauplii were added to the culture tank twice/day to reach a density of 3.8 ± 1.5 nauplii mL-1 from 3-28 DPH. From 13-31 DPH, P. crassirostris nauplii 75-100 µm (~Instar 3-4) were added to the culture tank twice/day to reach a density of 3.6 ± 1.7 nauplii mL-1.

Instar 1-2 Artemia sp. nauplii were added to the culture tank once/day to reach a density of 0.25 nauplii mL-1 from 19-46 DPH. The larvae were weaned onto a commercial diet of

250 µm sinking crumble (APBreed Top Dressed Otohime-A2, 50% protein, 13% fat,

3.5% fiber, 6.5% moisture, 15% ash, 2.0% Phosphorous, and min 250 mg/L

Astaxanthin) once metamorphosis was completed (~ 37 DPH). Larger crumble sizes of the same dietary formulation were gradually introduced as dictated by larval growth.

The successful feeding regime used in the 45-day melanurus wrasse larval growth and development study is depicted in Figure 2-2.

Larvae (mean ± SD, 8 ± 3) were sampled from 0-22, 25, 27-30, 37-40, and 45

DPH from the surface of the culture tank. Larvae were positioned laterally and flush on a Sedgewick Rafter Counting Chamber to provide scale and immediately photographed under a dissecting microscope as previously described. Larval photographs were digitally analyzed for yolk sac length and height, oil globule diameter, notochord length

(NL), standard length (SL), and upper and lower jaw length using ImageJ software v1.50i software (U. S. National Institutes of Health, Bethesda, Maryland, USA). Yolk sac

휋 volume was determined using the formula for a prolate spheroid: 푉 = ( )(푙)(ℎ2), where l 6 is yolk sac length and h is yolk sac height (Bagarinao, 1986). Oil globule volume was

휋 determined using the formula: 푉 = ( )(푑3), where d is the oil globule diameter 6

(Bagarinao, 1986). Notochord length, defined as the length from the anterior most point of the larvae to the posterior tip of the notochord, was measured for pre-flexion larvae.

Standard length, defined as the length from the anterior most point of the larvae to the end of the midlateral portion of the hypural plate, was measured for post-flexion larvae.

Mean lengths were calculated for all sampled larvae and plotted against time (DPH) to generate a growth curve. Mouth gape height was determined using the formula: 퐺퐻 =

√푈퐽퐿2 + 퐿퐽퐿2, where UJL = upper jaw length and LJL = lower jaw length (Wittenrich et al., 2007).

All larval photos were observed for the initiation of key developmental milestones including exogenous feeding, swim bladder inflation (SBI), and flexion. The number of larvae exhibiting SBI and flexion out of the total number of larvae sampled at each time point was recorded to attain a percentage. Initiation of exogenous feeding was characterized by the presence of a fully developed mouth, eyes, and an open primitive

digestive tract. Initiation of flexion was characterized by the elevation of the posterior notochord tip and beginning of hypural plate formation.

Results

Quarantine Protocol

The initial wrasse quarantine protocol, which included prophylactic antibiotic treatments, resulted in poor wrasse survival (<50%) throughout the quarantine period. A simplified quarantine protocol, eliminating the antibiotic treatments and focusing on maintaining water quality and providing a high quality and quantity diet, was then adopted for subsequent groups of wrasses resulting in increased survival (>80%).

Spawning Frequency and Quantity

F0 melanurus wrasse broodstock (1M:13F) began spawning two weeks post quarantine in August 2016 and ceased in March 2017 (Figure 2-3). Collected spawns

(n=99) ranged in quantity from 11-7,200 fertilized eggs (1,721 ± 1,642). Two large peaks in spawn quantity were observed during and several days following the full moon in December and January of 2016, however, there was no clear effect of lunar phase on spawn frequency or quantity for the remainder of the collection period. Spawning within the harem occurred daily at dusk, which is consistent with literature describing wild mealnurus wrasse spawning behavior (Kuwamura et al., 2000).

Larval Growth and Development Trial

Egg collection and development

Embryos collected the following morning were observed to be in the late neurula stage with clearly visible somites (Figure 2-4A). Fertilized eggs from captive broodstock measured 0.627 ± 0.013 mm in diameter (n=50). Total incubation time was approximately 15-21 hours (12-15 hours at 26.5 ± 1°C in broodstock system and the

remaining 3-6 hours at 24.80 ± 0.46°C in larval system) with hatching occurring between 12:00 and 15:00 the following day.

Larval growth and development

At 0 DPH, newly hatched melanurus wrasse larvae measured 1.590 ± 0.140 mm

NL, (n=10), and lacked developed eyes, mouth, and digestive tract. An elliptical yolk sac extended past the anterior of the larvae and reached back midway down the length of the notochord. Mean yolk sac volume measured 0.0609 ± 0.0228 µL, (n=10). The oil globule was positioned at the anterior most part of the yolk sac and measured 0.0011 ±

0.0001 µL (n=10) in volume. Small melanophores were present along the full length of the dorsal region of the larvae (Figure 2-4B).

At 1 DPH, larvae had absorbed 91.6% of their yolk sac with a remaining volume of 0.0051 ± 0.0037 µL, (n=10). The oil globule had been absorbed by 76.1% with a volume of 0.0003 ± 0.0001 µL, (n=10). Larvae measured 2.053 ± 0.094 mm NL, (n=10) and exhibited a short, straight, and undifferentiated gastrointestinal tract (GI) that remained closed. Small melanophores were reduced in number dorsally and appeared in small numbers ventrally. Formation of the mouth and eyes had commenced, and larvae did not actively swim but rather “hung” motionless in the water column (Figure 2-

4C).

At 2 DPH, larvae had very little yolk sac and oil globule remaining (0.2% and

4.4% of original volume, respectively). Larvae measured 2.194 ± 0.062 mm NL, (n=10) and now exhibited pigmented eyes, a partially open mouth, and an open posterior digestive tract. Melanophores were reduced in number, with only two appearing on the dorsal fin fold and two on the ventral fin fold of most larvae. Both sets of melanophores

were evenly spaced lengthwise on the fin folds. Most larvae remained immobile in the water column with short, slow, infrequent bouts of swimming.

At 3 DPH, larvae became more active with less time spent immobile and now measured 2.234 ± 0.139 mm NL, (n=14). Fully developed eyes, an open mouth, and absorbed yolk sac and oil globule indicated larvae could begin exogenous feeding.

Mouth gape height measured 172 ± 0.027 µm, (n=5). The digestive tract regions were now distinguishable as the foregut, midgut, and hindgut. P. crassirostris nauplii were identified in the guts of larvae confirming initiation of exogenous feeding. Melanophores present on 2 DPH grew in size and definition (Figure 2-4D).

At 7 DPH, larvae had reached a length of 2.748 ± 0.296 mm NL, (n=7). The foregut, midgut, and hindgut regions of the digestive tract were more clearly defined and greater in volume. Red-orange pigmentation began to appear over the anterior and posterior portions of the digestive tract, as well as the tip of the snout (Figure 2-4E).

At 11 DPH, SBI was first observed. Out of the larvae sampled, (n=10), 10% exhibited an inflated swim bladder. Notochord length averaged 3.526 ± 0.335 mm. The foregut, midgut, and hindgut regions of the digestive tract increased further in volume

(Figure 2-4F).

At 15 DPH, initiation of flexion and hypural plate formation was first observed.

Incidence of SBI was observed in 37.5% of larvae sampled. Notochord length averaged

4.866 ± 0.360 mm, (n=8). The complexity of the digestive tract drastically increased, now exhibiting a coiled midgut. Only the two dorsal spots and single posterior ventral spot remained on larvae (Figure 2-4G).

At 18 DPH, completion of flexion was observed in 12.5% of larvae, (n=8), characterized by hypural plate and caudal fin ray development. SBI was observed in

25% of larvae sampled. Mean standard length was determined to be 5.180 ± 0.422 mm,

(n=8). Increased red-orange pigmentation was localized around the coelomic cavity of larvae, tip of the snout, and posterior ventral spot (Figure 2-4H).

At 27 DPH, a single moribund larva measured 7.867 mm SL and three distinctly pigmented areas, appearing to be precursors to dorsal ocelli, were evenly spaced lengthwise down the dorsal fin. This larva did not have an inflated swim bladder.

From 34-36 DPH, larvae were observed displaying settlement behavior which culminated in burying themselves in the sand bed. Additional holes in the sand bed and missing larvae indicated the settlement of additional individuals.

From 37-43 DPH, larvae emerged from the sand bed fully metamorphosed with inflated swim bladders and a mean SL of 11.901 ± 0.081 mm, (n=2). Newly metamorphosed larvae had a prototypical fusiform body morphology with light yellow- green overall body color and rows of alternating lateral lines of black pigmented dots with broken horizontal lines of yellow-white (precursors of solid stripes). A single centered ocellated dorsal spot and single ocellated dorsal spot at the base of the caudal fin were also observed (Figure 2-4I).

At 45 DPH, two post-metamorphic larvae were sampled and measured (13.000 ±

0.375 mm). Yellow-green body color had intensified, stripes were darkened, and ocellated spots became more defined. An additional small anterior dorsal spot was observed (Figure 2-4J).

Swim bladder inflation was first observed at 11 DPH (3.847 ± 0.272 mm NL) in

10% of larvae sampled. From this time until 29 DPH (8.405 ± 1.651 mm SL), SBI ranged from 0-67%. Swim bladder inflation was variable and not observed in larvae sampled on 12, 14, 27, and 28 DPH. Initiation of flexion was first observed at 15 DPH

(4.866 ± 0.360 mm NL) with 25% of larvae sampled exhibiting flexion. On 16 and 17

DPH, 38% and 75% of larvae were observed to have initiated flexion, respectively. All larvae sampled for the remainder of the trial (≥18 DPH) exhibited some indication that the process of flexion had begun. Mean larval growth over the 45-day culture period was best described using the equation y=2.0264e0.0484x with an R2 value of 0.9618 and can be seen in Figure 2-5. Survival through metamorphosis was 0.54% yielding 17 juvenile wrasses. A total of 217 larvae were sampled throughout this trial.

Completion of Life Cycle of the Melanurus Wrasse

First generation (F1)

The same harem of F0 melanurus wrasses (1M:13F) that provided embryos for the larval growth and development trial, produced a separate spawn of 3,100 fertilized eggs in January 2017. Larvae from this spawn completed metamorphosis at 35 DPH with a 0.42% survival rate, resulting in 13 first generation (F1) juveniles. These F1s were grown out in the larval culture system for approximately three months and then transferred to a 200 L vat within a larger broodstock system in May 2017. During transfer at four months of age, it was noted that all F1’s possessed juvenile coloration

(three dorsal ocelli). In September 2017, an airlift egg collector was first deployed in the broodstock tank and a viable spawn was obtained from the eight-month-old F1 melanurus wrasses, confirming sexual maturity. Less than one week later, F1 melanurus wrasses were photographed for subsequent determination of total length

(TL). The harem now consisted of one male (1 posterior caudal ocellus) measuring

6.810 cm TL, one intermediate phase fish of unknown sex (2 dorsal ocelli) measuring

4.808 cm TL, and nine females (3 dorsal ocelli) ranging in size from 4.501-6.007 cm TL.

Mean TL for the entire population was calculated to be 5.171 ± 0.7234 cm TL, (n=11), with all individuals surpassing minimum market size (>3.8 cm). This harem was observed to spawn consistently for nine months, after which spawning data was no longer collected. Spawning frequency, spawn size (1,274 ± 571 total eggs) (n=72), and fertilization success (86.41±10.03 %) are depicted in Figure 2-6.

Second generation (F2)

An F1 spawn of 1,400 eggs (fertilization success = 88%) in late September 2017 was stocked in a 125 L larval tank and raised through metamorphosis. Metamorphosis was reached by six larvae between 40-42 DPH, resulting in 0.50% survival. This group was grown out for approximately five months and then transferred to a 200 L vat within a larger broodstock system in March 2018. By six months of age, most fish had reached approximate market size, ranging from to 3.437-4.541 cm (3.867 ± 0.416 TL, n=6) and displayed juvenile coloration.

Discussion

Development of high-quality broodstock is the foundation of any breeding endeavor (Calado et al., 2017; Olivotto et al., 2011). Post-quarantine melanurus wrasses adapted quickly to captive conditions. Broodstock readily consumed all commercial diets and within two weeks began spawning volitionally. The first two months of spawning yielded small spawn quantities, ranging from 23-1,353 fertilized eggs with a mean of 547 ± 380. Following two months in captivity, mean spawn quantity increased almost five-fold (2,251 ± 1,675), with collected spawns ranging from 93-7,267

fertilized eggs, and remained consistent for the following five months. After this period, spawn size gradually decreased and eventually ceased. Phenotypic sex change from female to male in several individuals was observed within the original wild harem over the course of several months prior to the decline in spawning. As protogynous hermaphrodites, maintaining the haremic structure of melanurus wrasses with respect to sex ratio is complex and in constant flux (Kuwamura et al., 2000). It is possible that the gradual shift in sex ratio from a female to male dominated population contributed to the decline and eventual cease of spawning within the population. The two observed large peaks in spawn quantity observed during and several days following the full moon in December and January of 2016 suggest a possible effect of lunar phase on spawn quantity, however, prolonged spawning data collection is required to better elucidate any influence due to lunar phase.

Overall spawning observations suggest that the melanurus wrasse can adapt quickly to a captive environment and will spawn volitionally without the need for hormone induction. As the environmental conditions of their native range are relatively static, changes in environmental parameters such as temperature or photoperiod were not needed to induce spawning in captive melanurus wrasses, greatly simplifying the reproduction process. The presence of multiple females in the spawning harem prevented evaluation of individual female fecundity and the number of females contributing to daily spawns. Therefore, it is important to note that spawning figures depict daily spawning characteristics of the entire harem and not individuals.

Pelagic larval duration and key developmental milestones were documented for the melanurus wrasse during this study. It is crucial to elucidate the timing of

developmental markers such as initiation of exogenous feeding, swim bladder inflation, flexion, settlement, and metamorphosis because they are often associated with high levels of mortality when specific culture parameters are not met (Moorhead and Zeng,

2010; Olivotto et al., 2017; Wittenrich et al., 2007).

The first, and perhaps most important, ontogenetic event is absorption of the larval yolk sac and oil globule reserves. Subsequent to this event, larvae must ingest and assimilate exogenous food items to sustain their energy demands (May, 1974;

Thresher, 1984; Yúfera and Darias, 2007). If proper live feed items are not available to the larvae at this point in development, metabolic demands will likely be unmet; resulting in an irreversible decline in larval health and early mortality. Under environmental parameters described, melanurus wrasse larvae had fully absorbed their yolk sac and oil globule reserves by 3 DPH and required the addition of P. crassirostris nauplii into the culture environment. It has been demonstrated that the larvae of several fish species preferentially selected food items that are 20-60% of their mouth gape

(Fernandez-Diaz et al., 1994; Ostergaard et al., 2005; Ronnestad et al., 2013), meaning that 3 DPH melanurus wrasse larvae (2.2 mm NL) with a mouth gape of approximately

172 µm would most likely select prey that was 34.4-103.2 µm in size. Rotifers and

Artemia sp. are the industry standard for feeding marine ornamental fish larvae.

However, B. rotundiformis rotifers and Instar I Artemia sp. range in length from approximately 60-200 and 400-500 µm, respectively (Calado et al., 2017), likely making these commonly produced live feed items too large for melanurus wrasse larvae at first feeding. Smaller live feed items, such as Parvocalanus sp. copepod Instar I nauplii that

can be as small as 30 µm, are thus required to meet the physical constraints of this species.

Swim bladder inflation is characterized by larvae gulping air at the air-water interface and forcing it through their pneumatic duct into the swim bladder, which assists in maintaining neutral buoyancy (Woolley and Qin, 2010). In physoclistous fishes, this pneumatic duct remains open for a limited period of time, after which, the duct will close, and the larvae will be unable to inflate their swim bladder (Woolley and Qin, 2010). The timing of this event is species dependent and identification of this critical developmental stage is essential for establishing successful larval culture protocols. The presence of an oily film on the water’s surface can increase surface tension and make it difficult for larvae to break through the air-water interface and successfully inflate their swim bladders (Woolley and Qin, 2010). Larvae that are unable to inflate and maintain buoyancy will expend excess energy for swimming and prey capture, often leading to poor growth and mortality (Chatain, 1989; Chatain and Ounais-Guschemann, 1990;

Woolley and Qin, 2010). Swim bladder inflation for melanurus larvae was first observed at 11 DPH. During this period, the air-water interface was kept clear of surface biofilms via manual removal, aeration, and increased water flow. Despite these efforts, percent swim bladder inflation remained low and inconsistent throughout development, demonstrating that this bottleneck remains pervasive for this species and further investigations are warranted to address it. Problems with swim bladder inflation are well documented with other marine species such as European seabass Dicentrarchus labrax and gilthead seabream Sparus aurata (Chatain, 1994, 1986; Daoulas et al., 1991;

Paperna, 1978; Vandeputte et al., 2009). The use of a surface skimmer could improve

removal of surface films, which was demonstrated to increase swim bladder inflation in gilthead sea bream (Chatain and Ounais-Guschemann, 1990). Increased turbulence can also increase surface accessibility for larvae, as shown with striped bass Morone saxatilis (Chapman et al., 1988). Photoperiod and light intensity are additional parameters to consider, as environmental requirements are generally species specific and crucial for successful swim bladder inflation in fish larvae (Woolley and Qin, 2010).

It is important to provide appropriate periods of light and darkness to allow for swim bladder inflation but effects on feeding must also be considered (Fielder et al.,

2002). A period of complete darkness is required for successful swim bladder inflation in several species such as striped trumpeter Latris lineata, Australian bass Macquaria novemaculeata, European seabass, and silver seabream Pagrus auratus (Battaglene and Talbot, 1990; Fielder et al., 2002; Trotter et al., 2003). The importance of a dark period for proper swim bladder inflation was also demonstrated in the culture of the ornate wrasse H. ornatissimus. Initial trials suggested that continuous light at 7 DPH inhibited swim bladder inflation in this species. Modifying the photoperiod to a 70% light and 30% dark daily cycle greatly improved swim bladder inflation success in the ornate wrasse (Baensch, 2016, personal communication). As a congener, it is likely that a dark period is also required for the melanurus wrasse to allow for successful swim bladder inflation. For all melanurus wrasse larval trials, photoperiod was kept at 16L:8D to provide an appropriate period of darkness to accommodate swim bladder inflation.

Flexion is an energetically taxing period for the development of larval fish. It is important to provide live feeds of the proper size and density to allow larvae to satisfy their increased metabolic demands. The rainbow wrasse Parajulis poecilopterus was

documented to initiate flexion at 6 mm TL and was completed at 8 mm TL (Kimura and

Kiriyama, 1993). The ornate wrasse began flexion at 20 DPH (4.4 mm TL) and was completed at 24 DPH (5.7 mm TL) (Baensch, 2017, personal communication). For the melanurus wrasse, this period was observed in larvae to begin at 15 DPH (4.866 ±

0.360 mm NL) and was first observed to be completed approximately three days later.

During this time, live feed densities were increased to accommodate increased energy demands of the larvae and flow rate was also increased to improve water quality.

Pelagic marine fish larvae complete their larval phase by exhibiting a behavior often called settlement (Victor, 1986a, 1986b). This is when larvae descend out of the open ocean and associate closely with the benthos. In aquaculture settings, most pelagic larvae display settlement behavior by reducing time spent in the open water column and gravitating closer to the sides and bottom of the culture tank, as well any additional structure. Cultured hogfish Lachnolaimus maximus larvae were observed to exhibit settlement behavior at 34 DPH and oriented strongly with the bottom of the tank

(Colin, 1982). It is hypothesized that wild Halichoeres larvae bury themselves in sand when initiating settlement and undergo metamorphosis while buried (Baensch, personal communication, 2017). This behavior has been confirmed under culture conditions in two Halichoeres species, the melanurus and ornate wrasse. During the melanurus wrasse larval trial described in this paper, sand was added to the culture tank at 25

DPH in preparation for settlement. Melanurus wrasse larvae began “settling” down into the sand between 34 and 37 DPH and emerged fully metamorphosed between 37 and

43 DPH. The presence of sand is hypothesized to act as a settling cue and lack of this

cue may have the potential to delay or inhibit settlement and subsequent metamorphosis, however this has not yet been confirmed experimentally.

Peer-reviewed literature regarding wrasse culture is generally limited to non- ornamental species such as the ballan wrasse Labrus bergylta (Hamre et al., 2013a;

Oie et al., 2017; Skiftesvik et al., 2013), humphead wrasse Cheilinus undulates (Hirai et al., 2013), and Tautog Tautoga onitis (Perry et al., 1998, 2001). To date, only ten species in the family Labridae have been cultured, with only half considered ornamental, including the Cuban hogfish Bodianus pulchellus (Ohs et al., 2018),

Hawaiian cleaner wrasse Labroides phthirophagus (Montalvo, personal communication,

2017; Sweet, 2016), bluestreak cleaner wrasse Labroides dimidiatus (Sweet, 2013), ornate wrasse H. ornatissimus (Baensch, personal communication, 2017; Sweet and

Pedersen, 2015), and melanurus wrasse H. melanurus. Culture information regarding ornamental wrasses has historically been limited to magazine and website articles, hobbyist breeding forums, social media reports, and conference presentations or abstracts. The information presented in this paper is the first of its kind for ornamental wrasse aquaculture and will help lay the groundwork for the culture of new wrasse species.

To the authors’ knowledge, this manuscript represents the first published report detailing the completion of the life cycle for an ornamental wrasse species and is a significant breakthrough in the marine ornamental aquaculture industry. Achieving

0.50% survival from hatching to metamorphosis represents a significant accomplishment and is encouraging for future culture efforts with this and other

Halichoeres species, given the short time and limited resources dedicated to

development of commercial production protocols. However, survival needs to continue to be increased to make the species a viable candidate for commercial aquaculture.

Successful captive production of several generations of this species shows great progress towards the development of reliable aquaculture protocols. Through the process of domestication, the opportunity exists to select for genes that confer desirable biological traits such as increased growth rates and disease resistance, decreased time to sexual maturity, and enhanced feed conversion ratio. Further research to optimize production protocols for this species is warranted as the melanurus wrasse is a popular wrasse species in the U.S. aquarium trade and the advancement towards its commercial production could provide consumers with an alternative option to those wild harvested.

Table 2-1. Water parameters measured throughout the 45-day H. melanurus larval rearing trial.

Parameter Value Temperature (°C) 24.80 ± 0.46 Salinity (g L-1) 35 ± 0 pH 8.04 ± 0.05 DO (mg L-1) 6.09 ± 0.41 -1 Alkalinity (mg L CaCo3) 234.2 ± 15.8 TAN (mg L-1) 0.00 - -1 NO2 (mg L ) 0.10 ± 0.05 - -1 NO3 (mg L ) 60 ± 20

Figure 2-1. H. melanurus sexual dimorphism; A) Juvenile/female with two dorsal ocelli and one caudal ocellus (enlarged middle ocellus); B) Female or male with one dorsal ocellus and one caudal ocellus; C) Terminal male with no ocelli, pink facial barring, and yellow patches around pectoral fins. Photo courtesy of author

Commercial Diet

Artemia

Instar 3-4 Nauplii

Instar 1-3 Nauplii

Algae

0 5 10 15 20 25 30 35 40 Days Post Hatch (DPH)

Figure 2-2. Feeding regime for 45-day H. melanurus larval rearing trial. Commercial Diet refers to Top Dressed Otohime (particle size A2-B1). Artemia sp. refers to newly hatched brine shrimp. Instar Nauplii refers to Parvocalanus crassirostris copepods at the noted Instar stages. Algae refers to Tetraselmis chuii.

Figure 2-3. Wild caught melanurus wrasse broodstock spawning frequency and number of fertilized eggs per spawn from August 2016-March 2017. Original sex ratio was 1M:13F but transitioned to being male dominated by March 2017. Lunar cycle is represented in the figures by black circles (full moon).

Figure 2-4. Larval development of Halichoeres melanurus; A) Late neurula stage embryos, approximately 14 hours post fertilization, diameter 0.627 ± 0.013 mm, n=50; B) 0 Days post hatch (DPH) newly hatched larva with large yolk sack and oil globule, 1.590 mm NL; C) 1 DPH with unpigmented eyes, an unopened mouth, yolk sack, and oil globule, 2.053 mm NL; D) 3 DPH with fully developed eyes, an open mouth, and absorbed yolk sac and oil globule indicating initiation of exogenous feeding, 2.234 mm NL; E) 7 DPH with separated posterior and anterior gastrointestinal tract, 2.748 mm NL;

Figure 2-4 Continued. F) 11 DPH with inflated swim bladder and increased pigmentation, 3.847 mm NL; G) 15 DPH with development of the hypural plates, beginning of flexion, and increased gut complexity, 4.866 mm NL; H) 18 DPH with completion of flexion and hypural plate formation, 5.180 mm SL; I) 37 DPH with completed metamorphosis post emergence from sand bed, increased pigmentation, and formation of ocellated spots on caudal fin and dorsal fins, completed formation of dorsal, caudal, and anal fins, 11.901 mm SL; J) 45 DPH with increased pigmentation and more defined lateral striping and ocellated spots, 13.265 mm SL. Photo courtesy of author.

Notochord Length Standard Length

Figure 2-5. Larval growth of H. melanurus over 45-day culture period. Notochord length was measured from the anterior of the snout to the posterior tip of the notochord pre-flexion. Standard length was measured from the anterior of the snout to the posterior end of the hypural plates post-flexion. All values are represented as mean ± SD. For observations on 0-2, 4-6, 8-12, DPH (n=10), 3 DPH (n=14), 7 DPH (n=7), 13 DPH (n=9), 14 DPH (n=11), 15-19 DPH (n=8), 20-22 (n=6), 25 DPH (n=3), 29, 37, 45 DPH (n=2), 27-28, 30 (n=1). Equation describing exponential trendline: 푦 = 2.0264푒0.0484푥 with an R2 value of 0.9618.

Figure 2-6. First generation melanurus wrasse spawning frequency, spawn size, and fertilization rate from September 2017-December 2017 with relation to the full moon cycle. Lunar cycle is represented in the figure by black circles (full moon).

CHAPTER 3 DEVELOPMENT OF EARLY LARVICULTURE PROTOCOLS FOR THE MELANURUS WRASSE Halichoeres melanurus

Introduction

The United States imports an estimated 7-11 million individual tropical marine fish from over 1,800 species and 125 families every year (Rhyne et al., 2012, 2017) to satisfy the demand of public aquariums, research institutions, private businesses, and over 2 million households (APPA, 2018). An alternative option to wild caught ornamental fish is the production of cultured counterparts. The marine ornamental aquaculture industry has grown considerably over the last few decades with the number of available captive bred species slowly increasing (Sweet, 2016). However, substantially less than 10% of marine species in the aquarium trade are commonly available from cultured sources, meaning the vast majority of species are still harvested directly from wild stocks across the globe (Calado et al., 2017; Wabnitz et al., 2003).

Currently, most cultured marine species exhibit demersal spawning strategies. Early life history traits that characterize this group include parental care and relatively large precocial larvae that result in high survival past metamorphosis (Calado et al., 2017;

Olivotto et al., 2011, 2017). Pelagic spawning species represent a minority of ornamental fish currently in production. Newly hatched larvae of these species are altricial (Gopakumar et al., 2009) with small mouth gapes that severely limit live feed options and survival past metamorphosis is low (Calado et al., 2017; Olivotto et al.,

2011, 2017).

Embryos produced from pelagic spawning species receive no parental care and environmental factors such as temperature greatly affect their rate of development and survival. For every fish species, there exists a temperature range corresponding to

native water temperatures, that will result in proper embryological development (Gray,

1928). Deviation from species specific temperature ranges can result in disruption of embryonic development often leading to mortality. It is well known that a strong inverse relationship exists between temperature and embryonic development time in fishes

(Beacham and Murray, 1990; Brown et al., 2011; Gray, 1928; Pauly and Pullin, 1988;

Pepin, 1991). Embryological development can be accelerated by increasing water temperature, which in turn decreases time to hatch and has been shown to result in smaller larvae (Brown et al., 2011; Gray, 1928). This occurs because a larger proportion of the yolk reserve is required to sustain the embryo, decreasing the proportion available for the formation of new tissues (Gray, 1928). Conversely, slower embryonic development can be achieved at a lower temperature, resulting in extended time to hatch and larger larvae (Brown et al., 2011; Gray, 1928). When attempting to culture pelagic spawning marine species, it is often challenging to provide first feed items of the proper size due to inherently small mouth gapes. It would be advantageous to elucidate a temperature range resulting in protracted incubation and thus enhanced embryological development leading to larger, more robust larvae. However, deviation too far outside of the species natural temperature range can result in decreased embryonic survival, an undesirable result when attempting to culture a species commercially.

Prior to the initiation of exogenous feeding, newly hatched larvae are exceedingly fragile and require appropriate environmental parameters such as microalgal density and light intensity for survival. These parameters are often species specific and if they fail to be met, mass mortalities can occur from increased stress

(Meeren et al., 2007; Pereira-Davison and Callan, 2018; Villamizar et al., 2011). Few studies have been performed evaluating the specific environmental parameters required for marine ornamental larvae during the initial developmental period between hatching and initiation of exogenous feeding, which is the first bottleneck in larviculture. Light is a critical environmental parameter that effects numerous aspects of larviculture such as hatching synchronicity, circadian system development, feeding, swim bladder inflation, and ultimately growth and survival (Barahona-Fernandes, 1979; Blanco-Vives et al.,

2010; Henne and Watanabe, 2003; Pereira-Davison and Callan, 2018; Puvanendran and Brown, 2002; Villamizar et al., 2009, 2011). Thresholds for light intensity have been postulated to be species specific and can vary with stage of development (Villamizar et al., 2011). Artificial lighting conditions used during culture can be far more intense than conditions larvae have evolved to develop under and have been shown to act as an environmental stressor and disrupt sense of orientation (Naas et al., 1996; Villamizar et al., 2011). Reduction in light intensity can be achieved through the addition of microalgae to the culture tank, often referred to as “greenwater” culture (Palmer et al.,

2007). Further research is needed to optimize these environmental parameters to increase larval survival to the exogenous feeding stage and ensure larvae are well positioned for the myriad of ontogenetic hurdles that must be overcome prior to metamorphosis.

Once larvae have fully absorbed their yolk sac and oil globule reserves, they must begin exogenous feeding to sustain increasing energy demands for proper growth and development (May, 1974; Thresher, 1984; Turingan et al., 2005; Yúfera and Darias,

2007). Many factors contribute to successful prey recognition and ingestion.

Recognition is thought to be largely based on prey appearance, locomotion, and olfaction (Fernandez-Diaz et al., 1994; Hunter, 1981; Lee et al., 2018; Ronnestad et al.,

2013; Turingan et al., 2005), as well as environmental conditions such as turbidity and light intensity. Greenwater is also used in aquaculture to manipulate turbidity. This practice has been shown to improve larval survival, growth, and feeding performance in several marine foodfish species such as barramundi Lates calcarifer, silver seabream

Pagrus auratusI, sand sillago Sillago ciliate, Atlantic cod Gadus morhua, cobia

Rachycentron canadum, and Atlantic halibut Hippoglossus hippoglossus (Faulk and

Holt, 2005; Meeren et al., 2007; Naas et al., 1992; Palmer et al., 2007). Larvae must then catch and ingest the identified prey item, which is dependent upon the larvae’s ability to visually track and capture their target as well as having an appropriate gape size to physically consume the prey (Turingan et al., 2005). Elucidating requisite microalgae concentrations that yield high survival and promote prey capture is an essential step in the development of aquaculture production protocols for any marine species.

First feeding is another significant bottleneck in marine larviculture, one that is often more difficult to overcome in pelagic spawning species because of their characteristically small and fragile larvae with inherently small mouths (Baensch and

Tamaru, 2009; Holt, 2003; Turingan et al., 2005). Newly hatched rotifers Brachionus spp. (100-340 µm in length, ≥ 60 µm in width) and Instar I Artemia spp. (400-500 µm in length) are live feed staples of the marine ornamental aquaculture industry that are commonly used as a first feed item for the larvae of most demersal spawning species

(Calado et al., 2017; Davis et al., 2018). These live feed types are relatively inexpensive

to produce and do not require intense labor. For gape-limited larvae, common live feeds such as rotifers may be simply too large or their nutritional profile inadequate. In recent years, alternative live feed types have been explored such as copepods, tunicates, ciliates, and dinoflagellates that are much smaller and exhibit different behavior, potentially affecting larval feeding response (Moorhead and Zeng, 2010; Olivotto et al.,

2011).

Although clearly defined culture protocols for these species are lacking and they are often more expensive to produce than rotifers and Artemia sp., they may be the only option for smaller, gape-limited, nutritionally demanding larvae (Moorhead and Zeng,

2010; Stottrup, 2000). Calanoid copepod species such as Parvocalanus spp. and

Pseudodiaptomus spp. have become more prominent in the marine ornamental aquaculture industry although few facilities produce them on a commercial scale.

Nauplii of these copepod species have been used successfully as a first feed item for altricial larvae that are unable to ingest rotifers (Calado et al., 2017; Cassiano et al.,

2011; Moorhead and Zeng, 2010; Stottrup, 2000). Compared to rotifers, copepod nauplii are inherently superior in their nutritional composition and do not require enrichment.

Copepods contain increased amounts of highly unsaturated fatty acids (HUFAs), especially eicosapentaenoic acid (EPA, 20:5n‐3) and docosahexaenoic acid (DHA,

22:6n‐3), which are essential for proper larval growth and development (Calado et al.,

2017; Hamre et al., 2013b; Hamre et al., 2008; Stottrup, 2000). The continued improvement of copepod culture protocols is essential to developing commercial production protocols for pelagic spawning species such as wrasses.

Wrasses, family Labridae, are very popular in the aquarium trade and are the second most imported family of marine ornamental fish into the United States (U.S.) by volume and first by number of species (Rhyne et al., 2012). Despite their immense popularity, there has been limited culture success with no commercial production of any ornamental species, mainly due to complexity and fluidity of mating social structure

(Robertson, 1981; Thresher, 1984, 1979; Warner and Robertson, 1978) and early life history characteristics such as reduced mouth gape, poor survival, and extended pelagic larval duration (Victor, 1986a). The high market demand and absence of captive production highlights the need for research to develop commercial aquaculture protocols (Murray and Watson, 2014). Primary literature evaluating environmental parameters for wrasse larviculture is scarce and limited to non-ornamental species such as the rainbow wrasse Parajulis poecilopterus (Kimura and Kiriyama, 1993), hogfish

Lachnolaimus maximus (Colin, 1982), humphead wrasse Cheilinus undulates (Hirai et al., 2013), tautog Tautoga onitis (Perry et al., 2001, 1998), and ballan wrasse Labrus bergylta (Hamre et al., 2013a; Skiftesvik et al., 2013). Information on ornamental wrasse culture is predominantly limited to online media and magazine articles, however recent scientific advances have been made with the Cuban hogfish Bodianus pulchellus (Ohs et al., 2018) and melanurus wrasse Halichoeres melanurus (Groover et al., 2018;

Barden et al., 2016).

By counting the number of daily increments between the center of the otolith and the mark corresponding to settlement, a study conducted by Victor (1986) determined that Halichoeres wrasses have relatively short larval durations averaging 26.5 days.

This indicates that Halichoeres wrasses have good potential for commercial production

due to relatively short larval durations compared to other wrasse species. The melanurus wrasse H. melanurus, first described by Bleeker (1851), is found widespread throughout the Western Pacific including Indonesia, Micronesia, and Samoa (Kuiter,

2010). Due to its small size (maximum 12 cm TL), distinctive markings and constant activity, it is in the top 20 most imported wrasse species into the U.S. for the aquarium industry with ~12,000 individuals imported in 2011 (www.aquariumtradedata.org).

Popularity in the industry coupled with a relatively short larval duration (mean 22.1 days;

Victor, 1986) highlight the economic potential for commercial production of the melanurus wrasse. The melanurus wrasse was first successfully cultured in 2015 at the

University of Florida Tropical Aquaculture Laboratory (Barden et al., 2016). Although only a few individuals survived past metamorphosis, this achievement laid the groundwork for the following research, which aimed to improve upon previous production methods. Before the ornamental aquaculture industry can begin producing ornamental wrasse species economically, it is crucial for larviculture parameters to be elucidated. Improved larval survival, growth, and feeding would ultimately increase culture efficiency and contribute to the development of reliable commercial aquaculture protocols.

Materials and Methods

Embryos from one F1 melanurus wrasse harem were used in experiments 3-1, 3-

2, and 3-6, while a combination of embryos from F0 and F1 harems were used in experiments 3-3, 3-4, 3-5, and 3-7.

Acquisition and Quarantine of Broodstock

Wild melanurus wrasses were sourced from commercial collectors in the

Philippines and Fiji and shipped by air freight to an ornamental fish wholesale company

in 2016 and 2017 (Segrest Farms, Gibsonton, FL, USA). The fish were then transported to the University of Florida Tropical Aquaculture Lab (UF-TAL) in Ruskin, FL where they were quarantined for 30 days. Wrasses were gradually acclimated to the salinity (30 ± 1 g L-1) and temperature (26 ± 1 °C) of the quarantine system. Saltwater used in all marine culture systems was prepared by mixing reverse osmosis water with synthetic sea salts until the desired concentration was achieved (Instant Ocean, Spectrum

Brands, Blacksburg, VA, USA). The ~1,500 L recirculating quarantine system was composed of three, 378 L, round, fiberglass holding tanks, with a dark interior, connected to a 378 L sump. System filtration included a fluidized sand bed, degassing column, and nitex mesh filters (200 and 100 µm). All tanks were covered with 0.635 cm mesh to keep fish securely contained. In the wild, melanurus wrasses bury themselves in the sand for protection (Kuwamura et al., 2000). To accommodate this natural behavior, a sand bed ~5 cm deep and covering ~ ¼ of the tank bottom was added. An assortment of PVC pipes was also provided on the tank bottom to increase habitat complexity and provide additional refuge. Throughout the duration of the quarantine period, wrasses were fed twice per day until apparent satiation. The diet consisted of a

0.840–1.410 mm sinking crumble (Reed Mariculture, Campbell CA, USA: APBreed Top

Dressed Otohime-C2, 49% protein, 14% fat, 3.5% fiber, 6.5% moisture, 15% ash, 1.5%

Phosphorous, and min 250 ppm Astaxanthin) and PE Frozen Mysis Shrimp (Mysis relicta) (Piscine Energetics, Vernon, British Columbia, Canada, 69.5% protein, 8.35% fat, 2.75% fiber, and 5.5% ash). In preparation for movement into the broodstock system, the salinity of the quarantine system was gradually increased via the addition of synthetic sea salt to 35 g L-1 by the end of the quarantine period. The following water

quality parameters were maintained throughout the quarantine period: 0 mg L-1 total ammonia nitrogen (TAN), 0 mg L-1 nitrite, < 40 mg L-1 nitrate, pH 8.0 ± 0.1, alkalinity

179.0 ± 17.9 mg L-1 calcium carbonate, and dissolved oxygen (DO) > 6.0 mg L-1. Once quarantine was completed, the fish were moved to the broodstock system. Fish began spawning regularly within two weeks, providing embryos used to culture first generation

(F1) melanurus wrasses. Twelve F1 melanurus wrasses were cultured to sexual maturity at UF-TAL (Groover et al. 2018), which served as broodstock for the embryo incubation and larviculture experiments described herein.

Broodstock Husbandry

The ~34,000 L broodstock system consisted of twelve, 2,650 L round, fiberglass, tanks with black walls and white bottoms. All tanks holding melanurus wrasse broodstock were covered with 0.635 cm mesh to prevent fish from jumping. A single airstone positioned at the tank bottom provided aeration. Filtration consisted of a bead filter, fluidized sand bed, protein skimmer, moving bed bioreactor with Kaldnes biomedia, and two 150-watt UV sterilizers. To accommodate the wrasses’ natural burrowing behavior, one 40 x 32 x 15 cm “sandbox” with at least 8 cm of thoroughly rinsed Quikrete® Pool Filter Sand (particle size 0.85-0.425 mm) was placed at the bottom of each tank. To provide shelter, PVC pipes of various sizes and artificial coral pieces were also furnished.

F1 melanurus wrasses were maintained in a separate ~9,000 L broodstock system comprised of twelve, 757 L rectangular concrete vats. Filtration for this system consisted of a fluidized sand bed, protein skimmer, and a single 80-watt UV sterilizer.

The 12 F1 melanurus wrasses were held in a single vat with a similar setup to that of

broodstock held in 2,650 L tanks with a “sandbox,” single airstone, PVC pipes, and a mesh cover.

The diet for all wrasse broodstock consisted of a 0.840–1.410 mm sinking crumble (APBreed Top Dressed Otohime-C2), PE Frozen Mysis Shrimp (M. relicta),

LRS Fertility Frenzy (Larry’s Reef Services, Advance, NC, USA, 15% protein, 3% fat,

1% fiber, 82% moisture, and 1 million CFU/gram microorangisms/probiotics), and LRS shad roe (American shad Alosa sapidissima, Larry’s Reef Services, Advance, NC, USA,

23.24% protein, 0.91% fat, 1.12% fiber, 74.58% moisture). Salinity was maintained between 33-36 g L-1 and measured every other day using a digital refractometer

(Milwaukee Instruments, Inc., Rocky Mount, NC). Water temperature was maintained using external heat pumps set to maintain a range of 25.5-27.5°C. Water quality parameters including DO, TAN, nitrite, nitrate, alkalinity, and pH were tested at least once every other week. DO was measured using a YSI ProODO meter (YSI

Incorporated, Yellow Springs, OH, USA). TAN, nitrite, and alkalinity were evaluated using standardized colorimetric protocols according to the manufacturer’s directions

(Hach, Loveland, CO, USA). Nitrate was tested using the API Nitrate Test Kit (Mars

Fishcare North America, Inc., Chalfont, PA, USA). A YSI EcoSense pH100A meter (YSI

Incorporated, Yellow Springs, OH, USA) was used to measure pH.

Egg Collection and Analysis

Water entering 2,650 L round fiberglass tanks created a circular flow. This directed movement assisted in the passive collection of buoyant embryos, which exited the tank via directed overflow and were then collected in 20 L buckets lined with 200 µm mesh (DiMaggio et al., 2017). Egg collectors were set daily before dusk then retrieved the next morning. A simple airlift collector lined with 200 µm mesh was used to

concentrate buoyant F1 melanurus embryos in the vat system. Spawn quantity was enumerated volumetrically for each experiment. Five 3 mL samples were taken from a container of homogenized embryos in a known volume of system water and the number of fertilized and unfertilized eggs were enumerated under a dissecting microscope. If multiple spawns were used in an experiment, embryos were combined and homogenized before enumeration. The mean number of fertilized and unfertilized eggs per mL was determined and multiplied by the total volume of water (mL) to calculate the total number of fertilized and unfertilized eggs in each spawn. A fertilization of at least

50% was required for a spawn to be utilized in an experiment.

Experimental Design of Embryo Incubation Experiments

Water temperature and its effect on melanurus wrasse embryo incubation time, percent survival, and larval size at hatch were evaluated across two experiments. For each experiment, water temperatures of 22, 25, and 28C were maintained in separate

89.9 cm x 42.5 cm x 14.9 cm water baths housed in a temperature-controlled room, maintained at 20  1C. Vigorous aeration from a single airstone positioned in the center of each bath sustained a uniform temperature throughout. Water bath temperatures were maintained using 100-300-watt aquarium heaters (Penn-Plax Inc.,

Hauppauge, NY, USA). A single Hobo Temperature Data Logger (Onset Computer

Corporation, Bourne, MA, USA) was submerged in each water bath, recording temperature every 15 minutes.

For each experiment, an airlift egg collector was set in the vat containing the harem of F1 melanurus wrasses at 16:00 (3.5 hours before dusk). The egg collector was monitored for eggs every hour starting at 17:00. At 20:00 (0.5 hours after dusk),

fertilized and unfertilized eggs were found in the collector. Spawn quantity and percent fertilization were determined as previously described.

Experiment 3-1 Effect of Temperature on Embryo Incubation Time and Larval Survival at Hatch

A single fertilized embryo (8-16 cell stage) was transferred to each well of fifteen,

24-well microplates (Corning, Tewksbury, MA, USA). Each well contained 1.5 mL of newly mixed salt water, matching the temperature and salinity of the broodstock system from which embryos originated. Upon completion of stocking at 22:00 (2 hours post collection), microplates (n=5/treatment) were randomly assigned for incubation in experimental water baths and left floating overnight with no light.

At 9:00 (11 hours post stocking, 13 hours post collection) the following morning, microplates were individually removed from the bath and embryos were inspected under a dissecting microscope. Each well within each microplate was inspected for embryo mortality and to ascertain if a hatched larva was present. Translucent embryos with visible development were deemed viable, while embryos with opaque cytoplasm were deemed nonviable. The number of viable and hatched embryos in each microplate were recorded every 1.5 hours until all viable embryos were hatched. Replicate microplates were evaluated in temperature-controlled environments similar to those of the experimental treatments to prevent fluctuations.

Experiment 3-2 Effect of Temperature on Larval Size at Hatch

A second trial was conducted to assess the effect of temperature on larval size at hatch. Fifty hand-counted fertilized embryos were stocked into 100 mL cups filled with seawater (n=4/treatment) then randomly assigned to each of the three temperature baths. All four replicates from each bath treatment were removed at a time point when

at least 50% of viable embryos in each temperature treatment hatched during the previous experiment (3-1). Replicates from the 28, 25, and 22°C baths were removed and evaluated 14, 20, and 26 hours post stocking, respectively. For each sampling point a minimum of ten hatched larvae were removed from each replicate with a 3 mL pipet, positioned laterally and flush on a Sedgewick Rafter Counting Chamber to provide scale, and immediately photographed under a trinocular dissecting microscope capable of image capture (Jenoptik, 40 ProgRes Capture Pro v2.8.8, Jupiter, FL, USA). Larval photographs were later analyzed for notochord length (NL) using ImageJ v1.50i software (U. S. National Institutes of Health, Bethesda, Maryland, USA). Notochord length was defined as the length from the anterior most point of the larvae to the posterior tip of the notochord.

Experimental Design of Larviculture Experiments

A series of experiments were conducted to evaluate effects of algae density, shading method, prey type and prey density on melanurus wrasse larval growth, survival, and feeding percentage. Experimental larviculture systems and methodologies were kept consistent among the five larval investigations and are described herein.

Deviations from these protocols are specifically noted under the subheading for each investigation.

Larviculture experiments were conducted in 13 L (working volume) tanks within a single system. Fiberglass larval tanks were cylindrical with a black interior and a single white stripe at the base to assist with visualizing larvae. Tanks were static with no water flow for the duration of the larval experiments. Light aeration was supplied by single airstone positioned centrally at the bottom of each tank and adjusted daily as needed to

maintain uniformity among replicates. Live Tetraselmis chuii microalgae used in experiments was batch cultured on site and harvested from carboys as needed. Algal cell densities (cells mL-1) were determined daily using a hemocytometer. Prior to introduction, algae was temperature acclimated to match conditions of the experimental larval tanks. Algae was added once to required treatments before the lights were turned on to reach desired cell densities prior to the addition of embryos.

Egg collectors for melanurus wrasse broodstock tanks were set before dusk the night prior to stocking experiments. The following morning, embryos were collected and evaluated for spawn quantity and percent fertilization. Tanks were randomly assigned to treatments. Experimental tanks were randomly stocked with 150 hand counted viable embryos per tank (11.5 larvae mL-1).

Larval growth and survival were assessed when larvae reached 3 DPH. The entire contents of each larval tank were transferred using a small diameter siphon and collected in a submerged 250 µm sieve, which allowed for concentration of larvae with little to no physical damage. Larvae were subsequently enumerated and photographed as previously described for later analysis. Water parameters including salinity, temperature, TAN, nitrite, nitrate, alkalinity, DO, and pH were tested in the larviculture system on the first day of each experiment and every tank was tested on the final day of each experiment. Water parameters were evaluated using methods previously stated.

The temperature of the static experimental tanks in the larviculture system was regulated through ambient temperature of the climate-controlled room. Water parameters for all larviculture experiments are presented in Table 3-1. Light intensity

was measured using a MW Portable Lux Meter (Milawakee Electric Tool Corporation,

Brookfield, WI, USA).

Experiment 3-3 Effect of Algal Density on Larval Growth and Survival to Three Days Post Hatch

The effect of algal density on larval growth and survival of melanurus wrasse larvae at 3 DPH was assessed using four algal density treatments of 0; 50,000;

100,000; and 200,000 cells mL-1 (n=4/treatment). Replicates were stocked temporally among two trials, one day apart, due to limited number of embryos. For each trial, a minimum of one replicate from each of the four experimental treatments was included.

Throughout the three-day trial, photoperiod was maintained at 16 hours light: 8 hours dark. Light intensity at the water’s surface for all treatments was 1188 ± 151 lux. Light intensity at the tank bottom for 0; 50,000; 100,000; and 200,000 cells mL-1 treatments measured 547 ± 83, 319 ± 53, 214 ± 24, and 84 ± 20 lux, respectively.

Experiment 3-4 Effect of Shading Method on Larval Growth and Survival to Three Days Post Hatch

The effect of shading method on melanurus wrasse larval growth and survival was assessed using three shading method treatments (n=5/treatment). Replicates were stocked temporally among two trials, one day apart, due to limited number of embryos.

For each trial a minimum of one replicate from each of the experimental treatments was included. One treatment shaded the tank with 50,000 cells mL-1 algae while the second treatment shaded the tank by covering the top with window screen. Both algae and window screen treatments resulted in a common light intensity of 300 lux at the tank bottom. In the third treatment, replicate tanks were covered entirely with an opaque plastic sheet that resulted in a reading of 0 lux measured at the tank bottom.

Throughout the three-day trial, photoperiod was maintained at 16 hours light: 8 hours

dark. The 50,000 cells mL-1 algae shading treatment was chosen because it was the lowest algal density that was shown to significantly improve larval survival in Experiment

3-3.

Experiment 3-5 Effect of Algal Density and Prey Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

The effect of algal density on proportion of larvae feeding at 3 DPH (first feeding) was assessed with three algal density treatments of 100,000; 300,000; and 500,000 cells mL-1 (n=5/treatment). Replicates were stocked temporally among two trials, one day apart, due to limited number of embryos. For each trial, a minimum of one replicate from each of the experimental treatments was included. A prey density of 5

Parvocalanus crassirostris copepod nauplii (< 75 µm) mL-1 was fed to all treatments.

This prey density was chosen based on previous melanurus wrasse culture success using this density. P. crassirostris nauplii were batch cultured at UF-TAL and fed a diet of live mircroalgae (Tetraselmis chuii, Tisochrysis galbana, and Chaetoceros muelleri).

A 16-hour light: 8-hour dark photoperiod was provided throughout the duration of the experiment using 32-watt fluorescent bulbs. Prior to 3 DPH, obliquely positioned ambient light was provided, resulting in a light intensity of 140 and 20 lux at the tank surface and bottom, respectively. Live feed items and algae were added once to the larval tanks prior to 9:00 when lights were turned on directly overhead at 3 DPH. Light intensity at the tank bottom for 100,000; 300,000; and 500,000 cells mL-1 treatments measured 324 ± 46, 28 ± 4, and 2 ± 1 lux, respectively. Feeding incidence was assessed by sampling 5-10 larvae in five minutes, one hour after P. crassirostris nauplii were added to the larval tank at 3 DPH. The larval tanks were fully harvested six hours after the addition of live feeds. At each time point, the sampled larvae were assessed

for evidence of prey capture by identifying the number of larvae with nauplii in their guts.

Transparent larvae were placed on a Sedgewick Rafter Counting Chamber and inspected for presence of P. crassirostris naupliar eye spots and exoskeletons in their gastrointestinal tracts, which were easily distinguished from green T. chuii algal cells.

Experiment 3-6 Effect of First Feed Type and Prey Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

The effect of first feed type on proportion of larvae feeding at 3 DPH (first feeding) was assessed with two prey treatments of unenriched Brachionus plicatilis (<75

µm) rotifers and P. crassirostris copepod nauplii (<75 µm) (n=5/treatment). A density of

5 live feed items mL-1 and an algal density of 100,000 cells mL-1 were used for both treatments. B. plicatilis rotifers were batch cultured at UF-TAL using a diet of live T. chuii and T. lutea microalgae. Rotifers and copepod nauplii were harvested, concentrated and introduced to the culture tanks in a similar volume. Methods for P. crassirostris culture, photoperiod, lighting, addition of algae and live feeds, and larval sampling and gut analysis adhere to those stated for the 3-5 study. Based on results of previous experiment 3-5, we chose to use an algal density of 100,000 cells mL-1. A density of 5 prey items mL-1 was chosen based on previous melanurus wrasse culture success using this density.

Experiment 3-7 Effect of Prey Density and Prey Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

The effect of prey density on proportion of larvae feeding at 3 DPH (first feeding) was assessed with three prey density treatments of 2.5, 5, and 10 mL-1 P. crassirostris copepod nauplii (<75 µm) (n=5/treatment). Replicates were stocked temporally among, two trials, one day apart, due to limited number of embryos. For each trial, a minimum of one replicate from each of the experimental treatments was included. An algal density

of 100,000 cells mL-1 was used for both treatments. Methods for P. crassirostris culture, photoperiod, lighting, addition of algae and live feeds, and larval sampling and prey capture analysis adhere to those stated for the 3-5 study. Based on results of previous experiments 3-5 and 3-6, we chose to use an algal density of 100,000 cells mL-1 and P. crassirostris copepod nauplii (<75 µm) as the live feed.

Statistical Analysis

Assumptions for linear models were tested prior to performance of statistical testing. Homogeneity of variance was evaluated using a Bartlett’s Test in addition to plotting residuals against predicted y-values. Normality of residuals was evaluated using a Shapiro Wilk test in addition to a q-q plot, plotting standardized residuals against theoretical quantiles. Independence of errors was achieved through randomization in experimental design. A one-way analysis of variance (ANOVA) was used to detect statistical differences among treatments in the following experiments: 3-1, 3-2, 3-3, and

3-4. Proportional survival data from experiment 3-3 was arcsine square root transformed to meet the assumptions of a linear model. A two-way ANOVA was used to detect statistical differences among treatments in experiment 3-5. When statistical differences were detected, a post hoc pairwise comparison was conducted using a

Tukey HSD test. A paired t-test was performed to detect statistical difference of the proportion of larvae feeding between 2 and 6 hours post feed introduction treatments in experiment 3-6. Experiment 3-7 data did not meet the assumptions of a linear model, therefore a generalized linear model (GLM) in the base package of RStudio

(family=poisson, V. 0.99.903 2015. RStudio: Integrated Development for R. RStudio,

Inc., Boston, MA), followed by a two-way ANOVA, was used to test for differences among the categorical predictor variables and any interaction between groups. A post

hoc pairwise comparison was performed using a Tukey HSD test. All statistical analyses were performed using JMP Pro v13.0.0 (SAS Institute, Cary, NC) unless otherwise stated. All data presented in this manuscript is depicted as the mean ± SD. A priori significance levels were set at P≤0.05.

Results

Experiments 3-1 and 3-2 Effect of Temperature on Incubation Time, Larval Size at Hatch, and Survival

Throughout the duration of the trial, water bath temperatures for 22, 25, and 28°C treatments measured 22.36  0.46, 25.10  0.29, and 28.86  0.41C, respectively.

Melanurus wrasse embryos hatched within a 3-hour time span from 21:00-24:00 and

15:00-18:00 when incubated at 22 and 25°C, respectively (Figure 3-1). Embryos hatched within a 4.5-hour time span from 10:30-15:00 when incubated at 28°C (Figure

3-1).

Larval survival at hatch did vary significantly among temperature treatments

(F2,12=42.8515, P<0.0001), specifically between 22 and 28°C (P<0.0001) and 22 and

25°C (P<0.0001) (Figure 3-2). Melanurus wrasse larval survival at hatch significantly decreased when embryos were incubated at 22°C (38 ± 12%) when compared to 25 (80

± 5) and 28°C (88 ± 10%) (Figure 3-2).

Melanurus wrasse larval notochord length (mm) at hatch was found to be significantly different among temperature treatments (F2, 9=19.3331, P=0.0006), with larvae in the 28°C treatment found to have a statistically shorter notochord length

(1.3101 ± 0.0264 mm) than those incubated in 22°C (1.3976 ± 0.0202 mm) (P=0.0004) and 25°C (1.3645 ± 0.0099 mm) (P=0.0101) treatments (Figure 3-3).

Experiment 3-3 Effect of Algal Density on Larval Growth and Survival to Three Days Post Hatch

Mean larval notochord length was not found to differ significantly among algal density treatments (F3,14=0.5068, P=0.6839). Mean larval notochord lengths were generally consistent across all treatments and measured 2.125 ± 0.055, 2.144 ± 0.186,

2.169 ± 0.108, and 2.225 ± 0.072 mm, for algal densities of 0, 50,000, 100,000, and

200,000 cells mL -1, respectively.

Larval survival was found to be significantly different among algal density treatments (F3,12=5.741, P=0.0113), with larval survival in the treatment with no algae

(12 ± 2%) significantly less (P≤0.0170) than survival in treatments with 50,000 (35 ±

15%), 100,000 (30 ± 5%), and 200,000 cells mL -1 (34 ± 11%) (Figure 3-4).

Experiment 3-4 Effect of Shading Method on Larval Growth and Survival to Three Days Post Hatch

Melanurus wrasse larval notochord length was found to be significantly different among shading method treatments (F2,12=9.8060, P=0.0030). Mean larval notochord length for the plastic treatment (2.021 ± 0.022 mm) was significantly (P≤0.0237) smaller than notochord lengths observed in both the algae (2.147 ± 0.053) and window screen treatments (2.112 ± 0.057) (Figure 3-5).

Larval survival was not found to be vary significantly among shading method treatments (F2,12=0.3076, P=0.7409) (Figure 3-6). When larvae were shaded with algae, window screen, and plastic, mean larval survival ranged only 8% among treatments with mean survival of 46.2 ± 16.8, 43.4 ± 16.0, and 51.8 ± 18.8%, respectively.

Experiment 3-5 Effect of Algal Density and Feed Availability Period on Proportion of Larvae feeding at Three Days Post Hatch

There was a significant effect of algal density on the proportion of larvae feeding at 3 DPH (F2=27.9961, P<0.0001). Mean larval feeding proportion significantly decreased (P<0.0001) in a 100,000 cells mL -1 culture environment when compared to the 300,000 cells mL -1 and 500,000 cells mL -1 (Figure 3-7). No significant interaction was found between algal density and feed availability period factors (F2=0.9915,

P=0.3857), nor was there a significant effect of feed availability period and proportion of larvae feeding (F1=0.3582, P=0.5551). Mean proportion of larvae feeding after 2 and 6- hour periods were similar within algal treatments; 100,000 cells mL -1 (18.16-15.76%),

300,000 cells mL -1 (36.57-39.78%), and 500,000 cells mL -1 (37.94-32.63%) (Figure 3-

7).

Experiment 3-6 Effect of First Feed Type and Feed Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

No larvae were found to have rotifers in their guts on 3 DPH after 2 and 6-hour feed availability periods. In larval tanks fed P. crassirostris nauplii (<75 µm) the proportion of larvae feeding ranged from 25.00-42.86% (27.74 ± 16.82%) and 33.33-

50.00% (31.67 ± 18.77%) after 2 and 6-hour feed availability periods, respectively.

Proportion of larvae feeding at 3 DPH for tanks fed P. crassirostris nauplii was not found to be significantly different between feed availability periods (t6=0.9610, P=0.3831).

Experiment 3-7 Effect of Prey Density and Feed Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

Proportion of larvae feeding varied significantly among prey density treatments

(H2=12.7986, P=0.0016). Mean proportion of larvae feeding statistically decreased

(P≤0.0060) at a feeding density of 2.5 nauplii mL-1 (14.41-23.30%) compared to 5.0

(23.76-58.27%) and 10.0 nauplii mL-1 (35.33-60.50%) (Figure 3-8). There was also a significant effect of feed availability period on proportion of larvae feeding after 2 and 6 hours (H1=8.0979, P=0.0044), with mean larval feeding proportion increasing by 61.69,

145.24, and 71.24% in 2.5, 5.0, and 10.0 nauplii mL-1 treatments, respectively (Figure 3-

8). A significant interaction was not detected between prey density and feed availability period factors (H2=0.3866, P=0.8242).

Water quality parameters for all studies are presented in Table 3-1.

Discussion

As in many other egg incubation studies, an inverse relationship was found between temperature and incubation time of melanurus wrasse embryos (Beacham and

Murray, 1990; Gray, 1928; Pauly and Pullin, 1988; Pepin, 1991). Embryos incubated at the highest temperature of 28°C completed hatching before those incubated at 25°C, which hatched before those incubated at 22°C. Delay of hatching time could be advantageous for small altricial larvae by allowing increased time for embryonic development, resulting in increased larval size at hatch (Gray, 1928). Decreased incubation temperature was shown to effectively increase larval length in five species of

Pacific salmon (Beacham and Murray, 1990). This trend was also observed in the melanurus wrasse when embryos incubated at 22°C produced larvae of significantly longer notochord lengths than those incubated at 25 and 28°C. However, incubation at

22°C also significantly decreased survival, possibly due to the temperature being at the lower end of the normal temperature range for this species. Incubation temperatures of

25 and 28°C yielded the highest percent survival at hatch, being closer to the natural temperature range of the melanurus wrasse. Based on these experimental results,

decreasing embryo incubation temperature to 22°C to achieve enlarged larvae does not appear to be a viable option given the severely decreased survival rate. Decreasing incubation temperature may only be feasible for demersal spawners as wrasses exhibit rapid development even when at the lower end of their temperature threshold. It is suggested that a temperature of approximately 25°C be used to incubate melanurus wrasse embryos to achieve preferable larval survival and size at hatch.

The addition of microalgae to the larval culture environment, commonly known as

“greenwater,” is a common practice in marine fish culture (Calado et al., 2017; Faulk and Holt, 2005; Naas et al., 1992; Palmer et al., 2007). Greenwater has been shown to provide many benefits to larviculture including light attenuation and increased nutrient uptake through indirect ingestion (Meeren, 1991). Algal density was not shown to have a significant effect on larval notochord length at 3 DPH. This could be because during the first three days of development, larval growth is more likely to be affected by parameters such as temperature, which remained consistent across treatments.

The presence of microalgae within the culture tank was shown to increase larval survival significantly at 3 DPH, prior to the initiation of exogenous feeding. With larvae only able to start ingesting algal cells at 3 DPH, it is unlikely that this would impact survival. A more probable cause of increased survival was the decrease in light intensity resulting from the introduction of algae. Some altricial larvae can be extremely photo- sensitive at this early stage in development due to lack of pigmentation and underdeveloped photo-receptors (Villamizar et al., 2011). Elevated light intensities have been shown to act as an environmental stressor resulting in mortality in sea bass

Dicentrarchus labrax (Barahona-Fernandes, 1979) and southern flounder Paralichthys

lethostigma (Henne and Watanabe, 2003). If the reduction in light intensity is the true driver behind the increased survival observed in this study, then algae introduction may not be necessary to achieve elevated survival. It could be advantageous to limit algal density in larval culture tanks early in development to reduce introduction of nutrient rich media, bacteria, and additional microorganisms such as ciliates that are associated with algae cultures. Therefore, although all three treatments containing algae significantly increased larval survival when evaluated at 3 DPH, an algal density of approximately

50,000 cells mL-1 is sufficient to reduce light intensity and increase larval survival while conserving algal resources and avoiding deterioration of water quality.

Once marine larvae reach the developmental milestone of exogenous feeding, increased algal density becomes paramount to enhancement of larval feeding performance and nutrition. The presence of microalgae within the culture environment has been shown to continually enrich live feeds and increase contrast to allow larvae to better visualize prey items, thereby increasing ability to capture prey (Calado et al.,

2017; Faulk and Holt, 2005; Palmer et al., 2007). Enhanced feeding performance has been achieved through increasing algal densities in culture for species such as halibut

Hippoglossus hippoglossus (Naas et al., 1992), Atlantic cod Gadus morhua (Meeren et al., 2007), barramundi Lates calcarifer, silver seabream Pagrus auratusI, sand sillago

Sillago ciliate (Palmer et al., 2007), and cobia Rachycentron canadum (Faulk and Holt,

2005). Greenwater culture techniques have been effectively employed in many marine ornamental species such as Pacific blue tang Paracanthurus hepatus (DiMaggio et al.,

2017), yellow tang Zebrasoma flavescens (Callan et al., 2018; Pereira-Davison and

Callan, 2018), and Milletseed Butterflyfish Chaetodon miliaris (Degidio et al., 2017), as

well as many non-ornamental wrasse species including Tautog Tautoga onitis (Perry et al., 1998), hogfish Lachnolaimus maximus (Colin, 1982), and the rainbow wrasse

Parajulis poecilopterus (Kimura and Kiriyama, 1993). Greenwater has also been used in the successful culture of several ornamental wrasse species, namely the ornate wrasse

Halichoeres ornatissimus (Baensch, 2017, personal communication), Hawaiian cleaner wrasse Labroides phthirophagus (Montalvo, 2017, personal communication), and the melanurus wrasse Halichoeres melanurus. Algal densities of 300,000 and 500,000 cells mL-1 were shown to significantly improve melanurus wrase feeding performance and are recommended when culturing first feeding larvae of this species.

When larval survival was evaluated at 3 DPH, the addition of T. chuii algae to the culture did not significantly differ from larval survival in a tank shaded with window screen when the same light intensity was maintained in both treatments (Figure 3-5).

This suggests that melanurus wrasse larvae do not require microalgae for the first three days of development, instead, decreased light intensity is important as they appear to be especially photo-sensitive. Based on these results, it is suggested that artificial shading, resulting in approximately 300 lux, is provided during first feeding to increase larval survival. Furthermore, elimination of algae prior to onset of exogenous feeding could also decrease introduction of microorganisms such as bacteria and ciliates, and prevent changes in water quality potentially manifesting in further increases in survival and growth throughout the culture period.

A natural light and dark photoperiod is essential for proper larval growth and development (Villamizar et al., 2011). Complete absence of light has been shown to decrease larval growth in European sea bass (Villamizar et al., 2009) and Senegal sole

Solea senegalensis (Blanco-Vives et al., 2010). In this study, mean notochord length of melanurus wrasse larvae decreased significantly when deprived of light during the first three days of development, which is consistent with other studies and suggests that some light is required for proper growth. It is thought that a natural photoperiod containing light and dark cycles are required for the establishment of a natural circadian rhythm, which during early larval development is thought to regulate the temporal co- ordination of many physiological processes (Vallone et al., 2007; Villamizar et al., 2009).

Disruption of this rhythm can negatively affect important aspects of larval development and performance (Villamizar et al., 2009).

To achieve the highest larval survival and growth with melanurus wrasse, it is recommended to shade larval tanks to approximately 300 lux while avoiding complete darkness from hatch until the initiation of exogenous feeding. Shading should be achieved using artificial means such as screening in place of algae to limit the potential of microbial introductions and maintain water quality.

Results of this study demonstrated that melanurus wrasse larvae were not able to ingest B. plicatilis rotifers (<75 µm) at first feeding under the specific culture parameters described, suggesting that an alternative first feed type and/or adjustment in culture parameters is required for this species. Prey size is a critical factor for larval feeding, especially with gape-limited larvae such as melanurus wrasses. However, the rotifers used in this experiment were of comparable size to copepod nauplii, which were ingested by the larvae, suggesting other factors may be deterring larvae from capturing rotifers. Aside from physical gape limitations, feeding success in marine fish larvae relies on a number of compounding factors such as visual and olfactory cues, swimming

behavior, and color (Calado et al., 2017; Hunter, 1981; Wittenrich et al., 2009). Rotifers may not satisfy one or more of these criteria and therefore may not be a suitable first feed prey item for melanurus wrasse larvae.

Larvae were able to ingest P. crassirostris Instar I and II nauplii (<75 µm) successfully at first feeding, achieving a feeding percentage of approximately 28 and

32% at 2 and 6 hours post feed introduction, respectively. This suggests P. crassirostris nauplii are an appropriate feed item for melanurus larvae at initiation of exogenous feeding. P. crassirostris nauplii likely resemble the natural prey of wild wrasse larvae

(McKinnon et al., 2003; Sampey et al., 2007), successfully eliciting a feeding response.

Up to a certain threshold, an increase in prey density has been shown to increase feeding success in several marine foodfish species such as southern bluefin tuna Thunnus maccoyii, yellowtail kingfish Seriola lalandi (Hilder et al., 2015), Japanese

Spanish mackerel Scomberomorus niphonius (Shoji and Tanaka, 2004), and greenback flounder Rhombosolea tapirina (Shaw et al., 2006). Feeding percentage of melanurus larvae at initiation of exogenous feeding was found to increase significantly when prey densities of 5.0 and 10.0 P. crassirostris copepod nauplii ml -1 were added to culture tanks when compared to 2.5 P. crassirostris copepod nauplii ml -1. This suggests that a prey density greater than 2.5 mL -1 is required to increase melanurus wrasse feeding percentage at this point in development. It is recommended that a prey density between

5.0 and 10.0 P. crassirostris copepod nauplii ml-1 should be used during first feeding of melanurus larvae. Feeding percentage was also found to significantly increase from 2-6 hours after the addition of P. crassirostris copepod nauplii, indicating that larvae became more adept at capturing nauplii the longer they had access to prey within the

tank. This emphasizes the need for continued elevated prey density in the culture tank, likely requiring multiple feedings throughout the day.

To the authors’ knowledge, this paper represents the first published research investigating early larval rearing culture parameters for an ornamental wrasse species.

These experiments contributed to an initial culture protocol for the melanurus wrasse that refined many crucial parameters, improving survival, growth, and feeding success.

Although further investigation into comprehensive culture parameters for this species is necessary, results from these early larviculture trials indicate that the melanurus wrasse is a promising candidate for commercial production. Furthermore, results from these experiments may have implications for the culture of other popular aquarium wrasses in the Halichoeres genus and potentially numerous other species in the Labridae family.

Table 3-1. Water quality parameters for H. melanurus larviculture experiments represented as mean ± SD.

Total Water Alkalinity Salinity DO Ammonia Nitrite Nitrate Experiment Temperature pH (mg L-1 (g L-1) (mg L-1) Nitrogen (mg L-1) (mg L-1) (°C) CaCo ) 3 (mg L-1) 3-3 25.46 ± 0.17 34.25 ± 0.12 8.06 ± 0.06 7.06 ± 0.20 179.55 ± 8.13 0.05 ± 0.09 0.05 ± 0.06 27.50 ± 10.0 3-4 25.51 ± 0.19 34.27 ± 0.05 8.08 ± 0.05 7.12 ± 0.06 180.12 ± 8.83 0.00 ± 0.00 0.00 ± 0.00 20.00 ± 0.00 3-5 25.52 ± 0.15 34.31 ± 0.17 8.10 ± 0.03 7.13 ± 0.06 183.21 ± 8.02 0.00 ± 0.00 0.08 ± 0.08 27.14 ± 9.94 3-6 25.42 ± 0.09 34.40 ± 0.00 8.10 ± 0.01 7.10 ± 0.01 182.97 ± 8.26 0.00 ± 0.00 0.10 ± 0.00 34.00 ± 9.66 3-7 25.46 ± 0.11 34.40 ± 0.00 8.10 ± 0.01 7.11 ± 0.02 184.68 ± 7.08 0.00 ± 0.00 0.07 ± 0.00 34.67 ± 9.15

Figure 3-1. Hatch (%) over time of H. melanurus embryos in response to incubation temperature (°C) (n=5).

100 b 90 b 80 70 60 a 50

40 Survival % Survival 30 20 10 0 22 25 28 Temperature (°C)

Figure 3-2. Survival (%) of H. melanurus embryos in response to incubation temperature (°C) represented as mean ± SD (n=5). Different lowercase letters indicate significant differences P ≤ 0.05.

1.45 a

1.4 a

1.35 b

1.3

1.25 Notochord Length (mm) Length Notochord

1.2 22 25 28 Temperature (°C)

Figure 3-3. Notochord length at hatch (mm) of H. melanurus larvae in response to incubation temperature (°C) represented as mean ± SD (n=4). Different lowercase letters indicate significant differences P ≤ 0.05.

b 50 45 b 40 b 35 30 25

Survival (%) Survival 20 a 15 10 5 0 0 50,000 100,000 200,000 Algal Density (cells mL-1)

Figure 3-4. Survival (%) of H. melanurus larvae at 3 DPH in response to algal density (cells mL-1) represented as mean ± SD (n=4). Different lowercase letters indicate significant differences P ≤ 0.05.

2.25 a 2.20 a 2.15 2.10 b 2.05 2.00

1.95 Notochord Length (mm) Length Notochord 1.90 1.85 Algae Screen Plastic Shading Method

Figure 3-5. Notochord length (mm) of H. melanurus larvae in response to shading method represented as mean ± SD (n=5). Different lowercase letters indicate significant differences P ≤ 0.05.

70 a 60 a a 50 40

30 Survival (%) Survival 20 10 0 Algae Screen Plastic Shading Method

Figure 3-6. Survival (%) of H. melanurus larvae in response to shading method represented as mean ± SD (n=5). Different lowercase letters indicate significant differences P ≤ 0.05. b b

50 b b b 45 40 b 35 a 30 25 a a 20 Feeding % Feeding 15 10 5 0 100,000 300,000 500,000 Algal Density (cells/mL) 2 Hour Feeding Period 6 Hour Feeding Period

Figure 3-7. Larval feeding (%) of H. melanurus larvae in response to algal density (cells mL-1) and prey (P. crassirostris nauplii <75 µm) availability period (# hours) represented as mean ± SD (n=5). Different lowercase letters above SD bars indicate significant differences (P ≤ 0.05) within algal density treatments between prey availability period treatments. Different lowercase letters above brackets indicate significant differences (P ≤ 0.05) among algal density treatments.

80 b b b b 70 60 a 50 a 40 b a

Feeding % Feeding 30 20 a 10 0 2.5 5 10 Prey Density (nauplii mL-1)

2 Hour Prey Availability Period 6 Hour Prey Availability Period

Figure 3-8. Larval feeding (%) of H. melanurus larvae in response to prey (P. crassirostris nauplii <75 µm) density (nauplii mL-1) and prey availability period (# hours) represented as mean ± SD (n=5). Different lowercase letters above SD bars indicate significant differences (P ≤ 0.05) within prey density treatments between prey availability period treatments. Different lowercase letters above brackets indicate significant differences (P ≤ 0.05) among prey density treatments.

CHAPTER 4 personEXPLORATION OF EARLY LARVICULTURE PROTOCOLS FOR THE YELLOW WRASSE Halichoeres chrysus

Introduction

The marine aquarium trade has a global presence with a multi-billion dollar value, supplying marine ornamental species to personal and public aquariums, research institutions, and private businesses primarily in the United States, Europe, Japan, and

China (Calado et al., 2017; Rhyne et al., 2012; Thornhill, 2012). Less than 10% of the estimated 1,800 species of marine ornamental fish in the trade are captive bred with the remainder harvested from coral reefs around the world (Calado et al., 2017; Rhyne et al., 2012; Wabnitz et al., 2003). Of the small percentage of species that are captive bred, only a quarter of them represent pelagic spawning marine ornamental species

(Calado et al., 2017; Moorhead and Zeng, 2010). Culture protocols for pelagic spawning species have been slow to progress due to a variety of challenging life history traits including complex social structures, small egg size, and altricial (Gopakumar et al.,

2009) larvae with small mouth gapes. Further investigation into the culture requirements for this under-represented group of fishes is required to improve survival to metamorphosis and produce more species at a commercial scale.

Temperature has been shown to be a critical environmental parameter that can drastically affect fish throughout embryonic and larval ontogeny. Incubating fish embryos are heavily influenced by temperature, with many studies demonstrating a strong inverse relationship between temperature and incubation time (Beacham and

Murray, 1990; Gray, 1928; Pauly and Pullin, 1988; Pepin, 1991). Intuitively, an increase in temperature can accelerate embryogenesis and shorten the incubation duration prior to hatching (Gray, 1928). Conversely, embryos incubated at temperatures below their

normal range may exhibit a delay in hatching allowing further time for development and physically larger larvae at hatch (Barden et al., 2013; Gray, 1928; Pepin et al., 1997).

For every species, there is a temperature range that is most appropriate for development of embryos, corresponding to the natural temperature range of their geographical distribution. Deviation too far outside of this natural range can disrupt proper development of embryos and result in increased mortality (Gray, 1928; Pepin,

1991). Historically, most incubation temperature studies have focused on foodfish species such as Atlantic cod Gadus morhua (Pepin et al., 1997) and several species of

Pacific salmon Oncorhynchus sp. (Beacham and Murray, 1990), while those focused on marine ornamental species are scarce (Barden et al., 2013). Most culture efforts target general temperature ranges corresponding to the geographic region of the species. For many demersal spawning species, such as clownfish, this strategy is sufficient due to early life history characteristics that produce robust precocial larvae that can quickly begin successful exogenous feeding and complications due to starvation are uncommon. It is crucial however, to identify this temperature range for pelagic spawning species with altricial larvae to maximize survival and potentially manipulate incubation regimes to increase size at hatch. Production of developmentally advanced larvae at hatch may increase survival rates during early larval culture, allowing for a more prolonged transition period between endogenous and exogenous feeding. This would certainly be advantageous for the commercial production of a species that struggles with this transition period, such as wrasses and the many other pelagic spawning marine ornamental species.

The time prior to the onset of exogenous feeding is a period when growth and survival should be maximized to increase chances of successful transition to live prey.

Marine larvae are especially fragile during this stage and culture parameters such as water flow, aeration, and lighting may have pronounced effects on survival. Numerous factors have been hypothesized to influence larval growth and survival during this period such as turbidity and light intensity, but most studies focus instead on factors affecting first feeding.

Following absorption of the yolk sac and oil globule reserves by the developing larvae, exogenous feeding must begin to sustain increasing energy demands for proper growth and development (May, 1974; Thresher, 1984; Turingan et al., 2005; Yúfera and

Darias, 2007). This developmental milestone is a critical step in the larviculture process and success at this juncture may foreshadow the overall success and survival of the cohort in culture. Many compounding factors have been shown to impact larval ability to recognize and capture prey leading to first feeding success. Prey must first be identified from visual or olfactory cues such as size, shape, movement, and smell (Dixson et al.,

2012; Fernandez-Diaz et al., 1994; Hunter, 1981; Lee et al., 2018; Ronnestad et al.,

2013; Turingan et al., 2005). Prey size is often the most limiting factor for marine altricial larvae because of their small mouth gapes (Baensch and Tamaru, 2009; Holt, 2003;

Turingan et al., 2005).

Currently, rotifers Brachionus sp. and Artemia sp. are the industry standard for feeding marine ornamental fish larvae. However, rotifers and Instar I Artemia sp. range in length from approximately 100-340 and 400-500 µm, respectively (Calado et al.,

2017; Davis et al., 2018), making these live feed items too large for the larvae of many

species of pelagic spawning marine ornamental fish at first feeding. Recently, alternative live feed types such as copepods, tunicates, ciliates, and dinoflagellates have been investigated for use during this challenging nutritional stage. These more novel prey items are typically much smaller and display varying swimming behavior, compared with rotifers, potentially resulting in improved larval feeding success

(Moorhead and Zeng, 2010; Olivotto et al., 2011).

Species of calanoid copepod, especially Parvocalanus sp., have increased in popularity in the marine ornamental aquaculture industry as a first feed item for altricial larvae that are unable to successfully feed on rotifers (Calado et al., 2017; Cassiano et al., 2011; Moorhead and Zeng, 2010; Stottrup, 2000). Advantages of using copepods in larviculture include small naupliar body size (≥ 30 µm), movement that elicits a larval feeding response, and superior nutritional composition including high levels of highly unsaturated fatty acids (HUFAs), especially eicosapentaenoic acid (EPA, 20:5n‐3) and docosahexaenoic acid (DHA, 22:6n‐3) (Calado et al., 2017; Hamre et al., 2013b; Hamre et al., 2008; Stottrup, 2000). Despite the many benefits of using copepods in larviculture, current culture methods for these organisms have yet to be optimized and are inefficient compared to those of rotifers, especially in respect to production densities, labor, and cost (Moorhead and Zeng, 2010; Stottrup, 2000).

The addition of microalgae to the culture environment, commonly known as

“greenwater,” is a common practice in both marine foodfish and ornamental aquaculture

(Calado et al., 2017; Degidio et al., 2017; DiMaggio et al., 2017; Faulk and Holt, 2005;

Meeren et al., 2007; Naas et al., 1992; Palmer et al., 2007; Perry et al., 1998). This technique has been shown to continually enrich live feeds, and increase contrast to

allow larvae to better visualize prey items, thereby increasing prey capture ability

(Calado et al., 2017; Faulk and Holt, 2005; Palmer et al., 2007). Greenwater protocols have also been utilized more recently in the successful culture of several species of ornamental wrasses (Groover et al., 2018; Ohs et al., 2018; Baensch, personal communication, 2017; Montalvo, personal communication), which is a largely unexplored family of marine ornamental fish in the aquaculture industry.

Of all marine ornamental fish imported annually into the U.S., Labrids comprise the second largest percentage by volume of individual fish and the highest number of species (228) (Rhyne et al., 2012). These imported species represent a significant portion of the total number of species within the family, estimated currently to contain

548 (Parenti and Randall, 2018), including both ornamental and non-ornamental specimens There are 60 genera within the Labridae family, the most speciose genus being Halichoeres with 80 species, followed by Cirrhilabrus with 58 species, and

Bodianus with 46 species (Parenti and Randall, 2018).

To date, there have been few successful culture efforts with members of the

Labridae family. A limited number of investigations have focused on species such as taugtog Tautoga onitis and the humphead wrasse Cheilinus undulatus that have value as food fish (Hirai et al., 2013; Perry et al., 1998, 2001). More recently, a number of studies have focused on the production and culture optimization of the Ballan wrasse

Labrus bergylta as this species has been explored for use as biocontrol of external parasites in the culture of salmonids (D’Arcy et al., 2012; Hamre et al., 2013a; Skiftesvik et al., 2013). Information regarding ornamental wrasse culture is even more scarce despite their immense popularity in the aquarium hobby. A gap analysis by Murray and

Watson (2014), designed to identify disparities in hobbyist demand for families of ornamental fish and current operational efforts to culture them, categorized Labridae as

“red”, indicating high demand in the trade with no operational culture initiatives.

The yellow wrasse Halichoeres chrysus, first described by Randall (1981), is one of the most popular wrasse species in the aquarium industry due to its bright yellow color and small size with a maximum length of 12 cm (Kuiter, 2010). They are found in small harems on shallow coastal reef habitats throughout the Western Pacific, ranging from Bali to Christmas Island in the Indian Ocean (Kuiter, 2010). Like most wrasses, the yellow wrasse is a protogynous hermaphrodite, with males generally being larger and more aggressive than females. Yellow wrasses are also sexually dimorphic, loosely characterized by the number of dorsal ocelli and presence of facial barring (Randall,

1980) (Figure 4-1). Juveniles are recognized by three evenly spaced black ocelli spread across the full length of the dorsal fin, with the middle black ocelli surrounded by a thin layer of white, and no visible barring on the face. Females traditionally display three black dorsal ocelli, devoid of a white ring around the center ocellus. Larger yellow wrasses with two dorsal ocelli (anterior and central) can be female or male (Randall,

1980). Terminal males only display one anterior dorsal ocellus, normally accompanied by green and pink facial barring. This species was the third most imported wrasse species into the U.S. by volume based on invoice data in 2008, 2009, and 2011 collectively, totaling 91,665 reported individuals (www.aquariumtradedata.org). There has been no successful culture of this species through metamorphosis to date, however there are characteristics of this species that show promise for future culture success.

By counting the number of daily increments between the center of the otolith and the

mark corresponding to settlement, a study conducted by Victor (1986) determined that the yellow wrasse has a short larval duration in the wild, averaging 26.1 days. The objective of this study was to elucidate parameters for yellow wrasse embryo incubation and larviculture that yield high hatching success, larval growth, and survival. This understanding would set the foundation for further research and contribute to development of commercial aquaculture protocols.

Materials and Methods

Acquisition and Quarantine of Broodstock

Yellow wrasses, mainly consisting of juveniles and small females, were collected by commercial fishers from coastal reef habitats in the Philippines in 2016 and 2017.

Wrasses were shipped to an ornamental fish wholesale company via air freight (Segrest

Farms, Gibsonton, FL, USA) then subsequently transported to the University of Florida

Tropical Aquaculture Laboratory (UF-TAL) in Ruskin, FL for 30 days of quarantine.

Wrasses were acclimated to the salinity and temperature of the quarantine system (30 ±

1 g L-1 and 26 ± 1 °C) then directly introduced to the ~1,500 recirculating system comprised of three, 378 L holding tanks and a 378 L sump. Round holding tanks were constructed of fiberglass, contained a dark interior, and were fitted with 0.635 mesh covers to securely contain fish. System filtration was contained within the sump and included a fluidized sand bed, degassing column, and nitex mesh filters (200 and 100

µm). To replicate the natural environment of the yellow wrasse, various PVC pipes were distributed around the tanks to increase habitat complexity and provide shelter. In addition, a ~5 cm deep sand bed covering ~1/4 of the tank bottom was added to promote natural burying behavior (Gerkema et al., 2000; Videler, 1986), which wild

Halichoeres wrasses utilize for shelter and protection.

Throughout the duration of the quarantine period, wrasses were fed twice per day until apparent satiation. The diet consisted of a 0.840–1.410 mm sinking crumble

(Reed Mariculture, Campbell CA, USA: APBreed Top Dressed Otohime-C2, 49% protein, 14% fat, 3.5% fiber, 6.5% moisture, 15% ash, 1.5% Phosphorous, and min 250 ppm Astaxanthin) and PE Frozen Mysis Shrimp (Mysis relicta) (Piscine Energetics,

Vernon, British Columbia, Canada, 69.5% protein, 8.35% fat, 2.75% fiber, and 5.5% ash).

The following water quality parameters were maintained throughout the quarantine period: 0 mg L-1 total ammonia nitrogen (TAN), 0 mg L-1 nitrite, < 40 mg L-1 nitrate, pH 8.0 ± 0.1, alkalinity 179 ± 17.9 mg L-1 calcium carbonate, and dissolved oxygen > 6.0 mg L-1. By the end of the quarantine period, salinity of the system had been gradually increased through the addition of synthetic sea salt in preparation for transition of wrasses into the 35 g L-1 broodstock system. Saltwater used in all marine systems was comprised of reverse osmosis water mixed with synthetic sea salts

(Instant Ocean, Spectrum Brands, Blacksburg, VA, USA).

Once 30 days had elapsed, 20 yellow wrasses (1 male, 2 intermediate phase, and 17 females) were moved to one 2,650 L tank within the broodstock system. These fish began spawning regularly within three weeks, providing viable embryos used in subsequent studies. Spawning frequency and fertilization success of this harem of yellow wrasses over a six-month period can be seen in Figure 4-7. This quarantine procedure was repeated with successive groups of wild caught yellow wrasses to establish three harems of yellow wrasse broodstock for subsequent experimentation.

Broodstock Husbandry

Yellow wrasses were held in 2,650 L round, fiberglass tanks with black walls and white bottoms, each securely fitted with 0.635 cm mesh. At the bottom of the tank, vigorous aeration was provided with a single airstone, various PVC pipes and artificial coral increased habitat complexity, and one 40 x 32 x 15 cm “sandbox” with at least 8 cm of thoroughly rinsed Quikrete® Pool Filter Sand (particle size 0.85-0.425 mm) was provided to promote burying behavior (Gerkema et al., 2000; Videler, 1986). The broodstock recirculating system had a total volume of ~34,000 L, consisting of 12 holding tanks and a sump. System filtration consisted of a bead filter, fluidized sand bed, protein skimmer, moving bed bioreactor with Kaldnes biomedia, and two 150-watt

UV sterilizers.

27.5°C. Dissolved oxygen (DO) was measured weekly using a YSI ProODO meter (YSI

Incorporated, Yellow Springs, OH, USA). Water quality parameters including TAN, nitrite, nitrate, alkalinity, and pH were tested at least once every other week. TAN, nitrite, and alkalinity were evaluated using standardized colorimetric protocols according to the manufacturer’s directions (Hach, Loveland, CO, USA). Nitrate was tested using the API Nitrate Test Kit (Mars Fishcare North America, Inc., Chalfont, PA, USA). A YSI

EcoSense pH100A meter was used to determine pH.

Yellow wrasse broodstock were fed 4-5 times per day until satiation with a diet consisting of a 0.840–1.410 mm sinking crumble (APBreed Top Dressed Otohime-C2),

PE Frozen Mysis Shrimp (M. relicta), LRS Fertility Frenzy (Larry’s Reef Services,

Advance, NC, USA, 15% protein, 3% fat, 1% fiber, 82% moisture, and 1 million

CFU/gram microorangisms/probiotics), and LRS shad roe (American shad Alosa sapidissima, Larry’s Reef Services, Advance, NC, USA, 23.24% protein, 0.91% fat,

1.12% fiber, 74.58% moisture).

Embryos from two harems of yellow wrasses were used for all subsequent egg incubation and larviculture studies. Harem #1 initially consisted of one male, two intermediate phase individuals of undetermined sex, and 17 females, but over time several females transitioned to intermediate phase and terminal phase males, changing the sex ratio. Spawning frequency and fertilization success of Harem #1 were recorded over a six-month period (Figure 4-2). Harem #2 was originally comprised of 23 female yellow wrasses when introduced to the broodstock tank. Over the next several weeks, several individuals were observed to change sex to male and the harem began producing viable embryos. Precise sex ratio at the time of spawning was not able to be determined.

Egg Collection and Analysis

In preparation for subsequent experimentation, separate egg collectors for

Harems 1 and 2 were positioned under the overflow of each broodstock tank to collect buoyant embryos circulating at the surface. Collectors consisted of a 20 L bucket lined with 200 µm mesh and were put in place several hours prior to the onset of dusk when yellow wrasses begin spawning (DiMaggio et al., 2017). Embryos were retrieved either several hours later or the following morning as dictated by experimental design.

Spawn quantity was enumerated volumetrically for each experiment. Five 3 mL samples were taken from a container of homogenized embryos, from Harems 1 and 2, in a known volume of system water and the number of fertilized and unfertilized eggs

were enumerated under a dissecting microscope. The mean number of fertilized and unfertilized eggs per mL was determined and multiplied by the total volume of water

(mL) to calculate the total number of fertilized and unfertilized eggs in each spawn.

Percent fertilization was calculated and a fertilization success of at least 50% was required for a spawn to be used in an experiment.

Experimental Design of Embryo Incubation Experiments

Water temperature and its effect on yellow wrasse embryo incubation time, percent survival, and larval size at hatch were evaluated across two experiments. The day of each experiment, egg collectors were affixed to broodstock tanks containing yellow wrasse harems at 16:00 (3.5 hours before dusk). Egg collectors were monitored for eggs every hour starting at 17:00. Embryos were observed in the collector at 18:00

(1.5 hours before dusk) and quickly harvested. Spawn quantity and percent fertilization were determined as previously described.

For each experiment, three water baths measuring 89.9 cm x 42.5 cm x 14.9 cm were housed in a temperature-controlled room, maintained at 20  1C. Separate water temperatures of 22, 25, and 28C were maintained with submersible aquarium heaters

(Penn-Plax Inc., Hauppauge, NY, USA). Temperature was recorded in each bath every

15 minutes by a submerged Hobo Temperature Data Logger (Onset Computer

Corporation, Bourne, MA, USA). To ensure temperature uniformity in each water bath for the duration of the trial, a single airstone positioned in the center of each bath provided ample water circulation.

Experiment 4-1 Effect of Temperature on Embryo Incubation Time and Larval Survival at Hatch

For this experiment, 15 individual, 24-well microplates (Corning, Tewksbury, MA,

USA) received one fertilized yellow wrasse embryo per well, with each well containing

1.5 mL of freshly mixed seawater matching the salinity of the source broodstock system.

Upon completion of stocking at 21:30 (2 hours post collection), microplates

(n=5/treatment) were randomly assigned to incubate in experimental water baths and left floating overnight without light to maintain a natural photoperiod.

The next morning at 8:30 (11 hours post stocking, 13 hours post collection), individual microplates were removed from the bath and embryos were inspected under a dissecting microscope in temperature-controlled environments to prevent temperature fluctuations within the wells. Embryos within each individual well were inspected for mortality and to determine if a hatched larva was present. Translucent embryos with visible development were deemed viable, while embryos with opaque cytoplasm were deemed nonviable. The number of viable and hatched embryos in each microplate were recorded every 1.5 hours until all viable embryos were hatched.

Experiment 4-2 Effect of Temperature on Larval Size at Hatch

A second trial was conducted to assess the effect of temperature on larval size at hatch. Yellow wrasse embryos were collected and evaluated at the same time point as in experiment 4-1. Cups (100 mL, n=4/treatment) where stocked with 50 hand-counted fertilized eggs then randomly assigned to each of the three temperature baths as described in experiment 4-1. All replicates for each temperature treatment were removed at the time point when at least 50% of viable embryos hatched during the previous experiment. Replicates from the 28, 25, and 22°C baths were removed and

evaluated 13.5, 18, and 25.5 hours post stocking, respectively. For each sampling point a minimum of ten hatched larvae were removed from each replicate with a 3 mL pipet, positioned laterally and flush on a Sedgewick Rafter Counting Chamber to provide scale, and immediately photographed under a trinocular dissecting microscope capable of image capture (Jenoptik, 40 ProgRes Capture Pro v2.8.8, Jupiter, FL, USA). Larval photographs were later analyzed for notochord length (NL) using ImageJ v1.50i software (U. S. National Institutes of Health, Bethesda, Maryland, USA). Notochord length was defined as the length from the anterior most point of the larvae to the posterior tip of the notochord.

Experimental Design of Larviculture Experiments

Five yellow wrasse early larviculture experiments remained consistent in overarching methodologies and system design components which are described herein.

Deviations from common methodologies and specific procedures for each experiment are described under the corresponding subheading. Larval survival, growth, and feeding performance were evaluated under varying culture conditions such as algal density, prey density, and prey type. These parameters were chosen based on potential to improve culture protocols through improved larval growth, survival, and feeding performance.

Yellow wrasse broodstock egg collectors were set several hours before dusk the night prior to stocking experiments. Embryos were collected and evaluated for spawn quantity and percent fertilization the following morning. A minimum fertilization rate of

50% was required for the spawn to be used in an experiment. Experimental tanks were randomly assigned treatments and stocked with 150 hand counted viable embryos per tank (11.5 larvae L-1).

Larviculture experiments were conducted in static 13 L (working volume), cylindrical, fiberglass tanks with a black interior and a single white stripe at the base to assist with visualizing larvae. Gentle aeration was supplied by a single airstone positioned centrally at the bottom of each tank and adjusted daily as needed to maintain uniformity among replicates. Larval tanks were located in a temperature-controlled room which maintained water temperature equal to that of ambient room temperature and ensured consistency among all treatments. Light intensity was measured using a MW

Portable Lux Meter (Milawakee Electric Tool Corporation, Brookfield, WI, USA). Water parameters including salinity, water temperature, dissolved oxygen (DO), TAN, nitrite, nitrate, alkalinity, and pH were tested in the larviculture system on the first day of each experiment and every replicate tank was tested on the final day of each experiment.

Water parameters were evaluated using methods stated previously. Water quality parameters for all larviculture experiments are presented in Table 4-1.

Live Tetraselmis chuii microalgae used in experiments was batch cultured on site and harvested from carboys as needed. Algal cell densities (cells mL-1) were determined using a hemocytometer. Prior to introduction, algae was temperature acclimated to match conditions of the experimental larval tanks. Algae was added once to required treatments to reach desired cell densities before the lights were turned on and prior to the addition of embryos.

Larval growth and survival were assessed when larvae reached initiation of exogenous feeding, which was shown to occur at 3 DPH under similar environmental conditions by initial larviculture trials. On 3 DPH, remaining larvae were collected via siphoning and concentrated in a submerged 250 µm sieve, which minimized physical

damage. Larvae were subsequently enumerated and photographed as previously described for later analyses.

Experiment 4-3 Effect of Algal Density on Larval Growth and Survival to Three Days Post Hatch

Four algal density treatments of 0; 50,000; 100,000; and 200,000 cells mL-1

(n=4/treatment) were investigated to elucidate effects on yellow wrasse larval growth and survival at 3 DPH. Throughout the three-day trial, photoperiod was maintained at 16 hours light: 8 hours dark. Light intensity at the water’s surface for all treatments was

1134 ± 149 lux. Light intensity at the tank bottom for 0; 50,000; 100,000; and 200,000 cells mL-1 treatments measured 539 ± 80, 322 ± 51, 218 ± 32, and 92 ± 23 lux, respectively.

Experiment 4-4 Effect of Shading Method on Larval Growth and Survival to Three Days Post Hatch

Three shading method treatments (n=5/treatment) were investigated to elucidate effects on yellow wrasse larval growth and survival. One treatment shaded the tank with

50,000 cells mL-1, which was chosen because it was the lowest algal density that was shown to significantly improve larval survival in Experiment 4-3. The second treatment shaded the tank by covering the top of the tank with window screen. Both algae and window screen treatments resulted in a common light intensity of 300 lux at the tank bottom. Light was blocked completely from entering the tank in the third treatment, by covering the tanks with an opaque plastic sheet, resulting in 0 lux. Throughout the three-day trial, photoperiod was maintained at 16 hours light: 8 hours dark.

Experiment 4-5 Effect of Algal Density and Prey Availability Period on Proportion of Larvae feeding at Three Days Post Hatch

Three algal density treatments of 100,000; 300,000; and 500,000 cells mL-1

(n=5/treatment) were investigated to elucidate effects of algal density on proportion of larvae feeding at first feeding. A prey density of 5 Parvocalanus crassirostris copepod nauplii (< 75 µm) mL -1 were used for all treatments. P. crassirostris nauplii were batch cultured at UF-TAL and fed a diet of live mircroalgae (Tetraselmis chuii, Tisochrysis lutea, and Chaetoceros muelleri). Overhead 32-watt fluorescent bulbs provided light during the 16-hour light: 8-hour dark photoperiod for the experiment. Prior to 3 DPH, obliquely positioned ambient light was provided, resulting in a light intensity of 140 and

20 lux at the tank surface and bottom, respectively. Algae and live feeds were introduced once to the larval tanks before the direct overhead lights were turned on at

9:00 at 3 DPH. Light intensity at the tank bottom for 100,000; 300,000; and 500,000 cells mL -1 treatments measured 320 ± 41, 26 ± 5, and 2 ± 1, respectively. One hour after introducing P. crassirostris nauplii to the larval tank, 5-10 larvae were sampled within a five-minute time span. Six hours after the addition of live feeds, the larval tanks were harvested completely. At each sampling point, larvae were assessed for evidence of feeding by identifying the proportion of larvae with ingested nauplii. Transparent larvae were placed on a Sedgwick Rafter slide and their digestive tract was inspected for the presence of P. crassirostris naupliar eye spots and exoskeletons, which were easily distinguished from green T. chuii algal cells. A density of 5 P. crassirostris nauplii mL -1 was chosen because this density has been previously used to successfully culture a congener wrasse species, the melanurus wrasse Halichoeres melanurus (Groover et al., 2018).

Experiment 4-6 Effect of Prey Density and Prey Availability Period on Proportion of Larvae feeding at Three Days Post Hatch

Three prey density treatments of 2.5, 5, and 10 mL-1 P. crassirostris copepod nauplii (<75 µm) (n=5/treatment) were investigated to elucidate effects of prey density on proportion of yellow wrasse larvae feeding at first feeding. Based on results of experiment 4-5, an algal density of 100,000 cells mL-1 and P. crassirostris copepod nauplii (<75 µm) were used for both treatments. Methods for P. crassirostris culture, photoperiod, lighting, addition of algae and live feeds, and larval sampling and prey capture analysis adhere to those stated for the 4-5 study.

Experiment 4-7 Effect of Prey Type and Prey Availability Period on Proportion of Larvae feeding at Three Days Post Hatch

Two prey treatments of Oithona colcarva copepod nauplii (<75 µm) and P. crassirostris copepod nauplii (<75 µm) (n=5/treatment) were investigated to elucidate effects of first feed type on proportion of yellow wrasse larvae feeding at first feeding. A prey density of 5 live feed items mL-1 was used for both treatments based on previous melanurus wrasse culture success (Groover et al., 2018) and an algal density of

100,000 cells mL-1 was used for both treatments following results of previous experiment 4-5. O. colcarva were batch cultured at UF-TAL using a diet of live T. chuii,

T. lutea, and C. muelleri microalgae. Methods for P. crassirostris culture, photoperiod, lighting, addition of algae and live feeds, and larval sampling and prey capture analysis adhere to those stated for the 4-5 study. Nauplii of both copepod species were harvested, concentrated, and introduced to the culture tanks in a similar volume.

Statistical Analysis

Assumptions for linear models were tested prior to execution of statistical testing.

Homogeneity of variance was evaluated using a Bartlett’s Test in addition to plotting

residuals against predicted y-values. Normality of residuals was evaluated using a

Shapiro Wilk test in addition to a q-q plot, plotting standardized residuals against theoretical quantiles. Independence of errors was achieved through randomization in experimental design. An a priori significance level of α ≤ 0.05 was used for all statistical tests. A one-way analysis of variance (ANOVA) was used to detect statistical differences among treatments in the following experiments: 4-1, 4-2, 4-3, and 4-4. When statistical differences were detected, a post hoc pairwise comparison was conducted using a Tukey HSD test. Data from experiments 4-5 and 4-6 did not meet the assumptions of a linear model; therefore, a two-way non-parametric test was used to detect statistical differences, followed by a Dunn’s test for multiple pairwise comparisons. A two tailed paired t-test was performed to detect statistical differences of the proportion of larvae feeding between prey type treatments in experiment 4-7. All data presented in this manuscript is depicted as the mean ± SD.

Results

Experiments 4-1 and 4-2 Effect of Temperature on Embryo Incubation Time, Larval Survival at Hatch, and Larval Size at Hatch

Yellow wrasse embryos hatched within a 4.5-hour time span from 18:30-23:00

(21.5-27 hours post stocking) and a 3-hour time span from 14:00-17:00 (16.5-19.5 hours post stocking) when incubated at 22 and 25°C, respectively. Embryos hatched within a 3-hour time span from 9:30-12:30 (12-15 hours post stocking) when incubated at 28°C (Figure 4-3).

Larval survival varied significantly among temperature treatments (F2,12=23.4114,

P<0.0001). Mean survival decreased at 22°C (38 ± 16%) (P=0.0002) compared to 25

(85 ± 5%) and 28°C (83 ± 13%) (Figure 4-4).

100

Yellow wrasse larval notochord length (mm) at hatch was found to be significantly different among temperature treatments (F2,9=9.2276, P=0.0066), specifically between 22 and 28°C (P=0.0051) treatments (Figure 4-5). Larvae hatching from embryos incubated at 22°C were larger (1.4536 ± 0.0694 mm) than those incubated at temperatures of 25°C (1.3908 ± 0.0621 mm) and 28°C (1.3328 ± 0.0769 mm) (Figure 4-5).

Experiment 4-3 Effect of Algal Density on Larval Growth and Survival to Three Days Post Hatch

Yellow wrasse larval notochord length was not found to be significantly different among algal density treatments (F3,9=9.787, P=0.4449) with an observed range of

2.157-2.187 mm.

Larval survival varied significantly among algal density treatments (F3=12.9540,

P=0.0047). Mean survival in the 0 cells mL-1 treatment (0.5 ± 1%) was statistically lower than the 50,000 cells mL-1 (36 ± 25%), 100,000 cells mL-1 (63 ± 8%), and 200,000 cells mL-1 (42 ± 13%) treatments (Figure 4-6). No significant differences were detected in mean survival among all treatments that were inoculated with algae (P≥0.0606).

Experiment 4-4 Effect of Shading Method on Larval Growth and Survival to Three Days Post Hatch

Yellow wrasse larval notochord length was found to be significantly different among light intensity treatments (F2,9=4.9753, P=0.0351). Mean notochord length increased in tanks shaded with algae (2.142 ± 0.018 mm) and window screen (2.121 ±

0.032 mm), compared to those not exposed to light (2.024 ± 0.084 mm; P=0.0383)

(Figure 4-7).

101

Larval survival was not found to vary significantly (F2,9=0.3921, P=0.6866) and was relatively consistent among the three shading treatments ranging from 26-32%

(Figure 4-8).

Experiment 4-5 Effect of Algal Density and Feed Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

A significant interaction (P>0.0866) was not detected between algal density and feed availability period factors, nor between algal density and proportion of larvae feeding, as well as between feed availability period and proportion of larvae feeding

(Figure 4-9). Across all treatments, mean proportion of larvae feeding remained low, ranging from 0.00-6.11%. Proportion of larvae feeding at 2 hours post feeding was 2.22

± 4.97, 0.00 ± 0.00, and 3.33 ± 7.46%, for the 100,000; 300,000; and 500,000 cells mL-

1 treatments, respectively. Proportion of larvae feeding at 6 hours post feeding was 3.26

± 3.46, 3.91 ± 6.23, and 6.11 ± 9.70%, for the 100,000; 300,000; and 500,000 cells mL-

1 treatments, respectively (Figure 4-9).

Experiment 4-6 Effect of Prey Density and Feed Availability Period on Proportion of Larvae Feeding at Three Days Post Hatch

No significant interactions (P>0.0909) were detected between prey density and feed availability period factors, nor between prey density and proportion of larvae feeding, as well as between feed availability period and proportion of larvae feeding

(Figure 4-10). Proportion of larvae feeding at 2 hours post feeding and 2.5, 5, and 10 nauplii mL-1 was 3.33 ± 7.45, 2.86 ± 6.39, and 9.44 ± 9.49%, respectively (Figure 4-10).

Proportion of larvae feeding at 6 hours post feeding and 2.5, 5, and 10 nauplii mL-1 was

3.47 ± 2.48, 6.40 ± 1.37, and 7.85 ± 3.78%, respectively (Figure 4-10).

102

Experiment 4-7 Effect of Prey Type on and Feed Availability Period on Proportion of Larvae feeding at Three Days Post Hatch

No yellow wrasse larvae were found to have either prey item in their gastrointestinal tracts after a 2-hour feed availability period. Proportion of larvae feeding after a 6-hour feed availability period was not found to be significantly different between prey type treatments (t8=0.3350, P=0.7463). In larval tanks fed P. crassirostris nauplii

(<75 µm) and O. colcarva nauplii (<75 µm) proportion of larvae feeding was 1.53 ± 1.0 and 1.29 ± 1.25% after a 6-hour feed availability period, respectively.

Mean water quality parameters for all studies are presented in Table 4-1.

Discussion

Temperature was shown to have an inverse relationship with yellow wrasse incubation time, in accordance with many other studies which have examined the same relationship (Beacham and Murray, 1990; Gray, 1928; Pauly and Pullin, 1988; Pepin,

1991). Temperatures of 25 and 28°C were shown to shorten yellow wrasse incubation time and decrease larval notochord length at hatch, hypothesized to be due to a reduced embryonic development period. Conversely, cooler incubation temperatures have been shown to effectively increase larval length in several foodfish species including five species of Pacific salmon (Beacham and Murray, 1990). While a cooler temperature of 22°C was shown to increase larval notochord length in yellow wrasses at hatch, this temperature also severely decreased survival by approximately 55% compared to survival achieved in 25 and 28°C. The natural temperature range of the species is ~24-27°C (Randall et al., 1990) suggesting that 22°C is at the lower end of this species’ thermal tolerance and too low to allow for proper embryonic development.

Based on the results of these studies, decreasing embryo incubation temperature to

103

22°C to achieve enlarged larvae and associated advantages at first feeding does not appear to be a viable option given the magnitude of the decrease in survival rate and relatively short incubation period. It is suggested that a temperature between 25 and

28°C be used to incubate yellow wrasse embryos to achieve acceptable larval survival and size at hatch.

“Greenwater” is a common practice in marine fish culture defined by the addition of microalgae to the larval culture environment (Calado et al., 2017; Faulk and Holt,

2005; Naas et al., 1992; Palmer et al., 2007). Benefits of greenwater culture are now thought to impact larvae prior to the onset of exogenous feeding. A study with cod

Gadus morhua indicated larvae were active filter feeders during early larval stages based on concentrations of algal cells in their gastrointestinal tracts being much higher than normal accumulations due to drinking rate (Meeren, 1991). This ingestion of algal cells has been shown to trigger digestive enzyme production in sea bass Dicentrarchus labrax, turbot Scophthalmus maximus, and halibut Hippoglossus hippoglossus larvae early in development, improving digestive function and subsequent growth (Cahu et al.,

1998; Reitan et al., 1997). Yellow wrasse larvae do not develop functional mouth parts until 3 DPH, therefore limiting the potential benefit of ingested algal cells this early in development. As a result, algal density was not shown to significantly effect yellow wrasse growth from 0-3 DPH.

Yellow wrasse larval survival was significantly increased by the presence of algae in the culture environment from 0-3 DPH, regardless of density. The tanks where no algae was introduced experienced almost complete mortality (~95.5%) by 3 DPH, while mean larval survival in tanks where algae was introduced ranged from ~36-63%

104

across treatments. It is hypothesized that the presence of algae was not the primary driver of survival, but rather the decrease in light intensity resulting from its introduction.

Increased light intensity has been shown to negatively affect photo-sensitive larvae of certain species (Villamizar et al., 2011) such as sea bass Dicentrarchus labrax

(Barahona-Fernandes, 1979) and southern flounder Paralichthys lethostigma (Henne and Watanabe, 2003). It appears that the yellow wrasse is also a light sensitive species requiring reduced light intensity to prevent stress and subsequent mortality.

This hypothesis was tested in a subsequent shading method study (Experiment

4-4). When algae and artificial shading methods were used to produce equal light intensity in the culture tanks, larval survival was not significantly impacted. This suggests that the yellow wrasse is photo-sensitive early in development and measures should be taken to reduce light intensity prior to the initiation of exogenous feeding. It is also important to note that as long as light intensity is reduced, there does not appear to be a requirement for the presence of microalgae in the culture tank before exogenous feeding. This could prevent alteration of water quality and reduce the potential for introduction of microorganisms such as bacteria and ciliates that are ubiquitous in algae cultures, likely improving larval survival during the first few days of development.

Study 4-4 also revealed that yellow wrasse growth was shown to decrease significantly when deprived of light during the first three days of development. This finding is consistent with other studies that examined the effect of complete darkness on the growth of European sea bass (Villamizar et al., 2009) and Senegal sole (Blanco-

Vives et al., 2010) where absence of light was shown to decrease larval growth. These findings demonstrate that a natural light and dark cycle is essential for proper larval

105

growth and development (Villamizar et al., 2011). Overall, these early larviculture trials have demonstrated that a natural photoperiod containing light and dark periods as well as reduced light intensity to approximately 300 lux through artificial means will improve yellow wrasse growth and survival prior to the initiation of exogenous feeding.

No effect of algal density, prey density, prey type, or feed availability period was detected on the proportion of yellow wrasse larvae feeding. In all larval feeding trials, feeding percentage never increased above 9%, suggesting that variables other than those tested in the current study may be affecting the ability of these larvae to successfully detect and capture the food items provided. It is also possible that the ranges chosen for tested variables were not appropriate to accommodate successful feeding behavior and may need to be revisited in further investigations.

Maternal effects that were not taken into account during these studies such as effects on egg size, as well as nutritional composition and volume of yolk sac and oil globule reserves could have impacted larval size at hatch and the transition period between endogenous and exogenous feeding (Hunter, 1981; May, 1974). Since no primary literature currently exists describing egg diameter or suitable proxies for egg quality for wild yellow wrasses, comparisons with embryos and larvae resulting from captive broodstock are difficult.

Temperature has been shown to impact larval feeding success, with cooler temperatures decreasing swimming speed, prey encounter frequency, and metabolic efficiency (Hunter, 1981). Although the temperature maintained throughout all larviculture trials (~25.5°C) fell within the natural temperature range of the yellow wrasse (~24-27°C; Randall et al., 1990), it’s possible that a higher temperature is

106

required to improve yellow wrasse feeding success, although this would likely increase the metabolic rate of the larvae necessitating increased feeding frequency. Trials examining the effect of temperature on yellow wrasse feeding success could elucidate information regarding this environmental parameter.

The addition of microalgae to the culture environment has been shown to improve survival and feeding performance in several foodfish species such as cobia

Rachycentron canadum (Faulk and Holt, 2005), silver seabream Pagrus auratus

(Palmer et al., 2007), halibut Hippoglossus hippoglossus (Naas et al., 1992), and

Atlantic cod Gadus morhua (Meeren et al., 2007). Limited primary literature exists describing culture methods for pelagic spawning marine ornamental species, however, of those published, most utilize the greenwater technique for larval culture. Milletseed butterflyfish Chaetodon miliaris feeding incidence improved from ~48 to 75% in greenwater (~500,000-800,000 cells mL-1) verses clear water. In the culture of yellow tang Zebrasoma flavescens, feed incidence was also found to increase four-fold in tanks containing 400,000-600,000 cells mL-1 microalgae when compared to clear water

(Pereira-Davison and Callan, 2018). Although feeding performance was not evaluated, the addition of microalgae to the culture environment proved paramount for the regal damselfish Neopomacentrus cyanomos, where larval survival was 0 and 20% in clear water and greenwater (60,000 cells mL-1), respectively (Setu et al., 2010). Feeding performance of yellow tang and milletseed butterflyfish was highest at algal densities comparable to or higher than those tested in yellow wrasse feeding trials, suggesting that a higher algal density >500,000 cells mL-1 could improve feeding success and should be investigated further.

107

Greenwater has also been employed in larviculture of non-ornamental wrasse species including the ballan wrasse Labrus bergylta (Ottesen et al., 2012), hogfish

Lachnolaimus maximus (Colin, 1982), Tautog Tautoga onitis (Perry et al., 2001, 1998), and the rainbow wrasse Parajulis poecilopterus (Kimura and Kiriyama, 1993), however parameters effecting feeding performance were not evaluated. To date, only five species of ornamental wrasses have been successfully cultured past metamorphosis; these are the ornate wrasse Halichoeres ornatissimus (Baensch, 2017, personal communication), Hawaiian cleaner wrasse Labroides phthirophagus (Montalvo, 2017, personal communication), bluestreak cleaner wrasse, Cuban hogfish (Ohs et al., 2018), and the melanurus wrasse Halichoeres melanurus (Barden et al., 2016; Groover et al.,

2018).

To the authors’ knowledge, this paper represents the first published research delineating captive spawning and larviculture in the yellow wrasse. These experiments created an initial culture protocol for the yellow wrasse that refined incubation temperature and larviculture culture parameters, improving survival and growth prior to first feeding. Based on experimental results, a temperature range between 25 and 28°C is recommended for embryo incubation and decreased light intensity (~300 lux) using artificial means prior to exogenous feeding. Feeding trials conducted with the yellow wrasse did not clearly define preferential culture conditions and resulted in poor feeding performance and larval survival.

Results from these investigations provide a strong foundation upon which to build yellow wrasse culture protocols. Additional research addressing critical bottlenecks such as first feeding are warranted for this for this valuable Labrid species. Through

108

continued efforts, it is likely that the yellow wrasse will be successfully cultured for the first time and creation of subsequent commercial production protocols will follow.

109

Table 4-1. Water quality parameters for H. chrysus larviculture experiments represented as mean ± SD.

Total Water Alkalinity Salinity DO Ammonia Nitrite Nitrate Experiment Temperature pH (mg L-1 (g L-1) (mg L-1) Nitrogen (mg L-1) (mg L-1) (°C) CaCo ) 3 (mg L-1) 4-6 25.43 ± 0.21 35.01 ± 0.09 8.08 ± 0.03 7.20 ± 0.18 177.22 ± 8.77 0.04 ± 0.05 0.04 ± 0.06 25.46 ± 9.34 4-7 25.54 ± 0.15 34.35 ± 0.03 8.10 ± 0.02 7.02 ± 0.21 181.88 ± 8.62 0.00 ± 0.00 0.00 ± 0.00 27.27 ± 10.09 4-8 25.40 ± 0.18 34.43 ± 0.11 8.10 ± 0.03 7.17 ± 0.09 183.21 ± 8.02 0.00 ± 0.00 0.06 ± 0.03 25.45 ± 9.34 4-9 25.39 ± 0.11 35.05 ± 0.05 8.05 ± 0.01 7.15 ± 0.08 180.12 ± 8.83 0.00 ± 0.00 0.09 ± 0.01 29.09 ± 10.44 4-10 25.47 ± 0.19 34.49 ± 0.03 8.09 ± 0.01 7.10 ± 0.12 184.99 ± 6.92 0.00 ± 0.00 0.05 ± 0.00 21.82 ± 6.03

110

Figure 4-1. H. chrysus sexual dimorphism; A) Female with three dorsal ocelli; B) Female or male with two dorsal ocelli (one anterior and one central); C) Terminal male with one anterior ocellus and green facial barring. Photo courtesy of author.

111

Figure 4-2. Wild caught H. chrysus broodstock spawning frequency and number of fertilized and unfertilized eggs per spawn from June-December 2017. Original sex ratio was one male: two, two spotted individuals of undetermined sex, and 17 females but transitioned to being male dominated over time. Lunar cycle is represented in the figure by black circles (full moon).

112

100 90 80 70 60 22°C 50 28°C 25°C

Hatch% 40 30 20 10 0 8:00 9:30 11:00 0:30 14:00 15:30 17:00 18:30 20:00 21:30 23:00 Time

Figure 4-3. Hatch (%) over time of H. chrysus embryos in response to incubation temperature (°C) (n=5).

100 b b 90 80 70 60 a

50 Survival (%) Survival 40 30 20 10 22 25 28 Temperature (°C)

Figure 4-4. Survival (%) of H. chrysus embryos in response to incubation temperature (°C) represented as mean ± SD (n=5). Different letters above bars denote statistically significant differences (P ≤ 0.05).

113

1.55 a 1.5 b 1.45 c 1.4

1.35

1.3 Notochord Length (mm) Length Notochord 1.25

1.2 22 25 28 Temperature (°C)

Figure 4-5. Notochord length at hatch (mm) of H. chrysus larvae in response to incubation temperature (°C) represented as mean ± SD (n=4). Different letters above bars denote statistically significant differences (P ≤ 0.05).

100 90 80 b 70 b b 60 50

Survival (%) Survival 40 30 20 10 a 0 0 50,000 100,000 200,000 Algal Density (cells mL-1)

Figure 4-6. Survival (%) of H. chrysus larvae at 3 DPH in response to algal density (cells mL-1) represented as mean ± SD (n=4). Different letters above bars denote statistically significant differences (P ≤ 0.05).

114

2.20 a a 2.15 b 2.10

2.05

2.00

Notochord Length (mm) Length Notochord 1.95

1.90 Algae Screen Plastic Shading Method

Figure 4-7. Notochord length (mm) of H. chrysus larvae in response to shading method represented as mean ± SD (n=5). Different letters above bars denote statistically significant differences (P ≤ 0.05).

45 a a 40 a 35 30 25

Survival (%) Survival 20 15 10 Algae Screen Plastic

Shading Method

Figure 4-8. Survival (%) of H. chrysus larvae in response to shading method represented as mean ± SD (n=5). Different letters above bars denote statistically significant differences (P ≤ 0.05).

115

20 a 15

a a 10 a a

Feeding % Feeding 5

0 100 300 500 -5 Algal Density (1,000 cells mL-1)

2 Hours Feeding Period 6 Hour Feeding Period Figure 4-9. Larval feeding (%) of H. chrysus larvae in response to algal density (cells mL-1) and prey availability period (# hours) represented as mean ± SD (n=5). Different letters above bars denote statistically significant differences (P ≤ 0.05) within algal density treatments between prey availability period treatments. 20 a

15 a a 10 a a

a Feeding % Feeding 5

0 2.5 5 10 -5 Prey Density (nauplii mL-1)

2 Hour Feeding Period 6 Hour Feeding Period Figure 4-10. Larval feeding (%) of H. chrysus larvae in response to prey density (nauplii mL-1) and prey availability period (# hours) represented as mean ± SD (n=5). Different letters above bars denote statistically significant differences (P ≤ 0.05) within prey density treatments between prey availability period treatments.

116

CHAPTER 5 CONCLUSIONS

Wild capture is the predominant source of animals for the marine ornamental aquarium industry. While many demersal spawning species are now commonly available as captive-bred, aquaculture of pelagic spawning ornamental fish is still in its relative infancy. Wrasses are the second most popular family of fishes in the marine aquarium trade by volume (Rhyne et al., 2012), yet, a mere ten species have been successfully cultured in captivity. Currently, only five species of ornamental wrasse have been successfully cultured to the juvenile stage, none of which are commercially produced. Information regarding the culture of ornamental wrasses has been mainly anecdotal, in the form of magazine articles and online media, describing the culture process without further refinement of methodologies. To advance ornamental wrasse culture, continued investigation of larviculture parameters for all larval stages is essential.

The melanurus wrasse Halichoeres melanurus and the yellow wrasse H. chrysus were chosen as candidate species from the family Labridae for the development of culture methodologies due to their popularity in the aquarium trade and short larval duration (Victor, 1986a). Critical culture aspects were assed for both species including broodstock spawning characteristics, egg incubation temperature, early larval culture considerations, and first feeding parameters. Broodstock husbandry, spawning, and larviculture methods were described for successful production of two generations of melanurus wrasse, effectively closing the life cycle of this species.

Larval development and culture requirements of the melanurus wrasse were described across a series of investigations. Fertilized eggs measured 0.627 ± 0.013 mm

117

in diameter and hatched approximately 15-21 hours later. Notochord length (NL) of newly hatched altricial larvae measured 1.590 ± 0.140 mm and lacked a functional mouth and gastrointestinal tract. Initiation of exogenous feeding, swim bladder inflation, and flexion were first observed at 3, 10, and 15 days post hatch (DPH), respectively.

Completion of metamorphosis was first observed at 37 DPH at a mean standard length of 11.851 ± 0.230 mm and larval growth appeared linear over the 45-day period.

Survival through metamorphosis was 0.54% yielding 17 juvenile wrasses. A separate group of first generation (F1) melanurus wrasses were cultured and reached sexual maturity in only eight months, a characteristic favorable for commercial production.

Second generation (F2) melanurus wrasses were then cultured from F1 embryos, effectively closing the life cycle of the species. This is the first published report detailing the completion of the life cycle for an ornamental wrasse species and is a significant breakthrough in the aquaculture industry, although continued research is needed to refine protocols and improve larval survival through metamorphosis.

To identify effective culture methods for the yellow wrasse and to further refine culture methods for the melanurus wrasse, a range of studies were conducted to evaluate parameters affecting embryo incubation, larval survival and growth prior to first feeding, and larval feeding success at initiation of exogenous feeding. For both wrasse species, an inverse relationship was found between water temperature and embryo incubation period with incubation time decreasing with increasing temperature from 22-

28°C. While cooler water temperatures resulted in statistically larger larvae at hatch, survival was negatively impacted. A temperature range between 25 and 28°C is recommended for embryo incubation of both wrasse species.

118

An examination of melanurus and yellow wrasse larvae prior to first feeding established that cultures containing algal densities of 50,000, 100,000, and 200,000 cells mL-1 increased larval survival compared to those devoid of algae. This correlation between addition of algae to the culture environment and increased larval survival was observed in both species and was further investigated to elucidate the mechanism behind the observed gains. These experiments demonstrated melanurus and yellow wrasse larval survival to be similar between treatments of the same light intensity (~300 lux) irrespective of the shading method, suggesting that the previously noted increase in survival was likely due to the decrease in light intensity and not from the algae itself.

When tanks were shaded with opaque plastic, resulting in 0 lux, larval growth was stunted in both species. Taken together, these results support a recommendation of decreased light intensity (~300 lux) using artificial means for early larviculture of

Halichoeres wrasses. This strategy will help to prevent degradation in water quality while improving larval survival prior to exogenous feeding.

Melanurus wrasse first feeding culture trials revealed that algal densities of

300,000 and 500,000 cells mL-1 resulted in elevated larval feeding success. Prey availability period did not significantly impact the proportion of larvae feeding. Rotifers

(<75 µm) were not able to be ingested by melanurus wrasse larvae under the described culture parameters and are not recommended as a first feed item. Ingestion of

Parvocalanous crassirostris nauplii (<75 µm) was significantly increased at prey densities ≥ 5.0 items mL-1 and use of this feeding regime has proven successful at culturing the melanurus wrasse through early larval bottlenecks (Groover et al., 2018).

119

Similar feeding trials conducted with the yellow wrasse resulted in poor feeding performance and larval survival. Results of these investigations did not clearly delineate preferential culture conditions and further investigation into first feeding culture parameters such as increased algal densities, alternate live feeds, feed attractants, prey densities, and water temperature should be pursued to increase yellow wrasse feeding success.

Information gathered from these trials will help to advance commercial aquaculture protocols for the melanurus and yellow wrasse and guide future aquaculture research with additional marine ornamental species. Continued investigation into larviculture parameters influencing growth, survival, and feeding success will help to create and refine methodologies which should lead to commercial protocols for captive propagation of Halichoeres wrasses.

120

LIST OF REFERENCES

APPA, 2018. American pet products association national pet owners survey 2017– 2018. Greenwich, CT.

Baensch, F., Tamaru, C.S., 2009. Spawning and development of larvae and juveniles of the rare blue mauritius angelfish, Centropyge debelius (1988), in the hatchery. J. World Aquac. Soc. 40, 425–439.

Bagarinao, T., 1986. Yolk resorption, onset of feeding and survival potential of larvae of three tropical marine fish species reared in the hatchery. Mar. Biol. 91, 449–459.

Barahona-Fernandes, M.H., 1979. Some effects of light intensity and photoperiod on the sea bass larvae (Dicentrarchus labrax) reared at the Centre Oceanologique de Bretagne. Aquaculture 17, 311–321.

Barber, C.V., Pratt, V.R., 1998. Poison and profits: cyanide fishing in the Indo-Pacific. Environ. Sci. Policy Sustain. Dev. 40, 4–9.

Barden, K.P., Cassiano, E.J., Hauville, M., Ramee, S., DiMaggio, M.A., 2016. First record of captive spawning and larval Rearing of the melanurus wrasse, Halichoeres melanurus, in: World Aquaculture Society - Aquaculture America. World Aquaculture Society.

Barden, K.P., Cassiano, E.J., Wittenrich, M.L., 2013. Linking temperature-mediated morphology with feeding performance and mortality through pelagic larval development of the ocellated dragonet Synchiropus Ocellatus, in: Aquaculture America. Nashville, TN.

Battaglene, S.C., Talbot, R.B., 1990. Initial swim bladder inflation in intensively reared Australian bass larvae, Macquaria novemaculeata (Steindachner) (Perciformes: Percichthyidae). Aquaculture 86, 431–442.

Beacham, T.D., Murray, C.B., 1990. Temperature, egg size, and development of embryos and alevins of five species of Pacific salmon: a comparative analysis. Trans. Am. Fish. Soc. 119, 927–945.

Blanco-Vives, B., Villamizar, N., Ramos, J., Bayarri, M.J., Chereguini, O., Sánchez- Vázquez, F.J., 2010. Effect of daily thermo- and photo-cycles of different light spectrum on the development of Senegal sole (Solea senegalensis) larvae. Aquaculture 306, 137–145.

Brown, C.A., Gothreaux, C.T., Green, C.C., 2011. Effects of temperature and salinity during incubation on hatching and yolk utilization of Gulf killifish Fundulus grandis embryos. Aquaculture 315, 335–339.

Cahu, C., Zambonino Infante, J., Péres, A., Quazuguel, P., Le Gall, M., 1998. Algal addition in sea bass (Dicentrarchus labrax) larvae rearing: effect on digestive

121

enzymes. Aquaculture 161, 479–489.

Calado, R., Olivotto, I., Oliver, M.P., Holt, J.G., 2017. Marine ornamental species aquaculture, First. ed. Wiley-Blackwell, West Sussex, UK.

Callan, C.K., Burgess, A.I., Rothe, C.R., Touse, R., 2018. Development of improved feeding methods in the culture of yellow tang, Zebrasoma flavescens. J. World Aquac. Soc.

Cassiano, E.J., Ohs, C.L., Weirich, C.R., Breen, N.E., Rhyne, A.L., 2011. Performance of larval Florida pompano fed nauplii of the calanoid copepod Pseudodiaptomus pelagicus. N. Am. J. Aquac. 73, 114–123.

Chapman, D.C., Hubert, W.A., Jackson, U.T., 1988. Influence of access to air and of salinity on gas bladder inflation in striped bass. Progress. Fish-Culturist 50, 23–27.

Chatain, B., 1994. Abnormal swimbladder development and lordosis in sea bass (Dicentrarchus labrax) and sea bream (Sparus auratus). Aquaculture 119, 371– 379.

Chatain, B., 1989. Problems related to the lack of functional swimbladder in intensive rearing of the seabass Dicentrarchus labrax and the sea bream Sparus auratus. Adv. Trop. Aquac. Work. .

Chatain, B., 1986. The swim bladder in Dicentrarchus labrax and Sparus auratus. Aquaculture 53, 303–311.

Chatain, B., Ounais-Guschemann, N., 1990. Improved rate of initial swim bladder inflation in intensively reared Sparus auratus. Aquaculture 84, 345–353.

Colin, P.L., 1982. Spawning and larval development of the hogfish Lachnolaimus maximus (Pisces: Labridae). Fish. Bull. 80, 853–862.

D’Arcy, J., Dunaevskaya, E., Treasurer, J.W., Ottesen, O., Maguire, J., Zhuravleva, N., Karlsen, A., Rebours, C., FitzGerald, R.D., 2012. Embryonic development in ballan wrasse Labrus bergylta. J. Fish Biol. 81, 1101–1110.

Daoulas, C., Economou, A.N., Bantavas, I., 1991. Osteological abnormalities in laboratory reared sea-bass (Dicentrarchus labrax) fingerlings. Aquaculture 97, 169– 180.

Davis, D.A., Derbes II, T.J., Head, M.E., 2018. Culture of small zooplankton for the feeding of larval fish. South. Reg. Aquac. Cent.

Dawes, J., 1998. International experience in ornamental marine species managment - Part 1: Perspectives.

Dee, L.E., Horii, S.S., Thornhill, D.J., 2014. Conservation and management of

122

ornamental coral reef wildlife: Successes, shortcomings, and future directions. Biol. Conserv. 169, 225–237.

Degidio, J.-M.L.A., Yanong, R.P.E., Watson, C.A., Ohs, C.L., Cassiano, E.J., Barden, K.P., 2017. Spawning, embryology, and larval development of the milletseed butterflyfish Chaetodon miliaris in the laboratory. N. Am. J. Aquac. 79, 205–215.

DiMaggio, M.A., Cassiano, E.J., Barden, K.P., Ramee, S., Ohs, C.L., Watson, C.A., 2017. First record of captive larval culture and metamorphosis of the Pacific blue tang, Paracanthurus hepatus. J. World Aquac. Soc. 48, 393–401.

Dixson, D.L., Pratchett, M.S., Munday, P.L., 2012. Reef fishes innately distinguish predators based on olfactory cues associated with recent prey items rather than individual species. Anim. Behav. 84, 45–51.

Faulk, C.K., Holt, G.J., 2005. Advances in rearing cobia Rachycentron canadum larvae in recirculating aquaculture systems: Live prey enrichment and greenwater culture. Aquaculture 249, 231–243.

Fernandez-Diaz, C., Pascual, E., Yufera, M., 1994. Feeding behaviour and prey size selection of gilthead seabream, Sparus aurata, larvae fed on inert and live food. Mar. Biol. 118, 323–328.

Fielder, D.S., Bardsley, W.J., Allan, G.L., Pankhurst, P.M., 2002. Effect of photoperiod on growth and survival of snapper Pagrus auratus larvae. Aquaculture 211, 135– 150.

Gerkema, M.P., Videler, J.J., de Wiljes, J., van Lavieren, H., Gerritsen, H., Karel, M., 2000. Photic entrainment of circadian activity patterns in the trpical Labrid fish Halichoeres chrysus. Chronobiol. Int. 17, 613–622.

Gopakumar, G., Ignatius, B., Santhosi, I., Ramamoorthy, N., 2009. Controlled breeding and larval rearing techniques of marine ornamental fishes. Asian Fish. Sci. 22, 797–804.

Gray, J., 1928. The growth of fish. Br. J. Exp. Biol. 6, 125–130.

Groover, E.M., DiMaggio, M.A., Alo, M.M., Degidio, J.L., Ramee, S.W., Lipscomb, T.N., Sipos, M.J., 2018. Greenwater culture and larval development of the melanurus wrasse Halichoeres melanurus, in: Aquaculture America. Las Vegas, NV.

Hamre, K., Nordgreen, A., Grøtan, E., Breck, O., 2013a. A holistic approach to development of diets for Ballan wrasse (Labrus berggylta) – a new species in aquaculture. PeerJ 1, e99.

Hamre, K., Srivastava, A., Ronnestad, I., Mangor-Jensen, A., Stoss, J., 2008. Several micronutrients in the rotifer Brachionus sp. may not fulfil the nutritional requirements of marine fish larvae. Aquac. Nutr. 14, 51–60.

123

Hamre, K., Yúfera, M., Ronnestad, I., Boglione, C., Conceição, L.E.C., Izquierdo, M., 2013b. Fish larval nutrition and feed formulation: knowledge gaps and bottlenecks for advances in larval rearing. Rev. Aquac. 5, S26–S58.

Henne, J.P., Watanabe, W.O., 2003. Effects of light intensity and salinity on growth, survival, and whole-body osmolality of larval Southern flounder Paralichthys lethostigma. J. World Aquac. Soc. 34, 450–465.

Hilder, P.I., Cobcroft, J.M., Battaglene, S.C., 2015. The first-feeding response of larval southern bluefin tuna, Thunnus maccoyii (Castelnau, 1872), and yellowtail kingfish, Seriola lalandi (Valenciennes, 1833), to prey density, prey size and larval density. Aquac. Res. 46, 2736–2751.

Hirai, N., Koiso, M., Teruya, K., Kobayashi, M., Takebe, T., Sato, T., Okuzawa, K., Hagiwara, A., 2013. Success of seed production of humphead wrasse Cheilinus undulatus with improvement of spawning induction, feeding, and rearing conditions, in: Rust, M., Olin, P., Bagwill, A., Fujitani, M. (Eds.), Proceedings of the 40th U.S.- Japan Aquaculture Panel Symposium. U.S. Department of Commerce, NOAA, Honolulu, Hawaii, pp. 107–111.

Holt, G.J., 2003. Research on culturing the early life stages of marine ornamental species, in: Cato, J.C., Brown, C.L. (Eds.), Marine Ornamental Species : Collection, Culture and Conservation. John Wiley & Sons, 251-254, p. 423.

Hunter, J.R., 1981. Feeding ecology and predation of marine fish larvae, Southerwest Fisheries Science Center: NPAA Fisheries Service.

Kimura, S., Kiriyama, T., 1993. Development of eggs, larvae and juveniles of the labrid fish, Halichoeres poecilopterus, reared in the laboratory. Japanese J. Ichthyol. 39, 371–377.

Kuiter, R.H., 2010. Labridae fishes: Wrasses, First Edit. ed. Kuiter, Rudie H, Seaford, Australia.

Kuwamura, T., Karino, K., Nakashima, Y., 2000. Male morphological characteristics and mating success in a protogynous coral reef fish, Halichoeres melanurus. J. Ethol. 18, 17–23.

Lee, I.S., Ohs, C.L., Broach, J.S., DiMaggio, M.A., Watson, C.A., 2018. Determining live prey preferences of larval ornamental marine fish utilizing fluorescent microspheres. Aquaculture 490, 125–135.

Livengood, E.J., Chapman, F.A., 2007. The ornamental fish trade: An introduction with perspectives for responsible aquarium fish ownership.

May, R.C., 1974. Larval mortality in marine fishes and the critical period concept, in: The Early Life History of Fish. Springer Berlin Heidelberg, Berlin, Heidelberg, pp. 3– 19.

124

McKinnon, A.D., Duggan, S., Nichols, P.D., Rimmer, M.A., Semmens, G., Robino, B., 2003. The potential of tropical paracalanid copepods as live feeds in aquaculture. Aquaculture 223, 89–106.

Meeren, T., 1991. Algae as first food for cod larvae, Gadus morhua L.: filter feeding or ingestion by accident? J. Fish Biol. 39, 225–237.

Meeren, T., Mangor-Jensen, A., Pickova, J., 2007. The effect of green water and light intensity on survival, growth and lipid composition in Atlantic cod (Gadus morhua) during intensive larval rearing. Aquaculture 265, 206–217.olivotto

Moorhead, J.A., Zeng, C., 2010. Development of captive breeding techniques for marine ornamental fish: A review. Rev. Fish. Sci. 18, 315–343.

Murray, J.M., Watson, G.J., 2014. A critical assessment of marine aquarist biodiversity data and commercial aquaculture: Identifying gaps in culture initiatives to inform local fisheries managers. PLoS One 9.

Naas, K., Huse, I., Iglesias, J., 1996. Illumination in first feeding tanks for marine fish larvae. Aquac. Eng. 15, 291–300.

Naas, K.E., Naess, T., Harboe, T., 1992. Enhanced first feeding of halibut larvae (Hippoglossus hippoglossus L.) in green water. Aquaculture 105, 143–156.

Ohs, C.L., Broach, J.S., Lee, I.S., Palau, A.T., 2018. Successful captive spawning and culture of Cuban hogfish. World Aquac. Soc. - Aquac. Am.

Oie, G., Galloway, T., Soroy, M., Holmvaag Hansen, M., Norheim, I.A., Halseth, C.K., Almli, M., Berg, M., Gagnat, M.R., Wold, P.A., Attramadal, K., Hagemann, A., Evjemo, J.O., Kjorsvik, E., 2017. Effect of cultivated copepods (Acartia tonsa) in first-feeding of Atlantic cod (Gadus morhua) and ballan wrasse (Labrus bergylta) larvae. Aquac. Nutr. 23, 3–17.

Olivotto, I., Chemello, G., Vargas, A., Randazzo, B., Piccinetti, C.C., 2017. Marine ornamental species culture: From the past to “Finding Dory.” Gen. Comp. Endocrinol. 245, 116–121.

Olivotto, I., Planas, M., Simões, N., Holt, G.J., Avella, M.A., Calado, R., 2011. Advances in breeding and rearing marine ornamentals. J. World Aquac. Soc. 42, 135–166.

Ostergaard, P., Munk, P., Janekarn, V., 2005. Contrasting feeding patterns among species of fish larvae from the tropical Andaman Sea. Mar. Biol. 146, 595–606.

Ottesen, O.H., Dunaevskaya, E., D’Arcy, J., 2012. Development of Labrus bergylta (Ascanius 1767) larvae from hatching to metamorphosis. Aquac. Res. Dev. 3, 127.

Palmer, P.J., Burke, M.J., Palmer, C.J., Burke, J.B., 2007. Developments in controlled green-water larval culture technologies for estuarine fishes in Queensland,

125

Australia and elsewhere. Aquaculture 272, 1–21.

Paperna, I., 1978. Swimbladder and skeletal deformations in hatchery bred Spams aurata. J. Fish Biol. 12, 109–114.

Parenti, P., Randall, J.E., 2018. A checklist of wrasses (Labridae) and parrotfishes (Scaridae) of the world: 2017 update. J. Ocean Sci. Found. 30, 11–27.

Pauly, D., Pullin, R.S. V., 1988. Hatching time in spherical, pelagic, marine fish eggs in response to temperature and egg size. Environ. Biol. Fishes 22, 261–271.

Pepin, P., 1991. Effect of temperature and size on development, mortality, and survival rates of the pelagic early life history stages of marine fish. Can. J. Fish. Aquat. Sci. 48, 503–518.

Pepin, P., Orr, D.C., Anderson, J.T., 1997. Time to hatch and larval size in relation to temperature and egg size in Atlantic cod (Gadus morhua). Can. J. Fish. Aquat. Sci. 54, 2–10.

Pereira-Davison, E., Callan, C.K., 2018. Effects of photoperiod, light intensity, turbidity and prey density on feed incidence and survival in first feeding yellow tang (Zebrasoma flavescens) (Bennett). Aquac. Res. 49, 890–899.

Perry, D.M., Mercaldo-Allen, R., Burgh, S., 2001. Growth and culture of larval and juvenile tautogs in a closed recirculating-seawater system. N. Am. J. Aquac. 63, 300–305.

Perry, D.M., Mercaldo-Allen, R., Kuropat, C.A., Hughes, J.B., 1998. Laboratory culture of tautog. Progress. Fish-Culturist 60, 50–54.

Puvanendran, V., Brown, J.A., 2002. Foraging, growth and survival of Atlantic cod larvae reared in different light intensities and photoperiods. Aquaculture 214, 131– 151.

Randall, J.E., 1980. Two new Indo-Pacific Labrid fishes of the genus Halichoeres, with notes on other species of the genus. Pacific Sci. 34, 415–432.

Randall, J.E., Allen, G.R., Steene, R.C., 1990. Fishes of the Great Barrier Reef and Coral Sea. University of Hawaii Press, Honolulu, Hawaii.

Reitan, K.I., Rainuzzo, J.R., Oie, G., Olsen, Y., 1997. A review of the nutritional effects of algae in marine fish larvae. Aquaculture 155, 207–221.

Rhyne, A.L., Tlusty, M.F., Schofield, P.J., Kaufman, L., Morris, J.A., Bruckner, A.W., 2012. Revealing the appetite of the marine aquarium fish trade: The volume and biodiversity of fish imported into the United States. PLoS One 7.

Rhyne, A.L., Tlusty, M.F., Szczebak, J.T., Holmberg, R.J., 2017. Expanding our

126

understanding of the trade in marine aquarium animals. PeerJ 5.

Robertson, D.R., 1981. The social and mating systems of two labrid fishes, Halichoeres maculipinna and H. garnoti, off the Caribbean coast of Panama. Mar. Biol. 64, 327– 340.

Ronnestad, I., Yúfera, M., Ueberschär, B., Ribeiro, L., Saele, O., Boglione, C., 2013. Feeding behaviour and digestive physiology in larval fish: current knowledge, and gaps and bottlenecks in research. Rev. Aquac. 5, S59–S98.

Rubec, P.J., Cruz, F., Pratt, V., Oellers, R., McCullough, B., Lallo, F., 2001. Cyanide- free net-caught fish for the marine aquarium trade. Aquarium Sci. Conserv. 3, 37– 51.

Sampey, A., McKinnon, A.D., Meekan, M.G., McCormick, M.I., 2007. Glimpse into guts: overview of the feeding of larvae of tropical shorefishes. Mar. Ecol. Prog. Ser. 243– 257.

Setu, S.K., Ajith Kumar, T.T., Balasubramanian, T., Dabbagh, A.R., Keshavarz, M., 2010. Breeding and rearing of regal damselfish Neopomacentrus cyanomos (Bleeker, 1856): The role of green water in larval survival. World J. Fish Mar. Sci. 2, 551–557.

Shaw, G.W., Pankhurst, P.M., Battaglene, S.C., 2006. Effect of turbidity, prey density and culture history on prey consumption by greenback flounder Rhombosolea tapirina larvae. Aquaculture 253, 447–460.

Shoji, J., Tanaka, M., 2004. Effect of prey concentration on growth of piscivorous Japanese Spanish mackerel Scomberomorus niphonius larvae in the Seto Inland Sea, Japan. J. Appl. Ichthyol. 20, 271–275.

Skiftesvik, A.B., Bjelland, R.M., Durif, C.M.F., Johansen, I.S., Browman, H.I., 2013. Delousing of Atlantic salmon (Salmo salar) by cultured vs. wild ballan wrasse (Labrus bergylta). Aquaculture 402–403, 113–118.

Stottrup, J.G., 2000. The elusive copepods: their production and suitability in marine aquaculture. Aquac. Res. 31, 703–711.

Sweet, T., 2016. The state of the marine breeders’ art 2017 (A CORAL special report). CORAL.

Sweet, T., 2013. Captive-breeding: State of the art 2014. CORAL.

Sweet, T., Pedersen, M., 2018. The state of the marine breeder’s art, 2018 (A CORAL special report). CORAL 38–47.

Sweet, T., Pedersen, M., 2015. The state of the marine breeders’ art 2016 (A CORAL special report). CORAL.

127

Thornhill, D.J., 2012. Ecological impacts and practices of the coral reef wildlife trade. Defenders Wildl. 187.

Thresher, R.E., 1984. Reproduction in reef fishes. T.F.H. Publication, Inc Ltd., Neptune City, NJ.

Thresher, R.E., 1979. Social behavior and ecology of two sympatric wrasses (Labridae: Halichoeres spp.) off the coast of Florida. Mar. Biol. 53, 161–172.

Tlusty, M.F., 2002. The benefits and risks of aquacultural production for the aquarium trade. Aquaculture 205, 203–219.

Townsend, D., 2011. Sustainability, equity and welfare: A review of the tropical marine ornamental fish trade. SPC Live Reef Fish Inf. Bull. 20, 2–12.

Trotter, A.J., Battaglene, S.C., Pankhurst, P.M., 2003. Effects of photoperiod and light intensity on initial swim bladder inflation, growth and post-inflation viability in cultured striped trumpeter (Latris lineata) larvae. Aquaculture 224, 141–158.

Turingan, R.G., Beck, J.L., Krebs, J.M., Licamele, J.D., 2005. Development of feeding mechanics in marine fish larvae and the swimming behavior of zooplankton prey: Implications for rearing marine fishes, in: Lee, Cheng‐Sheng O’Bryen, Patricia J. Marcus, N.H. (Ed.), Copepods in Aquaculture. Blackwell Publishing, Ames, Iowa, USA, pp. 119–132.

Vallone, D., Lahiri, K., Dickmeis, T., Foulkes, N.S., 2007. Start the clock! Circadian rhythms and development. Dev. Dyn. 236, 142–155.

Vandeputte, M., Dupont-Nivet, M., Haffray, P., Chavanne, H., Cenadelli, S., Parati, K., Vidal, M.O., Vergnet, A., Chatain, B., 2009. Response to domestication and selection for growth in the European sea bass (Dicentrarchus labrax) in separate and mixed tanks. Aquaculture 286, 20–27.

Victor, B.C., 1986a. Duration of the planktonic larval stage of one hundred species of Pacific and Atlantic wrasses (family Labridae). Mar. Biol. 90, 317–326.

Victor, B.C., 1986b. Larval settlement and juvenile mortality in a recruitment-limited coral feef fish population. Ecol. Monogr. 56, 145–160.

Videler, J.J., 1986. Sleep under sand cover of the labrid fish Coris julis. Phylogeny of Sleep 145–147.

Villamizar, N., Blanco-Vives, B., Migaud, H., Davie, A., Carboni, S., Sánchez-Vázquez, F.J., 2011. Effects of light during early larval development of some aquacultured teleosts: A review. Aquaculture 315, 86–94.

Villamizar, N., García-Alcazar, A., Sánchez-Vázquez, F.J., 2009. Effect of light spectrum and photoperiod on the growth, development and survival of European

128

sea bass (Dicentrarchus labrax) larvae. Aquaculture 292, 80–86.

Wabnitz, C., Taylor, M., Green, E., Razak, T., 2003. From ocean to aquarium: the global trade in marine ornamental species. UNEP-WCMC, Cambridge, UK.

Warner, R.R., Robertson, D.R., 1978. Sexual patterns in the labroid fishes of the western Caribbean, I: the wrasses (Labridae). Smithson. Contrib. to Zool. 1–27.

Wittenrich, M.L., Rhody, N.R., Turingan, R.G., Main, K.L., 2009. Coupling osteological development of the feeding apparatus with feeding performance in common snook, Centropomus undecimalis, larvae: Identifying morphological constraints to feeding. Aquaculture 294, 221–227.

Wittenrich, M.L., Turingan, R.G., 2011. Linking functional morphology and feeding performance in larvae of two coral-reef fishes. Environ. Biol. Fishes 92, 295–312.

Wittenrich, M.L., Turingan, R.G., Creswell, R.L., 2007. Spawning, early development and first feeding in the gobiid fish Priolepis nocturna. Aquaculture 270, 132–141.

Woolley, L.D., Qin, J.G., 2010. Swimbladder inflation and its implication to the culture of marine finfish larvae. Rev. Aquac. 2, 181–190.

Yúfera, M., Darias, M.J., 2007. The onset of exogenous feeding in marine fish larvae. Aquaculture 268, 53–63.

129

BIOGRAPHICAL SKETCH

Elizabeth M. Groover spent more than half her life living in New Hampshire, always intrigued by the marine life inhabiting its “expansive” 18 miles of coastline.

Knowing her career path would somehow involve the ocean and its inhabitants, she pursued a Bachelor of Science degree in marine biology and a minor in aquaculture and aquarium science at Roger Williams University in Rhode Island. After graduating from

RWU in 2015, Elizabeth traveled half way around the world and embarked on a six- month internship with Biota Marine Life Nursery in Palau. Elizabeth was awarded a graduate assistantship at the University of Florida Tropical Aquaculture Laboratory in

August 2016 and began her master’s program under the guidance of Dr. Matthew

DiMaggio. In the summer of 2018, Elizabeth graduated with her Master of Science degree in fisheries and aquatic sciences.

130