from oocyte to larva

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The long snout enables the fish to reach filamentous algae ...... representatives of seahorses (Syngnathidae) with insemination in the ven- tral brood pouch of the ...
Russian Academy of Sciences Severtsov Institute of Ecology and Evolution Russian-Vietnamese Tropical Research and Technological Center

Natal’ya G. Emel’yanova Dimitri A. Pavlov

FROM OOCYTE TO LARVA: HORMONAL INDUCTION OF OOCYTE MATURATION AND INITIAL DEVELOPMENT OF ORNAMENTAL CORAL REEF FISHES

Edited by Dmitry S. Pavlov Academician of Russian Academy of Sciences, Department of Ichthyology, Biological Faculty of Moscow State University, Moscow, Russia

KMK SCIENTIFIC PRESS Moscow 2012

Natal'ya G. Emel'yanova, Dimitri A. Pavlov. From Oocyte to Larva: Hormonal Induction of Oocyte Maturation and Initial Development of Ornamental Coral Reef Fishes. Moscow: KMK Scientific Press. 2012. 170 p. The culture of ornamental coral reef fish species conserves natural reef resources by offering alternatives to wild capture and develops a new source of organisms for the aquarium trade. Hormonal stimulation of maturation of sex cells facilitates obtaining large numbers of eggs and larvae available for subsequent rearing and allows researchers to collect valuable knowledge about life histories of the species to improve the management of natural stocks. In the monograph, detailed information on hormonal stimulation of oocyte maturation and ovulation, morphological changes of oocytes during maturation, ultrastructure of gametes, assessment of egg and sperm quality, embryonic and larval development, and transition of the larvae to exogenous feeding is presented for three model species from the families Acanthuridae (scopas tang Zebrasoma scopas) and Pomacentridae (threespot damselfish Dascyllus trimaculatus and scissortail sergeant Abudefduf sexfasciatus), which are used in aquaria trade. In addition, a brief description of the experiments on hormonal stimulation of oocyte maturation and ovulation is given for 16 other ornamental coral reef fish species. The results are discussed based on recent achievements in developmental biology and marine aquaculture. The book is including 38 black and white and 21 color illustrations intended for ichthyologists, zoologists, marine aquaculturists, and students of the universities or colleges.

Н.Г. Емельянова, Д.А. Павлов. От ооцита до личинки: гормональная стимуляция созревания ооцитов и начальное развитие декоративных коралловых рыб. М.: Товарищество научных изданий КМК. 2012. 170 с. Искусственное разведение декоративных коралловых рыб является альтернативой изъятия живых особей из природы, подрывающего естественные популяции, и создаёт дополнительный источник организмов для аквариумной индустрии. Гормональная стимуляция созревания половых клеток способствует получению большого числа икры и личинок, которые могут использоваться для последующего культивирования, и позволяет накапливать сведения о жизненном цикле рыб с целью разработки стратегии управления естественными популяциями. В монографии детальная информация по гормональной стимуляции созревания и овуляции ооцитов, морфологическим изменениям ооцитов в процессе созревания, ультраструктуре гамет, оценке качества яиц и спермы, эмбрионально-личиночному развитию и переводу личинок на экзогенное питание приведена для трёх модельных видов, являющихся объектами морской аквариумистики: Zebrasoma scopas (Acanthuridae), Dascyllus trimaculatus и Abudefduf sexfasciatus (Pomacentridae). Дано краткое описание экспериментов по гормональной стимуляции созревания и овуляции ооцитов у 16 других декоративных коралловых видов рыб. Результаты обсуждаются в связи с современными достижениями в области биологии развития и морской аквакультуры. Книга, включающая 38 черно-белых и 21 цветную иллюстрацию, предназначена для ихтиологов, зоологов, специалистов в области морской аквакультуры, студентов и аспирантов университетов и ВУЗов. Reviewers: M.I. Shatunovskii and E.V. Mikodina ISBN 978-5-87317-831-5

© IPEE RAS, 2012 © KMK SCIENTIFIC PRESS, 2012

TO THE MEMORY OF PROFESSOR GEORGIJ G. NOVIKOV

Georgij G. Novikov, professor of the Department of Ichthyology, Biological Faculty of Moscow State University, Moscow, Russia, head of the Laboratory of Fish Ontogeny. Over many years until his death (in 2007) he was a manager of the investigations of the Coastal Department of the Russian-Vietnamese Tropical Research and Technological Center. This book was prepared according to his initiative. We are grateful to Georgij G. Novikov for a possibility of the joint work and his continuous support of our study.

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CONTENTS Introduction ................................................................................................................. 8 Chapter1. Reproductive biology of fishes from the families Acanthuridae and Pomacentridae: a brief review ........................................................................ 12 1.1. Acanthuridae ...................................................................................................... 12 1.2. Pomacentridae .................................................................................................... 13 Chapter 2. Hormonal stimulation of maturation and ovulation of oocytes ....... 18 2.1. Methods of hormonal stimulation: a brief review .............................................. 18 2.2. Schemes of hormonal stimulation and morphological changes of oocytes in coral reef fish species ................................................................................................ 22 2.2.1. Zebrasoma scopas ........................................................................................ 22 2.2.1.1. Biological parameters and external morphology of the gonads .............. 22 2.2.1.2. Double injections .................................................................................... 23 2.2.1.3. Single injections ...................................................................................... 26 2.2.1.4. Size composition of oocytes ................................................................... 26 2.2.1.5. Morphological changes of oocytes during maturation ............................ 31 2.2.1.6. Temporal variation of oocyte maturation ................................................ 35 2.2.2. Dascyllus trimaculatus ................................................................................. 35 2.2.2.1. Biological parameters and external morphology of gonads .................... 36 2.2.2.2. Double injections .................................................................................... 37 2.2.2.3. Size composition of oocytes ................................................................... 38 2.2.2.4. Morphological changes of oocytes during maturation ............................ 42 2.2.3. Abudefduf sexfasciatus ................................................................................. 44 2.2.3.1. Biological parameters and external morphology of the gonads .............. 44 2.2.3.2. Double and single injections ................................................................... 44 2.2.3.3. Size composition of oocytes ................................................................... 47 2.2.3.4. Morphological changes of oocytes during maturation ............................ 47 2.2.4. Conclusions .................................................................................................. 50 2.3. Some data on hormonal stimulation and maturation of oocytes in other coral fishes ......................................................................................................................... 54 2.3.1. Biological parameters of the fishes .............................................................. 54 2.3.2. Materials on induced ovulation .................................................................... 55 2.3.3. Features of gametes ...................................................................................... 57 Chapter 3. Morphology and ultrastructure of gametes ....................................... 59 3.1. Oocyte envelopes ............................................................................................... 59 3.1.1. Oocyte envelopes in teleost fishes: a brief review ........................................ 59 3.1.2. Oocyte envelopes in coral reef fish species .................................................. 59 3.1.2.1. Zebrasoma scopas ................................................................................... 59 3.1.2.2. Dascyllus trimaculatus ............................................................................ 62 3.1.2.3. Abudefduf sexfasciatus ............................................................................ 64 3.1.3. Conclusions .................................................................................................. 64 3.2. Spermatozoa ...................................................................................................... 66

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3.2.1. Spermatozoa in teleost fishes: a brief review ............................................... 65 3.2.2. Spermatozoa in coral reef fish species .......................................................... 67 3.2.2.1. Zebrasoma scopas ................................................................................... 67 3.2.2.2. Dascyllus trimaculatus ............................................................................ 68 3.2.2.3. Abudefduf sexfasciatus ............................................................................ 69 3.2.3. Conclusions .................................................................................................. 70 Chapter 4. Assessment of sperm quality ............................................................... 72 4.1. Methods of assessment of sperm quality in teleost fishes: a brief review .......... 72 4.2. A method of assessment of sperm quality in coral reef fish species ................... 74 4.3. Conclusions ........................................................................................................ 80 Chapter 5. Assessment of egg quality .................................................................... 84 5.1. Methods of assessment of egg quality in teleost fishes: a brief review .............. 84 5.2. Morphological changes in the eggs and assessment of their quality in coral reef fish species ................................................................................................................ 88 5.2.1. Zebrasoma scopas ........................................................................................ 88 5.2.1.1. Morphology of oocytes after stripping ..................................................... 88 5.2.1.2. Morphological changes in non-inseminated oocytes ................................ 88 5.2.1.3. Morphological changes in eggs of different quality after insemination ... 89 5.2.1.4. Fertility of oocytes after their in vivo storage in the female's ovarian cavity ..................................................................................................................... 94 5.2.1.5. Fertility of oocytes after their in vitro storage in ovarian fluid ................ 96 5.2.1.6. Fertility of oocytes after their short-term storage in marine water ........... 98 5.2.2. Dascyllus trimaculatus ................................................................................ 98 5.2.2.1. Fertility of oocytes after their in vitro storage in ovarian fluid ............... 98 5.2.2.2. Fertility of oocytes after their short-term storage in marine water .......... 98 5.2.3. Abudefduf sexfasciatus ............................................................................... 100 5.2.3.1. Fertility of oocytes after their in vivo storage in the female's ovarian cavity ................................................................................................................. 100 5.2.3.2. Fertility of oocytes after their in vitro storage in ovarian fluid ............. 102 5.2.3.3. Fertility of oocytes after their short-term storage in marine water ........ 102 5.3. Conclusions ...................................................................................................... 102 Chapter 6. Development of eggs and transition of larvae to exogenous feeding .... 108 6.1. Culture of marine fish species: a brief review .................................................. 108 6.2. Culture of coral reef fish species ...................................................................... 112 6.2.1. Zebrasoma scopas ...................................................................................... 112 6.2.2. Dascyllus trimaculatus ............................................................................... 115 6.2.3. Abudefduf sexfasciatus ............................................................................... 117 6.3. Conclusions ...................................................................................................... 120 Concluding remarks ................................................................................................ 123 References .............................................................................................................. 129 Color Plates ............................................................................................................. 150 Acknowledgments .................................................................................................. 171

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ÑÎÄÅÐÆÀÍÈÅ Предисловие .............................................................................................................. 8 Глава 1. Биология размножения рыб семейств Acanthuridae и Pomacentridae: краткий обзор ......................................................................... 12 1.1. Acanthuridae ...................................................................................................... 12 1.2. Pomacentridae .................................................................................................... 13 Глава 2. Гормональная стимуляция созревания и овуляции ооцитов ......... 18 2.1. Методы гормональной стимуляции: краткий обзор ...................................... 18 2.2. Схемы гормональной стимуляции и морфологические изменения ооцитов у коралловых рыб ..................................................................................... 22 2.2.1. Zebrasoma scopas ........................................................................................ 22 2.2.1.1. Биологические параметры и морфология гонад ................................. 22 2.2.1.2. Двукратные инъекции ........................................................................... 23 2.2.1.3. Однократные инъекции ........................................................................ 26 2.2.1.4. Размерный состав ооцитов ................................................................... 26 2.2.1.5. Морфологические изменения ооцитов в процессе созревания ......... 31 2.2.1.6. Временная изменчивость ооцитов в процессе созревания ................ 35 2.2.2. Dascyllus trimaculatus ................................................................................. 35 2.2.2.1. Биологические параметры и морфология гонад ................................. 36 2.2.2.2. Двукратные инъекции ........................................................................... 37 2.2.2.3. Размерный состав ооцитов ................................................................... 38 2.2.2.4. Морфологические изменения ооцитов в процессе созревания ......... 42 2.2.3. Abudefduf sexfasciatus ................................................................................. 44 2.2.3.1. Биологические параметры и морфология гонад ................................. 44 2.2.3.2. Двукратные и однократные инъекции ................................................. 44 2.2.3.3. Размерный состав ооцитов ................................................................... 47 2.2.3.4. Морфологические изменения ооцитов в процессе созревания ......... 47 2.2.4. Заключение ................................................................................................. 50 2.3. Некоторые данные по гормональной стимуляции и созреванию ооцитов других коралловых рыб ....................................................................................................... 54 2.3.1. Биологические показатели рыб ................................................................. 54 2.3.2. Данные по индукции овуляции ................................................................. 55 2.3.3. Особенности гамет ..................................................................................... 57 Глава 3. Морфология и ультраструктура гамет .............................................. 59 3.1. Оболочки ооцитов ............................................................................................ 59 3.1.1. Оболочки ооцитов у костистых рыб: краткий обзор ............................... 59 3.1.2. Оболочки ооцитов у коралловых рыб ....................................................... 59 3.1.2.1. Zebrasoma scopas ................................................................................... 59 3.1.2.2. Dascyllus trimaculatus ............................................................................ 62 3.1.2.3. Abudefduf sexfasciatus ............................................................................ 64 3.1.3. Заключение ................................................................................................. 64 3.2. Сперматозоиды ................................................................................................. 65 3.2.1. Сперматозоиды костистых рыб: краткий обзор ....................................... 65 3.2.2. Сперматозоиды коралловых рыб ............................................................... 67 3.2.2.1. Zebrasoma scopas ................................................................................... 67 3.2.2.2. Dascyllus trimaculatus ............................................................................ 68

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3.2.2.3. Abudefduf sexfasciatus ............................................................................ 69 3.2.3. Заключение ................................................................................................. 70 Глава 4. Оценка качества спермы ...................................................................... 72 4.1. Методы оценки качества спермы костистых рыб: краткий обзор ............... 72 4.2. Метод оценки качества спермы у коралловых рыб ....................................... 74 4.3. Заключение ....................................................................................................... 80 Глава 5. Оценка качества яиц ............................................................................. 84 5.1. Методы оценки качества яиц костистых рыб: краткий обзор ...................... 84 5.2. Морфологические изменения яиц и оценка их качества у коралловых рыб .. 88 5.2.1. Zebrasoma scopas ........................................................................................ 88 5.2.1.1. Морфология ооцитов после отцеживания из тела самки .................. 88 5.2.1.2. Морфологические изменения в неосеменённых ооцитах .................. 88 5.2.1.3. Морфологические изменения в яйцах разного качества после осеменения ......................................................................................................... 91 5.2.1.4. Фертильность ооцитов после их задержки в овариальной полости самки (in vivo) .................................................................................................... 94 5.2.1.5. Фертильность ооцитов после их выдерживания в овариальной жидкости (in vitro) .............................................................................................. 96 5.2.1.6. Фертильность ооцитов после их кратковременного выдерживания в морской воде .................................................................................................... 98 5.2.2. Dascyllus trimaculatus ................................................................................. 98 5.2.2.1. Фертильность ооцитов после их выдерживания в овариальной жидкости (in vitro) .............................................................................................. 98 5.2.2.2. Фертильность ооцитов после их кратковременного выдерживания в морской воде .................................................................................................... 98 5.2.3. Abudefduf sexfasciatus ............................................................................... 100 5.2.3.1. Фертильность ооцитов после их задержки в овариальной полости самки (in vivo) .................................................................................................. 100 5.2.3.2. Фертильность ооцитов после их выдерживания в овариальной жидкости (in vitro) ............................................................................................ 102 5.2.3.3. Фертильность ооцитов после их кратковременного выдерживания в морской воде .................................................................................................. 102 5.3. Заключение ..................................................................................................... 102 Глава 6. Развитие яиц и переход личинок на экзогенное питание ............. 108 6.1. Культивирование морских рыб: краткий обзор ........................................... 108 6.2. Культивирование коралловых рыб ............................................................... 112 6.2.1. Zebrasoma scopas ...................................................................................... 112 6.2.2. Dascyllus trimaculatus ............................................................................... 115 6.2.3. Abudefduf sexfasciatus ............................................................................... 117 6.3. Заключение ..................................................................................................... 120 Заключительные комментарии ............................................................................. 123 Список литературы ............................................................................................... 129 Цветные иллюстрации .......................................................................................... 150 Благодарности ....................................................................................................... 171

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Introduction Although reefs cover less than one quarter of one percent of the marine environment, they are considered to be amongst the most biologically rich and productive ecosystems on Earth, often described as the “rainforest of the seas”. Coral reefs support over 4000 species of fish (or a third of the world’s marine fish species), about 800 species of reef-building corals, and a great number of other invertebrates and sponges. Although most fish kept in aquariums are from freshwater, the acquisition of marine ornamental fish has greatly increased representing a popular and rapidly expanding trend in the hobby to establish marine reef mini-ecosystems within the aquarium. Between 1.5 and 2 million people worldwide are believed to keep marine aquaria. The trade which supplies this hobby with live marine animals is a global multi-million dollar industry, worth an estimated US$200– 330 million annually. Marine ornamental fish make up approximately 85% of the trade in live coral reef species for aquaria. All countries of the European Union and, to a lesser extent, Japan are the market for ornamental fish; however, the United States is the single largest importer of ornamental fish in the world (Chapman et al. 1997; Baquero 1999; Larkin and Degner 2001; Wabnitz et al., 2003; Livengood and Chapman, 2007). Collection of species for the trade is predominantly from the Philippines, Fiji, Indonesia, Brazil, Vietnam, the Maldives, Sri Lanka, the Red Sea, and Hawaii. Currently, as many as 30 million coral reef fish belonging to 1000 species are collected annually to supply private and public aquaria around the world. Of these about 8 million go to the US, 8 million to Europe and the rest to countries such as Japan, Australia, and South Africa (Wood, 2001). Damselfish (Pomacentridae) make up almost half of the trade, with species of angelfish (Pomacanthidae), surgeonfish (Acanthuridae), wrasses (Labridae), gobies (Gobiidae), and butterflyfish (Chaetodontidae) accounting for approximately another 25–30 percent. The most traded species are the blue-green damselfish (Chromis viridis), the clown anemonefish (Amphiprion ocellaris), the whitetail dascyllus (Dascyllus aruanus), the sapphire devil (Chrysiptera cyanea), and the threespot dascyllus (Dascyllus trimaculatus) (Wabnitz et al., 2003). It is estimated that 90–99% of the trade in ornamental marine fish is provided by wild collection. Nearly all the specimens come from coral reefs. These ecosystems are under considerable stress from a range of direct and indirect human and “natural” impacts including changes to the global climate which have caused widespread coral bleaching and mortality. Almost 90% of the coral reefs in the Philippines and Indonesia (as well as Cambodia, Singapore, Taiwan, Vietnam, Malaysia, and China) are threatened. Many 8

of the most threatened areas, some of which have few protected areas, are also those areas most exploited by the marine aquarium trade. The industry that supports the trade in marine ornamentals worldwide is mostly unlicensed and unregulated (Brown, 1997; Tullock, 1998; Wood, 2001). There are several key differences between the food fishing industry and the marine ornamental industry that will require a new approach to both research and management. These differences are not only related to the differing collection methods of each industry, but to the varying life history traits and biological characteristics of targeted fish. Over-collection of targeted organisms and destructive fishing practices, such as cyanide fishing and breaking coral to obtain stunned fish, may threaten the biodiversity of fish on a local level and can result in local extinctions of rare or endemic species. Additionally, the selective removal of certain species of fish from a particular reef may lead to unnatural shifts in community structure and function (Hodgson et al., 2003). The greatest losses of wild-caught aquarium fish may occur during the handling period between capture, local exporting/importing warehouses, and transportation docks. While in transit, fish may be subjected to physical injury, extreme changes in water quality conditions, water temperature fluctuations, and indiscriminate exposure to toxic chemicals used as prophylactic treatments for disease control (Livengood and Chapman, 2007). Reports indicate that mortality of all marine species captured for the trade may be 50% or higher during the process of removing fish from the reef to the often primitive holding facilities prior to export. An additional 30% mortality can then occur at each step in the transport chain, culminating in what may be over 90% mortality before retail purchase (Rubec et al., 2001). Increasing pressures on natural populations of coral reef animals due to their expanding popularity in the aquarium trade has stimulated interest in the culture of marine tropical fish. This culture conserves natural reef resources by offering alternatives to wild capture and develops a new source of organisms for the aquarium trade. In the closed systems, the juveniles and market-size fish of a wide variety of species can be produced year round. Furthermore, rearing aquarium fish will lead to the production of hardier species, which fare better in captivity and survive longer. In addition, cultivation of ornamental fish can be much more feasible compared to that of fish for human consumption. For example, in 2000, 1 kg of aquarium fish from the Maldives was valued at almost US$500, whereas 1 kg of reef fish harvested for food was worth only US$6. Increased rarity often implies higher prices. Individuals of two rare species, the yellow-faced angelfish, Pomacanthus xanthometapon, and the bluegirdled angelfish, 9

Pomacanthus navarchus, fetch prices in the range of hundreds of dollars in the US. The peppermint angelfish Centropyge boylei may command a price as high as US$10 000. The deep reef species, Tinker’s butterflyfish, Chaetodon tinkeri, can sell for up to US$1000 per pair due to the difficulty in collecting the fish living at depths of 27 to 135 m. (Wabnitz et al., 2003). Artificial reproduction of ornamental coral reef fishes will also allow researchers to collect valuable information about their life history to improve the management of natural stocks and our understanding of how these organisms respond to human impacts (Olivotto et al., 2011). However, aquaculture is growing very slowly due to economic and biological constraints to rearing ornamental marine fishes in captivity. Cultured fish account for one to two percent of the global trade. There are now over 100 species of marine fish that have been successfully bred in captivity, but very few of these species are as yet, produced in commercial quantities. Only about 40 of the species currently traded can be farm raised and are commercially available. A list of the species bred for the marine aquarium trade is represented by Wabnitz et al. (2003). The species successfully bred at the largest marine ornamental fish hatchery in the world (ORA) include 17 species of anemonfish (Pomacentridae), 11 species of Pseudochromidae, 8 species of Gobiidae, 7 species of Syngnathidae, and 4 species of Apogonidae (Oceans, Reefs and Aquariums, 2008). The two key bottlenecks that currently limit expansion of the marine ornamental industry are the control of captive maturation and spawning and the identification of appropriate first-feed items for marine ornamental fish larvae (Ostrowski and Laidley, 2001). At present, natural spawning in captivity is almost exclusively applied to obtain offspring of marine ornamental fishes. For the beginning of natural reproduction, a prolonged adaptation of the fish to artificial conditions is required using complex and expensive systems with water recirculation and control of environmental parameters. For example, Chrysiptera parasema (Pomacentridae) needs a three-month period to start spawning (Olivotto et al., 2003), and the first spawning is registered in seven months after capture of adult exemplars of ornamental fish species from the families Pomacanthidae (pygmy angelfish Centropyge argi) and Labridae (spotfin hogfish Bodianus puchellus, bluehead wrasse Thalassoma bifasciatum, and clown wrasse Halichores maculipinna) (Holt and Riley, 2000). Hormonal stimulation of maturation is an alternative method for obtaining mature sex products in ornamental fish species. Up to now, hormonal induction of oocyte maturation and ovulation almost has not been conducted for marine ornamental fish species, but it is applied for several cold and 10

warm water marine species used for human consumption and representing the objects for aquaculture. A goal of this book is the description of the schemes of hormonal stimulation of oocyte maturation and ovulation for coral reef ornamental fish species with different reproductive strategies using the model objects. We describe the features of gametogenesis and gamete structure that is important for the understanding of general and specific (environmentally induced) processes of development of sex cells in the fishes from different taxa. An important part of the work on hormonal treatment of the fish is the assessment of the quality of sex products. In this book, the new and modified methods are suggested for the assessment of egg and sperm quality in marine fishes. In addition, the experiments on egg incubation and rearing of larvae are carried out. The study was conducted in the Coastal Department of the Russian–Vietnamese Tropical Research and Technological Center (Nha Trang, Vietnam) from 2002 to 2008. In addition, the experiments were done at the Mariculture Station of the Institute of Aquaculture no. 3 (Nha Trang, Vietnam) (Color Plates, Fig. 1). The fishes for investigations were collected in the Nha Trang Bay of the South China Sea in the coastal areas of the northern and southern parts of Hon Tre Island (12°10'–12°14' N; 109°14'–109°21' E). This area was included into the Marine Protected Area of the Nha Trang Bay since 2002 (Le Doan Dung, 2007). The fishes were sampled by fishermen using simple diving equipment (hookah diving) and drive-in nets (up to 50 m in length, mesh size less than 1.5 cm). Live fishes were transported to the laboratory for several hours. At the Tropical Center, the fishes were kept in the tanks 4 m3 in volume in the recirculated system (water exchange rate 4–8 m3/h) with the biofilters. For the short-term keeping of the fish (for several days) during the experiments on hormonal stimulation of sexual maturation and ovulation, they were placed in the cages (80 x 80 x 100 cm) installed in the tanks. The salinity was between 33 and 35‰, and the water temperature was 25–29°C. The model objects for the study are as follows: scopas tang Zebrasoma scopas from the family Acanthuridae, threespot damselfish Dascyllus trimaculatus, and scissortail sergeant Abudefduf sexfasciatus from the family Pomacentridae (Color Plates, Fig. 2). The representatives of the former species possess pelagic eggs, and the fishes from two latter species deposit demersal adhesive eggs protected by the male during embryonic development inside of egg envelopes. D. trimaculatus, Abudefduf spp., and Z. flavescens are included in the top ten most traded species of ornamental fish according to datasets derived for the United States, the EU, and worldwide. 11

Chapter 1.Reproductive biology of fishes from the families Acanthuridae and Pomacentridae: a brief review Fishes are the vertebrate taxon with the highest diversity of life styles. This diversity by all sorts of mating systems, from the spawning in large groups to strict monogamy is observed in coral fishes. The majority of these fishes spawn pelagic eggs with no-care or demersal eggs protected by parents, but the fishes brooded the eggs and live-bearers can be found in several families (Jan, 2000). A mode of reproduction is crucial for developing of the methods of induced maturation of sex products of the species, and the features of oocytes and spermatozoa maturation, gamete structure, and embryonic and larval development can be only explained based on the knowledge on the reproductive biology of the species. A brief review of the reproductive patterns of the representatives of the families Acanthuridae and Pomacentridae, and, in particular, the species chosen as the model objects, is given in this chapter. 1.1. Acanthuridae The Surgeonfish of the family Acanthuridae belong to the suborder Acanthuroidei of the order Perciformes. The representatives of the family are widely distributed on the coral reefs of lower latitudes of the Pacific, Indian, and Atlantic oceans. The family includes three subfamilies (Acanthurinae, Prionurinae, and Nasinae), six genera, and approximately 80 species (Randall, 2002). The representatives of the subfamily Acanthurinae are characterized by a deep compressed body usually of an oval shape and a scalpedlike sliding spine on each side of the caudal peduncle. The spine is very sharp and is used by the fish for protection from predators as well as a way of establishing itself with other fish. All species of Zebrasoma, the majority of the species of the genus Acanthurus, and a part of the species of the genus Naso are herbivorous. Other species are planktivorous, and all species of the genus Ctenochaetus are detritivorous. The eggs and larvae are pelagic. The genus Zebrasoma is a relatively small group consisting of seven species that are characterized by their round laterally compressed bodies and pointed snouts. The long snout enables the fish to reach filamentous algae in reef interstices that are out of the reach of other Acanthurids. The representatives of the genus are found in every tropical ocean and sea except the Atlantic. The fish are popular in the aquarium trade because of their color, shape, and propensity to eat nuisance algae in the aquarium. 12

The species of the genus Zebrasoma are herbivorous, and, therefore, have an important role in the ecosystems. For example, in Hawaii, 80% of the catch of ornamentals consists of herbivorous fishes (primarily Zebrasoma), and reductions in the abundance of herbivores can cause algal overgrowth of corals (Tissot and Hallacher. 2003). The scopas tang Zebrasoma scopas also known as the brown scopas tang or twotone tang is distributed in the Indo-Pacific: East Africa, including the Mascarene Islands to the Tuamoto Islands, north to southern Japan, south to Lord Howe and Rapa islands at the water temperature between 25 and 28°C. Adult individuals of scopas tangs usually do not exceed 20 cm, but the maximum reported size is 40.0 cm of standard length (SL). The fish is light yellow or yellow dorsally, and brown or black ventrally. These colors gradually get darker from front to back. The body is covered with fine, intricate, light blue markings. The tail is a solid brown color. Juveniles are a little more attractive, with larger dorsal and anal fins and a slightly purplish body color. There are no visible differences between males and females. The fish occurs in coral-rich areas of lagoon and seaward reefs from a depth of 1 to 60 m. Juveniles are generally solitary fish and never stray far from the cover of the reef. Adults roam wide expanses of the reef and can be found singly, in pairs, and in large groups of the same or mixed species of tangs (Myers, 1991; Randall, 2002). The behavior and spawning of scopas tang from the Indo-Pacific is described by Robertson (1983). At both Aldabra and Palau islands, adults defended feeding territories against conspecific exemplars and representatives of Zebrasoma and Acanthurus. At Aldabra, all territories were occupied by groups of two–three fish while those at Palau were defended by one–four fish. Multi-individual groups at both sites consisted of a male and one or more smaller females. Single territorial fish at Palau were of both sexes. Nonterritorial fish were seen in compact schools of 50–75 individuals that fed in intertidal areas. The spawning was observed throughout the lunar cycle except for the few days immediately after new and full moon. At Aldabra, pair matings were observed in nine cases on ebb tides between 11:00 and 18:00 h. All the spawnings were observed in subtidal habitat normally occupied by the species. At Palau, over 50 pair matings were recorded, as well as a single group (four participants) spawning. The spawning of scopas tang in the groups (from eight to ten individuals) was also observed in the Society Islands (Randall, 1961). At Palau, each pair resumed feeding together in a territory on the substrate directly below the spawning site. The pair matings occurred in subtidal areas down to depths of at least 10 m and in intertidal areas as much as 300 m from the outer reef 13

edge. No large-scale spawning migrations by scopas tang living inshore from reef edge were evident. A change in coloration (slight paling of the head region and orange irises) of the spawning pairs sometimes was observed during courtship or spawning. During a pair spawning, the two fish began leaving the substrate and slowly rising up to 2 m above it. The male usually initiated such rising. They hung in the water column for a few seconds and then returned to the substrate. Milt was released at the apex of pair movements, each of which ended 1–4 m above the bottom. As in other reef fishes releasing planktonic eggs, there was a tendency to move up of the bottom closer to the surface to release eggs away from egg predators that live on reefs. The spawning mainly in the dusk at the localities with strong currents directed to the open sea represents an additional common adaptation of fishes with pelagic eggs to prevent predation.

1.2. Pomacentridae Damselfishes of the family Pomacentridae belong to the suborder Labroidei of the order Perciformes. Within Pomacentridae, Allen (1991) separated four subfamilies: Amphiprioninae, Chrominae, Lepidozyginae, and Pomacentrinae. The number of the genera and species reaches 28 and 350, respectively (Nelson, 1994; Robertson, 1998). According to Allen (1991), the genera Acanthochromis, Chromis, and Dascyllus should be placed together in the subfamily Chrominae on the basis of shared structural characteristics, and the genus Abudefduf belongs to the subfamily Pomacentrinae. Recent data (Quenouille et al., 2004) on the mitochondrial and nuclear DNA sequences reconsider previous systematics of damselfishes. Four principal clades, which are not corresponded to the subfamilies of Allen (1991) are recognized. The monophyly of the Chrominae and Pomacentrinae supsignificant ported by Allen (1991) is rejected. Clade 2 including Chromis +Dascyllus is highly signifcant, and the relative phylogenetic separation and basal placement of this clade in the Pomacentridae provides strong support for Allen’s (1991) general notion of the Chrominae. The species of the genus Abudefduf comprise Clade 3, which is similar to Clade 2 in the structure and phylogenetic history. In addition, the analysis does not support the monophyly of the genus Amphiprion. Pomacentridae is a prodigious family of small, often brightly colored, diurnally active shallow water fishes. The family occurs worldwide in subtropical and tropical waters and is considered one of the most important families of fishes inhabiting coral and rocky reefs. They are also common in other shallow-water habitats including sand and rubble patches, silty em14

bayments, harbours, and seagrass beds. Damselfishes are one of the most numerous groups occurring on tropical reefs, in terms of both number of species and number of individuals. Solitary fishes, pairs, or aggregations are equally common, depending on the species. Most species are highly territorial, especially during reproductive periods. Spawning occurs throughout most of the year at tropical localities. Eggs are deposited in clusters on the bottom and are guarded by the male. The eggs are elliptical in shape and are attached to the substrate by adhesive filaments. Hatching occurs within 2 to 7 days. Most species have a pelagic larval stage lasting between about 10 to 50 days, except Acanthochromis polyacanthus, which bypasses this stage entirely. Food habits are variable. The larger species tend to feed mainly on algae, many others are omnivorous, consuming algae and small invertebrates, while others rely on planktonic items. Species of the genus Amphiprion and some species of the genus Dascyllus are symbiotic with large sea anemones. Damselfishes are an extremely important group of tropical and sub-tropical marine fishes kept in marine aquariums. Anemone fishes are the leading group in this respect. Of the nine described Dascyllus species three are widely used in the aquarium hobby: threespot dascyllus (known also as threespot damselfish or domino damselfish) Dascyllus trimaculatus, four-striped (blacktail) humbug Dascyllus melanurus, and three-striped (white-tailed) damsel Dascyllus aruanus. A protogyny (ontogenetic sex change from female to male sex products) is reported for some representatives of the genus (Asoh and Yoshikawa, 2003; Asoh, 2005). Based on the results of histological analysis, threespot dascyllus is a gonochoristic species (Asoh and Yoshikawa, 2003) with not functional protogyny (Asoh and Kasuya, 2002). According to the phylogeographical and molecular systematics study (McCafferty et al., 2002), the ability to change sex (from functional females to males) arose once in the ancestor to the entire genus, and was lost in the ancestor of the D. trimaculatus complex. Threespot dascyllus inhabits coral and rocky reefs at a depth from 1 to 55 m and reaches maximum size of 11.0 cm SL. The species is distributed in the Indo-Pacific: Red Sea and East Africa to the Line and Pitcairn islands, north to southern Japan, south to Sydney, Australia. It is not found in the Hawaiian and Marquesan islands. The coloration of the juveniles is overall black with scale centers bluish with white blotch on the forehead and a prominent white spot on the upper side; all fins are black except the transparent pectoral and outer portion of soft dorsal rays. In a mature exemplar, the marks will most likely disappear or leave just a remnant spot on the 15

side. The juveniles are often commensal with large sea anemones, sea urchins, or small coral heads. The adults occur in small to large aggregations. Stomach contents include algae, copepods, and other planktonic crustaceans. The threespot dascyllus is among the easiest of all marine fish to keep in aquariums. The fish can eat all kinds of live, frozen and flake foods, and algae. As the majority of other representatives of the family Pomacentridae, threespot dascyllus possesses demersal eggs, which are deposited on a solid substrate. A description of the spawning of this species has not been found in the literature, but such data are available for three-striped damsel D. aruanus (Mizushima et al., 2000). A very distinct semilunar spawning cycle was found in a population on the coral reefs of Sesoko Island, Okinawa. Spawning occurred from June to September only in the early morning, during a period of 2–4 days immediately before or around the time of the new and full moon. When multiple females spawned in the same nest (up to four females), they usually spawned on the same morning or within two consecutive days. It was observed, that urogenital papillae of females projected for about 1 h after spawning. During the spawning period, 33% of marked females (n = 105) spawned from one to six times. The spawning intervals of the females spawned more than once were 13–59 days (mode = 14). Males protected eggs deposited on the substrate for 2.5 days until hatching. Hatching occurred just after sunset, i.e., at the high tide of spring tide; the strong ebb current then would rapidly disperse the newly hatched larvae offshore and allow them to escape from reef dwelling predators. Females tended to synchronize spawning in male’s nest. Another study that supports a semilunar spawning periodicity in pomacentrids is based on the dynamics of gonadosomatic index (GSI) and ovarian histology in brackish damsel Pomacentrus taeniometopon, which repeats spawning twice a month (Pisingan et al., 2006). Large exemplars of the genus Abudefduf are sometimes seen in markets, but the family is not considered as an important food fish. Of the twenty valid species of the genus, approximately nine species, including scissortail sergeant A. sexfasciatus, make their way into the aquarium trade. The fish can be maintained quite nicely on flake food, and are often offered brine shrimp, algae, and frozen foods. Because of their aggressive nature, they often can be kept with other large fish with similar temperaments. Scissortail sergeant is a non-migratory marine fish living on inshore and offshore coral or rocky reefs at a depth between 1 and 20 m. It is distributed in the Indo-Pacific: Red Sea to Pinda, Mozambique, and the Tuamoto Islands; north to southern Japan; south to Lord Howe and Rapa islands. It is 16

not recorded from the Hawaiian Islands (Allen, 1991). The fish feed on zoobenthos, zooplankton, and algae (Sano et al., 1984). Scissortail sergeant reaches 22 cm total length (TL). Its coloration is white with five black bands on the body and a dark longitudinal stripe on both lobes of the caudal fin. The reproductive and social biology is described in sergeant major A. saxatilis in “The Living Seas” aquarium 62 meters in diameter and 8 meters deep (Prappas et al., 1991). During courtship and spawning, both sexes show color dimorphism. The breeding males and females exhibited a deep bluish color with the bars very difficult to discern. During courtship, males had a conspicuous “white mask” on the head region. Sergeant majors go trough a series of behavioral actions associated with reproduction. Activities include the following events: (1) nest preparation, (2) courtship, (3) spawning, and (4) brood care. Nests were usually prepared on sheltered and vertical, horizontal, or overhanging surfaces, within artificial reef structures or natural rock surfaces. Some males defended eggs on more than one site, if the sites were in close proximity. After a suitable territory was chosen by the male and the territory established, nest preparation began with the clearing of algae, gravel, and other debris. During spawning and brood care, territories were vigorously guarded against interspecific intruders. The courtship included the “signal jumps”. The “signal jump” is the male’s attempt to attract a female to the spawning site. The male swims rapidly upward in an undulating manner and then returns to the point where the behavior began. If the male was successful, the ripe female would respond by returning to the nesting site, and the eggs were deposited and inseminated simultaneously. The female deposits the eggs by pressing her urogenital papilla to the nest surface and the male swims after the female pressing his urogenital papilla against the eggs to complete insemination. After a single spawning session, which ranged from 45 to 90 min, the female is chased away and the male remains to protect the brood. The males would guard the spawn (during the night and day), fanning the eggs and removing non-viable eggs from the nest to promote normal development of the remaining brood. Egg size was 0.6 × 1.0 mm, and the shape of pink or red colored eggs was elliptical. The average number of eggs in the mass was 27 000. It was common for a male to spawn consecutively as well as intermittently with two or three females. At a site, a male spawned with five females and guarded two separate nesting areas 0.75 meters apart. Embryos hatched on the average in six days. The larvae reach 2.4 mm in length approximately 36 hours after hatching.

17

Chapter 2. Hormonal stimulation of maturation and ovulation of oocytes 2.1. Methods of hormonal stimulation: a brief review At present, development of aquaculture is impossible without the control of the reproductive function in fishes. This control is necessary to obtain high quality gametes and progeny. For several fish species, hormonal stimulation of oocyte maturation (to obtain fertilized eggs and larvae for subsequent cultivation) can be used to prevent elimination of offspring or spawners from natural water bodies. For example, the culture of such species as freshwater eels (Anguilla spp.), yellowtail and greater amberjack (Seriola spp.), groupers (Epinephelus spp.), and bluefin tuna (Thunnus thynnus) is based almost entirely on the collection of juveniles from nature (Ottolenghi et al., 2004). Similar situation is usual for the majority of coral reef fishes. Sometimes, the reproduction of fishes can be controlled by means of manipulation of environmental parameters, such as photoperiod, water temperature, water depth and volume in the tanks, presence of spawning substrate etc. (Whitehead et al., 1978; Moksness and Pavlov, 1996). However, artificial environment often is responsible for inhibition of sexual maturation (Verigin et al., 1975; Makeyeva et al., 1994), and, in some cases, imitation of the environmental conditions usual for sexual maturation of the fish in nature (certain social or environmental parameters including water depth or pressure) is impossible or not profitable. Thus, hormonal therapy is applied to control reproduction of cultivated species, in particular, to induce or synchronize maturation and ovulation of oocytes. For some species, hormonal stimulation is the only way to obtain fertilized eggs; for other species, exogenous hormones are used to increase egg production and to facilitate culture practice (Goren et al., 1995; Haffray et al., 2005). Hormonal stimulation of oocyte maturation and ovulation was developed and applied in fish culture long before the understanding of biological mechanisms of these processes. Independently on each other, Von Ihering (1937) and Hasler (1939) showed that injections of pituitary extracts can lead to oocyte maturation and ovulation in fishes. In Russia, the method of hypophysation was developed and applied in the hatcheries by Gerbil’skii (1941). For many years, this method of regulation of the reproductive function in fishes remained unique. At the initial step of the hypophysation method, grinded pituitaries or pituitary extracts obtained from the spawners during a season of reproduction were injected to adult individuals of the same or other fish species (Von Ihering, 1937; Gerbil’skii, 1941; Migita et al., 1952; 18

Ball, 1954; Bezdenezhnykh, 1956). Modernization of the hypophysation method is connected with searching the most effective ways of injection of pituitary preparations (in the cerebrum ventricles, intramuscularly, intraperitoneally, or in the gonads), methods of their preservation (fresh or acetonised), and searching of pituitaries characterized by universal action to different fish species (homo- and heteroplastic injections) (Steffens, 1957; Konradt, 1961; Vinogradov et al., 1966; Makeyeva and Verigin, 1971; Lemanova, 1974; Burlakov, 1985; Brzuska, 1987; Glubokov, 1993). Pituitary extract of carp is widely distributed, and purified gonadotropic hormone (GtH) has been applied in several countries as a consequence of development of the method of protein detachment (Donaldson, 1973; Yaron, 1995). Phylogenetic studies (Li and Ford, 1998; Querat et al., 2000) demonstrated that the piscine GtHs, previously known as GtH I and GtH II, are directly related to the tetrapod follicle stimulating hormone (FSH) and luteinizing hormone (LH), respectively. This concept was further confirmed based on the chemical traits of the GtHs and their differential action during the reproductive cycle (Swanson and Dittman, 1997). Sex steroid hormones have been also used for stimulation of sexual maturation in fishes (Burlakov et al., 1985; Sorensen, 1986; Richter and Van der Hurk, 1987). However, many gonadotropins, androgens, and estrogens are species specific, and their application is not accompanied by intensive ovulation. Human chorionic gonadotropin (HCG) is widely used; it is structurally homologous to LH at a large degree, but its effect on oocyte maturation in many objects of culture is negligible (Barannikova, 1975; Verigin et al., 1975; Donaldson and Hunter, 1983; Zaki, 1989; Ludwig et al., 2002). Subsequent investigations have shown that the highest link of the regulation of the reproductive system is represented by hypothalamic releasing hormones (gonadoliberins). In the last two decades, after the discovery of gonadotropin-releasing hormones GnRH (Schally, 1978) and the synthesis of highly active agonists of GnRH (Goncharov, 1984; Crim and Bettles, 1997), spawning induction therapies shifted from the use of GtHs. It is connected with GnRHas action at the higher level of the hypothalamushypophysis-gonads axis; they can induce more integrated stimulation of reproductive processes by means of direct or indirect stimulation of other hormones involved into oocyte maturation, e.g., growth hormone (Le Gac et al., 1993), insulin-like growth factors (Negatu et al., 1998), prolactin (Weber et al., 1995), and thyroid hormones. Investigation of the structure of GnRHs shows that they are represented by decapeptides, which differ in the animals of different classes in aminoacid composition (Glubokov, 1993). In vertebrates, including fishes, at least 14 GnRH forms are found (Maňanós 19

et al., 2009). The following forms of gonadoliberins prevail in fishes: mammal (mGnRH), sea bream (sbGnRH), salmon (sGnRH), and chicken II (cGnRH II) (Mylonas and Zohar, 2007). Releasing hormones used for hormonal therapy are represented by synthetic analogs because native hormones are destructed by peptides very rapidly (Zohar et al., 1990). Substitutions of the GnRH decapeptide at position 6 with a dextrorotatory (D) amino acid and at position 10 with an ethylamide group produce superactive agonists, which are resistant to enzymatic degradation (Goren et al., 1990; Ulloa-Aguirre and Timossi, 2000), thus being cleared from circulation much slower than native GnRHs (Gothilf and Zohar, 1991; Haffray et al., 2005). GnRHas remain in the blood for a longer time than native hormones and stimulate more intensive release of LH from the brain. In addition, based on the modification of polarity and tertiary structure, some of GnRHas show higher affinity to the pituitary GnRH receptors (Habibi et al., 1989; Pagelson and Zohar, 1992). Owing to a large resistance to enzymatic decomposition and high receptor affinity, the gonadotropinreleasing hormone agonists (GnRHas) activity can be increased at 30–100 times in comparison to the activity of native GnRHs connecting with the induction of LH release (Peter et al., 1988; Zohar et al., 1989; Crim and Bettles, 1997). The synthetic analogs of mammalian (mGnRHa) and salmonid (sGnRHa) hormones are widely used in the fish culture practice (Mylonas and Zohar, 2007). The investigation of the mechanisms of hypothalamic neurosecretory control of fish reproduction shows that gonadotropin releasing inhibition factor (dopamine) is located in the hypothalamus (Chang and Peter, 1983; Barannikova, 1984; Godukhin and Motlokh, 1992; Glubokov, 1993; Barannikova et al., 2005). This factor inhibits the processes of synthesis and release of gonadotropins from hypophysis. Therefore, the substances preventing the release of dopamine or antagonists of dopamine are used together with releasing hormones in the practice of fish culture (Lam, 1982; Epler et al., 1989; Poponov et al., 1990; Glubokov et al., 1991; Glubokov, 1993). At the initial step, the dopamine antagonists (DA) were applied during hormonal therapy of freshwater fishes. In these species, dopamine inhibits the basal release of LH and reduces or inhibits GnRH-induced LH release (Peter and Yu, 1997). The use of dopamine antagonists (e.g., domperidone, pimozide, reserpine, metoclopramide, etc.) prevents inhibiting action of gonadotropocytes of the hypophysis and increases stimulating effect of GnRHa on LH release. The data on species specific dopaminergic regulation of gonadotropocytes in fishes can be found. For example, a role of dopamine in the regulation of maturation and ovulation is substantially lower in salmonids than in cyp20

rinids (Van der Kraak et al., 1986). The dopaminergic sensibility of oocytes of several fish species (in particular, rainbow trout Oncorhynchus mykiss =Parasalmo mykiss) decreases during the breeding season (Glubokov et al., 1991). Inhibiting role of dopamine is registered in freshwater fishes (e.g., in catfishes and cyprinids) (Saligaut et al., 1999; Silverstein et al., 1999; Yaron et al., 2003). In marine fishes, dopaminergic regulation is found in mullets (Mugil spp.) (Glubokov et al., 1994; Aizen et al., 2005), but, most likely, it is absent in the majority of valuable commercial fish species (Copeland and Thomas, 1989; King et al., 1994; Zohar et al., 1995; Mikodina and Strebkova, 1998; Prat et al., 2001; Kumakura et al., 2003). At present, hormonal manipulations with the use of GnRHa/DA combination are applied mainly for cyprinids (Yaron, 1995; Mikolajczyk et al., 2003, 2004), catfishes (Silverstein et al., 1999; Brzuska, 2001; Wen and Lin, 2004), and mullets (Glubokov et al., 1994; Aizen et al., 2005). Today both GnRHas and GtHs are used extensively in spawning induction therapies. One of the main advantages of the use of GtH preparations is that they act directly at the level of the gonads and can be effective even if pituitary LH stores are low, or the pituitary gonadotrophs are not responsive to GnRHa. In such situations, GnRHa may not be effective at all or may require a long time to elicit a response. However, the major drawbacks of the use of hypophysation are (1) the potential for transmission of diseases from donor to recipient fish and (2) the variation of LH content in donor pituitaries. The latter may vary according to body weight, sex, and age of donor fish, the time of year the pituitaries were collected, and the period of time elapsed from the death of the fish to the collection and preservation of the pituitary (Yaron, 1995). Nevertheless, hypophysation as a method to induce oocyte maturation, ovulation, and spawning is still employed in freshwater aquaculture, especially in developing countries. During two last decades, implantable sustained-release delivery systems for GnRHas are widely used for the control of reproductive processes in cultured fishes (see reviews by Zohar, 1996; Mylonas and Zohar, 2001, 2007). The interest in developing GnRHa-delivery systems stems from both the reproductive physiology of fish and the need to optimize broodstock management practices. For example, although GnRHas resist enzymatic degradation in the blood circulation compared to the native peptides, their maximum half-life does not exceed 23 min in vivo (Gothilf and Zohar, 1991). Such a short time of the circulation is a reason of the absence of ovulation (in some cases) after injections of GnRHas, especially after single injections (Mikolajczyk et al., 2003; Kaminski et al., 2004). In addition, single or double GnRHa injections, most often, induce ovulation of 21

only the first oocyte batch in the species with multiple spawning (Carrillo et al., 1995; Zohar and Mylonas, 2001; Levavi-Sivan et al., 2004), and additional injections are required to obtain subsequent batches (Pankhurst et al., 1986; Mylonas et al., 2003). The repeated injections are accompanied by additional time and labor work; additional handling can lead to damage of the fish and has a negative effect on oocyte maturation (Pankhurst and Van Der Kraak, 1997). Therefore, the application of implantable sustained-release delivery systems for GnRHas has an advantage in comparison to traditional practice of hormonal injections. In some species (e.g., in group-synchronous multiple-spawning shi drum Umbrina cirrosa), both injections of GnRHa and GnRHa-delivery systems can be used, and the former method can be even more suitable. Multiple cycles of oocyte maturation in this species can be induced only after gradual injections (Mylonas et al., 2004) that can be connected with discontinuous type of oogenesis. The interval between subsequent spawning reaches approximately seven days (Mylonas et al., 2003). The completion of vitellogenesis and recruitment of new oocyte generations can be associated with a low LH level in plasma, but oocyte maturation is accompanied by abrupt increase of this level, as it is described in other fish species (Peter and Yu, 1997). Thus, the application of GnRHas injections in fish culture will be continued due to comparative simplicity and a low price of this method in comparison to the use of GnRHa-delivery systems. Artificial stimulation of oocyte maturation and ovulation can be accompanied by the obtaining of eggs of a low quality due to different reasons, including overdoses of hormonal preparations (Makeyeva et al., 1987; Lahnsteiner and Patarnello, 2003). Therefore, optimal doses of hormones should be empirically determined for each species. The differences in the quality of eggs obtained with the use of different hormones are not revealed (Chen, 2005). Therefore, the main criteria for the application of a method of hormonal therapy are the number of females reacting to the hormonal action and degree of synchronization of ovulation in certain individuals (Wen and Lin, 2004).

2.2. Schemes of hormonal stimulation and morphological changes of oocytes in coral reef fish species 2.2.1. Zebrasoma scopas 2.2.1.1. Biological parameters and external morphology of the gonads In total, 445 females and males of Z. scopas were used in the experiments. A part of the fishes were subjected to biological analysis. The following param22

Table 1. Biological parameters of Zebrasoma scopas Sex (number of exemplars) Females (26) Males (13)

Length, cm

Weight, g

TL 10.0–15.5 13.2 ± 1.5

SL 8.3–13.6 11.3 ± 1.2

G 26.0–99.0 60.0 ± 17.6

10.5–16.5 14.1 ± 1.6

9.3–15.0 12.3 ± 1.4

31.0–110.0 75.4 ± 21.4

G1

GSI, %

24.0–88.0 53.4 ± 16.5

g 0.05–5.35 1.41 ± 1.12

0.17–9.66 2.69 ± 2.21

29.0–100.0 69.3 ± 20.2

0.04–0.48 0.24 ± 0.14

0.14–0.76 0.34 ± 0.22

Note: Above the line, range of the data; below the line, mean and standard deviation. GSI = g/G1 × 100, where g – gonad weight and G1 – weight of gutted fish.

eters was determined: total length (TL), standard length (SL), total body weight (G), weight of the gutted fish (G1), and gonad weight (g) (Table 1). In the male, the urogenital opening is located between the pelvic and anal fins (Color Plates, Fig. 3a). In the female, the genital opening is located in the same area (Color Plates, Fig. 3b). Before the completion of oocyte maturation, the genital opening of the female has the size similar to the size of the urogenital opening of the male, or the former is slightly larger. Therefore, owing to the absence of other secondary sex characters, the sex of the fish is difficult to indicate. Before ovulation, the genital opening of the female increases, and the genital area becomes convex (Color Plates, Fig. 3c). Such females can be easily discriminated from the males. Before ovulation, the genital opening is covered by a narrow membrane, which is disrupted just before release of oocytes; and a new membrane appears again after that. The gonads are represented by the short paired ovaries fused at their caudal parts approximately at one-third of their lengths; their coloration, more often, is yellow (Color Plates, Fig. 4a). The testes are also fused in their caudal parts (Color Plates, Fig. 4b). 2.2.1.2. Double injections The spawners of Z. scopas kept in the laboratory for several days do not mature without hormonal stimulation, and the processes of oocyte resorption are observed in the oocytes of older generation. The hormonal preparation used for stimulation of oocyte maturation and ovulation was represented by LH-RH-a, in particular, surfagon (des Gly10(D-Ala)6LHRH) produced in Russia. For this and other fish species, the preparations were injected into the abdominal cavity of the fish under the pectoral fin. In preliminary experiments, surfagon was injected together with (neuroleptic) eglonil (including sulpiride, a blocker of receptors of dopamine). In double injections, ovulated oocytes were obtained using different summing 23

doses of surfagon (from 30 to 80 µg/kg of body weight), and the application of eglonil had no effect on ovulation. Therefore, eglonil was not used in subsequent experiments conducted on Z. scopas. To take into account these data, the schemes of hormonal stimulation were developed according to the following directions: (1) determination of the effective and lowest doses of surfagon; (2) assessment of the effect of the interval between injections I and II on the time of oocyte maturation and ovulation; and (3) assessment of a possibility of the application of single injections for hormonal stimulation. During subsequent experiments, the minimum doses of surfagon were equal to 5 and 15–20 µg/kg for the injections I and II, respectively. Based on positive results obtained in these experiments, the doses applied in other experiments were substantially lower: 2 + 8 µg/kg. The fishes were observed just after the transfer to the laboratory, and the exemplars preliminary indicated as males were separated from the females (based on the slight differences in morphology of the genital area). The females were kept separately from the males, and some of the females were tagged individually by means of cutting of certain rays of the dorsal and anal fins. In the tagged fishes, oocytes were obtained at different time intervals by the method of biopsy. In many females, “natural” ovulation (as a result of oocyte maturation in the sea) was registered just after their transfer to the laboratory. In such cases, oocytes were stripped from the body, and the females were used for subsequent hormonal injections. “Natural” ovulation, as a rule, was observed just after the transfer of the fish to the laboratory, and only in several cases in 3 or even 8 h. The preliminary experiments showed that in the males subjected to hormonal stimulation spermiation was observed rarely; it was registered in several males after injection II. However, sperm obtained from crushed testes was successfully activated by marine water (Emel’yanova et al., 2006); thus, the males were not injected in subsequent experiments. To determine onset of ovulation following by hormonal injections, all females were periodically observed in 7–10 h after injection II. Just before ovulation, (as it was mentioned above) a convex light-yellow membrane was seen in the genital area of the female. After a pressure on the abdomen, the membrane prevented the release of ovulated oocytes. Therefore, the membrane was carefully pricked by a needle, and oocytes were stripped. This operation, most likely, had no effect on the maturation of subsequent oocyte batches: the membrane regenerated in several hours. The injection I was applied, most often, in the day of the transfer of the fish to the laboratory (in 2–9 h), and, in several cases, on the following day (in 24

18 h). The interval between injections I and II ranged between 12 and 24 h, and it was equal to 6 h in only one experiment. The results of the experiments with low doses of surfagon (2 + 8 µg/kg) are represented in Figs. 1a–1e. Ovulation is registered in almost all fishes subjected to hormonal injections. In the majority of individuals (approximately 80%), the total duration of the interval between injection I and ovulation reaches 33–39 h (most often, 36 h). In some females, this interval is lower reaching 25–30 h (Fig. 1c) or larger (up to 42 h). The difference, most likely, is connected with different endocrine sensibility of certain individuals. The interval from “natural” to induced ovulation ranges widely: from 26 to 52 h reaching 41–42 h in the majority of females. Summing dose of surfagon (within the range applied in this study) has no effect on the duration of the interval between injection I and ovulation. The increase of the interval between injections I and II leads to decrease of the interval between injection II and ovulation. It is especially pronounced if the intervals between injections I and II differ at two times reaching 12 and 24 h. In this case, the intervals between injection II and ovulation are 24 and 12 h, respectively (Figs. 1a and 1b). Thus, the total duration of the interval between injection I and ovulation remains the same (36 h). The interval from the transfer of the fish to the laboratory to the beginning of hormonal injections can be increased at least to 18 h: oocyte maturation and ovulation are observed (in one case) in 34 h after injection I (Fig. 1d) or (in another case) in 27 h after injection I. As it is mentioned above, at the intervals between injections I and II 12 and 24 h, summing time from injection I and ovulation remains the same. Therefore, the effect of lower intervals between injections I and II on oocyte maturation and ovulation was tested. In the group of the females (n = 5) with the interval between injections I and II 6 h, induced ovulation is registered only in one female in 43 h after injection I (Fig. 1e). By this time, maturation of the oocytes of older generations was not completed in other females. However, in the control group of the females with the interval between the injections 12 h, ovulation is registered in 36 h after injection I. Thus, the interval between the injections reaching less than 12 h can not be recommended for egg production. It is important to note, that periodical sampling of oocytes (biopsy) from the genital opening, as a rule, has no effect on oocyte maturation and ovulation and duration of these processes. Batch fecundity (actual fecundity after the stripping of oocytes) in the females of different size ranges from 1160 to 28 793 oocytes (n = 14). The lowest values of fecundity (1160 and 2050 oocytes) are registered for the 25

Synchronous ovulation Asynchronous ovulation

20

10

0

n=7

II

I

30

50

40

60

70

(a) I 20

10

0

n=8

II 30

50

40

60

70

(b) n=9 I 0

II 20

10

30

50

40

60

70

(c) I 0

10

n = 11

II 20

30

40

50

60

70

(d) I 0

n=5

II 10

20

30

40

50

60

70

(e) Fig. 1. Schemes of hormonal stimulation of Zebrasoma scopas: double injections of surfagon (injection I, 2 µg/kg + injection II, 8 µg/kg). (a) Interval between injections I and II 12 h; (b) interval between injections I and II 24 h (in both cases, ovulation is observed in 36 h after injection I); (c) ovulation is observed in 25–30 h after injection I; (d) injection I is applied in 18 h after the transfer of the fish to the laboratory; (e) interval between injections I and II 6 h (ovulation is observed in a female in 43 h after injection I). Arrows in the beginning of the scheme denotes “natural” ovulation in the fish (as a result of oocyte maturation in nature) after their transfer to the laboratory. The time before injection I is calculated from the transfer of the fish to the laboratory. n – number of females. Схемы гормональной стимуляции Zebrasoma scopas: двукратные инъекции сурфагона (инъекция I, 2 мкг/кг + инъекция II, 8 мкг/кг). (а) Интервал между инъекциями I и II 12 ч; (b) интервал между инъекциями I и II 24 ч (овуляция в обоих случаях через 36 ч после инъекции I); (c) овуляция через 25–30 ч после инъекции I; (d) инъекция I через 18 ч после доставки рыб в лабораторию; (e) интервал между инъекциями I и II 6 ч (овуляция лишь у одной самки через 43 ч после инъекции I). Стрелка в начале схемы обозначает «естественную» овуляцию (в результате созревания ооцитов в природе) после доставки рыб в лабораторию. Время до инъекции I отсчитано от доставки рыб в лабораторию. n – число самок.

26

Oocyte maturation

n = 10

I 10

0

20

30

40

50

60

70

(a)

n=9

I 0

10

20

30

40

50

60

70

(b)

Fig. 2. Schemes of hormonal stimulation of Zebrasoma scopas: single injections of surfagon with the doses (a) 20 µg/kg and (b) 10 µg/kg. Other designations as in Fig. 1. Схемы гормональной стимуляции Zebrasoma scopas: однократные инъекции сурфагона дозами (a) 20 мкг/кг и (b) 10 мкг/кг. Остальные обозначения как на рис. 1.

females 32 and 40 g in weight. In the females with the average body weight 71 g, the average fecundity reaches 12 912 oocytes. 2.2.1.3. Single injections The single injections with the doses of surfagon equal to the summing doses applied at the double injections (20–25 or 10 µg/kg) do not lead to oocyte ovulation (Figs. 2a and 2b). The females were checked in 20–30 h after the injection with a periodicity 1.5–2 h. Oocyte maturation comes to the end, and hydrated transparent cells with a lipid droplet are seen (see section 2.2.1.5). The development of oocytes does not depend on the presence or absence of “natural” ovulation in the females, but it is induced by the hormonal stimulation. The interval from the injection to the completion of oocyte maturation is approximately similar to the interval from injection I to the completion of oocyte ovulation at the double injections. At the single injections with the doses of surfagon 4 and 2 µg/kg similar to the doses applied for injection I at the double injections, the completion of oocyte maturation is not registered. Following by the injections with these doses, maximum oocyte diameter reaches 520 and 450 µm, respectively. 2.2.1.4. Size composition of oocytes Competent application of hormonal injections is impossible without the control of oocyte development. This information can be obtained during the analysis of oocyte diameter distributions and oocyte morphology. 27

To study size composition of oocytes and their morphological features during maturation, oocyte samples were obtained in vivo by means of the probe (Color Plates, Fig. 1e). Oocytes were sampled before injections I and II, as well as with the intervals 3 or 4 h from injection I to ovulation. The analysis of oocyte size composition was conducted with the image analyzing system including Nikon Eclipse E-200 microscope, analogous black and white video camera, and a computer. The objects were measured using ImageJ software. Morphology of oocytes obtained by the method of biopsy during the period of maturation was assessed after their treatment in Serra solution (Makeyeva and Emel’yanova, 1990) (for opaque oocytes) or in physiological solution (for transparent oocytes). The oocyte diameter distribution in the females shows the dominance of the cells of the lowest size and a presence of cells of intermediate size between the cells of the minimum and maximum diameter indicating the occurrence of continuous type of oogenesis and indeterminate fecundity (Götting, 1961; Oven, 1976). In intact females (before injection I), oocytes of older generation, most often, possess diameter approximately 350 µm (from 300 to 400 µm) (Fig. 3a). Such oocytes have completed the period of vitellogenesis, they possess maturational competence (ability to respond to hormonal therapy), and they are able to ovulation after hormonal stimulation. The intact individuals with larger oocytes up to 500 µm in diameter (at the period of maturation) are registered rarely. Before injection II (sometimes independently on the interval between injections I and II ranging from 12 to 24 h), oocyte diameter of the older age group can be increased. These oocytes reach 450 µm in diameter (Fig. 3b), rarely 400, 500, or even 550 µm in diameter. After induced ovulation, the size composition of oocytes is similar to that in intact females. A number of residual oocytes, which are not subjected to ovulation due to various reasons, are found in the ovaries. The size of such cells is substantially larger than the size of other oocytes (Fig. 3c). Diameter of ovulated oocytes ranges from 569 to 680 µm, on average 632 µm (n = 322). The assessment of size composition of oocytes from different females with a periodicity of 4 h shows that diameter of oocytes of the largest size group can be increased slightly or even insignificantly before injection II. This feature is observed at different intervals (from 12 to 24 h) between injections I and II. For example, oocytes of the largest size group obtained from a female before injection II (in 12 h after injection I) possess diameter 350– 400 µm, which is similar to that in intact individuals (Fig. 4). Significant increase of oocyte diameter is observed in several hours (in this female, in 12 h) after injection II. Sometimes the largest oocytes are separated from smaller cells on the histograms. In this female, such separation is registered 28

from 28 h after injection I to ovulation. Onset of ovulation is registered in 32 h after injection I, and ovulation is completed in 36 h. The size composition of all oocytes with maturational competence (before injection II) is compared in the females with different intervals between injections I and II. As it is mentioned above, the interval between injection I and ovulation is similar in the majority of individuals independently on the interval between injections I and II. Therefore, it can be suggested, that both oocyte size composition and the states of oocytes depend on the inter60 50 40 30 20 10 0

Number of oocytes, %

50 40 30 20 10 0 50 40 30 20 10 0 50

150

250

350

450

Oocyte diameter, µm

Fig. 3. Oocyte diameter frequency distribution in the ovary of a female of Zebrasoma scopas: (a) before injection I; (b) before injection II; (c) after egg ovulation. Распределение ооцитов по диаметру в яичнике Zebrasoma scopas: (a) перед инъекцией I; (b) перед инъекцией II; (c) после овуляции.

29

val between the injections. Oocyte compositions are compared in two groups of females (four individuals in each) with the intervals between the injections 15 and 22 h. Oocytes 400–450 µm in diameter appear before injec70

70

60

60

30 20

10

10

60

50

65 0

55 0

45 0

35 0

25 0

70

15 0

0

70

50

0

65 0

55 0

n = 158

65 0

40

20

60

8h

50 40

n = 225

40 30

20

20

10

10

65 0

55 0

45 0

35 0

70

25 0

0

70

15 0

0 50

28 h

50

30

50

150

250

n = 193

350

450

550

650

60

20

10

10

16 h

n = 236

50

65 0

55 0

45 0

35 0

25 0

70

15 0

0

70

50

0

60

36 h

50

40

40

30

30

20

20

10

10

0

65 0

30

55 0

40

20

60

n = 245 32 h Onset of ovulation

50

45 0

30

n = 280

35 0

40

15 0

12 h Injection II

50

25 0

60

50

24 h

50

30

45 0

50

65 0

60

55 0

40

n = 291

45 0

4h

50

55 0

45 0

35 0

25 0

15 0

70

50

0

70

35 0

10

0

25 0

20

10

35 0

30

20

60

Number of oocytes, %

40

25 0

30

n = 217

20 h

50

15 0

40

n = 204

15 0

0h Injection I

50

n = 164

0 150 200 250 300 350 400 450 500 550 600 650 700 050 100

150 200 250 300 350 400 450 500 550 600 650 700 050 100

Oocyte diameter, µm

Fig. 4. Dynamics of maturation and ovulation of oocytes in a female of Zebrasoma scopas. The time is calculated from injection I. Динамика созревания и овуляции ооцитов у самки Zebrasoma scopas. Время отсчитано от инъекции I.

30

60

a

50

n = 1173

40

20 10

b

50

700

650

600

550

500

450

400

350

300

250

200

150

50

0 60

100

Number of oocytes, %

30

n = 872

40 30 20 10

600

700

650

500

600

550

400

500

450

300

400

350

200

300

250

100

200

150

50

0

100

0

700

Oocyte diameter, µm

Fig. 5. Oocyte diameter in the ovary of a female of Zebrasoma scopas before injection II: (a) 15 h from injection I, (b) 22 h from injection I. Диаметр ооцитов в яичнике самки Zebrasoma scopas перед инъекцией II: (a) 15 ч после инъекции I, (b) 22 ч после инъекции I.

tion II in two groups (Figs. 5a, 5b). Average oocyte diameters in all oocytes larger 350 µm (average diameter of oocytes completed vitellogenesis and characterized by maturational competence) in two groups are as follows: (1) 384 µm (SD 39, n = 71) and (2) 387 µm (SD 25, n = 92). The difference is not significant (p = 0.536). Similar results are obtained in the comparison of oocyte composition in two groups of females (two individuals in each) with the intervals between injections I and II 12 and 24 h. Average oocyte diameters in all oocytes larger 350 µm in two groups are as follows: (1) 419 µm (SD 40, n = 50) and (2) 417 µm (SD 59, n = 44). The difference is not significant (p = 0.427). Thus, the change of the interval between injections I and II does not lead to a substantial change of oocyte diameter before injection II. Nevertheless, increasing interval between the injections should lead to increase of the number of oocytes at later phases of maturation before injection II (see below). However, average diameter of oocytes remains the same because the number of oocytes at advanced developmental stages is low. Substantial increase of oocyte diameter is registered only after injection II. 31

2.2.1.5. Morphological changes of oocytes during maturation In intact females, oocytes of the size class 300–400 µm, on average, 350 µm (completed vitellogenesis and characterized by maturational competence) are represented by cells with the nucleus located in the center. Numerous small lipid droplets surround the nucleus representing a wide ring (Fig. 6a). Following by injection I, migration of the nucleus to the animal pole begins, and lipid droplets begin to fuse. Just before injection II (in the majority of oocytes with maturational competence), the nucleus is located at a comparatively large distance from the animal pole. Lipid droplets become slightly larger, and they are located around the nucleus representing a narrower ring (Figs. 6b–6d). In addition, oocytes including substantially enlarged lipid droplets (from several to two droplets or even only one droplet, as a result of fusion of the droplets) together with smaller lipid droplets can be found. Thus, morphological development of oocytes occurs asynchronously. This asynchronous oocyte development, most often, is observed in the females, which already possess oocytes at the period of maturation during collection of the fish in the sea. After injection II, fusion of lipid droplets continue in the majority of oocytes with maturational competence (Figs. 6e, 6f, 7); and, finally, only one lipid droplet appears (Fig. 6g). The nucleus is seen near the lipid droplet at a distance from the animal pole. Following by the appearance of only one lipid droplet, intensive process of homogenization (fusion) of yolk granules is observed. However, the majority of yolk granules are still present, and they are responsible for a dark coloration of the oocyte in transmitted light (Fig. 6h). In such oocytes, the nucleus is not seen. Oocyte diameter increases due to hydration of the cell (Figs. 6h–6j). The most intensive increase of oocyte diameter is registered during homogenization of yolk accompanied by active hydration (Figs. 6i, 6j, 7b, 7c). Just before ovulation, oocytes become transparent (Fig. 6k). Diameter of ovulated oocytes is almost twice as large as diameter of oocytes in the end of vitellogenesis. A single injection of surfagon with a dose 10 µg/kg leads to the completion of maturation. If the dose reaches 2 µg/kg, maturation goes to the stage of the appearance of only one lipid droplet. Fusion of yolk granules does not occur or it is at the initial step, and oocytes are totally opaque. Maximum diameter of the cells does not exceed 450 µm (on average 416 µm) even in 36 h after the injection when ovulation is registered in the control group of the fish subjected to double injections. A single injection of surfagon with a dose 4 µg/kg leads to initial homogenization of yolk. Single largest oocytes with semitransparent cytoplasm can be seen in 36 h after the injection, but oocytes with only one lipid 32

Fig. 6. Morphology of oocytes of Zebrasoma scopas: (a) before injection I; (b, c, d) before injection II; (e–j) after injection II; (k) ovulated oocyte. (a–g) Oocytes are treated in Serra solution; (h–k) oocytes in vivo. 1, nucleus; 2, lipid droplet; 3, fusing yolk granules; 4, cortical alveole; 5, micropyle. Морфология ооцитов Zebrasoma scopas: (a) перед инъекцией I; (b, c, d) перед инъекцией II; (e–j) после инъекции II; (k) овулировавший ооцит. (a–g) Ооциты просветлены в жидкости Серра; (h–k) ооциты in vivo. 1, ядро; 2, жировая капля; 3, сливающиеся желточные гранулы; 4, кортикальная альвеола; 5, микропиле.

33

Fig. 7. Fragments of oocytes of Zebrasoma scopas after injection II: (a) less than ten yolk droplets; (b, c) fusion of yolk granules and hydration of oocytes. (a) Oocytes are treated in Serra solution; (b, c) oocytes in vivo. Фрагменты ооцитов Zebrasoma scopas после инъекции II: (a) менее 10 жировых капель; (b, c) слияние желточных гранул и гидратация ооцитов. (a) Ооциты просветлены в жидкости Серра; (b, c) ооциты in vivo.

droplet and opaque cytoplasm are the most numerous. The destruction of these oocytes is observed in 43 h after the injection. Thus, the minimum dose of surfagon suitable for the completion of maturation should exceed 4 µg/kg. Small individuals 10.0–11.0 cm TL and approximately 30 g in weight, most likely, mature for the first time. They possess a small number of oocytes with maturational competence. Following by hormonal injections, these oocytes complete maturation, and some of them are able to ovulation. The diameter of ovulated oocytes (~ 450 µm) is lower than that in repeat spawners. The number of oocytes in the beginning of vitellogenesis is also comparatively small. For histological analysis, the ovarian fragments were fixed in Bouin’s solution. Subsequent treatment of the material was conducted according to generally accepted method (Roskin and Levinson, 1957). The histological sections of the ovarian fragments of intact females (before hormonal stimulation) show the cells at different phases of the periods of vitellogenesis and previtellogenesis that is usual for the fishes with continuous type of oogenesis. The oocytes in the beginning of the maturation period are rarely seen. The oocytes completed vitellogenesis with maturational competence are characterized by a presence of the nucleus located in the center of the cell surrounded by a layer of small lipid droplets (Color Plates, Fig. 5a). A 34

layer of cortical alveoli is very narrow, and it is almost not detected in the peripheral zone of the cytoplasm. Following a certain time after injection II, the state of the largest oocytes has changed. The oocytes with the nucleus located at the animal pole (Color Plates, Fig. 5b) are registered extremely rarely. Such oocytes are never seen on the cells in vivo obtained by the method of biopsy even after their treatment in Serra solution. This feature can be connected with a short time of the occurrence of the nucleus at the animal pole. A part of oocytes of older generation can possess only one lipid droplet located in the central part of the cell near the nucleus (Color Plates, Fig. 5c). In these oocytes, intensive process of fusion of yolk granules is observed. This process is accompanied by hydration of the cell leading to homogenization of yolk. After ovulation, empty follicles are seen in the ovary, and the majority of oocytes of older generation are represented by the cells completed vitellogenesis (as in the ovaries of intact individuals) (Color Plates, Fig. 5d). In intact females kept in the tanks for several days without feeding, oocyte resorption begins rapidly: resorption of the largest oocytes is registered in 3– 6 days after the transfer of the fish to the laboratory (Color Plates, Fig. 5e). During the spawning season, numerous cells at different developmental stages, including spermatozoa, are found in the testes of the males (Color Plates, Fig. 5f). 2.2.1.6 Temporal variation of oocyte maturation As it is mentioned above, the interval between injection I and ovulation reaches approximately 36 h in the majority of the females, but it can be different in certain individuals. Therefore, the rate of morphological changes in oocytes can be also different. The intervals between injections I and II can be changed as well. To represent the chronology of oocyte maturation, the interval between injection I and ovulation (independently on its absolute duration) is accepted as 100% (Table 2). The absolute values (in hours) of onset of morphological changes are given in the text below. These values are obtained for the variant with the interval between injections I and II reaching 15 h; ovulation is registered in 30 h after injection I. The most notable morphological changes are registered in ocytes after injection II. A representative morphological criterion of the changes is the fusion of lipid droplets. In the majority of maturing oocytes, fusion of numerous lipid droplets with the appearance of several (≤ 10) large droplets is observed in 1–2 h after injection II. As a rule, single oocytes with only one lipid droplet appear at the same time. A large number of oocytes with only one lipid droplet appear, most often, in 3 h after injection II (in 18 h 35

Table 2. Chronology of the processes of oocyte maturation (in percents of the interval from injection I to ovulation) and diameter of oocytes in Zebrasoma scopas

Morphological features of the oocyte Less than ten lipid droplets Only one lipid droplet Onset of homogenization of yolk Completion of homogenization of yolk Appearance of ovulated oocytes

Temporal parameters of the changes, % mean lim 50 36–69 60 42–75 70 64–78 80 – 90 –

Oocyte diameter, µm mean 371 441 480 547 632

lim 342–401 365–515 404–557 500–586 569–680

n 82 71 95 80 322

after injection I). Single oocytes with this morphology can appear before injection II both earlier (in 12 h (33%) or 15 h (50%) after injection I) or later (in 24 h (67%) after injection I). (Injection II is conducted in 12, 15, and 24 h after injection I, respectively). In 6 h after injection II (in 21 h after injection I), homogenization of yolk granules begins, but the oocytes are opaque, and only small areas of the oocyte become transparent. In 9 h after injection II (in 24 h after injection I; ~ 80%), the oocytes become substantially more transparent, and their diameter increases. In 27 h after injection I (~ 90% in various variants), transparent oocytes, which are able to ovulation, are observed. Mass ovulation, as a rule, is registered in 3 h after the appearance of first ovulated oocytes. As it is mentioned above, the increase of the interval between injections I and II leads to shortening of the interval between injection II and ovulation. After injection I (with a dose of surfagon 2 µg/kg), morphological development of oocytes can continue until the appearance of oocytes with only one lipid droplet and opaque cytoplasm. During a longer interval between injections I and II, the oocytes at this phase of development, most likely, become more numerous, that leads to shortening of the interval between injection II and ovulation. The increase of the number of these oocytes is not accompanied by larger average oocyte diameter: the increase of oocyte diameter is insignificant until onset of yolk homogenization. 2.2.2. Dascyllus trimaculatus 2.2.2.1. Biological parameters and external morphology of gonads In total, 327 individuals were used for the experiments on hormonal stimulation. A part of the fish was subjected to biological analysis (Table 3). The ovaries of adult exemplars of D. trimaculatus are paired and fused in their caudal parts; they possess a light yellow coloration (Color Plates, Fig. 4c). The testes are also paired and fused in their caudal parts (Color Plates, 36

Table 3. Biological parameters of Dascyllus trimaculatus (designations as in Table 1) Sex (number of exemplars) Females (41) Males (24)

Length, cm TL SL 8.5–13.0 7.5–11.0 10.5 ± 1.5 8.9 ± 1.1 8.5–13.0 7.5–11.2 10.5 ± 1.6 9.3 ± 1.5

G 20.8–80.0 40.1 ± 21.5 17.6–86.0 38.6 ± 16.2

Weight, g G1 20.0–68.0 36.4 ± 14.8 16.9–80.0 36.4 ± 17.9

g 0.09–3.95 1.00 ± 0.24 0.01–0.69 0.26 ± 0.13

GSI, % 0.37–8.27 2.48 ± 1.23 0.06–1.50 0.74 ± 0.48

Fig. 4d). A small papilla is located in the urogenital area of the individuals of both sexes. In intact spawners, the urogenital papilla of the male and genital papilla of the female are almost similar: the papilla of the female is only slightly shorter and wider than the papilla of the male. Several hours before ovulation, the papilla of the female substantially increases forming a small ovipositor approximately 3 mm in length. These females can be easily separated from the males. As in other representatives of the family Pomacentridae, the size of the papilla of D. trimaculatus, most likely, rapidly decreases after egg deposition. As it is known, the papilla of related species, D. aruanus, is protruding for 1 h after the spawning (Mizushima et al., 2000). 2.2.2.2. Double injections For hormonal injections, surfagon together with eglonil or only surfagon were applied. Based on the preliminary results, ovulated oocytes can be obtained after double injections of surfagon with eglonil. The summing doses of the preparations were 30–35 µg/kg and 25 mg/kg of body weight, respectively. Ovulation was registered in 40–43 h after injection I (in 25–28 h after injection II). Based on these data, lower doses of the preparations were used for subsequent experiments: 5 µg/kg of surfagon + 5 mg/kg of eglonil for injection I and 15 µg/kg of surfagon + 15 mg/kg of eglonil for injection II. The following treatments were applied: (1) injection of surfagon and eglonil during the day of the transfer of the fish to the laboratory; (2) injection of surfagon and eglonil on the following day from the transfer of the fish to the laboratory; and (3) injection of only surfagon in 2–4 h after the transfer of the fish to the laboratory. In some fishes, the oocyte samples were obtained by the method of biopsy for the control of the state of oocytes. Ovulated oocytes are obtained from a part of the females in each experiment after the injections of surfagon + eglonil both in the day of the transfer of the fish to the laboratory and on the following day (in 34.5 h after this transfer). The proportion of individuals with ovulated oocytes in different experiments does not exceed 50%. These results differ from 37

those described above for Z. scopas characterized by the proportion of fishes with ovulated oocytes reaching 100%. The difference is connected with the specific development of oocytes in D. trimaculatus: in a part of the fish, vitellogenesis is not completed in the oocytes of older generation, and, thus, they do not possess maturational competence (see below). The interval between injections I and II ranged from 12 to 17 h. In the variant represented in Fig. 8a, ovulation is registered in 33.5–40.0 h after injection I (in 17.5–24.0 h after injection II). In different experiments, maximum interval between injection I and ovulation reaches 43 h, and the most usual interval is 40 h. The interval between injection II and ovulation ranges from 17 to 28 h, most often, from 22 to 24 h. It is important to note, that ovulation is registered only in the females not subjected to biopsy. If oocytes are sampled in vivo, only maturation of oocytes is observed, and this maturation is completed approximately by the time of ovulation in other females from the same experiment (Fig. 8b). The absence of ovulation in mature oocytes, most likely, is connected with the damage of the genital papilla by the probe. The hormonal injections conducted on the following day (in more than 24 h) after the transfer of the fish to the laboratory show that the interval from injection I to ovulation is similar to that in the case when the fishes were injected just after their transfer (Fig. 8c). The application of surfagon without eglonil has no effect on the proportion of females with ovulated eggs. In several females, ovulation is registered in 38.5–42.0 h after injection I (in 21.5–30.0 h after injection II) (Figs. 8d, 8e). As in the injections of surfagon together with eglonil, maximum interval between injection I and ovulation reaches 43 h. A trend to the increase of the interval between injection II and ovulation at decreasing interval between injections I and II is registered. As it is mentioned above, similar pattern is observed in Z. scopas. The number of ovulated oocytes stripped from the females with body weight between 30 and 45 g (n = 10) ranges from 320 to 4325, on average, 2092. In three individuals with body weight 42, 50, and 60 g, the actual fecundity is substantially larger: 15 763, 20 138, and 15 068 oocytes, respectively. In several males, spermiation is registered after injection I or after injection II when both surfagon + eglonil or only surfagon are applied. Sperm of intact males obtained from crashed testes are activated in marine water. Therefore, in the majority of experiments, the males were separated from the females (based on a small difference in the shape of the papilla), and the males were not injected.

38

2.2.2.3. Size composition of oocytes The size composition of intact females (from which ovulated eggs are obtained as a result of hormonal stimulation) shows that two large groups of oocytes are present in the ovaries. The first group includes previtellogenous and vitellogenous oocytes 50–350 µm in diameter, and the second group is represented by vitellogenous oocytes 450–650 µm in diameter. In Ovulation (number of females, %) Final maturation (number of females, %) I 20

10

0

50%

II 30

n=8 50

40

60

70

(a) I 20

10

0

33%

II 30

n=6

50

40

60

(b) I 20

10

0

30

n=5 20%

II 50

40

70

60

70

(c) I

50%

II 20

10

0

30

n = 10 50

40

60

70

(d) I 0

(e)

10

n = 10

40%

II 20

30

40

50

60

70

Hours

Fig. 8. Schemes of hormonal stimulation of Dascyllus trimaculatus: (a–c) injections (I + II) of surfagon + eglonil; (d, e) injections (I + II) of surfagon; (b) the samples of oocytes are obtained during oocyte maturation by the method of biopsy. The time before injection I is calculated from the transfer of the fish to the laboratory. n – number of females. Proportions of females with ovulated (or matured) oocytes (in percents from the total number of females) are indicated. Схемы гормональной стимуляции Dascyllus trimaculatus: (a–c) инъекции (I + II) сурфагона + эглонила; (d, e) инъекции (I + II) сурфагона; (b) инъекции самкам, у которых брали пробы ооцитов методом биопсии. Время до инъекции I отсчитано от доставки рыб в лабораторию. n – число самок. Указаны доли самок с овулировавшими или только созревшими ооцитами (в процентах от общего числа самок).

39

40

n = 50

(a)

30 20 10 0 50

50

150

250

350

450

550

650

40

750

(b)

n = 35

30 20 10

Number of oocytes, %

0

40 50

150

250

350

450

550

650

n = 276

750

(c)

30 20 10 0 50

50

150

250

350

450

550

650

n = 128

40

750

(d)

30 20 10 0

30

50

150

250

350

450

550

650

n = 95

750

(e)

20

10

0 150 200 250 300 350 400 450 500 550 600 650 700 750 800 050 100

Oocyte diameter, µm

Fig. 9. Oocyte diameter in the samples of the ovaries of Dascyllus trimaculatus: (a–c) a female (1) with ovulated oocytes after the hormonal stimulation; (d, e) a female (2) with immature oocytes after the hormonal stimulation. Female 1: (a) before injection I; (b) ovulated oocytes; (c) after ovulation. Female 2: (d) before injection I; (e) after injection II at the time estimated for ovulation. n – total number of oocytes. Диаметр ооцитов в пробах яичников самoк Dascyllus trimaculatus: (a–c) самка (1) с ооцитами овулировавшими после гормональной стимуляции; (d, e) самка (2) с ооцитами несозревшими после гормональной стимуляции. Самка 1: (a) перед инъекцией I; (b) овулировавшие ооциты; (c) после овуляции. Самка 2: (d) перед инъекцией I; (e) после инъекции II в расчетное для наступления овуляции время. n – общее число ооцитов.

40

the latter group, the cells 500–600 µm in diameter prevail (Fig. 9a). A hiatus between the groups includes two size classes. It is obvious, that oocytes 450–650 µm in diameter possess maturational competence. The diameter of ovulated oocytes is 550–750 µm, and oocytes 600–700 µm in diameter prevail (Fig. 9b). After stripping of ovulated oocytes, vitellogenous oocytes up to 350 µm in diameter are found in the ovaries (Fig. 9c); sometimes, oocyte diameter can be slightly lower or larger. Following by injections I and II, ovulation is not registered if intact females possess oocytes less than 400 µm in diameter (Figs. 9d, 9e). In such cells, vitellogenesis is not completed, and they do not possess maturational competence. 2.2.2.4. Morphological changes of oocytes during maturation In intact females, oocytes of the largest size group are represented by the cells with the nucleus located in the center of the cell or slightly displaced to the animal pole. Numerous lipid droplets are distributed within a wide ring surrounding the nucleus (Figs. 10a, 10b). As a rule, the lipid droplets are slightly larger in the oocytes with displaced nucleus than in the cells with the nucleus in the center. Following by injection I, migration of the nucleus towards the animal pole begins (or continues), and fusion of lipid droplets is observed. Just before injection II, the nucleus is located at a comparatively large distance from the animal pole, lipid droplets become larger, and they are distributed within a narrower ring around the nucleus (Figs. 10c, 10d); several largest lipid droplets can be seen (Fig. 10d). During subsequent maturation, oocytes with one or two large lipid droplets and several small lipid droplets appear (Figs. 10e–10h). Fusion of lipid droplets continues during subsequent development of the cells (Fig. 10i). These events are accompanied by insignificant increase of oocyte diameter as a result of hydration. The yolk granules become substantially larger, most likely, as a result of their partial fusion or hydration. Ovulated oocytes, most often, possess only one lipid droplet, and they are comparatively transparent (Fig. 10j). At histological sections, the majority of oocytes of intact females are represented by the cells filled with yolk. The nucleus is located in the center of the cell, and it is surrounded by numerous small lipid droplets (Color Plates, Fig. 6a). The cells in the beginning of vitellogenesis, and previtellogenous oocytes are also seen. Oocytes in the beginning of vitellogenesis, approximately 100 µm in diameter, are characterized by the appearance of lipid droplets near the nucleus. In several hours after injection II, intensive fusion of lipid droplets is observed. By the appearance of two, three, or (rarely) one large lipid droplets (together with several small lipid droplets), the nucleus is well 41

expressed, and it is surrounded by lipid droplets (Color Plates, Fig. 6b). The localization of the nucleus at the animal pole is not registered. In ovulated oocytes, two or three large lipid droplets (instead of only one droplet) can be seen. In the ovary of the female fixed in two days after ovulation, previtellogenous oocytes, oocytes in the beginning of vitellogenesis, and residual oo2

1

a

b

c

d

e

f

g

h

i

j

1

2

Fig. 10. Maturation of oocytes of Dascyllus trimaculatus: (a, b) before injection I; (c, d) before injection II; (e–i) after injection II; (j) ovulated oocyte. (a–i) Oocytes are treated in Serra solution; (j) oocyte in vivo. 1, nucleus; 2, lipid droplet. Созревание ооцитов Dascyllus trimaculatus: (a, b) перед инъекцией I; (c, d) перед инъекцией II; (e–i) после инъекции II; (j) овулировавший ооцит. (a–i) Ооциты просветлены в жидкости Серра; (j) ооцит in vivo. 1, ядро; 2, жировая капля.

42

cytes not subjected to ovulation (filled with yolk and, sometimes, at the state of resorption) are seen (Color Plates, Fig. 6c). Empty follicles are not found at the sections as a result of their rapid resorption. During the spawning period, numerous spermatozoa together with cysts with sex cells at earlier developmental stages are seen in the testes of the males (Color Plates, Fig. 6d). 2.2.3. Abudefduf sexfasciatus 2.2.3.1. Biological parameters and external morphology of the gonads In total, 540 exemplars were used in the experiments, and some of them were subjected to biological analysis (Table 4). The paired gonads (both ovaries and testes) of A. sexfasciatus are fused in their caudal parts (Color plates, Figs 4e, 4f). Both ovaries and testes, most often, are asymmetrical. Sexual dimorphism is almost totally absent, but certain differences in the structure of the papilla located in the urogenital area can be observed. In the females with oocytes before the period of maturation or at the initial phase of this period, the genital papilla is weakly expressed, and its shape is similar to that of the urogenital papilla of the male. The papilla of such female is slightly shorter and wider than the papilla of the male. Before ovulation, the size of the genital papilla of the female is increased forming a small ovipositor approximately 5 mm in length. These females can be easily distinguished from the males. Table 4. Biological parameters of Abudefduf sexfasciatus (designations as in Table 1) Sex (number of exemplars) Females (25) Males (14)

Length, cm TL SL 9.5–17.0 8.5–14.0 13.0 ± 2.0 10.9 ± 1.6 10.1–15.0 8.5–12.5 12.1 ± 1.7 10.1 ± 1.9

G 22.0–114.0 53.9 ± 24.9 24.0–70.0 39.4 ± 17.3

Weight, g G1 20.5–108.0 49.4 ± 22.7 22.0–62.0 37.4 ± 15.9

g 0.30–7.96 2.01 ± 1.93 0.02–1.55 0.51 ± 0.43

GSI, % 0.88–11.71 3.75 ± 2.74 0.09–2.83 1.26 ± 0.73

2.2.3.2. Double and single injections Both double (5–7 + 15–20 µg/kg of body weight) and single (20 µg/kg) injections of surfagon without eglonil were applied. The interval between injections I and II ranged from 12 to 24 h. Three treatments were used: (1) double injections in the day of the transfer of the fish to the laboratory; (2) double injections on the following day from the transfer of the fish to the laboratory, and (3) single injections. In some individuals, oocytes were sampled by the method of biopsy. Several males were injected using the schemes applied for the females. However, the sperm from crashed testes could be activated in marine water (Pavlov and Emel’yanova, 2006), and, therefore, the majority of males were not injected. 43

As a result of hormonal injections conducted in several hours from the transfer of the fish to the laboratory (Figs. 11a–11e) or on the following day (Fig. 11g), ovulated oocytes are obtained (as in D. trimaculatus) only in a part of the females (less than 57%). It is connected with incomplete vitellogenesis in oocytes of older generation in a part of intact individuals. In the variants represented in Figs. 11a–11e, ovulation is observed in 40.0– 47.5 h after injection I (in 21.0–31.0 h after injection II). The range of the interval between injection I and ovulation can be larger (39.0–51.0 h), but most often it is equal to 42–43 h. It should be noted, that ovulation is registered in several females subjected to biopsy (Figs. 11d, 11e) that is different from the situation observed in D. trimaculatus. In some females of A. sexfasciatus subjected to biopsy, only maturation of oocytes (without ovulation) is registered; and oocyte maturation is completed approximately at the time of ovulation in other females (Fig. 11e). The absence of ovulation of matured oocytes (as in D. trimaculatus) is connected with the damage of the genital papilla as a result of insertion of the probe. The injections conducted in more than a day after the transfer of the fish to the laboratory show that maturation of oocytes is completed in a part of the fishes (Fig. 11f). In all these exemplars, the samples of oocytes have been obtained by biopsy. Therefore, ovulation, most likely, can be induced on the following day after the transfer of the fish to the laboratory. As in D. trimaculatus and Z. scopas, a trend towards the increase of the interval between injection II and ovulation at lower interval between injections I and II is registered. The single injections show that ovulated oocytes can be obtained in certain individuals. For example, ovulation is registered in 48 h after the injection (Fig. 11g). At single injections, the interval between injection I and ovulation can reach 64 h. The numbers of ovulated oocytes obtained from three females of A. sexfasciatus 73, 75, and 83 g in weight were 19 836, 12 800, and 15 330, respectively (on average, 15 989). In smaller females of D. trimaculatus (average body weight 50 g), average fecundity (16 990 oocytes) is higher (Emel’yanova et al., 2009a) that is connected with larger size of oocytes in A. sexfasciatus. In the males subjected to double and single injections of surfagon or surfagon + eglonil, spermiation is not registered. However, in a part of individuals of D. trimaculatus, spermiation is observed after double injection of the same preparations or even after only injection I (Emel’yanova et al., 2009a). The difference can be connected with the features of hormonal regulation of spermiation in the species. 44

Ovulation (number of females, %) Protracted ovulation (number of females, %) Final maturation (number of females, %) I

22%

II

0

10

20

30

40

n=9

50

70

60

(a) I 10

0

57%

II 20

30

40

50

n=7 70

60

(b) 36%

II

I 0

10

20

30

40

50

n=8 70

60

(c) I 10

0

50%

II 20

30

40

n=4

50

70

60

(d) I 0

n=2

II 10

20

30

40

50

70

60

(e) II

I 0

10

20

30

40

50

n=5 60

20% 70

(f) n=4

I 0

10

20

30

40

50

60

25% 70

(g) Fig. 11. Schemes of hormonal stimulation of Abudefduf sexfasciatus: (a–e) double injections (I + II) in the day of the transfer of the fish to the laboratory; (f) double injections on the following day after the transfer of the fish to the laboratory; (g) single ingection. (d, e, f) The samples of oocytes are obtained during oocyte maturation by the method of biopsy. The time before injection I is calculated from the transfer of the fish to the laboratory. n – number of females. Proportions of females with ovulated (or matured) oocytes (in percents from the total number of females) are indicated. Схемы гормональной стимуляции Abudefduf sexfasciatus: (a–e) двукратные инъекции (I + II) в день доставки рыб в лабораторию; (f) двукратные инъекции на следующий день после доставки рыб в лабораторию; (g) однократная инъекция. (d, e, f) Инъекции самкам, у которых брали пробы ооцитов методом биопсии. Время до инъекции I отсчитано от доставки рыб в лабораторию. n – число самок. Указаны доли самок с овулировавшими или только созревшими ооцитами (в процентах от общего числа самок).

45

2.2.3.3. Size composition of oocytes In intact females of A. sexfasciatus (from which ovulated oocytes are obtained later as a result of hormonal stimulation), large vitellogenous oocytes prevail, and a hiatus is observed between the larger and smaller cells (Figs. 12a, 12d) that is usual for the fishes with discontinuous oogenesis (Oven, 1976, 2004). If the largest oocytes of intact females reach 700–950 µm in diameter (less often, 600–800 µm in diameter with the modal group 700–800 µm in diameter) (Figs. 12a, 12d), they, most often, are subjected to maturation and ovulation after the hormonal stimulation. Following by injection II, a more distinct separation of larger oocytes (later subjected to ovulation) from smaller cells is observed (Fig. 12b). If the diameter of larger oocytes in intact females is less than 700 µm, they do not possess maturational competence (as a consequence of incomplete vitellogenesis), and ovulation is not induced after the hormonal stimulation (Figs. 12e, 12f). The diameter of ovulated oocytes ranges from 750 to 1100 µm, more often, from 900 to 1000 µm (Fig. 12c). After ovulation, the ovaries include mainly previtellogenous and small vitellogenous oocytes, as well as single residual oocytes (Fig. 12g). In some individuals, the majority of vitellogenous oocytes are less than 400 µm in diameter, but a small number of the cells up to 500 µm or even 700 µm in diameter can be observed. 2.2.3.4. Morphological changes of oocytes during maturation Morhological changes of oocytes of A. sexfasciatus are in general similar to those in D. trimaculatus. In intact females, oocytes of the larger size group are represented by oval cells with the nucleus located in the center of the cell or slightly displaced to the animal pole. Numerous lipid droplets are located around the nucleus representing a wide ring (Figs. 13a, 13b). As a rule, in the oocytes with displaced nucleus, lipid droplets are larger due to their partial fusion. Following by injection I, the processes of migration of the nucleus to the animal pole and fusion of lipid droplets begin (or continue). Just before injection II, the nucleus is located at a distance from the animal pole, and lipid droplets become larger locating within a comparatively narrow ring around the nucleus; the largest lipid droplet can be seen (Fig. 13c). Following by injection II, migration of the nucleus to the animal pole continues, and lipid droplets become larger as a result of their fusion (Figs. 13d, 13e). By the appearance of only one large lipid droplet, the nucleus is still seen (Fig. 13f), and it disappears later (Fig. 13g). These changes are associated with a slight increase of oocyte diameter as a result of hydration. Yolk granules become larger, especially at the vegetative pole, 46

30

40

(a)

(d)

n = 57

n = 28

30 20 20 10

50 40

0

0

0 95 95 0

85 85 0

0

0

n = 148

75

65

0

55

0

45

35

25

0

(f)

75 0

50

0

0

50

95

10

75

n = 48

85

0

0 65

0 55

0

0 45

(c)

35

0 25

15

0

0 30

50

0 60

65 0

10

55 0

10

n = 139

45 0

20

35 0

20

(e)

25 0

30

15

50

30

15 0

0

50

95

10

0

0 85

0

0

0

0

n = 44

75

65

55

45

(b)

35

0 25

15

0

0 40

Number of oocytes, % 50

0 40

0

10

20

30 20

10

10

0

95

0

85

0

75

0

65

0

55

0

0

45

35

25

0

0

100 200 300 400 500 600 700 800 900 1000

0

50

0

15

0

50

95

(g)

10

0

85

75

0

0

30

65

0

55

0

0

0

45

35

15

25

0

0 100 200 300 400 500 600 700 800 900 1000 1100

50

0

n = 137

20

10

Oocyte diameter, μm

95 0

85 0

75 0

65 0

55 0

45 0

35 0

25 0

15 0

0 100 200 300 400 500 600 700 800 900 1000

50

0

Fig. 12. Oocyte diameter in four females of Abudefduf sexfasciatus (females 1, 2, and 4 had ovulated oocytes after the hormonal stimulation; female 3 remained immature after the hormonal stimulation). Female 1: (a) before injection I; (b) before injection II; (c) ovulated oocytes. Female 2: (d) before injection I. Female 3: (e) before injection I; (f) after injection II at the time estimated for ovulation. Female 4: (g) after ovulation and stripping of ovulated oocytes. n – total number of oocytes. Диаметр ооцитов четырёх самок Abudefduf sexfasciatus (от самок 1, 2 и 4 овулировавшие ооциты получены после гормональной стимуляции; самка 3 не созрела после гормональной стимуляции). Самка 1: (a) перед инъекцией I; (b) перед инъекцией II; (c) овулировавшие ооциты. Самка 2: (d) перед инъекцией I. Самка 3: (e) перед инъекцией I; (f) после инъекции II в расчетное для наступления овуляции время. Самка 4: (g) после овуляции и отцеживания овулировавших ооцитов. n – общее число ооцитов.

47

most likely, as a result of their partial fusion and hydration. Ovulated oocytes are comparatively transparent, with (most often) only one lipid droplet located at the vegetative pole (Fig. 13h). A substantial hydration of oocytes during their maturation in A. sexfasciatus and D. trimaculatus is not registered due to the deposition of demersal eggs. The diameter of ovulated oocyte is approximately 100 µm larger than the diameter of the oocyte filled with yolk.

Fig. 13. Maturation of oocytes of Abudefduf sexfasciatus: (a, b) before injection I; (c) before injection II; (d–g) after injection II; (h) ovulated oocyte. (a–f) Oocytes are treated in Serra solution; (g, h) oocytes in vivo. 1, nucleus; 2, lipid droplet. Созревание ооцитов Abudefduf sexfasciatus: (a, b) перед инъекцией I; (c) перед инъекцией II; (d–g) после инъекции II; (h) овулировавший ооцит. (a–f) Ооциты, просветлённые в жидкости Серра; (g, h) ооциты in vivo. 1, ядро; 2, жировая капля.

48

At histological sections of ovarian fragments of intact females oocytes filled with yolk prevail. They are characterized by an oval shape and the nucleus locating in the center and surrounding by small lipid droplets and spherical yolk granules. The oocytes at earlier phases of the periods of vitellogenesis and previtellogenus are also seen (Color Plates, Fig. 7a). In the beginning of vitellogenesis, lipid droplets appear near the nucleus. The diameter of such cells is approximately 150 µm. In the ovaries of females after ovulation, empty follicles remain, but they are subjected to resorption very rapidly. Both previtellogenous oocytes and oocytes at initial phases of vitellogenesis are seen. In addition, not numerous residual oocytes (not subjected to ovulation) can be observed (Color Plates, Fig. 7b). In the oocyte filled with yolk, follicular cells reach the largest height at the animal pole and in the area adjacent to the animal pole (Color Plates, Fig. 7c) that is connected with the development of the chorion. In ovulated oocyte, filaments of the chorion are located only at the animal pole (see section 3.1.2.3). Thus, the substances of the filaments are synthesized at the animal pole and around the animal pole. During the spawning season, numerous spermatozoa are observed in the testes, and the cysts with cells at earlier developmental stages can be seen (Color Plates, Fig. 7d). 2.2.4. Conclusions Based on the results of the experiments on hormonal stimulation of oocyte maturation and ovulation in Z. scopas, D. trimaculatus, and A. sexfasciatus, double injections of surfagon both with and without dopamine antagonists (DA) can be applied. The latter scheme is more simple and feasible. As is known, the combination GnRHa/DA is applied, most often, for freshwater fishes (De Lee et al., 1988; Epler et al., 1989; Godukhin and Motlokh, 1992; Yaron, 1995; Brzuska, 2001; Mikolajczyk et al., 2004; Wen and Lin, 2004). The use of this combination is connected with the composition of GnRH including both liberins stimulating the action of gonadotropocytes of the hypophysis and dopamine, a static factor inhibiting this action (Rosen, 1994). However, induced maturation in such fishes as sturgeons (Acipenseridae), several salmonids (Salmonoidei), perch Perca fluviatilis, tench Tinca tinca, and silver carp Hypophthalmichthys molitrix is possible without DA (Goncharov, 1984, 1991; Erdahl and McClain, 1987; Kouril et al., 1991; Travkina and Vinogradova, 1991; Makeyeva et al., 1994). Such a possibility, most likely, is connected with a low level of efficiency 49

of dopaminergic regulation of gonadotropocytes of the hypophysis in these fishes (Van der Kraak et al., 1986; Moretti et al. 1999; Poortenaar and Pankhurst, 1999) or even with the absence of this regulation, which is registered in some marine fishes (Mylonas and Zohar, 2007). Summing doses of the preparation used for the injections to Z. scopas, D. trimaculatus, and A. sexfasciatus can range widely, but the application of low doses is more feasible. For hormonal stimulation of Z. scopas, the doses 2 µg/kg + 8 µg/kg (for injections I and II, respectively) of surfagon are optimal; lower doses lead to unstable results or to the absence of ovulation. For the representatives of the family Pomacentridae, the summing dose 20 µg/kg can be applied, but the lower doses have not been tested. The interval between the injections can be changed within a certain range. The increase of the interval between injections I and II leads to a shorter interval between injection II and ovulation, but the total interval from injection I to ovulation remains without substantial changes. This feature is observed in all three studied species (especially in Z. scopas). Thus, the total time to ovulation is connected mainly with the summing dose of the preparation and with the duration of its action. In Z. scopas, the occurrence of “natural” ovulation (induced by natural environment in the sea) registered just after the transfer of the fish to the laboratory has no effect on the incidence and time of occurrence of subsequent ovulation induced by hormonal stimulation. In A. sexfasciatus, ovulation is possible both at double and single injections of surfagon. In the latter case, a longer interval between injection I and ovulation is observed. A possibility of the application of single injections for D. trimaculatus from the same family (Pomacentridae) can be suggested. In Z. scopas, single injection leads to oocyte maturation without ovulation that is different from the situation described above in A. sexfasciatus. In the majority of fish species, induced ovulation is possible only after double injections, but single injections can be applied for several species (Barannikova et al., 1989; Epler and Bieniarz, 1989; Makeyeva et al., 1994). Thus, species specific endocrinological sensitivity is important for development of the schemes of hormonal stimulation. In three studied species, the differences in the interval between injection I and ovulation are small: most often, this interval reaches 36 h in Z. scopas with pelagic eggs and 40–43 h in two species of the family Pomacentridae with demersal eggs. Ovulated oocytes in two latter species can be obtained on the following day after the transfer of the fish to the laboratory. As is known, hormonal therapy can be applied for the tropical fish species only within one or two days after their transportation from natural environment; 50

and the processes of oocyte resorption are registered afterwards (Colin et al., 2003). According to the classification of Götting (1961), two main types of oogenesis (discontinuous and continuous) can be distinguished in fishes. In the fishes with discontinuous oogenesis, vitellogenous oocytes are separated from the reserve fund during the spawning season; in the fishes with continuous oogenesis, a gradual transition between oocytes of all developmental phases is registered. However, vitellogenous oocytes are not separated from oocytes of the reserve fund before the spawning season or during the largest part of the spawning season in the majority of fishes from the seas of low latitudes (Oven, 1976, 2004). Based on asynchronous growth of oocytes in these fishes, which is more pronounced during vitellogenesis, some authors suggest more detailed schemes of oogenesis (Lisovenko, 1985, Alekseev and Alekseeva, 1996). In particular, the species with discontinuous-asynchronous vitellogenesis are separated from the representatives with continuous oogenesis (Alekseev and Alekseeva, 1996). According to discontinuous-asynchronous vitellogenesis, oocytes of the fund, which is used during coming spawning event (not separated from the reserve fund), form the groups with different size and condition of the cells. The fractions of oocytes appear during vitellogenesis, before onset of maturation. Size distribution of oocytes before injection I in the females of D. trimaculatus and A. sexfasciatus (characterized by subsequent induced ovulation) shows that (among vitellogenous oocytes) the cells of the smaller size groups do not prevail, and the cells of certain size classes are absent. Such a size distribution of oocytes represents a characteristic of discontinuous oogenesis (Oven, 1976). After the spawning, oocytes at the phases of incomplete oogenesis remain in the ovaries. In various females, these cells can possess different diameter; and therefore, different time is required for completion of vitellogenesis and acquisition of maturational competence. Owing to a prolonged reproduction season in the tropical representatives of the family Pomacentridae, the recruitment of the current oocyte fund from the reserve fund takes place during the release of oocyte batches. As is known, Dascyllus albisella spawns over the entire year with the interval between release of distinct oocyte fractions from 5 to 7 days (Asoh and Yoshikawa, 2003). In the conditions of the aquarium, Amphiprion perideraion spawns for 8 months with the interval from 8 to 27 days between egg depositions (Ho et al., 2007). To take into account the opinion of Oven (1976, 2004), the type of oogenesis in D. trimaculatus and A. sexfasciatus seems discontinuous with fractional oocyte deposition. However, based on Alekseev and Alekseeva (1996), the development of oocytes in these species is continuous with discontinuous-asynchro51

nous vitellogenesis. Nevertheless, we suggest that the type of oogenesis in these species is discontinuous that is supported by the histograms and histological sections: certain size groups of vitellogenous oocytes are absent that is not usual for the species with continuous development of sex cells. The realization of vitellogenous oocytes is accompanied by their recruitment from the reserve fund. Therefore, the type of oogenesis of D. trimaculatus and A. sexfasciatus differs from the typical discontinuous oogenesis described in many papers (e.g., Makeyeva, 1992; Alekseev and Alekseeva, 1996). To take into account the size composition of oocytes in D. trimaculatus after ovulation and comparatively small diameter of ovulated oocytes, the interval between depositions of oocyte batches in this species, most likely, is shorter than in A. sexfasciatus. The size composition of oocytes in Z. scopas shows that the cells of smaller size groups prevail (that differs from the pattern observed in the representatives of the family Pomacentridae), and the oocytes of intermediate size classes are seen. Thus, the type of oogenesis is continuous (Götting, 1961; Oven, 1976), oocytes with maturational competence are always present in the ovaries during the spawning season, and the eggs can be always obtained from the majority of females. In the majority of females of this species, induced ovulation was registered (as a rule) in 41–42 h after the “natural” ovulation. Therefore, the interval between releases of oocyte batches in nature, most likely, can reach approximately two days. Similar interval (from one to two days) is reported for several warm-water fish species with continuous type of oogenesis (Oven, 1976, 2004). Owing to the features of oogenesis in two representatives of the family Pomacentridae, oocytes with maturational competence can be found (during the spawning season) not always, but with a certain periodicity. If such oocytes are absent, the hormonal stimulation would be unsuccessful. As it is mentioned above, oocytes of older generation in intact females (before injection I) of the representatives of the families Pomacentridae and Acanthuridae possess the nucleus in the center of the cell surrounded by small lipid droplets. Following by injection I, the nucleus is slightly displaced to the animal pole, lipid droplets become larger due to their partial fusion (or, rarely, total fusion), and oocyte diameter increases insignificantly. Based on a gradual increase of oocyte size during maturation (especially in Z. scopas), fusion of lipid droplets and hydration of oocytes occur synchronously. According to Alekseev and Alekseeva (1996), oocyte diameter remains the same during the increase of the size of lipid droplets, and it becomes larger only during homogenization of yolk accompanied by oocyte hydration. 52

As it shown for several objects of aquaculture, following by injection I, oocyte polarization is reported and the nucleus migrates to the animal pole near the area of micropyle directly under the oocyte envelope; this movement is completed before injection II (Verigin et al., 1975; Lemanova and Sakun, 1975; Kazanskii et al., 1978; Duvarova, 1981; Makeyeva and Emel’yanova, 1990). A different pattern is registered in Z. scopas, D. trimaculatus, and A. sexfasciatus: movement of the nucleus continues after injection II synchronously with fusion of lipid droplets (Emel’yanova et al., 2006, 2009a, 2009b). In maturing oocytes of these species, the nucleus is seen by the time of the appearance of only one lipid droplet that is reported also for other marine fish species (Alekseev and Alekseeva, 1996; Oven, 2004).

2.3. Some data on hormonal stimulation and maturation of oocytes in other coral fishes This part of the study was conducted with the aim to obtain gametes from different coral fish species for subsequent artificial insemination and incubation of the eggs for identification of eggs and larvae of different species. These data are included in the Atlas of the Eggs and Larvae of Coastal Fishes of South Vietnam (Shadrin et al., 2003). In this section, the first preliminary data on hormonal stimulation of oocyte maturation in several fish species, as well as the data on morphological parameters of the fishes and some features of their gametes are included. We hope that these materials can be useful for subsequent development of the methods of hormonal stimulation and for better understanding of reproductive biology of the species. 2.3.1. Biological parameters of the fishes The main biological parameters of the studied species are given in Table 5. 2.3.2. Materials on induced ovulation Ovulated oocytes from 16 ornamental coral fish species (with the exclusion of three model objects) are obtained (Table 6). Among them, there are representatives of five families of the order Perciformes, a family from the order Scorpaeniformes, and a family from the order Tetraodontiformes. In the majority of the studied fish species, ovulation is induced by means of injections of surfagon + eglonil, less often, by means of only surfagon. For two species (P. dickii and D. zebra), both variants were tested, and ovulation was registered in both cases. Thus, as it is mentioned above, synthetic 53

Table 5. Biological parameters of the fishes Species

TL, cm

SL, cm

Abudefduf bengalensis

14.5 (13.5–15.0)14.3

12.3 (11.5–13.0)12.3

Dascyllus reticulatus

(6.3–7.7)6.8 (7.0–9.0)7.8

(5.8–6.4)6.1 (6.5–7.8)7.0

Neopomacentrus anabatoides

8.2 9.0 (7.5–10.0)8.6 (7.8–11.0)9.3

6.2 7.0 (6.0–8.0)7.3 (6.0–9.0)7.7

Acanthurus nigrofuscus Acanthurus triostegus

(11.0–20.5)15.8 (15.3–19.0)17.2 (13.0–18.5)14.7 (14.0–15.5)14.6

(9.5–16.5)13.0 (11.5–14.0)13.0 (11.0–16.5)12.7 (12.0–13.5)12.5

Thalassoma lunare Thalassoma hardwicke Stethojulis bandanensis

(14.2–19.0)15.5 (17.0–24.0)19.5 (10.2–14.5)12.3 15.0 (10.5–11.7)11.3 11.2

(12.0–15.0)12.7 (13.5–19.0)15.3 (9.2–12.5)10.6 13.0 (9.5–10.5)10.3 9.5

Siganus spinus

(18.8–19.0)18.9 (15.5–19.0)16.9

(16.0–16.2)16.1 (13.2–16.0)14.4

Parupeneus multifasciatus

(14.1–16.5)15.2 (15.7–18.0)17.0

(17.0–18.6)17.9 (17.7–21.8)19.8

(12.0–14.0)13.2 (13.0–19.5)15.3

(84.0−90.0)87.0 (49.0−160.0)103

Plectroglyphidodon dickii

G, g

G1, g

g, g

Number of exemplars

Pomacentridae 94.0 (76.0–97.0)86.7

85.0 (72.0–92.0)81.7

3.5 –

1 3

(9.2–12.8)10.9 (11.0–20.0)15.5

(9.0–12.0)10.0 (10.5–18)14

(0.1–0.4)0.3 (0.1–0.3)0.3

4 4

5.8 9.0 (10.5–21.5)15.5 (11.0–21.0)16.0

– 0.6 (0.10–0.20)0.17 (0.01–0.03)0.02

1 1 6 3

(25.0–168.0)96.5 (47.5–101.0)76.7 (46.0–144.0)73.1 (59.0–91.0)68.6

(0.5–6.0)3.3 (0.3–4.4)2.7 (0.4–3.0)1.8 (1.6–6.4)2.9

2 4 10 5

(24.5–49.0)30.8 (29.0–75.0)47.9 (15.0–40.0)27.5 49.0 (20.0–20.0)20.0 18.5

(0.1–0.8)0.4 – (0.03–1.00)0.52 – (0.6–0.8)0.7 –

6 9 8 1 4 1

(86.0–88.0)87.0 (50.0–85.0)65.4

(1.3–1.5)1.4 (1.6–3.0)2.2

2 7

(60.2–81.8)73.7 (78.9–130.0)102.7

(49.4–76.6)68.2 (75.4–123.8)98.3

(1.20–2.40)1.80 (0.03–0.80)0.23

7 4

(11.0–11.5)11.1 (11.8–16.0)13.0

Scorpaenidae (45.0–58.0)49.2 (55.0–136.0)76.8

(39.0–54.0)43.8 (50.0–128.0)72.0

5 8

(16.0−16.5)16.3 (13.4−19.8)16.4

(13.0−13.5)13.3 (11.0−17.8)13.7

(88.0−98.0)93.0 (51.0−170.0)107.0

(0.60–7.00)2.30 (0.01–0.07)0.04 (0.80−0.88)0.8 4 (0.04−0.15)0.8 9

(11.7–14.0)12.5 (10.0–11.0)10.3 (14.5–16.5)15,2 (12.0–15.0)13.5

(10.0–11.5)10.5 (8.0–9.5)8.7 (12.5–14.5)13,3 (10.7–13.0)11.9

6.7 10.0 (12.5–25.0)18.4 (13.0–25.5)18.0 Acanthuridae (38.0–195.0)116.5 (51.5–120.0)89.6 (50.0–156.0)82.1 (65.0–100.0)72.0 Labridae (26.0–51.0)32.5 (32.0–104.5)56.8 (16.0–47.0)29.9 52.0 (15.0–24.0)20.3 21.0 Siganidae (93.0–97.0)95.0 (58.0–93.0)68.9 Mullidae

Dendrochirus zebra Scorpaenopsis possi

Monacanthus ciliatus Cantherines pardalis

Monacanthidae (32.0–57.0)44.4 (20.0–31.0)24.5 (51.0–82.0)71.3 (39.0–73.0)56.0

(30.2–46.0)39.5 (18.4–27.0)22.2 (51.0–82.0)62.7 (35.0–82.0)58.5

(0.3–3.2_)1.7 (0.10–0.40)0,25 (0.70–2.00)1.35 (0.20–0.60)0.41

2 6

7 3 3 2

Note: Above the line, females; below the line, males. In the parentheses, limits of the values; after the parentheses, mean value; the dashes mean the absence of the data; other designations as in Table 1.

analogs of releasing hormones without DA can be applied at least for some species. A comparison of the interval between injection I and ovulation in different fish species shows that (in general) it is lower in the fishes with pelagic eggs (Acanthuridae, Labridae, Scorpaenidae, and Mullidae) than in the fishes with demersal eggs (Pomacentridae, Monacanthidae, and Siganidae). If the interval between injection I and ovulation is substantially lower than that in other representatives of related species or in the representatives of the same species, a more derived oocyte condition in intact females (before injection I) is registered. For example, in intact females of Plectroglyfidodon dickii, oocytes (in two first variants) were in the beginning of the period of maturation, the nucleus was displaced at one-third of the distance from the oocyte center to the animal pole, and lipid droplets were enlarged insignificantly. Therefore, only one 54

Table 6. Doses of the preparations and intervals between injections I and II and between injection I and ovulation in different ornamental coral fish species Family (order) and species

Doses of surfagon Interval, h (µg/kg) + eglonil (mg/kg) I II I – II I – ovulation Pomacentridae (Perciformes) Abudefduf bengalensis 10 + 10 25 +1 0 16 37 Dascyllus reticulatus 7+7 15 + 15 20 38 Neopomacentrus anabatoides 10 + 10 20 + 15 18 26. Plectroglyphidodon dickii 20 12 15 + 15 12 5+5 20 + 15 23 36–50 10 20 18–22 34–36 Acanthuridae Acanthurus nigrofuscus 15 + 10 45 + 15 15 25 Acanthurus triostegus 10 + 10 35 + 15 14 33–37 5 + 10 20 + 15 12 32 Labridae Thalassoma lunare 10 + 10 30 + 15 21 30–31 Thalassoma hardvicke 5 + 10 25 +15 14 40 Stethojulis bandanensis 7+7 15 +15 20 31 Halichoeres marginatus 5+5 15 +15 23 34 Siganidae Siganus spinus‫٭‬ 5+5 15 + 15 15 50–62 Mullidae Parupeneus multifasciatus 10 20 17 24–27 Scorpaenidae (Scorpaeniformes) Dendrochirus zebra 10 + 10 20 + 10 14 25 10 25 14 25 Scorpaenopsis possi 8+8 20 + 15 14 36–40 Monacanthidae (Tetraodontiformes) Monacanthus ciliatus 20 43–65 Cantherines pardalis 5+5 20 + 15 17 39–42

Note: The exemplars of Siganus spinus were injected by surfagon (15 µg/kg) for the third time in 23 h after injection II (in 38 h after injection I); ovulation was registered in 11.5–23.5 h after injection III. The dashes mean the absence of the data.

injection was applied, and ovulation was registered in 12 h. On the contrary, ovulation in Siganus spinus is observed only after injection III. This injection was conducted based on the condition of oocytes sampled in 23 h after injection II (the time of ovulation in the representatives of many species): the oocytes were at a good condition, but the nucleus located in the center of the cell, and lipid droplets were comparatively large. In the species with demersal eggs, such condition was observed before injection II. 2.3.3. Features of gametes Pomacentridae. The demersal eggs possess an oval shape, and the chorion is represented by a bunch of filaments at the animal pole. The type of oo55

genesis is discontinuous. In P. dickii, the oocytes possess an oval-pear shape that is different from the oocyte shapes in other representatives of the family. Ovulated oocytes are semi-transparent with only one lipid droplet. In A. bengalensis, oocytes possess a crimson coloration; in other studied species of the family (with the exception of A. sexfasciatus), oocytes have light yellow coloration. Sperm is activated in marine water, and spermatozoa are mobile at least for several minutes. Acanthuridae. Ovulated oocytes are pelagic, of rounded shape, transparent, without coloration, with a light yellow lipid droplet. The actual batch fecundity of A. triostegus is between 33 918 and 271 560 oocytes. In a part of the males, spermiation is registered after injection II. The type of oogenesis is continuous. Spermatozoa of the representatives of the genus Acanthurus retain motility for approximately two minutes after activation (at 25°C). Labridae. Ovulated oocytes are spherical; they are transparent, without coloration, with a lipid droplet. The type of oogenesis is continuous. All species of the family are protoginous hermaphrodites; therefore, the males are larger than the females. Sperm obtained from crashed testes is weakly activated in marine water, and spermatozoa retain motility for less than 3 min at 25°C. Siganidae. Ovulated oocytes are spherical; they are slightly adhesive and transparent, without coloration, with several lipid droplets. The females are characterized by a large actual fecundity (the weight of ovulated oocytes is 5–10 g in a female 15–16 cm TL and 56–70 g in weight). The type of oogenesis is discontinuous. Following by the injections, a large number of sperm can be obtained, and sperm is activated in marine water. Mullidae. Ovulated oocytes are pelagic, they are spherical and transparent, without coloration, with a lipid droplet. In Parupeneus multifasciatus, actual batch fecundity ranges from 1537 to 26 423 oocytes, on average, 7929 oocytes (n = 18 females) (Pavlov et al., 2011). The type of oogenesis is continuous. Sperm obtained from crashed testes is immobile after activation in marine water; only single cells possess weak agitated movements for several seconds. Scorpaenidae. Oocytes ovulate into the ovarian cavity, and they are included into gelatinous matrix produced by somatic cells. Actual batch fecundity of Dendrochirus zebra is approximately 5620 oocytes. Following by the pressure to the abdomen of the female, ovulated oocytes can not be stripped. Therefore, oocytes were obtained directly from the ovaries. To conduct artificial insemination in D. zebra and Scorpaenopsis possi, gelatinous matrix was partly removed. The type of oogenesis is continuous with egg deposition in batches. Ovulated oocytes are pelagic; they are spherical, 56

without coloration, with a lipid droplet in D. zebra and without lipid droplets in S. possi. In D. zebra, sperm obtained from crashed testes is activated in marine water. Spermatozoa retain motility for approximately 1.5 h at 24°C. In S. possi, the duration of progressive movements of activated spermatozoa from crashed testes is less then 6 min, but this duration in the sperm obtained from the urinary bladder (used for sperm storage) is at least 15 min. Investigation of the reproductive biology in three coral reef fish species from the family Scorpaenidae (Scorpaenopsis possi and Sebastapistes cyanostigma from the subfamily Scorpaeninae and Dendrochirus zebra from the subfamily Pteroinae) demonstrates special adaptations that are absent in the majority of teleost fishes. Among them, a hypertrophied urinary bladder in males, that is used for sperm storage, morphology of ovaries with stroma and germinative tissue located in the exovarian cavity and separated from the ovarian wall, and, specialized secretory epithelium of the inner ovarian wall for producing a floating gelatinous matrix surrounding ovulated eggs, peduncular structure of oocytes, extremely narrow egg envelope, differentiation of cytoplasm and yolk in ovulated oocytes before their activation, and very small perivitelline space in developing eggs (Pavlov and Emel’yanova, 2007; Pavlov et al., 2010) Monacanthidae. Ovulated oocytes of Cantherines pardalis are spherical, with a weak adhesiveness, they are transparent, with several (4–12) lipid droplets: 3–4 largest lipid droplets and several smaller droplets. Oocytes possess a greenish coloration. Actual fecundity of a female is 25 168 oocytes. Ripe sperm is weakly activated in marine water; spermatozoa retain motility for 5–30 s. The sperm from the ejaculate is activated better in the presence of oocytes. In both studied species, sperm from the crashed testes is not activated in marine water.

57

Chapter 3. Morphology and ultrastructure of gametes 3.1. Oocyte envelopes 3.1.1. Oocyte envelopes in teleost fishes: a brief review In teleost fishes, primary and secondary egg envelopes can be distinguished based on their origin (Ivanov, 1956; Ginzburg, 1968; Laale, 1980; Makeyeva, 1992). The primary egg envelope (zona radiata) develops as a result of synthetic activity of cytoplasmic organelles of the oocyte, but secondary egg envelope (chorion) is produced by the cells of follicular epithelium. In several fish species, follicular epithelium, most likely, participates in the development of the zona radiata (Hosokawa, 1985; Emel’yanova and Makeyeva, 1989). The zona radiata is found in all fishes, but the chorion is observed mainly in the species with adhesive eggs. A large diversity of egg envelopes is connected with both systematic position of the fish and ecological features of reproduction and embryonic development (Kryzhanovskii, 1949; Rass, 1953; Ivankov and Kurdyaeva, 1973; Makeyeva and Mikodina, 1977; Mikodina and Makeyeva, 1980; Ivankov, 1987). A function of the primary egg envelope is the protection of the egg from mechanical actions. The secondary egg envelope is used mainly for the attachment of the eggs to the substratum due to its adhesiveness connecting with the presence of mucopolysaccharides. The width of the zona radiata is variable in the fishes with different reproductive biology (Makeyeva, 1992; Drozdov and Ivankov, 2000; Oven, 2004). The narrowest egg envelope is usual in pelagic eggs developed in the water mass. In the fishes with adhesive demersal eggs, the zona radiata is comparatively wide. The difference in morphology of the zona radiata is well expressed based on its ultrastructure. In the majority of marine and freshwater fish species, the zona radiata includes two or three layers with a certain ultrastructure (Emel’yanova, 1979; 1997, 1999, 2003; Laale, 1980; Grierson and Neville, 1981; Hart and Donovan, 1983; Brusle, 1985; Prokhorchik et al., 1988; Morrison, 1990; Fausto et al., 2004; Emel’yanova et al., 2009a). In teleost fishes, the micropyle is located at the animal pole of the egg. It includes the micropylar funnel and micropilar canal, a small depression in the egg envelope and the canal, or only the canal; two latter variants are often found in marine fishes (Riehl and Gotting, 1974; Mikodina, 1979, 1987; Hosokawa, 1985; Hirai, 1988; Li et al., 2000; Mekkawy and Osman, 2006). 58

3.1.2. Oocyte envelopes in coral reef fish species 3.1.2.1. Zebrasoma scopas For electron microscopy, ovulated oocytes and fragments of testes were fixed in 2.5% glutaraldehyde mixed with 2% paraformaldehyde made in phosphate buffer, pH 7.4, with the addition of 2.5% NaCl and 1.5% sucrose. Then the samples were postfixed in 1% OsO4. Subsequent mounting of the samples for scanning electron microscopy (SEM) and transmission electron microscopy (TEM) was done according to generally accepted methods (Weakley, 1981).

Fig. 14. Ultrastructure of egg envelope in Zebrasoma scopas. SEM: (a) surface of egg envelope of the oocyte; (b) micropyle. TEM: (c, d) egg envelope. 1, pore; 2, internal layer of the zona radiata; 3, external layer of the zona radiata; 4, chorion; 5, mitochondrion; 6, endoplasmic reticulum. Ультраструктура яйцевой оболочки Zebrasoma scopas. Сканирующая электронная микроскопия (СЭМ): (a) поверхность яйцевой оболочки ооцита; (b) микропиле. Трансмиссионная электронная микроскопия (ТЭМ): (c, d) яйцевая оболочка. 1, пора; 2, внутренний слой zona radiata; 3, наружный слой zona radiata; 4, хорион; 5, митохондрия; 6, эндоплазматическая сеть.

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In SEM preparations, oocyte surface is perforated by rounded pores developed during the final phases of maturation (Fig. 14a). The diameter of the pore ranges from 0.06 to 0.32 µm, on average 0.19 µm on average 1.98 µm (SD 0.07, n = 83). The distance between the neighboring pores is 0.81– 4.15 µm (SD 0.67, n = 113). At the animal pole of the oocyte, the micropyle is represented by the canal with the rounded opening 4.45–5.12 µm in diameter, on average 4.81 µm (SD 4.71, n = 11). The canal consists of several concentric layers. The internal part of the canal is more narrow, 2.54–3.22 µm in diameter, on average 2.84 µm (SD 0.31, n = 6) (Fig. 14b). The oocyte surface is comparatively smooth. The particles of different shape and size, most likely, of extraneous origin can be seen. In TEM preparations, the envelope of ovulated oocyte is represented by a narrow zona radiata approximately 2 µm in width and (most likely) a weakly expressed chorion approximately 0.1 µm in width (Fig. 14c). The zona radiata, most likely, is composed of two layers, but the widths of the layers are extremely different. Therefore, almost all zona radiata is represented by the internal layer approximately 2 µm in width. This layer is composed of homogenous substance of intermediate electron density, and it includes from five to seven bands (located parallel to the plasmalemma) of larger electron density alternating with the areas with lower electron density. In the substance of the internal layer, distinct microvilli and remains of the canals (previously including micro- and macrovilli) can be seen. The external layer of the zona radiata is very thin, approximately 0.1 µm in width; its electron density is similar to that in the internal layer. Above this layer, an additional layer of similar width, but with a high electron density (chorion) is registered (Fig. 14c). Friable fibrils can be seen on the surface of this layer (Fig. 14d). At the surface layer of the peripheral cytoplasm, numerous mitochondria, vesicles, and shortened cisterns of the smooth endoplasmic reticulum are seen. 3.1.2.2. Dascyllus trimaculatus Ovulated oocytes of D. trimaculatus are spherical, with the villous chorion at the animal pole (Fig. 15a). The chorion is represented by a dense bunch Fig. 15. Ultrastructure of oocytes of Dascyllus trimaculatus. SEM: (a) a view of the oocyte; (b) a part of the animal pole with filaments of the chorion; (c) oocyte surface with pores. TEM: (d) development of the chorion in the follicular cells; (e) egg envelope at the animal pole; (f) a fragment of the oocyte with the zona radiata, filaments of the chorion are absent in this area. 1, zona radiata; 2, fragments of the chorion in the follicular cells; 3, filaments of the chorion; 4, narrow layer above the zona radiata; 5, a fragment of a cortical alveole in the peripheral part of the oocyte.

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Ультраструктура ооцитов Dascyllus trimaculatus. СЭМ: (a) общий вид ооцита; (b) часть анимального полюса с ворсинками хориона; (c) поверхность ооцита с порами. ТЭМ: (d) формирование хориона в фолликулярных клетках; (e) яйцевая оболочка на анимальном полюсе; (f) фрагмент ооцита с zona radiata, ворсинчатый хорион в этой области отсутствует. 1, zona radiata; 2, формирование хориона в фолликулярных клетках; 3, ворсинки хориона; 4, узкий слой над внешней частью zona radiata; 5, фрагмент кортикальной альвеолы в периферическом слое ооцита.

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of filaments of different length and width (Fig. 15b). The oocyte surface without chorion is smooth, with rounded pores (Fig. 15c). The diameter of the pore ranges from 0.04 to 0.20 µm, on average, 0.14 µm (SD 0.03, n = 97). The distance between neighboring pores is 0.36–1.91 µm, on average 0.84 µm (SD 0.35, n = 99). The completion of development of the chorion occurs in the follicular cells just before ovulation (Fig. 15d). The egg envelope of the ovulated oocyte is composed of the zona radiata and chorion (at the animal pole) (Fig. 15e). The width of the zona radiata is 7–8 µm at the animal pole and 4–5 µm at the vegetative pole. The zona radiata consists of the substance of intermediate electron density including alternating bands (with a small difference of electron density) located parallel to the oocyte surface. The number of the bands, most often, reaches 10–12. The external part of the zona radiata (approximately 1 µm in width) is characterized by larger electron density similar to this density in the filaments of the chorion (Fig. 15e). The most external layer of the zona radiata, approximately 0.1 µm in width, is not always seen at ultrathin sections. This layer is often comparatively friable, and its electron density is similar to that in the main part of the zona radiata. The filaments of the chorion are absent within a small area around the animal pole (Fig. 15f). 3.1.2.3. Abudefduf sexfasciatus In ovulated oocyte, the egg envelope is represented by the zona radiata and chorion. As in D. trimaculatus, the chorion is located only at the animal pole. It is represented by the bunch of filaments of different length and width (Figs. 16a, 16b). The oocyte surface without chorion is smooth, with rounded pores (Fig. 16c). The diameter of the pore ranges from 0.10 to 0.26 µm, on average, 0.18 µm (SD 0.04, n = 73). The distance between neighboring pores is 0.66–2.14 µm, on average 1.23 µm (SD 0.27, n = 84). At ultrathin sections, the zona radiata consists of the substance of intermediate electron density (Fig. 16d). The width of this zone is 8–12 µm at the animal pole and 4–6 µm at the vegetative pole. Alternating bands (most often, up to 12 bands) slightly different by electron density and located parallel to the oocyte surface are seen in the zona radiata. Depending on the section plane, filaments of the chorion are seen as rounded or elongated structures (Fig. 16e). The external part of the zona radiata approximately 1.5 µm in width can possess a larger electron density representing an impression of the additional layer.

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Fig. 16. Ultrastructure of oocytes of Abudefduf sexfasciatus. SEM: (a) a view of the oocyte; (b) a part of the animal pole with filaments of the chorion; (c) oocyte surface with pores. TEM: (d) zona radiata, filaments of the chorion are absent in this area; (e) transversal section of filaments of the chorion. Ультраструктура ооцитов Abudefduf sexfasciatus. СЭМ: (a) общий вид ооцита; (b) часть анимального полюса с ворсинками хориона; (c) поверхность ооцита с порами. ТЭМ: (d) zona radiata, ворсинчатый хорион в этой области отсутствует; (e) поперечные срезы ворсинок хориона.

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3.1.3. Conclusions In three model species of ornamental coral reef fishes, average diameter of pores at the surfaces of ovulated oocytes is more or less similar. The largest diameter of pores (on average, 0.19 µm) is registered in pelagophil Z. scopas, a species with the smallest egg diameter (in comparison to this diameter in two representatives of the family Pomacentridae). We suppose that a lower diameter of pores in the species of the family Pomacentridae can be connected with a special morphology of the egg envelopes with the chorion located only at the animal pole. The development of the substances of the chorion is restricted to the part of the oocyte adjacent to the animal pole. Other Follicular cells surrounding the vegetative zone of the oocyte do not participate in the synthesis of the chorion and possess comparatively small size; the diameter of macrovilli entering the egg envelope just before ovulation (most likely) is also small. In two representatives of the family Pomacentridae, larger oocytes of A. sexfasciatus posess larger pores than smaller oocytes of D. trimaculatus. In three model species, maximum distance between neighboring pores (1.98 µm) is registered in Z. scopas, and the minimum distance (0.84 µm) in D. trimaculatus. Comparatively similar values of the diameter of the pore and the distance between the pores are reported for the larger eggs of several cold water Antarctic fish species (Mikodina and Pukova, 2001). For the pelagophils from the White and Black seas with the eggs of different diameter, similar values of pore diameters and distances between them are registered (Mikodina, 1987). Based on the opinion of the author, these characteristics can not be used for identification of the eggs. Among three model species, the micropyle is observed only in Z. scopas: in the representatives of the family Pomacentridae, the micropile is masked by filaments of the chorion. In Z. scopas, the diameters of the external and internal openings of the micropylar canal are different. Similar pattern is observed in other marine and freshwater fishes. Nevertheless, the canal with the same diameter of the openings can be found in many fish species. The diameter of the canal is important for the process of insemination (Murata, 2003). The zona radiata is the narrowest in (pelagophil) Z. scopas, and it is substantially wider (especially at the animal pole) in the representatives of the family Pomacentridae with demersal eggs. Thus, the maximum strength of the egg envelope is achieved in the area of the attachment of the egg to the substratum. The zona radiata in Z. scopas and D. trimaculatus consists of two layers. The same structure, most likely, is usual for A. sexfasciatus. However, the difference between the densities of the external and internal layers in the latter species is lower. In the majority of marine 64

and freshwater fish species, the zona radiata is composed of two or three layers (Anderson, 1967; Riehl, 1977; Mikodina, 1978; Prokhorchik et al., 1988; Kim et al., 2002; Fausto et al., 2004). In three model species, the internal layer of the zona radiata includes alternating bands with slightly different densities, which are parallel to the oocyte surface. Their number is low in Z. scopas and substantially larger in D. trimaculatus and A. sexfasciatus, the species with larger eggs and wider egg envelopes. A similarity in the ultrastructure of the internal layer of the zona radiata is observed in many marine fish species that is partly connected with their reproductive biology (Mikodina, 1987; Emel’yanova, 1997, 2003; Kim et al., 2002).

3.2. Spermatozoa 3.2.1. Spermatozoa in teleost fishes: a brief review In the majority of teleost fishes, spermatozoa belong to the type with flagellum. The ultrastructure of spermatozoa is described in many publications (Ginzburg, 1968; Billard, 1970; Fribourg et al., 1970; Nicander, 1970; Turdakov, 1972; Todd, 1976; Afzelius, 1978; Drozdov et al., 1981; Emel’yanova and Makeyeva, 1985; 1991a, 1991b; Mattei, 1991; Drozdov and Ivankov, 2000; Shadrin and Emel’yanova, 2007; Emel’yanova et al., 2009a; etc.). Both general features of spermatozoon morphology and specific adaptations connected with taxonomic position and reproductive biology are reported. Substantial differences in spermatozoon morphology are registered in the fishes with external and internal insemination (Ginzburg, 1968; Makeyeva, 1992). The differences are connected with the size of the cells, shape of the head and middle piece, number of mitochondria, etc. A common feature of spermatozoa of teleost fishes is the absence of acrosome found in the sperm of cartilaginous fishes and lower teleost fishes. Spermatozoa of the fish with internal insemination are characterized by the presence of elongated head and well developed mitochondrial apparatus in the middle piece that is important for a long life of the cells in the genital tract of the female. Similar spermatozoon structure is found in the representatives of seahorses (Syngnathidae) with insemination in the ventral brood pouch of the male (Hara and Okiyama, 1998; Kornienko and Drozdov, 2001). Nevertheless, some species with internal insemination possess spermatozoa with a simple structure (rounded head and small middle piece), as it is observed in false kelpfish Sebastiscus marmoratus (Mattei, 1991). Spermatozoon morphology of teleost fishes with external insemination is more variable. In particular, the shape of the head is substantially different 65

in marine and freshwater fishes from various taxonomic groups (Mattei, 1991; Drozdov and Ivankov, 2000; Hayakawa and Munehara, 2004; etc.). The differences in the structure of the middle piece are connected mainly with the number of mitochondria. For example, in cyprinids, a connection between reproductive biology and the area of mitochondria sections is reported. The latter parameter is correlated with the duration of sperm motility. In the species reproduced in the water current, the area of mitochondria sections and duration of sperm motility are lower than in the fishes spawned in stagnant water (Emel’yanova ana Makeyeva, 1985). In addition to the amount of mitochondrial material, the differences in the morphology of centriolar complex and the size of the intranuclear invagination of the head are registered. The proximal and distal centrioles can locate perpendicular (Siluridae) or almost perpendicular (Cyprinidae) to each other, they form a blunt angle (Bagridae) or even parallel to each other, and the proximal centriole can be located slightly above the distal one (basal plate) (Pomacentridae) (Emel’yanova and Makeyeva, 1998; Shadrin and Emel’yanova, 2007). Two centriolar complexes are found in channel catfish Ictalurus punctatus and brown bullhead Amiurus nebulosus (Ictaluridae). In the former species, the proximal centrioles are modified into the ring-shaped structures, and they are absent in the latter species (Emel’yanova and Makeyeva, 1991a, 1991b). The centriolar complex of wels Silurus glanis is located in the cytoplasm of the middle piece in the area of a small invagination of the head (less then 1/3 of the head diameter). In the representatives of the family Bagridae, the intranuclear invagination reaches approximately 2/3 of the head length (Emel’yanova and Makeyeva, 1991b), and in Parupeneus spilurus (Mullidae), this invagination is even larger (Gwo et al., 2004). The flagellum of the spermatozoon is composed of the axoneme surrounded by cytoplasmic membrane and plasmalemma. The axoneme, most often, consists of nine peripheral and two central duplets of microtubules. In spermatozoa of several fish species, the axoneme is without the central microtubules (as in the representatives of the order Anguilliformes), or only one central microtubule is observed (as in several species of the family Cichlidae) (Bern and Avtalion, 1990; Mattei, 1991). The peripheral duplets of microtubules can be electron transparent or (more rarely) electron dense (as in spermatozoa of the representatives of the families Clupeidae and Engraulidae (Mattei, 1991). Among spermatozoa of vertebrates, only fishes possess three types of gametes classified based on the presence of flagella and their number: spermatozoa with one flagellum (as in the majority of fishes), with two flagella 66

(Ariidae, Apogonidae, Ictaluridae, and Malapteruridae), and without flagellum (aba Gymnarchus niloticus (Gymnarchidae), representatives of Mormiridae, etc.) (Baccetti, 1985; Mattei, 1991). Thus, the general model of the spermatozoon structure of fishes (as in mammals) can not be suggested (Mattei, 1991). A polymorphism of spermatozoa is found in the representatives of the family Cottidae (both in the species with internal and external insemination). During spermatogenesis, both normal spermatozoa participating in fertilization and abnormal spermatozoa unable to fertilization but ensuring the success of this process are produced. The abnormal spermatozoa (paraspermatozoa), in fact, are represented by aberrant spermatids developing as a result of incomplete cytokinesis of the second meiotic division (Hayakawa and Munehara, 2004). In Syngnathus schlegeli (Syngnathidae), both typical spermatozoa with elongated heads and atypical spermatozoa with larger heads of different shapes are found in the ejaculate. Based on the opinion of the authors (Watanabe et al., 1998), a function of the latter cells (with rotatory movements) is the mixing of the typical spermatozoa densely packed in the brood pouch of the male. Thus, the appearance of additional or even unique features of spermatozoa structure in some fish species is connected mainly with specific patterns of their reproductive biology. 3.2.2. Spermatozoa in coral reef fish species 3.2.2.1. Zebrasoma scopas The spermatozoon includes the head of almost spherical shape approximately 1.8 µm in diameter and comparatively small middle piece (0.8 x 1.1 µm); the length of the flagellum reaches on average 29 µm. The location of the flagellum is comparatively symmetrical (Figs. 17a, 17b). The head of the spermatozoon is surrounded by the nuclear membrane and plasmalemma. Chromatin material of the head is tightly packed, but distinct particles can be distinguished. A small invagination reaching 1/3 of the head length with the centriolar complex is observed in the cytoplasm in the base of the head (Figs. 17c–17e). The centrioles are perpendicular to each other, and the proximal centriole is located above the distal one (Fig. 17f). The centriolar complex is located slightly asymmetrically. At transversal sections of the proximal centiole above the triplets of microtubules, the protrusions of amorphous electron dense material (most likely, centriolar appendages) are seen. The same material sometimes is found near the lateral surfaces of the longitudinal section of the basal plate. In the cytoplasm of the middle piece, from two to five (more often, three) sections of mitochondria are observed. 67

Fig. 17. Spermatozoon ultrastructure of Zebrasoma scopas. SEM: (a, b) a view of spermatozoa. TEM: (c–e) longitudinal sections of spermatozoa; (f) centriolar complex; (g, h) transversal sections of flagella. 1, middle piece; 2, mitochondrion; 3, proximal centriole; 4, distal centriole; 5, axoneme. Ультраструктура сперматозоидов Zebrasoma scopas. СЭМ: (a, b) общий вид сперматозоидов. ТЭМ: (c–e) продольные срезы сперматозоидов; (f) центриолярный комплекс; (g, h) поперечные срезы жгутиков. 1, средняя часть; 2, митохондрия; 3, проксимальная центриоль; 4, дистальная центриоль; 5, аксонема.

The axoneme of the flagellum is composed of nine peripheral and two central duplets of electron transparent microtubules. The axoneme is surrounded by the narrow or (sometimes) comparatively wide cytoplasmic membrane (Figs. 17g and 17h). 3.2.2.2. Dascyllus trimaculatus The spermatozoon consists of the head, small middle piece, and flagellum approximately 28 µm in length (Figs. 18a, 18b). The bean-shaped head is well expressed at ultrathin sections (Fig. 18c). Chromatin is densely packed, but distinct particles can be seen. The size of the head along the longer and shorter axes is approximately 2.5 × 0.8 µm. The centriolar complex is located asymmetrically in the middle piece in a small invagination inside of the head. The centrioles are almost parallel to each other; the proximal 68

Fig. 18. Spermatozoon ultrastructure of Dascyllus trimaculatus. SEM: (a, b) a view of spermatozoa. TEM: (c) longitudinal section of a spermatozoon; (d) transversal sections of flagella. 1, proximal centriole; 2, distal centriole; 3, mitochondrion. Ультраструктура сперматозоидов Dascyllus trimaculatus. СЭМ: (a, b) общий вид сперматозоидов. ТЭМ: (c) продольный срез сперматозоида; (d) поперечные срезы жгутиков. 1, проксимальная центриоль; 2, дистальная центриоль; 3, митохондрия.

centriole is located slightly above the distal one. The middle piece is small: approximately 1.0 × 1.5 µm. Several sections of mitochondria (most often, from two to four, but sometimes more) are seen (Fig. 18c). The axoneme of the flagellum is composed of nine peripheral and two central duplets of electron transparent microtubules. The axoneme is irregularly surrounded by the cytoplasmic membrane. Therefore, lateral lobes can be seen at the transversal sections of the flagellum (Fig. 18d). 3.2.2.3. Abudefduf sexfasciatus The spermatozoon is consists of the bean-shaped head (at ultrathin sections), small middle piece, and flagellum approximately 25 µm in length (Figs. 19a–19c). Chromatin is densely packed, but distinct particles can be seen. The size of the head along the longer and shorter axes is approxi69

Fig. 19. Spermatozoon ultrastructure of Abudefduf sexfasciatus. SEM: (a, b) a view of spermatozoa. TEM: (c–e) longitudinal sections of spermatozoa. 1, centriolar complex; 2, mitochondrion; 3, transversal section of the flagellum. Ультраструктура сперматозоидов Abudefduf sexfasciatus. СЭМ: (a, b) общий вид сперматозоидов. ТЭМ: (c–e) продольные срезы сперматозоидов. 1, центриолярный комплекс; 2, митохондрия; 3, поперечный срез жгутика.

mately 2.6 × 0.8 µm. As in D. trimaculatus, the centriolar complex is located in the cytoplasm in a small invagination of the head. The centrioles are almost parallel to each other, and the proximal centriole is located above the distal one (Figs. 19d, 19e). The middle piece is comparatively small (1.2 × 1.3–1.8 µm), and several sections of mitochondria are seen (Fig. 19d). The axoneme of the flagellum is composed of nine peripheral and two central duplets of electron transparent microtubules. The axoneme is irregularly surrounded by the cytoplasmic membrane. Therefore, one or two lateral lobes can be seen at the transversal sections of the flagellum. 3.2.3. Conclusions In three model species, the smallest spermatozoa are observed in Z. scopas, and the largest spermatozoa are registered in A. sexfasciatus. Therefore, 70

the size of the spermatozoon is correlated with the size of ovulated oocytes. The shape of the spermatozoon head in the representatives of the families Acanthuridae and Pomacentridae is also different: spherical and bean-like, respectively. The particles observed in the densely packed chromatin of the model species are also reported in other fishes (Brusle, 1981; Stein, 1981), but they often are not expressed (Emel’yanova and Makeyeva, 1985; Mattei, 1991). The functional role of the particles is not clear. The differences in the size and shape of the middle piece of the spermatozoon in three species can be connected with the structure of the head and with morphology and location of the centriolar complex. In Z. scopas, the centrioles are located perpendicular to each other, and the proximal centriole is situated above the distal one; the butt of the former centriole is directed to the upper lateral part of the distal centriole. In the representatives of the family Pomacentridae, the centrioles are almost parallel to each other, and the proximal centriole is located above the distal one. Substantial differences in the location of the centrioles in the centriolar complex are described in many fish species (Afzelius, 1978; Emel’yanova and Makeyeva, 1985; 1991a, 1991b, 1998; Mattei, 1991; Gwo, 1995). This location, most likely, is important for the movement of the flagellum, and it can be connected with a degree of asymmetry of attachment of the flagellum to the head. The order Perciformes, including the objects of this study, is the largest order of the vertebrates with 150 families (Nelson, 1994). In the representatives of the order, spermatozoon morphology is exteremly variable: from a spherical head and a small middle piece with single mitochondrion in Сlinidae to elongated head with well developed middle piece in the representatives of the family Embiotocidae (with internal insemination). In the representatives of the family Apogonidae, the spermatozoon is supplied by two flagella (that is unique for the order). Despite a substantial diversity of spermatozoon morphology within the order, spermatozoa of the perciform type prevail. This type is characterized by a spherical head with the invagination in its basal part and the centrioles located outside of the invagination. The mitochondria are distributed in the base of the flagellum. The axoneme is characterized by the electron dense section of a microtubule in four duplets. According to Mattei (1991), the occurrence of spermatozoa of the perciform type in the species can be important for phylogenetic relationships within the order. It should be noted, that spermatozoon ultrastructure of the model species is slightly different from that in the perciform type. These species differ in the shape of the head (at least in the representatives of the family Pomacen71

tridae), location of the centriolar complex of the middle piece in the invagination of the head (but not outside of the head), and the presence of electron transparent microtubules in the axoneme of the flagellum. These features of spermatozoon structure, most likely, are directly connected with the specificity of reproduction.

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Chapter 4. Assessment of sperm quality 4.1. Methods of assessment of sperm quality in teleost fishes: a brief review To reach a high fertilization rate, an assessment of gamete quality is required before insemination in vitro. The development of quantitative objective methods for the assessment of sperm quality represents an important problem. In the process of insemination in large eggs, such as those of salmonids, the duration of motility is such that spermatozoon are not able to swim around even half of the circumference of the egg (Kime et al., 2001). Thus, even minor decreases in the motility of spermatozoa could have a profound effect on their ability to fertilize an egg. Assessment methods of sperm quality based on the fertilization rate of eggs are widely used in aquaculture. However, for the comparison of different sperm samples it is necessary to use ovulated oocytes from the same female. In addition, egg and sperm quality can not be identified if the tests are carried out at different times, and the sperm–egg ratio should be strictly controlled during the experiments. As is already known, there is a strong positive correlation between fertilization rate and sperm motility (Rurangwa et al., 2001). Until recently, a semi-quantitative method for the assessment of this motility has been applied. According to this method, sperm motility is assessed based on a certain scale (Ginzburg, 1968; Turdakov, 1972; etc.). For example, such a scale is presented in a review of Trippel (2003): (0) all spermatozoa are inactive; (1) < 5% of motile cells; (2) 5– 29% of motile cells; (3) 30–79% of motile cells; (4) 80–95% of motile cells; and (5) > 95% of motile cells. According to this scale, the inactive spermatozoa include both actually immobile cells and cells with vibrating movement. The method is subjective, and statistical treatment of the data is impossible. Several other parameters used for quality evaluation of sperm including spermatozoa morphology, cell membrane resistance, mitochondria viability and functionality, and metabolic activity (see review by Cabrita et al., 2009) are rarely applied in fish culture practice. The methods of computer-assisted sperm analysis (CASA) assessing real velocities of different cells are developed from the 1990s. The methodology used until recently in ichthyology and aquaculture has come from humanfertility clinics. It involves videotaping spermatozoa movement via a microscope and subsequent analysis using computer programs. The programs, which can automatically follow a trajectory of each cell, are widely applied. The CASA methods in fish are used to investigate the influence of the environ73

ment and ecological conditions on sperm characteristics (Elofsson et al., 2003a, 2003b; Alavi and Cosson, 2004), to optimize conditions for artificial insemination (Christ et al., 1996; DeGraaf et al., 2004), to select males with a high sperm quality for establishing broodstocks (Kime et al., 2001), to conduct experiments on sperm storage, including cryopreservation (Wayman et al., 1998; Suquet et al., 2000; Rurangwa et al., 2001; Terrence and Tiersch, 2001), as well as in studies devoted to the pollution effect of a water environment on populations (Rurangwa et al., 1998; Schoenfuss et al., 2002; Van Look and Kime, 2003; Lahnsteiner et al., 2004). A system for the computer analysis of human sperm has been developed by Video TesT (St. Petersburg, Russia); it includes a microscope with a phase-contrast device, a standard microcellular camera for sperm, a color analogous system to transfer images to the computer, and special software (Videotest-sperm 2.1, 2004). Videotaping of sperm motility is conducted at a rate of 50 frames/s, and the files are saved in AVI format. The duration of videotaping is 1 s. The system involves two methods of automated analysis: “Motility” and “Morphology”. All information, including approximately ten parameters, appears automatically on the screen of the computer. The trajectories of spermatozoa movements can be seen on a videotape, and the colors of the tracks are different according to the gradation of sperm motility established by the Word Health Organization (WHO). Similar systems are known from other countries. For example, “Hobson Sperm Tracker” software can simultaneously track up to 200 spermatozoa and generate 14 parameters of their movement (Kime et al., 2001). The sperm of most teleost fishes differs from that of mammals in the following aspects: (1) the trajectories of fish spermatozoa are generally more complex than in mammals; (2) active movements of spermatozoa last for a short time (in the majority of species, less than 2 min); (3) spermatozoa do not have acrosomes and penetrate into the egg through micropyle (Kime et al., 2001). Therefore, in all studies devoted to the computer analysis of sperm motility in fish conducted up to now, the methodologies used for the assessment of human sperm fertility require some adaptations. Owing to their complexity and high prices, the present systems for CASA can not be used in the majority of ichthyological laboratories. A method for the computer analysis of fish sperm that does not require considerable finances and involves easily available free software (Pavlov, 2006) is described in the following section. This software is a plugin for the free National Institutes of Health software ImageJ (http:// www.uhnresearch.ca/wcif). Another plugin for JmageJ software to analyze sperm motility in fishes is developed by Wilson-Leedy and Ingermann (2007), and it is available at http://rsb.info.nih.gov/ij/plugins/casa.html. 74

4.2 A method of assessment of sperm quality in coral reef fish species The following abbreviations are used in two following sections: ALH, amplitude of the lateral spermatozoon head displacement; BCF, beat cross-frequency of the flagellum; LIN, linearity (LIN = VSL/VCL x 100); MOT, proportion of motile spermatozoa with progressive movement; VAP, velocity average path, smoothed path velocity of the spermatozoon head; VCL, velocity curvilinear; VSL, velocity straight line. The equipment used for the analysis of sperm motility includes a microscope and video camera connected to a computer. Videotaping is conducted at a total magnification of approximately x400. It is essential to obtain a stabilized image of the preparation with activated sperm within several seconds. For better identification of the distinct cells by the computer program, the cells should be slightly out of focus, and they should look dark on a light background. The method involves the use of three computer programs: a program for capturing images and montages of movies Pinnacle studio, a program for image analysis ImageJ, and Microsoft Excel (Fig. 20). Pinnacle studio. In the Capture mode, an image is captured from the analogous or digital video; the image will be seen in the upper left window of the screen. The capture quality should be set to high. The recording time depends on the period during which the cells remain within the microscopic field. The periodicity of the videotaping depends on the dynamics of spermatozoa activity and the total duration of their motility. The AVI files are saved automatically in the folder, which should be chosen in the lower right window, DV Camcorder. In the edit mode find the required file (for example, the file “min 1”, recorded in 1 min after sperm activation). Then drag the icon of the file into the lower Movie Window and choose the length of the clip (for example, 1 s). Using the button in the upper right part of the Movie Window, separate the part of the clip that should be analyzed (for example, the first 13 frames). In the Make Movie mode, save the file in an uncompressed AVI format. The number of such files should be equal to the number of videotapes recorded for the analysis of spermatozoa motility in the sample. For the subsequent transformation of pixel values into microns, an image of a part of the scale of the object micrometer should be recorded (with the same magnification) immediately before or after the registration of sperm motility. 75

“Pinnacle studio” Capture of an image and recording of the videotapes with duration not less than 1 s and periodicity depending on the total period of sperm motility.

Saving of the videotapes with the same duration of recording as uncompressed AVI files.

“ImageJ” Import of the uncompressed AVI file into ImageJ; the file is opened as a sequence of frames (stack). Smoothing and thresholding of the images to obtain binary black and white pictures.

Using the plugin MTrack2 to analyze sperm motility.

Copying the results (the length of the track of each spermatozoon along the curvilinear trajectory and the distance between the start and the end points of the track) into the Microsoft Excel.

“Microsoft Excel” Conversion of pixels into microns and calculation of the curvilinear velocity of sperm movement along the real trajectory (VCL) and straight line velocity of their movement between the start and end points of the track (VSL).

Statistical and graphical treatment of the data.

Fig. 20. A scheme of computer analysis of sperm motility in fishes. Схема компьютерного анализа подвижности спермы рыб.

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ImageJ. Open an uncompressed AVI file with the videotaped movements of spermatozoa: File/Import/AVI. The file is opened as a sequence of frames (stack). This sequence should be smoothed (Process/Smooth), and binary black and white images can be obtained (Process/Binary/Threshold). The spermatozoa should be black on a white background (Fig. 21). If the cells are white and the background is black, the images should be inverted (Edit/Invert). Sperm motility is analyzed by means of the plugin MTrack2 (Plugins/Particle counting/MTrack2). Four main parameters should be set in the control

Fig. 21. Spermatozoa of a male Zebrasoma scopas, 16 min after sperm activation: (a) raw image (heads and flagella of spermatozoa are seen); (b) black and white image after processing the videotape. Сперматозоиды самца Zebrasoma scopas через 16 мин после активации: (a) исходное изображение, видны головки и жгутики сперматозоидов; (b) чёрно-белое изображение после обработки видеозаписи.

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Fig. 22. Sperm curvilinear velocity (VCL) in two males of Zebrasoma scopas depending on the time after sperm activation and duration of in vitro sperm storage (up to 28 h) at 4.5°C. Frequency distribution of sperm velocity is presented. Each box is composed of five horizontal lines that display the 10th, 25th, 50th, 75th, and 90th percentiles of a variable (the rectangle includes from the 25th to 75th percentiles). All values above the 90th percentile and below the 10th percentile are plotted separately as circles. Скорость перемещения сперматозоидов двух самцов Zebrasoma scopas по криволинейной траектории (VCL) в зависимости от продолжительности хранения спермы in vitro (до 28 ч) при 4.5°C и времени после её активации. Проиллюстрировано распределение скорости перемещения сперматозоидов: каждый бокс включает 5 горизонтальных линий – 10, 25, 50 (медиана), 75 и 90% данных; от 25 до 75% данных заключены в прямоугольник; все значения за пределами 10 и 90% данных воспроизведены в виде точек.

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window, Object Tracker: (1) minimum object size (by default, the size of the object means its area in pixels), (2) maximum object size (pixels), (3) maximum velocity (pixels), and (4) minimum track length (frames). After obtaining binary images, even not well-defined objects become black. Therefore, the range of the cell size can be substantially wider compared to this range in the initial raw image. By replaying the tape frame by frame, the moving spermatozoa can be identified. Then the minimum and maximum areas of the cells can be determined (Analyze/Analyze Particles). The final results obtained after the computer sperm analysis may depend on the maximum velocity of spermatozoa (parameter 3) set in the window, Object Tracker. This parameter represents a maximum distance (in pixels) that a spermatozoon covers between two subsequent frames. If the value of this parameter is too small, the fastest spermatozoa will be excluded from the analysis; if the value is too large, the number of erroneous calculations increases: the label can be transferred from one cell to another adjacent cell during the alteration of frames. However, from a certain maximum-velocity value set in the window Object Tracker, the distance of the straight line spermatozoa movement (between the start and end points of the track) re-

Fig. 23. Sperm curvilinear velocity (VCL) in two males of Dascyllus trimaculatus depending on the time after sperm activation and duration of in vitro sperm storage (up to 7 h) at 4.5°C. Designations of the boxes as in Fig. 22. Скорость перемещения сперматозоидов двух самцов Dascyllus trimaculatus по криволинейной траектории (VCL) в зависимости от продолжительности хранения спермы in vitro (до 7 ч) при 4.5°C и времени после её активации. Обозначения как на рис. 22.

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mains comparatively stable. At the same time, the distance of spermatozoa movement along the real curvilinear trajectory becomes larger. To determine the maximum velocity of spermatozoa, the sequence of frames should be analyzed, and the cells with more rapid movement followed. For each cell as such, select its trajectory between two subsequent frames (Straight line selection) and measure the length of the trajectory (Analyze/Measure). The maximum value (in pixels) can be set into the Object Tracker window. The minimum track length (frames) (parameter 4) is equal to the number of frames in the opened sequence (stack). By the four parameter settings, the option Display Path Lengths in the lower part of the window, Object Track-

Fig. 24. Sperm curvilinear velocity (VCL) in two males of Abudefduf sexfasciatus depending on the time after sperm activation and duration of in vitro sperm storage(up to 20 h) at 4.5°C. Designations of the boxes as in Fig. 22. Скорость перемещения сперматозоидов двух самцов Abudefduf sexfasciatus по криволинейной траектории (VCL) в зависимости от продолжительности хранения спермы in vitro (до 20 ч) при 4.5°C и времени после её активации. Обозначения как на рис. 22.

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er, can be chosen to determine the characteristics of sperm motility. If the option Show Labels is selected, a new window with the same sequence of frames appears on the screen, and the labels appear automatically on each cell (Fig. 21b). The trajectories of each cell can be seen if the Show Paths option is selected. The Results window, which opens automatically, shows the distance of the movement of each spermatozoon along the real curvilinear trajectory (Length, VCL) and the linear distance between the start and end points of this trajectory (Distance traveled, VSL). To determine the ratio between the pixels and microns import an uncompressed AVI file with the image of a part of the scale of the object micrometer to the ImageJ program and calibrate this image (Analyze/Set Scale). Microsoft Excel. The values pasted into Microsoft Excel can be transformed from pixels to microns. Then the distances that spermatozoa cover per second (curvilinear velocity, VCL, and straight line velocity, VSL) can be determined. Graphical presentation of the data and their statistical treatment can be done both in this and in other programs. The examples of the application of the method for three model species are given in Figs. 22, 23, 24.

4.3. Conclusions From several methods for automatically assessment of sperm motility in fishes used up to now, each have their specific advantages and disadvantages, but in all cases, an adaptation from its primary design for human clinics is necessary (Kime et al., 2001). The main advantage of the method described above is its realization without complex equipment and expensive software. Using this method, sperm tracks can be calculated, most likely, for an unlimited number of cells in the microscopic field. The main parameters of spermatozoa activity (VCL, VSL, LIN, and MOT) can be determined. To calculate the latter parameter, the characteristics of inactive cells should be identified. As a rule even (at the first blush) inactive spermatozoa are able to weak agitated movements or slow movement with the fluid drift. The minimum velocity parameter of the object can not be set into the control window of the MTrack2 plugin; almost all spermatozoa are automatically registered as motile. According to the classification of WHO, spermatozoa with VCL less than 5 µm/s belong to cells with non-progressive movements. Based on the fish classification used for CASA (Lahnsteiner et al., 1996), spermatozoa with a velocity less than 5 µm/s, from 5 to 20 µm/s, and more than 20 µm/s belong to inactive, partly motile, and fully motile 81

categories, respectively. Other authors refer spermatozoa with an average smoothed path velocity of the spermatozoon head (VAP) less than 20 µm/s to inactive (Burness et al., 2005). According to our observation, spermatozoa with VSL less than 3 µm/s can be preliminary referred to inactive. The automatic calculation of the spermatozoa number can be conducted both in the whole microscopic field and in any selected area. Therefore, the concentration of spermatozoa in the ejaculate can be determined using microcellular cameras for sperm where the cells actually form a monolayer. Such parameters as VAP and several other parameters (for example, ALH or BCF) can not be determined by the method described above. However, the two latter parameters are used for CASA in fish comparatively rarely. In the present systems for CASA in fish, an objective x20 with a phase contrast device is used more often, and the total magnification is not large reaching x50 (Van Look and Kime, 2003) and x100 (Burness et al., 2005), or, in several studies, x250 and x400 (Trippel, 2003). The tape is recorded more often from 0.5 s (Burness et al., 2005) to 15 s (Kime et al., 2001; Van Look and Kime, 2003) and 1 min (Trippel, 2003). According to the method described above, the phase contrast device can be absent, and the total magnification is approximately x400. A large magnification leads to higher precision of the measurements, but spermatozoa leave the microscopic field rapidly, and the videotape should be analyzed for a short time (approximately 0.5 s). For synchronous activation of all spermatozoa, a double dilution of sperm is applied in many of the studies. For the initial dilution, the sperm is mixed with an extender having an osmotic concentration similar to that in the sperm fluid. For the second dilution, the solution is diluted in the activated medium. For example, sperm of red drum Sciaenops ocellata, obtained from crushed testes, was diluted in Hank’s balanced salt solution (200 mOsm/kg) or in a NaCl solution (400 mOsm/kg), and was then activated by marine water (800 mOsm/kg) (Wayman et al., 1998). The sperm of Siberian sturgeon Acipenser baeri was better activated at a final dilution of 1 : 600. It was diluted in an extender preventing sperm activation (at a ratio of 1 : 20) and then in an activated solution (at a ratio of 1 : 30) (Billard et al., 1999). The sperm of goldfish Carassius auratus was diluted in the extender (1 : 100) and then activated in distilled water reaching a final dilution of 1 : 2000 (Schoenfuss et al., 2002). The sperm of three model species was diluted once in marine water, which, most likely, affected the results; the movement of spermatozoa was not synchronous in different microscopic fields. Owing to the rapid decrease in sperm motility in the majority of fish species, a fully stabilized image without fluid drift and a focused microscope 82

should be obtained as quickly as possible. During an analysis of sperm motility in Atlantic cod Gadus morhua, a drop of diluted sperm was placed into a hemacytometer, covered by a coverslip, and focused for 20–30 s (Trippel, 2003). According to Kime et al (2001), a stabilization of an image and focus (using the multitest slide, 12-well, ICN, Basingstoke, UK, with the depth of the well approximately 0.0116 mm) can be reached ideally within 5 s of mixing the sperm with the water. Rapid focusing is especially important for fish such as salmonids with the total duration of sperm motility reaching 20–30 s. However, in many marine fishes, spermatozoa retain activity for several minutes, and rapid focusing is not necessary The duration of progressive movements of more than 50% of spermatozoa in Z. scopas, D. trimaculatus, and A. sexfasciatus at 25°C is 3–20, 6–9, and 5–11 min after sperm activation, respectively (Pavlov, 2006; Pavlov and Emel’yanova, 2006). This comparatively long period of sperm motility, most likely, is connected with the comparatively prolonged period of fertilizing ability in the eggs of marine fish: their activation is connected mainly with the penetration of a spermatozoon into the micropyle (see sections 5.2.1.2 and 5.2.1.3). In the majority of fishes spawned in fresh water (salmonids, cyprinids, and silurids), the values of VCL and VSL are similar, and the average initial value of VCL just after sperm activation is approximately 110 µm/s (Kime and Tveiten, 2002). The velocity of spermatozoa movement (VCL) in marine fishes is lower reaching in Z. scopas, D. trimaculatus, and A. sexfasciatus in 1 min after sperm activation 15.3–74.5, 58.4– 92.2, and 12.7–21.6 µm/s, respectively. In Atlantic cod, VCL reaches on average 75 µm/s, but it can be as large as 1000 µm/s (Trippel and Neilson, 1992). The characteristics of sperm motility differ in spotted wolffish Anarhichas minor; the average values of VCL and VSL are 50 and 12 µm/s, respectively (LIN = 24%), which is connected with the movement of spermatozoa along the winding trajectories that are substantially different from straight line tracks (Kime and Tveiten, 2002). A special pattern of sperm motility in spotted wolffish is associated with internal insemination of eggs, which has been also described for other representatives of the family Anarhichadidae (Pavlov, 2004). The spermatozoa of the studied coral-reef fish species possess a comparatively low initial velocity, retain activity for at least 5 min, and the sperm can be stored at 4.5°C for several hours (Pavlov and Emel’yanova, 2006). Based on the features of sperm motility, substantial differences between the pelagophil, Z. scopas, and guarding litophils from the family Pomacentridae are absent. However, the differences in the initial sperm velocity are registered in A. sexfasciatus and D. trimaculatus. They can be connected 83

with the special features of reproductive ecology, which represent a subject for further investigations. Thus, CASA is a useful tool for an objective assessment of sperm quality in fish. However, a subjective factor, which can affect the final results, is connected with the selection of the value of maximum velocity of spermatozoa that should be set in the control window of the computer program. The errors in the measurements are assessed more often during a comparison of the trajectories of cell movement calculated automatically and manually by means of a frame-by-frame analysis of the videotape (e.g., Davis et al., 1992). One of the goals of future works is to development mathematical methods for a quantitative assessment of errors during the analysis of sperm motility.

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Chapter 5. Assessment of egg quality 5.1. Methods of assessment of egg quality in teleost fishes: a brief review Egg quality in the broad interpretation of this term depends on the intrinsic properties of eggs and environment when the egg develops after fertilization (Bromage et al., 1994). Egg quality in the strict sense can be determined as an ability of ovulated oocytes to fertilization and subsequent development of viable offspring (Kjørsvik et al., 1990; Bonnet et al., 2007). According to latter definition, the properties of the egg depend on the genotypes of the female and male, as well as on the physiological events that occur inside of the egg. The features of these events are connected mainly with the conditions of oocyte development during oogenesis. In the aquaculture practice, the conditions of keeping of spawners (water quality, feeding regime, and stress inducing factors), as well as different types of manipulations with spawners (e.g., induction of oocyte maturation and ovulation by means of control of environmental parameters or using hormonal injections) can be reflected in egg quality (Ginzburg, 1968; Schreck et al., 2001; Thorsen et al., 2003; Pavlov et al., 2004; Gagliano and McCormick, 2007). A storage of ovulated oocytes in the body cavity (in salmonids) or in the ovarian cavity (in the majority of other teleost fishes) is often the main reason for decreasing oocyte fertility (Korovina, 1986; Bromage et al., 1994; Rizzo et al., 2003; Pavlov et al., 2004; Bonnet et al., 2007). In addition, the ability to fertilization and normal embryonic development decreases after a short-time (for several hours) storage of ovulated oocytes in ovarian fluid or physiological solution before insemination (Ginzburg, 1968; Stoss, 1983; Rizzo et al., 2003; Pavlov et al., 2004). In the conditions of hatcheries, incubation of poor quality eggs leads to inefficient exploitation of equipment and unreasonable expenditures of labor and time. To decrease these expenditures and to monitor the condition of the broodstock, the assessment of egg quality should be conducted at the earliest stages of egg development. The quality of inseminated pelagic eggs of marine fishes is usually assessed based on their ability to float or sink in marine water. However, floating eggs are not always composed of only fertilized eggs. In addition, fertilized eggs can be characterized by a poor quality exhibiting abnormal development. The egg size is not a reliable indicator of their quality, but a low variability by size in the eggs obtained from a female suggests that the probability of their normal development is increased. Based on other meth85

ods for the assessment of egg quality with the use of cytological or biochemical methods, the results can not be obtained quickly. Therefore these methods are almost not applied in aquaculture (see reviews: Kjørsvik et al., 1990; Brooks et al., 1997; Pavlov et al., 2004; Rideout et al., 2004; Lahnsteiner et al., 2009). Morphology of oocytes at initial developmental stages, most likely, can represent a reliable indicator of their quality. In the ovulated oocyte of teleost fish, the cytoplasm is more or less proportionally distributed over the yolk surface, and cortical alveoli are located inside of the cytoplasm beneath the cyptoplasmic membrane. The cortical reaction includes the chain of initial changes of the cortical layer of the egg including exocytosis of cortical alveoli. Following by the action of activating factor (for example, a contact between the membrane of a spermatozoon and cytoplasmic membrane of the oocyte), calcium binding in the cytoplasm releases as a wave from the animal to vegetative pole of the egg. The wave is accompanied by the release of content of cortical alveoli. The alveolus moves close to the cytoplasmic membrane, and colloid substance releases into the perivitelline space between this membrane and zona radiata followed by the destruction of the alveolus membrane (Ginzburg, 1968; Gilkey, 1981). In fertilized eggs of marine fishes, the cortical reaction is not always completed: cortical alveoli are registered in cleaved eggs of several species (see reviews: Kjørsvik et al., 1990; Brooks et al., 1997; Pavlov et al., 2004). These observations suggest that the dynamics of cortical reaction can be connected with egg quality. However, the data on the assessment of oocyte quality in marine fishes just after ovulation, during cortical reaction, and in the process of subsequent swelling of the egg are restricted. Morphology of cells at the stages of cleavage is a reliable indicator of their quality that is widely used. For several marine fish species, a positive correlation between the proportion of eggs exhibiting normal cleavage and hatching success of embryos, survival of larvae (see reviews: Shields et al., 1997; Pavlov et al., 2004), or even juveniles (Kjørsvik et al., 2003) is registered. In the experiments conducted on the eggs of Atlantic halibut Hippoglossus hippoglossus (Shields et al., 1997) and haddock Melanogrammus aeglefinus (Rideout et al., 2004), a negative correlation between the proportion of eggs with deviations from normal development at the stages from 8 to 32 blastomeres (including asymmetrical positions of cells) and percentage of hatched embryos is found. According to the results of other investigators, the eggs of Atlantic halibut (Vallin and Nissling, 1998) and Atlantic cod Gadus morhua (Thorsen et al., 2003) with asymmetric blastomere arrangement exhibit normal subsequent development. Based on the 86

results of Rani (2005), a correlation between the proportion of Atlantic cod eggs with abnormalities at the stages of four or eight blastomeres and survival of embryos in the middle of embryogenesis or at hatching is absent. In the eggs of loach Misgurnus fossilis, the size of blastomeres is variable from initial developmental stages, and this feature is regarded as mainly normal, but not pathological (Doronin, 1985). A substantial variation of the size of blastomeres in the normally developing egg is described for Atlantic cod (Makhotin, 1982; Rani, 2005), and common wolfish Anarhichas lupus (Pavlov, 1986). Therefore symmetrical arrangement of blastomeres is not a good indicator of egg quality. However, morphology of oocytes released from the female’s body, as well as morphology of eggs during initial embryonic development, remain the most reliable criteria of egg quality. The application of these criteria is illustrated in the following sections.

5.2. Morphological changes in the eggs and assessment of their quality in coral reef fish species 5.2.1. Zebrasoma scopas 5.2.1.1. Morphology of oocytes after stripping After stripping of oocytes from the body of a female, normal oocytes are transparent, of regular rounded shape, 555–672 µm in diameter, on average, 631 µm (SD 32, n = 16). Diameter of the egg after swelling is 606–685 µm, on average, 654 µm (SD 18, n = 40). Average diameter of the lipid droplet is 156 µm. The micropyle is seen at the surface of the egg envelope, and cortical alveoli are located in the cytoplasmic layer of the oocyte (Fig. 25a). Diameter of a cortical alveolus ranges from 1.1 to 5.5 µm, on average, 2.9 µm (SD 0.8, n = 200). The following abnormalities are registered in the oocytes after their stripping from the female’s body. The inclusions in the cytoplasm are seen (Fig. 25b), or cortical alveoli are absent (Fig. 25c). In the latter case, a perivitelline space is formed between the yolk surface and oocyte envelope (zona radiata). A presence of the oocytes with opaque content means coagulation of the yolk (Figs. 25d–25f). The size of such oocytes can be substantially larger than the size of normal sex cells. These abnormal oocytes were stored in the ovarian cavity of the female for some time (several hours or even several days) after ovulation. The oocytes of smaller size with folders on their envelopes are registered in some egg batches (Fig. 25g). In addition, the structures of unknown origin can be seen on the envelope (Figs. 25h, 25i). 87

5.2.1.2. Morphological changes in non-inseminated oocytes Egg quality was assessed in 4–5 h after insemination (or after placing of the oocyte in marine water). In normal fertilized eggs, the stages of middle- or small-cell morula are observed by this time. The eggs were separated into following categories: (1) with normal cleavage, (2) with abnormal cleavage, (3) with uncleaved cytoplasmic protrusion (blastodisc), and (4) without blastodisc. Morphological changes in non-inseminated oocytes after their placing in marine water are shown in Fig. 26. Many cortical alveoli are seen in the cytoplasmic layer of ovulated oocyte (Fig. 26a). A narrow space, which

Fig. 25. Morphology of oocytes of Zebrasoma scopas after their stripping from the females: (a) normal oocyte, the arrows show cortical alveoli and micropyle; (b) oocyte with inclusions inside of the cytoplasm; (c) cortical reaction took place in the ovarian cavity of the female; (d–f) oocytes damaged as a result of their prolonged storage in the ovarian cavity of the female after ovulation; (g) oocyte of a small diameter with folders on the envelope; (h, i) oocytes with the structures of unknown origin on the envelopes. Строение ооцитов, отцеженных у самок Zebrasoma scopas: (a) нормальный ооцит, кортикальные альвеолы и микропиле (отмечены стрелками); (b) ооцит с включениями в цитоплазме; (c) кортикальная реакция прошла в овариальной полости самки; (d–f) ооциты, поврежденные в результате продолжительной задержки в овариальной полости самки после овуляции; (g) ооцит небольшого диаметра со складчатой оболочкой; (h, i) ооциты со структурами неизвестного происхождения на оболочке.

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Fig. 26. Distribution of cortical alveoli and formation of the perivitelline space in non-inseminated ovulated oocytes of Zebrasoma scopas: (a, a1) oocyte before its placing into the water (0 h); (b, b1) uncompleted cortical reaction, 4 h from placing of the oocyte into the water, single cortical alveoli, uncleaved blastodisc; (b2) uncompleted cortical reaction, 4 h from placing of the oocyte into the water, parthenogenetic cleavage; (c, c1) partial cortical reaction, aggregation of the cytoplasm at the animal pole is absent, 1 h from placing of the oocyte into the water; (d, d1), partial cortical reaction, 2 h from placing of the oocyte into the water; (e, e1), partial cortical reaction, 4 h from placing of the oocyte into the water. Распределение кортикальных альвеол и формирование перивителлинового пространства в неосемененных овулировавших ооцитах Zebrasoma scopas: (a, a1) ооцит перед помещением в воду (0 ч). Неполная кортикальная реакция: (b, b1) 4 ч после помещения ооцита в воду, единичные кортикальные альвеолы, недробящийся бластодиск; (b2) то же, партеногенетическое дробление. Частичная кортикальная реакция, агрегация цитоплазмы на анимальном полюсе отсутствует: (c, c1) 1 ч после помещения ооцита в воду; (d, d1) 2 ч; (e, e1) 4 ч.

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can be seen in Fig. 26a1 between the yolk surface and zona radiata, is an artifact appearing during microscopy or so-called “presumptive perivitelline space” described by Ginzburg (1968). Uncompleted cortical reaction is registered in a part of oocytes after their placing in marine water. During this reaction, the number of cortical alveoli decreases substantially over the first minutes after keeping of the oocyte in marine water, single cortical alveoli remain in 4 h (Fig. 26b), and the blastodisc appears (Fig. 26b1). During subsequent storage of the oocyte, the cleavage of the blastodisc is not registered. Thus, these oocytes belong to category 3. In such oocytes, completed cortical reaction with the disruption of all cortical alveoli is registered only in several cases. Parthenogenetic cleavage is observed in single oocytes (Fig. 26b2). In such oocytes, the cells of the blastodisc are substantially different in size, and the surface of the blastodisc is uneven due to a low adhesion between adjacent blastomeres. Partial cortical reaction is observed in other oocytes. A small number of cortical alveoli are subjected to exocytosis over the first minutes after placing of the oocyte in marine water. The number of cortical alveoli reduces substantially for 1–2 h (Figs. 26c, 26c1, 26d, 26d1). In 4 h after placing of the oocyte in marine water, the width of the perivitelline space at the animal pole of the oocyte reaches approximately 70 µm, many intact cortical alveoli remain, and aggregation of the cytoplasm at the animal pole is not observed (Figs. 26e, 26e1). Such oocytes belong to category 4. In non-inseminated eggs from different females, the ratio between the oocytes of categories 3 and 4 ranges widely: the oocytes of a category can be totally absent. 5.2.1.3. Morphological changes in eggs of different quality after insemination In 5 min after insemination, completed cortical reaction is registered in a part of the eggs (Fig. 27a). These eggs begin to cleave, and, therefore, they belong to categories 1 or 2. Uncleaved blastodiscs are formed in single eggs with completed cortical reaction. These eggs belong to category 3. Uncompleted cortical reaction occurs in other eggs (Fig. 27b). Some of these eggs begin to cleave; and, therefore, they belong to category 2. In the majority of eggs with uncompleted cortical reaction, the blastodisc is formed, but the cleavage is not observed. Such eggs belong to category 3. In a part of oocytes, cortical alveoli remain intact for the first minutes after placing of the egg in marine water (Fig 27c), and partial cortical reaction is registered in 2 h. These cells belong to category 4. 90

Fig. 27. Exocytosis of cortical alveoli in the eggs of Zebrasoma scopas of various quality in 5 min after insemination: (a) completed cortical reaction; (b) uncompleted cortical reaction; (c) cortical reaction is not registered, partial cortical reaction will occur later. Экзоцитоз кортикальных альвеол в яйцах Zebrasoma scopas разного качества через 5 мин после осеменения: (a) полная кортикальная реакция; (b) неполная кортикальная реакция; (c) кортикальная реакция не прошла, впоследствии наблюдается частичная кортикальная реакция.

In the eggs that undergo cleavage and, therefore, belong to categories 1 and 2, the duration of exocytosis of cortical alveoli (from disruption of the first alveoli to disappearance of almost all alveoli) is from 12 to 27 s (Color Plates, Fig. 8a). However, the time from insemination of the oocyte to the beginning of cortical reaction (latent period of cortical reaction) can vary in different eggs: from 11 to 267 s. In many eggs from egg batches with poor quality (inseminated in 3 h after in vitro storage), cortical alveoli remain intact for several hours after the insemination. In some eggs, uncompleted cortical reaction is observed: the number of cortical alveoli decreases abruptly in 5 min after insemination, from more than 100 alveoli per 0.03 mm2 in the ovulated oocyte to 30–50 alveoli per 0.03 mm2. This number decreases over the first hour after insemination and then stabilizes at a low level reaching approximately ten alveoli per 0.03 mm2. In these eggs, abnormal cleavage of the blastodisc is registered. Thus, they belong to category 2. Partial cortical reaction is observed in the inseminated but unfertilized oocytes: the number of cortical alveoli decreases up to 60 per 0.03 mm2 in 2 h after placing of oocytes in the water. These oocytes belong to category 4. The relationship between the proportion of eggs with normal cleavage and the number of eggs with completed cortical reaction is not significant (Fig. 28a). However, the relationship between the proportion of all (normally and abnormally) cleaved eggs and the number of eggs with completed cortical reaction or summing percentage of eggs with completed and uncompleted cortical reaction (registered in 5 min after insemination) is substantially stronger and significant (Figs. 28b, 28c). The relationship between the features of cortical reaction and subsequent egg morphology is illustrated in Table 7. 91

92

0

a

20

60

80

Completed CR, %

40

y = 0,2275x + 36,744 r 2 = 0,0653 p > 0.05

100

120

Normal + abnormal cleavage, %

Normal cleavage, %

0

20

40

60

80

100

120

0

b 20

60

80

Completed CR, %

40

y = 0,492x + 47,061 r 2 = 0,5743 p = 0.0001

100

120

0

20

40

60

80

100

120

0

c 40

60

80

100

Completed + uncompleted CR, %

20

y = 1,0021x - 3,9161 r 2 = 0,7129 p < 0.0001

120

Fig. 28. Relationships (a) between the proportion of eggs of Zebrasoma scopas with normal cleavage and proportion of eggs with completed cortical reaction (CR), (b) between the proportion of eggs with normal and abnormal cleavage and proportion of eggs with completed cortical reaction, and (c) between the proportion of eggs with normal and abnormal cleavage and proportion of eggs with completed and uncompleted cortical reaction. The data for the egg batches from 20 females. Çàâèñèìîñòü (a) äîëè íîðìàëüíî äðîáÿùèõñÿ ÿèö Zebrasoma scopas îò äîëè ÿèö ñ ïîëíîé êîðòèêàëüíîé ðåàêöèåé (CR), (b) äîëè íîðìàëüíî è àíîìàëüíî äðîáÿùèõñÿ ÿèö îò äîëè ÿèö ñ ïîëíîé êîðòèêàëüíîé ðåàêöèåé è (c) äîëè íîðìàëüíî è àíîìàëüíî äðîáÿùèõñÿ ÿèö îò äîëè ÿèö ñ ïîëíîé è íåïîëíîé êîðòèêàëüíîé ðåàêöèåé. Äàííûå äëÿ èêðû îò 20 ñàìîê.

0

20

40

60

80

100

120

Normal + abnormal cleavage, %

Table 7. Features of cortical reaction and morphology of eggs of Zebrasoma scopas Cortical reaction Completed -“-“Uncompleted -“Partial

Proportion of cortical alveoli broken in 2 h after insemination, % 100 100 100 ~ 90 ~ 90 ~ 40

Fertilization

Categories of eggs

+ + – + – –

1 – normal cleavage 2 – abnormal cleavage 3 – uncleaved blastodisc 2 – abnormal cleavage 3 – uncleaved blastodisc 4 – without blastodisc

5.2.1.4. Fertility of oocytes after their in vivo storage in the female’s ovarian cavity In the experiments on in vivo storage of oocytes, small oocyte batches were stripped and inseminated just after ovulation and in certain time after ovulation. In a part of the eggs stored in the ovarian cavity of the female for 2 or 4 h after ovulation, the borders between blastomeres are not distinct (from the initial cleavages) due to incomplete cell divisions (Figs. 29a, 29b). Owing to divisions of nuclei without cytotomy, multinuclear syncytia appear in some eggs (Fig. 29c). In the eggs from several females, the syncytium is formed in the central part of the blastodisc, but cell divisions continue at its periphery (Fig. 29d). The proportion of eggs with such abnormality can reach 90%. A poor adhesion between adjacent cells is registered in many eggs (Figs. 29e–29i). As a result, large intercellular spaces appear in the blastodisc up to the total separation of the cells from each other. This abnormality can be accompanied by incomplete cell divisions, desynchronization of cleavages, or formation of syncytia. At the stage of middle-cell morula, degradation of the cells can be seen both at the periphery (Fig. 29j) and in the central region (Fig. 29k) of the blastodisc. The layer of periblast (yolk syncytial layer) with large nuclei is seen clearly. This layer spreads over the surface of the yolk and can be seen beyond the border of the blastoderm. In the eggs with the abnormalities, total degradation of cells is registered at the steps of cleavage and blastulation, before onset of gastrulation. The proportions of normally cleaved eggs in the control series (oocytes are stripped and inseminated just after ovulation) and in the series stored in vivo for 2 and 4 h are 83% (SD 18), 30% (SD 29), and 35% (SD 33), respectively (Color Plates, Fig. 9). In the eggs from the majority of females, the storage of ovulated oocytes in vivo for 2 h leads to the significant decrease of the number of eggs with normal cleavage and to increas93

Fig. 29. Abnormalities at the step of cleavage after in vivo oocyte storage in Zebrasoma scopas: (a) unclear borders between the cells due to incomplete divisions at the stage of 8 blastomeres, age 2 h 00 min from insemination (at 25°C); (b) the same abnormality at the stage of 32 blastomeres, age 2 h 40 min; (c) appearance of the syncytium in a sector of the blastodisc, age 2 h 30 min; (d) appearance of the syncytium in the central region of the blastodisc, age 3 h 30 min; (e) weak adhesion between the blastomeres, incomplete cell divisions, age 1 h 45 min; (f) weak adhesion between the blastomeres, desynchronization of cleavage, age 2 h 00 min; (g) weak adhesion between the blastomeres, appearance of the syncytium, age 3 h 00 min; (h, i) weak adhesion between the blastomeres, total separation of the cells from each other, age 3 h 30 min; (j) degradation of cells at the periphery of the blastodisc, age 4 h 00 min; (k) degradation of cells in the central region of the blastodisc, age 4 h 00 min. Arrows show the margin of the periblast (yolk syncytial layer). Аномальное развитие яиц Zebrasoma scopas на этапе дробления после задержки овулировавших ооцитов в овариальной полости самки (in vivo): (a) нечёткие границы между клетками вследствие неполных клеточных делений на стадии 8 бластомеров, возраст 2 ч 00 мин после осеменения (при 25°C); (b) аналогичное нарушение на стадии 32 бластомеров, возраст 2 ч 40 мин; (c) формирование синцития в одном из секторов бластодиска, возраст 2 ч 30 мин; (d) формирование синцития в центральной части бластодиска, возраст 3 ч 30 мин; (e) ослабление адгезии между бластомерами, неполные клеточные деления, возраст 1 ч 45 мин; (f) ослабление адгезии между бластомерами, десинхронизация дробления, возраст 2 ч 00 мин; (g) ослабление адгезии между бластомерами, формирование синцития, возраст 3 ч 00 мин; (h, i) ослабление адгезии между бластомерами, полное обособление клеток друг от друга, возраст 3 ч 30 мин; (j) деградация клеток на периферии бластодиска, возраст 4 ч 00 мин; (k) деградация клеток в центральной части бластодиска, возраст 4 ч 00 мин. Стрелками обозначен край перибласта.

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ing proportions of abnormally cleaved and unfertilized (with uncleaved blastodisc or without blastodisc) eggs. In the eggs from the majority of females, the proportion of normally cleaved eggs decreases substantially after in vivo storage of oocytes for 4 h. 5.2.1.5. Fertility of oocytes after their in vitro storage in ovarian fluid Following by in vitro storage (in ovarian fluid) of ovulated oocytes for 3 h at 25°C, a large part of eggs develops abnormally. In some eggs, a large number of cortical alveoli remain in 10 min after insemination (Fig. 30a), and exocytosis of the majority of cortical alveoli is observed in 1 h (Fig. 30b). Owing to desynchronization of cleavage, six cells can be seen instead of eight blastomeres (Fig. 30c). During later development, termination of divisions is registered in several cells. In other cells, the grooves of cleavage divide the cells incompletely (Fig. 30d). In 4 h after insemination, the borders between the cells of the blastodisc disappear (Fig. 30e), and a protrusion with homogenous cytoplasm has formed (Fig. 30f). Several cortical alveoli can remain intact at the stage of two blastomeres (Fig. 30g) and at following stages. At the stage of 32 blastomeres, the grooves of cleavage can divide the cells incompletely, or they are absent. Therefore, the divisions of nuclei are not accompanied by cytotomy. The drop-shaped spaces appear in the blastodisc showing onset of cell degradation (Fig. 30h). Desynchronization of cleavage leading to the formation of cells of different size and incomplete grooves of cleavage are also observed (Fig. 30i). Following by in vitro storage of oocytes at 7°C for 3h, the changes in the yolk structure are registered in abnormally cleaved eggs. These changes are accompanied by uneven coloration of yolk content. The size of the cells is substantially different, and the cells are separated from each other at the initial stages of cleavage due to total absence of reciprocal adhesion (Figs. 30j–30l). The average proportions of eggs with normal cleavage in the control series (oocytes are stripped and inseminated just after ovulation) and after in vitro storage of oocytes for 2 and 4 h at 25°C are 88% (SD 7), 79% (SD 14), and 34% (SD 29), respectively (Color Plates, Fig. 10a). The differences are significant. In vitro storage of oocytes for 2 h at 7°C leads to an abrupt decrease of their quality: the proportion of eggs exhibiting normal cleavage is only 15% (SD 22) (Color Plates, Fig. 10b). After in vitro storage of oocytes for 4 h at the same temperature, normally cleaved eggs are absent.

95

Fig. 30. Development of eggs of Zebrasoma scopas after in vitro storage of oocytes in ovarian fluid for 3 h at (a–i) 25°С and (j–l) 7°С. Temperature 25°С, egg no. 1: (a) age 10 min from insemination; (b) age 1 h; (c) age 2 h; (d) age 3 h; (e) age 4 h; (f) age 5 h. Temperature 25°С, egg no. 2: (g) 2 blastomeres, age 1 h 30 min from insemination (the image is focused to cortical alveoli); (h) 32 blastomeres, age 2 h 40 min, the arrows show the division of the cell without cytotomy and a drop-like free space in the blastosdisc. Temperature 25°С, egg no. 3: (i) formation of the syncytium, age 2 h 40 min from insemination. Temperature 7°С: (j–l) 4 h after insemination. Развитие яиц Zebrasoma scopas после выдерживания овулировавших ооцитов in vitro в овариальной жидкости в течение 3 ч при (a–i) 25°С и (j–l) 7°С. Температура 25°С, яицо № 1: (a) возраст 10 мин после осеменения; (b) возраст 1 ч; (c) возраст 2 ч; (d) возраст 3 ч; (e) возраст 4 ч; (f) возраст 5 ч. Температура 25°С, яицо № 2: (g) 2 бластомера, возраст 1 ч 30 мин после осеменения (фокус наведён на кортикальные альвеолы); (h) 32 бластомера, возраст 2 ч 40 мин, стрелки обозначают деления клеток, не сопровождающиеся цитотомией, и каплеобразные свободные пространства, формирующиеся в бластодиске. Температура 25°С, яицо № 3: (i) формирование синцития, возраст 2 ч 40 мин после осеменения. Температура 7°С: (j–l) 4 ч после осеменения.

96

5.2.1.6. Fertility of oocytes after their short-term storage in marine water The oocytes stripped from the female’s body were stored in marine water for a short time before insemination. In experiment 1, the eggs from two females were used. Average number of eggs exhibiting normal cleavage in the control series (insemination of oocytes just after their stripping) is 77%. Following by in water storage of oocytes before insemination for 5 and 10 min, the proportions of normally cleaved eggs are 63 and 41%, respectively (Color Plates, Fig. 11a). In experiment 2, the average proportion of normally cleaved eggs obtained from five females decreases from 83% in the control series to 8% after in water storage of oocytes before insemination for 30 min (Color Plates, Fig. 11b). The difference is significant (p = 0.043). After the storage of oocytes in the water before insemination for 60 and 90 min, normal cleavage is registered only in a small part of eggs from female no. 5 (19 and 7%, respectively), and it is absent in the eggs from other females. In experiment 3, ovulated oocytes of poor quality obtained from five females were used. In the control series, normally cleaved eggs are absent, and the average proportion of abnormally cleaved eggs is 80%. The storage of oocytes in the water for 40 min before insemination leads to a comparatively small decrease (on the average, at 23%) of the proportion of abnormally cleaved eggs (Color Plates, Fig. 11c). The differences in the numbers of eggs with abnormal cleavage between the series are not significant (p > 0.05). 5.2.2. Dascyllus trimaculatus 5.2.2.1. Fertility of oocytes after their in vitro storage in ovarian fluid The quality of ovulated oocytes was assessed after their in vitro storage (in ovarian fluid) at 25 and 5°C before insemination. Oocytes from two females of different quality were used in the experiment: the oocytes from female 1 were stripped immediately after ovulation, and the oocytes from female 2 were obtained in several hours after ovulation. The groups of oocytes (nine groups from each female) were placed in Petri dishes and stored in vitro for 2, 4, 6, and 8 h at 25 and 5°C. The control groups included the oocytes inseminated just after their stripping from the females (duration of in vitro storage = 0). In the control series, the proportions of normally cleaved eggs from females 1 and 2 are 85 and 35%, respectively. A decrease of these proportions is registered even after in vitro storage of ovulated oocytes for 2 h: at 25°C, up to 46 and 28%, respectively (Color Plates, Fig. 12a); and at 5°C, up to 21 and 3%, respectively (Color Plates, Fig. 12b). In general, the damage of ovulated oocytes after in vitro storage at 5°C is observed earlier than the damage after the storage at 25°C. 97

5.2.2.2. Fertility of oocytes after their short-term storage in marine water After storage of oocytes in water for 60 min before insemination, the abnormalities can be seen from the first cycles of cell division. They are as follows: a week adhesion between blastomeres, separation of several blastomeres from the main group of cells, and blockage of subsequent cleavage (Figs. 31a–31c). In several eggs (< 5%) from the experimental series, the blastodisc is represented by two separated groups of cells (Fig. 31d). In a part of inseminated but unfertilized oocytes, exocytosis of the majority of cortical alveoli is observed (all alveoli are destroyed in several eggs), and the blastodisc appears. The cleavage of the blastodisc is not registered (Fig. 31e). These oocytes belong to category 3. In another part of inseminated but unfertilized oocytes, some of cortical alveoli are subjected to exocytosis during several hours, and the perivitelline space appears in 5 h after placing of the oocyte into water; aggregation of the cytoplasm at the ani-

Fig. 31. Morphological changes in the eggs of Dascyllus trimaculatus after in vitro storage of oocytes in the water before insemination for 60 min: (a–d) abnormal cleavage, 4 h 30 min after insemination; (e) unfertilized oocyte with uncompleted cortical reaction, uncleaved blastodisc, 5 h 30 min after storage of the oocyte in the water; (f) unfertilized oocyte with partial cortical reaction, 5 h 30 min after storage of the oocyte in the water. Морфологические изменения в яйцах Dascyllus trimaculatus после выдерживания овулировавших ооцитов в морской воде в течение 60 мин перед осеменением: (a– d) аномальное дробление, 4 ч 30 мин после осеменения; (e) неоплодотворённый ооцит с неполной кортикальной реакцией, недробящийся бластодиск, 5 ч 30 мин после выдерживания ооцитов в морской воде; (f) неоплодотворённый ооцит с частичной кортикальной реакцией, 5 ч 30 мин после выдерживания ооцитов в морской воде.

98

mal pole is not observed, and the structure of the yolk becomes not homogenous (Fig. 31f). These oocytes belong to category 4. The storage of ovulated oocytes from a female in the water for 30 min before insemination does not lead to the decrease of the proportion of eggs with normal cleavage: this proportion in the control and experimental series is similar (~ 60%) (Color Plates, Fig. 13a). The storage of ovulated oocytes from another female in the water for 40 min before insemination is not accompanied by the substantial decrease of the proportion of normally cleaved eggs. A notable decrease of this proportion (from 96% in the control series to 46 and 40% in the experimental series) is registered after storage of oocytes in the water before insemination for 50 and 60 min, respectively (Color Plates, Fig. 13b). 5.2.3 Abudefduf sexfasciatus 5.2.3.1 Fertility of oocytes after their in vivo storage in the female’s ovarian cavity The eggs with normal cleavage of the blastodisc are illustrated in Figs. 32a–32c. Morphological abnormalities of the eggs at the step of cleavage after the storage of ovulated oocytes in the female’s ovarian cavity for 5 h after ovulation are as follows: uneven cleavage of the cells (Fig. 32d), a low adhesion between the cells (Fig. 32e), and indistinct cleavage grooves separating neighboring cells incompletely (Fig. 32f). These abnormalities are accompanied by a delay of the developmental rate. At the stage of smallcell morula, adhesion between cells of the blastodisc and cytoplasmic layer of the yolk is weak. As a result, epiboly of the yolk by the blastoderm does not begin (Fig. 32g). In unfertilized oocytes of A. sexfasciatus, exocytosis of only a small part of cortical alveoli occurs. As a result, the zona radiata is separated from the cytoplasmic layer of the yolk only in several hours after placing of the oocyte into water (Fig. 32h). In experiment 1, the proportion of eggs (from seven females) with normal cleavage ranges from 26 to 93% in the control group (Color Plates, Fig. 14a). These groups was designated as control (0 h after ovulation). However, comparatively low initial egg quality in several females indeed can be connected with the stripping of the first egg batch in a certain time after ovulation. Following by the storage of ovulated oocytes in the female’s ovarian cavity, the proportion of normally cleaved eggs (from females 1, 2, and 3 with the eggs of the best initial quality) decreases on the average from 87 (0 h) to 27% (5 h); and normally cleaved eggs from females 4, 6, and 7 are absent after in vivo storage of oocytes for 5 h. In experiment 2, the proportion of normally cleaved eggs for egg batches (from females 8, 99

Fig. 32. Morphological changes of the eggs of Abudefduf sexfasciatus after in vivo storage of ovulated oocytes for 5 h: (a) normal cleavage, 32 blastomeres, age 3 h 30 min from insemination (at 25°C); (b) normal cleavage, 64 blastomeres, age 4 h 00 min; (c) normal cleavage, small-cell morula, age 6 h 00 min; (d) abnormal cleavage, age 3 h 30 min; (e, f) abnormal cleavage, age 4 h 00 min; (g) abnormal development (a weak adhesion between cells of the blastodisc and the cytoplasmic layer of the yolk), age 6 h 00 min; (h) unfertilized oocyte, age 6 h 00 min. Морфологические изменения в яйцах Abudefduf sexfasciatus после задержки овулировавших ооцитов в овариальной полости самки (in vivo) в течение 5 ч: (a) нормальное дробление, 32 бластомера, возраст 3 ч 30 мин после осеменения (при 25°C); (b) нормальное дробление, 64 бластомера, возраст 4 ч 00 мин; (c) нормальное дробление, мелкоклеточная морула, возраст 6 ч 00 мин; (d) аномальное дробление, возраст 3 ч 30 мин; (e, f) аномальное дробление, возраст 4 ч 00 мин; (g) аномальное дробление (ослабление адгезии между клетками бластодиска и цитоплазматическим слоем желтка), возраст 6 ч 00 мин; (h) неоплодотворённый ооцит, возраст 6 ч 00 мин.

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9, and 10 with a good initial egg quality) in vivo stored for 0, 5, and 10 h is on average 90, 74, and 35%, respectively (Color Plates, Fig. 14b). Normally cleaved eggs from females 11 and 12 with a poor initial egg quality are absent after in vivo storage of ovulated oocytes for 5 h. 5.2.3.2. Fertility of oocytes after their in vitro storage in ovarian fluid After in vitro storage of ovulated oocytes, the morphological abnormalities at the step of cleavage are similar to those observed after in vivo oocyte storage. Assessment of fertility of ovulated oocytes of A. sexfasciatus shows that (at 25°C) a substantial decrease of the proportion of normally cleaved eggs (on the average from 97 to 59% in the control and experimental groups, respectively) is registered after in vitro storage for 4 h (Color Plates, Fig. 15a). The damage of oocytes stored in vitro at 27 and 25°C occurs at approximately the same time (after 4–5 h of the storage). Fertility of oocytes decreases faster at 5°C than at 25°C. After in vitro storage of oocytes for 2 h at 5°C, the proportion of normally cleaved eggs is on average 55%, and such eggs are absent after the storage for 4 h (Color Plates, Fig. 15b). 5.2.3.3 Fertility of oocytes after their short-term storage in marine water When the oocytes of a high quality are used (with the proportion of normally cleaved eggs 96%), a small decrease of the proportion of eggs exhibiting normal cleavage (up to 88–89%) is registered after the storage of oocytes in the water before insemination for 15–20 min (Color Plates, Fig. 16a). In another experiment, the proportion of normally cleaved eggs in the control series is 40%. The decrease of this parameter up to 19% is registered after storage of oocytes in the water before insemination for 20 min (Color Plates, Fig. 16b).

5.3. Conclusions In the majority of teleost fishes spawned in fresh water, ovulated oocytes are activated under the influence of water irrespective on the presence of spermatozoa. In non-inseminated activated eggs, the cytoplasm is concentrated at the animal pole, and the blastodisc appears. The blastodisc does not undergo divisions, or it can exhibit parthenogenetic cleavage. In several fish species, unfertilized eggs remain alive until hatching of embryos in fertilized eggs (e.g., Soin, 1953; Pavlov et al., 2004). In marine fishes, normal activation of ovulated oocytes, most often, is induced after the penetration of a spermatozoon into the micropyle. Non-inseminated oocytes of marine fishes from various ecological groups can be separated into two 101

main groups: the oocytes activated (incompletely or, sometimes, completely) in marine water with subsequent appearance of the blastodisc, which does not undergo cleavage (category 3) and the oocytes that are not activated just after their placing in marine water exhibiting partial cortical reaction during subsequent storage in the water (category 4). A presence of two oocyte groups among unfertilized eggs is reported for common wolfish (Pavlov and Moksness, 1994; Pavlov et al., 2004), Atlantic halibut (Bromage et al., 1994), Atlantic cod (Thorsen et al., 2003), and Z. scopas (Pavlov and Emel’yanova, 2008a). The dynamics of cortical reaction in non-inseminated eggs, most likely, can be used for the assessment of their potential quality: a large proportion of eggs from category 4 with partial cortical reaction indicate a poor quality of the egg batch. Cortical alveoli not subjected to exocytosis inside of the blastodisc are observed in the oocytes of rainbow trout stored in vivo (Nomura et al., 1974). The presence of cortical alveoli in some eggs (inside of the cells or beyond of the blastodisc) at initial developmental stages is reported for Atlantic cod (Kjёrsvik et al., 1990) and common wolfish (Pavlov and Moksness, 1996). The vacuoles observed inside of the intercellular spaces or at the periphery of the blastodisc of Atlantic halibut (Shields et al., 1997), most likely, are also represented by cortical alveoli. A presence of cortical alveoli outside of the blastodisc (registered in the eggs of Z. scopas with uncompleted cortical reaction) means that a narrow cytoplasmic layer is located in the sites of their localization between the yolk surface and plasmatic membrane. Thus, aggregation of the cytoplasm did not come to the end. A relationship between exocytosis of cortical alveoli and cytoplasm aggregation at the animal pole in the egg of Fundulus heteroclitus is discussed by Brummett and Dumont (1981). According to the authors, these processes are closely related to each other, and the breakage of cortical alveoli can represent a physiological stimulus leading to aggregation of the cytoplasm at the animal pole. Following by fertilization, completed cortical reaction is observed in normal oocytes of marine fishes. As is known, a certain period of time (latent period of cortical reaction) is usual between the action of activating agent and exocytosis of first cortical alveoli (Ginzburg, 1968). A delay of the breakage of cortical alveoli around the site of the contact between the spermatozoon and plasmatic membrane can stimulate penetration of the spermatozoon into the cytoplasm of the oocyte (Wolenski and Hart, 1987). A large variability of the latent period of cortical reaction in Z. scopas (from several seconds to 4.5 min), most likely, is not connected with different time of penetration of the spermatozoon into the micropyle because of high 102

sperm concentrations used in the experiments. Such a large variability of this period, most likely, can be explained by different physiological conditions of oocytes. In normal oocyte of Z. scopas, the duration of exocytosis of cortical alveoli is less than 30 s. Similar to this process in Fundulus heteroclitus (Brummett and Dumont, 1981), it is not linear: several alveoli remain intact for a comparatively long time irrespective of their localization in the oocyte in relation to the micropyle. The total duration of exocytosis of cortical alveoli in Z. scopas is lower than in Fundulus heteroclitus (40–90 s) (Brummett and Dumont, 1981) and medaka Orizias latipes (53 s) (Iwamatsu et al., 1992). Parthenogenetic cleavage is described for several freshwater and anadromous fish species (Soin, 1968), but it is observed rarely in non-inseminated eggs of marine fishes. It is registered in single eggs of Atlantic cod (Thorsen et al., 2003) and Z. scopas (Pavlov and Emel’yanova, 2008a). Thus, an occurrence of cleavage suggests that the egg is fertilized: i.e., a spermatozoon penetrated into the egg and the fusion of female and male pronuclei took place. In the previous investigations conducted on the eggs of Atlantic halibut and haddock, any deviation from the morphological picture characterized by equal blastomere size with clear intercellular borders and symmetrical position of the cells was regarded as an abnormality. A correlation between the proportion of eggs in the batch with “abnormal” development of a certain type and percentage of embryos hatched from egg envelopes was negative (Shields et al., 1997; Rideout et al., 2004). However, the attempts to study the effect of the abnormality of a certain type on survival of embryos were unsuccessful. According to Rani (2005), a correlation between the proportion of “abnormal” eggs of Atlantic cod and survival of embryos was not significant. Thus, certain deviations from normal development revealed at initial stages of cleavage not necessary lead to the morphological abnormalities during subsequent ontogeny. In Z. scopas, the abnormalities that lead to total degradation of cells of the blastodisc and 100% egg mortality are as follows (cause > consequence): (1) mitotic cleavage of nuclei accompanied by incomplete cytotomy > appearance of cells with indistinct margins; (2) mitotic divisions of nuclei with almost total absence of cytotomy > appearance of cells of different size and multinuclear syncytia; (3) a poor adhesion between cells > partial or total separation of the cells from each other; (4) desynchronization of cleavage > the number of cells is not multiple of four during the period of synchronous cleavages of cells, appearance of cells substantially ranging in size. The abnormalities of different types can be often found together that is also reported for other fish species (Shields et al., 1997; Pavlov et al., 2004; Rideout et 103

al., 2004; Rani, 2005). The abnormalities in cell divisions after in vivo and in vitro storage of oocytes, in general, are similar. However, in the latter case, the abnormalities of the types 1 and 4 can be seen from the first divisions that lead to a rapid degradation of cells. Destructive changes are registered after in vivo storage of oocytes of Z. scopas for 2 h (Pavlov and Emel’yanova, 2008a), and substantial decrease of fertility is registered after in vivo storage of oocytes of A. sexfasciatus for 5 h (Emel’yanova et al., 2009b). Thus, oocytes of marine tropical fishes with demersal eggs, most likely, are slightly more resistant to overripening in the female’s ovary than oocytes of fishes with pelagic eggs. A short period of fertility of ovulated oocytes is also reported for other warm water marine and freshwater fish species. Its duration in carp Cyprinus carpio is 1.5–2.0 h (Korovina, 1986), in silver carp Hypophthalmichthys molitrix < 1 h (Makeyeva et al., 1987), in stiped bass Morone saxatilis ~ 1 h (Stevens, 1966), and in Prochilodus marggravii ~1 h (Rizzo et al., 2003). Among the three model species, oocytes of A. sexfasciatus are the most resistant to in vitro storage. The egg quality of Z. scopas and A. sexfasciatus reduced at a higher degree after in vivo storage of oocytes than after their in vitro storage for the same time. In the experiments conducted on neotropical freshwater fish Prochilodus marggravii at 26°C (Rizzo et al., 2003), the destruction of oocytes is also registered earlier after their in vivo storage than after in vitro storage. Similar pattern is reported for European catfish Silurus glanis (Linhart and Billard, 1995). The oocytes of three model species damaged substantially slowly at a high temperature (25°C) than at a low (7°C) temperature (Pavlov and Emel’yanova, 2008a; Emel’yanova et al., 2009a, 2009b). Similar results are obtained for Prochilodus marggravii with in vitro oocyte storage at 26 and 18°C (Rizzo et al., 2003), and for vundu Heterobranchus longifilis with oocyte storage at 21–22 and 3–5°C (Nguenga et al., 2004). However, the destruction of oocytes in European catfish is more rapid after their in vitro storage at 8 and 25°C than at 19°C (Linhart and Billard, 1995). Thus, in vitro stored oocytes of teleost fishes, most likely, retain fertility for a longest period at a certain optimal temperature close to the temperature at spawning. According to Korovina (1986) and Makeyeva et al. (1987), a main reason for poor quality of oocytes after their storage in the ovarian cavity of the female is asphyxia due to the absence of connection between the oocyte and blood vessels of the theca. So-called auto-activation leading to the release of the content of a part of cortical alveoli and appearance of the perivitelline space is known for in vitro stored (in ovarian fluid or physio104

logical solution) oocytes of both freshwater and marine fishes (Stoss, 1983). This process, most likely, is a possible reason for a poor fertility of oocytes. Decreasing fertility of ovulated oocytes after their in vivo storage is accompanied by biochemical changes. Lower concentration of amino acids (Lahnsteiner, 2000) and changes in the levels of RNA messengers for 39 genes (Aegerter et al., 2005) are registered in rainbow trout, and the decrease of total energetic reserves is reported for in vivo stored oocytes of carp C. carpio (Boulekbache et al., 1989). As it is proposed (Rizzo et al., 2003), the abnormalities in the migration of organelles during the concentration of cytoplasm at the animal pole of the egg are possible due to the changes in the spatial location of microfilaments of the cytoskeleton. The changes of similar nature, most likely, represent the reasons of abnormalities in the process of mitosis at initial stages of cleavage. In particular, chromosomal abnormalities (aneuploidy, triploidy, and tetraploidy) are observed in the overripened oocytes of European catfish (Várkonyi et al., 1998). In addition, triploid prelarvae are found among the progeny obtained from overripened oocytes of rainbow trout (Aegerter and Jalabert, 2004). Similar abnormalities, most likely, occur in the eggs of three model species after in vivo or in vitro storage of oocytes at 25–27°C. In vitro storage of oocytes at a low temperature (5–7°C) leads to the appearance of abnormalities in the structure of both cytoplasm and yolk and to the blockage of penetration of spermatozoa into the majority of oocytes. Based on the results of experiments conducted on Z. scopas, D. trimaculatus, and A. sexfasciatus, the number of eggs exhibiting normal cleavage is not decreased after the storage of ovulated oocytes in marine water for 5, 40, and 15 min, respectively (Pavlov and Emel’yanova, 2008b). Pelagic eggs of Z. scopas are distributed with water current, and, therefore, they should be inseminated just after their release from the female’s body. This feature, most likely, is connected with a short time during which the oocytes retain fertility in the water. In the representatives of the family Pomacentridae deposited the eggs on the substratum, a more prolonged interval before the contact between gametes is possible, and fertilization rate can depend substantially on the duration of fertilizing ability of gametes in the water. A small number of ovulated oocytes of Z. scopas retained fertility in marine water for 90 min (see Color Plates, Fig. 11b). However, exocytosis of a large number of cortical alveoli is observed in non-inseminated oocytes by this time (Pavlov and Emel’yanova, 2008a). Therefore, the penetration of a spermatozoon into the cytoplasm of the oocyte and fertilization of a small part of oocytes, most likely, is possible after the beginning of cortical 105

reaction and separation of zona radiata from the cytoplasmic membrane of the oocyte. According to Tavolga (1950), swelling of the oocyte in marine fish Bathygobius soporator is completed in 20 min after its placing into water, but the egg can be fertilized for 3 h after onset of swelling. Based on the opinion of Laale (1980), egg swelling in this species is not accompanied by the closure of the micropylar canal, and perivitelline fluid is not characterized by agglutinating properties. Even in several freshwater fish species, oocytes placed in the water retain fertility for several minutes. For example, fertilization of a part of oocytes (>10%) of European catfish is possible after storage of oocytes in fresh water for 4 min (Linhart and Billard, 1995). Thus, the mechanisms of blockage of penetration of excess of sperm into the eggs of teleost fishes require further investigations. Our results suggest that in studied fish species fusion of gametes can occur in several minutes after their release into water. A probability of the penetration of a spermatozoon into the micropyle and subsequent fertilization increases with the increase of the duration of a contact between gametes that is supported in the experiments on the insemination of the eggs in starred sturgeon Acipenser stellatus, lacustrine brown trout (Ginzburg, 1968), common wolfish (Pavlov, 1994; Pavlov et al., 2004), and turbot Psetta maxima (Suquet et al., 1995). Therefore, the duration of a contact between gametes should be taken into account for the estimation of an optimal sperm to egg ratio. According to “dry” method of insemination, the eggs are mixed with sperm, and the activity of spermatozoa is induced by means of addition of water (Ginzburg, 1968). A modification of this method for the fish spawned in fresh water is the use of salt solutions to enlarge duration of sperm motility and delay cortical reaction in oocytes (Billard, 1988). Gametes of marine fishes retain fertility in marine water for at least several minutes. Therefore, the dry method of insemination can not be effective in many cases. It can lead to damage of the oocytes during their mixing with sperm and to ineffective expenditures of sperm. In fishes with adhesive eggs, this method can lead to the appearance of large conglomerates of eggs in which they can die during incubation due to a poor gas exchange inside of the clutch. To inseminate large egg batches, ovulated oocytes of marine fishes can be placed in the vessel with marine water mixed with sperm. Owing to a long period (several minutes) of gamete fertility in marine water, the addition of sperm after placing of oocytes into water is also possible. For insemination of large numbers (several thousand) of adhesive oocytes of the representatives of the family Pomacentridae, they should be dispersed proportionally on the pieces of gauze placed in the vessel with marine water mixed with sperm. 106

Chapter 6. Development of eggs and transition of larvae to exogenous feeding 6.1. Culture of marine fish species: a brief review The first attempt of artificial reproduction of marine fish was conducted by Norwegian former ship’s officer G.M. Dannevig who organized a cod hatchery in southern Norway in 1880s. He established a broodstock, transferred the fish to a spawning basin, collected fertilized eggs for their incubation in the laboratory, and the yolk-sac larvae were released to the sea. The fate of these larvae remained unknown, and further progress in marine fish culture was connected with a discovery (in 1960s) of brackish water rotifer Brachionus plicatilis as an effective live food for marine fish larvae. However, significant survival of the larvae was achieved later (in late 1970s) when the important role of microalgae and highly unsaturated n-3 fatty acids (n3 HUFA) in the diet was proved. The production of viable juveniles is still the main constraint for culture of marine fish species. Three cultivation concepts can be defined (Olsen et al., 2004): larval feeding in (1) large closed natural-like systems; (2) relatively large mesocosms, enclosures, or outdoor tanks; and (3) relatively small tanks. The intensive rearing method (concept 3) represents the current international standard for marine larvae rearing. The challenges of developing suitable technology to rear marine fish are multidisciplinary. The production chain involves several steps, and the knowledge about the following fundamental issues is required: (1) general biology of fish species; (2) chemical and physical environmental requirements; (3) nutritional requirements; (4) requirements to microbial composition of the environment. The processes of water treatment and hatchery operation are in general similar for both food and ornamental fishes (Fig. 33). However, virtually all hatcheries will have their own procedures and infrastructure characteristics, and additional lines for production of special live feed can be added for some ornamental fish species. Marine fish larvae are very fragile, and they are sensitive to physical disruption, especially at early developmental stages. Therefore they are maintained in stagnant or semi-stagnant conditions in the early feeding stage. The size of the tanks used for rearing is also important: small tanks can not be suitable due to damage of larvae as a result of their mechanical contacts 107

with the walls; in addition, water characteristics are usually more stable in larger tanks. The larvae of many marine fish species normally ingest microalgae before they start feeding on bigger prey. The addition of algae has an obvious impact on the water quality, light environment, and the contrast between background color and prey color. It is still discussed whether the uptake of microalgae is active or simply a consequence of water-drinking activity. During the early feeding period larvae, most likely, act as filter feeders. Larger organisms are actively consumed, and a sequence of feeding objects from rotifers to Artemia nauplii, juvenile Artemia, and formulated feed is usual for intensive culture of marine fish larvae. The optimum food concentration is a very important factor for each species: at low prey densities larvae can not find food organisms, and at high prey concentrations, the larval digestion efficiency is reduced and defecation rate increases that can substantially reduce water quality.

Fig. 33. Schematic illustration of water processing and production process lines during intensive rearing of marine fish larvae (Olsen at al., 2004). Схема, иллюстрирующая подготовку воды и производственные линии необходимые для интенсивного культивирования морских рыб (Olsen at al., 2004).

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Feeding protocol for each marine fish species is unique, and it depends much on the size of start-feeding larva and the size of its mouth. The size of larvae of the present or potential objects for aquaculture can differ up to ten times (Fig. 34). The larvae of some marine species (e.g., wolfish) can be directly transferred to formulated feed just after start feeding. However, the size of start feeding larvae is much smaller, and the morphological condi-

Fig. 34. Relative size of the larvae at the transition to exogenous feeding of several fish species considered as present or possible objects for aquaculture: (a) scopas tang Zebrasoma scopas, (b) scissortail sergeant Abudefduf sexfasciatus, (c) Atlantic cod Gadus morhua, (d) common wolffish Anarhichas lupus, (e) Arctic charr, dwarf form Salvelinus alpinus. The drawings are represented from the following papers: Makhotin et al., 1984; Shadrin et al., 2003; Kjёrsvik et al., 2004; Pavlov and Osinov, 2008. Размер личинок некоторых видов рыб – существующих или возможных объектов аквакультуры – при переходе на экзогенное питание: (a) Zebrasoma scopas, (b) Abudefduf sexfasciatus, (c) треска Gadus morhua, (d) зубатка Anarhichas lupus, (e) арктический голец, карликовая форма Salvelinus alpinus. Иллюстрации заимствованы из следующих источников: Makhotin et al., 1984; Shadrin et al., 2003; Kjёrsvik et al., 2004; Pavlov and Osinov, 2008.

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tion at the transition to exogenous feeding is substantially less advanced in the majority of marine fish species, especially in ornamental coral fishes. It is the reason why they can consume only special live organisms of a small size (less then 100 µm) making the whole technology extremely complex and expensive. In addition, special incubation systems should be designed for certain developmental stages, as it is done for yolk-sac larvae of Atlantic halibut (Fig. 35). However, a complex technology can be feasible due to high prices of ornamental fishes at the market.

Fig. 35. Incubator for Atlantic halibut Hippoglossus hippoglossus yolk-sac larvae (2.8 m3 in voume). The seawater inlet is from the lowest valve under the cone. Low-salinity water is added through the orifices in the vertically mounted pipeline to create a continuous salinity gradient from the lowest part of the inner pipeline to the surface. An outlet sieve is located close to the surface, and the second valve at the bottom is for taking out dead larvae (Holm et al., 2004). Система для содержания предличинок атлантического палтуса Hippoglossus hippoglossus (объём 2.8 м3). Морская вода подаётся из нижнего патрубка, расположенного под конической частью. Вода низкой солёности подаётся из отверстий патрубка, установленного вертикально, для создания постоянного градиента солёности от нижней конической части до поверхности воды. Слив воды, защищённый мелким газом, установлен близко к поверхности воды; второй патрубок, расположенный в конической части, служит для удаления мёртвых предличинок (Holm et al., 2004).

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The following main problems should be solved for successful rearing of the larvae: (1) special design of the tanks including the systems of water supply and oxygenation; (2) optimum concentration of microalgae in the tanks; (3) optimum time for the beginning of feeding; (4) species composition of live feed (including organisms of natural microzooplankton, Brachionus sp., and Artemia) and the optimum size of prey for the larvae of different age; (5) optimum densities of the larvae and their prey and change of these parameters with age; (6) retention time of the rotifers in the larval tanks, in particular at the early stages of first feeding; (7) effect of the methods of n-3 HUFA enrichment of rotifers on the survival and growth of the larvae.

6.2. Culture of coral reef fish species The experiments were conducted at the Mariculture Station of the Institute of Aquaculture no. 3 (Nha Trang, Vietnam). The marine water intake of the facility located at a distance of 40 m from the shore at a depth of 20 m from the water surface. The water was subjected to mechanical cleaning and UV sterilization. In addition, a biofilter 2 m3 in volume with crumbled corals was installed in the laboratory. The laboratory included four tanks (each 3m3 in volume) for spawners and eight tanks (each 0.3 m3 in volume) for eggs and larvae. Several larger tanks (1–2 m3 in volume) with upwelling water flow were also tested for rearing of the larvae. The water temperature ranged from 25.0 to 27.5°С. Larvae were reared at combined illumination: natural light (from the windows of the laboratory) and continuous light from fluorescent lamps installed over each tank at a distance of 1 m from the water surface. Approximately up to 20% of water was replaced in each tank daily. The air diffusers with small pores and a weak air pressure were used. Oxygen concentration in the tanks with larvae exceeded 5.0 mg/l, but sometimes it dropped up to 4.5 mg/l. The “green water” technique (using microalgae Nannochloropsis oculata) was applied. The size of the alga cell was from 2.4 to 3.2 µm, on average 2.8 µm. The color of the water in the tanks was light green. The larvae were fed on rotifers Brachionus plicatilis cultured on microalgae and yeast and enriched by Selco. In addition, the larvae were fed on natural zooplankton collected with plankton net, mesh size 20 µm. The 111

length of a rotifer ranged from 122 to 272 µm (the width twice as low as the length), and the length of crustaceans from zooplankton ranged from 144 to 412 µm. Before feeding of larvae, rotifers and zooplankton were filtered through gauze, mesh size 130 µm. The illustrations of embryonic and larval development were made on live individuals using the analogous and digital cameras. 6.2.1. Zebrasoma scopas In total, the eggs from 14 females (n ~ 180 000) were used for the experiments. For artificial insemination, ovulated oocytes were placed into trays with marine water and diluted sperm. The eggs were washed from sperm in 5 min after insemination and placed in the tanks with stagnant water. Initial density of larvae was up to four individuals per liter. Concentration of feeding organisms in the water ranged from 5 to 25 exemplars/ml in different experiments. As far as we know, embryonic and larval development of Z. scopas has not been described, but a newly hatched larva and a larva at the age of a day after hatching are illustrated by Shadrin at al. (2003). Embryonic and larval development is represented in Fig. 36 and Color Plates, Fig. 17. The oocytes of a female are transparent, of regular rounded shape, 0.56–0.67 mm in diameter, on average, 0.63 mm (SD 0.03, n = 16). Diameter of the egg after swelling is 0.61–0.69 mm, on average, 0.65 mm (SD 0.02, n = 40). Average diameter of the lipid droplet is 0.16 mm. The micropyle is seen at the surface of the egg envelope, and cortical alveoli are located in the cytoplasmic layer of the oocyte. Diameter of a cortical alveolus ranges from 1.1 to 5.5 µm, on average, 2.9 µm (SD 0.8, n = 200). Mass hatching of the embryos (on average, 1.5 mm TL) is registered at the age 22–23 and 27 h after insemination at 27 and 25°C, respectively. In three Fig. 36. Embryonic development of Zebrasoma scopas at 25°C. (a) Egg swelling, age 40 min from insemination; (b, c) 2 blastomeres, age 1 h 20 min; (d) 4 blastomeres, age 1 h 40 min; (e) 8 blastomeres, age 2 h 00 min; (f) 16 blastomeres, age 2 h 20 min; (g) 32 blastomeres, age 2 h 40 min; (h) 64 blastomeres, age 3 h 00 min; (i) middle-cell morula, the periblast layer is outside of the blastodisc, age 3 h 30 min; (j) small-cell morula, age 6 h 00 min; (k) beginning of gastrulation, age 9 h 00 min; (l) formation of the embryonic ring and embryonic shield, age 9 h 30 min; (m) beginning of organogenesis, epiboly of the blastoderm 70% of the yolk surface, age 11 h 40 min; (n) eye bubbles, closure of the yolk plug, 14 h 00 min; (o) first somites in the body of the embryo, age 16 h 00 min; (p) 6 somites, age 18 h 00 min; (q) 28 muscular segments in the body, otoliths in the auditory capsules, weak pulsations of the heart tubule, age 23 h; (r) first melanophores on the head, movements of the embryo, age 24 h. The arrows indicate the periblast layer at morula stages, otoliths in the auditory capsule, and the heart tubule.

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Эмбриональное развитие Zebrasoma scopas при 25°C. (a) Набухание яйца, возраст 40 мин после осеменения; (b, c) 2 бластомера, возраст 1 ч 20 мин; (d) 4 бластомера, возраст 1 ч 40 мин; (e) 8 бластомеров, возраст 2 ч 00 мин; (f) 16 бластомеров, возраст 2 ч 20 мин; (g) 32 бластомера, возраст 2 ч 40 мин; (h) 64 бластомера, возраст 3 ч 00 мин; (i) морула среднеразмерных клеток, граница перибласта за пределами бластодиска, возраст 3 ч 30 мин; (j) мелкоклеточная морула, возраст 6 ч 00 мин; (k) начало гаструляции, возраст 9 ч 00 мин; (l) формирование зародышевого кольца и зародышевого щитка, возраст 9 ч 30 мин; (m) начало органогенеза, эпиболия бластодермы 70% поверхности желтка, возраст 11 ч 40 мин; (n) глазные пузыри, замыкание желточной пробки, возраст 14 ч 00 мин; (o) первые сомиты в теле эмбриона, возраст 18 ч 00 мин; (q) 28 миомеров, отолиты в слуховых капсулах, слабая пульсация сердечной трубки, возраст 23 ч; (r) первые меланофоры на голове, движения эмбриона, возраст 24 ч. Стрелками обозначены граница перибласта на стадии морулы, отолиты в слуховой капсуле и сердечная трубка.

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days from hatching (at 27°C), zooplankton and rotifers were placed into the tanks with the larvae. By this time, the jaws of the larva are slightly mobile, the yolk is resorbed, and intestinal peristalsis is registered. The food objects, which are slightly larger than algae cells of “green water” (most likely, the largest cells of the algae), are seen in the intestines of some larvae. By the age four days, the larvae possess a strong positive reaction to light. They are concentrated near the water surface, and the darts of the larvae to prey are observed. The feeding items approximately 50 µm in size and smallest rotifers are observed in the intestines of some larvae. Mass mortality of the larvae was registered by the age five days from hatching. It was caused by a low concentration of available food and oxygen deficit. 6.2.2. Dascyllus trimaculatus In addition to comparatively small tanks (0.3 m3 in volume) used for rearing of the larvae, two tanks with recirculated water supply (1 and 3m3 in volume) were modified to obtain proportional water flow from the bottom to the surface. The air diffusers with small pores and a weak air pressure were applied in small tanks. In the large tanks, aeration of the water was not used to prevent mechanical damage of larvae, and water exchange rate was 1.0–1.5 l/min. In total, the eggs from 15 females (n ~ 180 000) were used for incubation and subsequent rearing of the larvae. Almost all eggs were obtained after stripping of the females subjected to hormonal stimulation. However, natural spawning was observed in several females injected previously by surfagon and kept in the tanks together with the males (5–7 females + 2 males in a tank). Several females deposited the eggs on the walls of the flowerpots installed in the tanks, and the eggs were fertilized. The time of natural spawning was similar to the time of stripping of ovulated oocytes from other injected females. For artificial insemination, ovulated oocytes obtained by the method of hormonal stimulation were placed into a syringe with the opening 2–4 mm in diameter, and then they were proportionally distributed on the pieces of gauze (mesh size 0.5–1.0 mm) in a vessel with marine water (2 l in volume) and diluted sperm (obtained from crashed testes of the males). In 10 min after artificial insemination, the pieces of gauze with attached eggs were hanged in a tank with a comparatively large water exchange rate (4 l/min) and intensive aeration (Color Plates, Fig. 18). The keeping of eggs of fishes from the family Pomacentridae in darkness stimulates hatching of embryos (e.g., Olivotto et al., 2003). Therefore, the tanks were covered by a dark polyethylene film several hours before expected hatching. The initial density of larvae in the small tanks was up to four individuals per liter, and 114

Fig. 37. Embryonic development of Dascyllus trimaculatus at 27°C. (a) Egg swelling, cytoplasmic disc at the animal pole, age 40 min from insemination; (b) 2 blastomeres, age 1 h 15 min; (c) 4 blastomeres, age 1 h 50 min; (d) 32 blastomeres, age 4 h 30 min; (e) middle-cell morula, age 5 h 30 min; (f) small-cell morula, age 9 h 30 min; (g) beginning of gastrulation, epiboly of the blastoderm 30% of the yolk surface, age 10 h 40 min; (h) segmentation of the body, large melanophores on the preanal and postanal parts of the body, age 22 h; (i) appearance of the crystalline lenses in the eye vesicles, the yolk sac is fully surrounded by the body of the embryo, turnover of the embryo, age 1 day 2 h; (j) intensive movements of the embryo, age 1 day 8 h; (k) enlarged melanophores on the head and in the ventral row, age 2 days 4 h; (l) substantial resorption of the yolk, stage before hatching, age 3 days 0 h. Эмбриональное развитие Dascyllus trimaculatus при 27°C. (a) Набухание яйца, цитоплазматический диск на анимальном полюсе, возраст 40 мин после осеменения; (b) 2 бластомера, возраст 1 ч 15 мин; (c) 4 бластомера, возраст 1 ч 50 мин; (d) 32 бластомера, возраст 4 ч 30 мин; (e) морула среднеразмерных клеток, возраст 5 ч 30 мин; (f) мелкоклеточная морула, возраст 9 ч 30 мин; (g) начало гаструляции, эпиболия желтка бластодермой на 30% его поверхности, возраст 10 ч 40 мин; (h) формирование сомитов, крупные меланофоры в преанальной и постанальной частях тела, возраст 22 ч; (i) появление хрусталиков в глазных пузырях, желточный мешок полностью окружён телом эмбриона, поворот зародыша, возраст 1 сут. 2 ч; (j) интенсивные перемещения эмбриона, возраст 1 сут. 8 ч; (k) крупные меланофоры на голове и в вентральном ряду, возраст 2 сут. 4 ч; (l) значительная резорбция желтка, стадия перед вылуплением, возраст 3 сут. 0 ч.

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it reached 40 individuals per liter in the large tanks. Feeding of larvae by zooplankton and rotifers started on the second day after hatching. Concentration of feeding organisms in the water ranged from 5 to 25 exemplars/ml in different experiments. To our knowledge, the embryonic and larval development of D. trimaculatus has not been described before. Embryonic development of D. trimaculatus is illustrated in Fig. 37. The diameter of ovulated oocyte is 0.55–0.75 mm, and the cells 0.60–0.70 mm in diameter prevail; average diameter of the lipid droplet is 0.19 mm. A cluster of adhesive filaments is located at the animal pole. At the stage of late organogenesis, the embryo completes the turnover process when the head is turned from facing the attachment end (animal pole) to the free end of the egg envelope (vegetative pole) (Figs. 37h, 37i). This process is described in other species of the family Pomacentridae (Olivotto et al., 2003; Yasir and Quin, 2007; Shadrin and Emel’yanova, 2007). At 27°C, mass hatching of embryos is observed in 78–82 h after insemination of the eggs. The average total length of a newly hatched embryo is 2.1 mm, the diameter of the yolk sac is approximately 0.29 mm, and the diameter of the lipid droplet is 0.15 mm (Color Plates, Fig. 19a). The numbers of preanal and postanal segments are 8 and 24, respectively. Melanophores are seen along the dorsal and ventral parts of the body. Free embryos are distributed in the water column and possess a weak positive phototaxis. At the age 1 day after hatching, the yolk sac is almost resorbed, the lipid droplet is still seen, the depth of the dorsal part of the fin fold (most likely, having a hydrostatic function) increases, and pulsations of the heart locating at the horizontal plane inside of the pericardial cavity are observed (Color Plates, Fig. 19b). By the age 2 days after hatching, the free embryos 2.2–2.3 mm TL are ready to exogenous feeding (Color Plates, Fig. 19c). At this stage, the mouth has a terminal position, and the lipid droplet is almost totally resorbed. The average gape (the distance between the upper and lower jaws at maximum opening of the mouth) of the larva is 109 µm. By the age 4 days, the lower jaw of the larvae slightly protrudes in the anterior direction, differentiation of the digestive system continues (Color Plates, Fig. 19d), and the width of the fin fold is substantially increased (Color Plates, Fig. 19e). By the age 6–8 days, an aggregation of melanophores is seen above the anterior part of the intestine in the area of the liver (Color Plates, Figs. 19f, 19g). On the dorsal part of the body, melanophores are small with a sparse distribution; but melanophores in the ventral row are larger. The food objects up to 150 µm in length are found in the intestines of the larvae. The larvae are able to fast swimming and abrupt darts. By the age 10 days, comparative size of 116

the head increases (Color Plates, Fig. 19h). By the age 15 days, the muscular buds are seen in the anlages of the dorsal and anal fins of the larva more than 3 mm TL, and the rays appear in the anlage of the caudal fin (Color Plates, Fig. 19i). Black pigment is distributed mainly in the area of the opercular plate, and the black spots are seen on the flanks over the lateral line. Large melanophores are located in the ventral postanal part of the body and in the ventral part of the caudal fin. The larvae are still distributed in the water column. The proportion of eggs (obtained from different females) with normal cleavage ranged from 70 to 95%; and the proportion of free embryos after hatching was 90% from the number of normally developing eggs. Mass mortality of larvae registered at the age of 4–6 days after hatching was connected with the problem of their transition to exogenous feeding: owing to a large size of prey, only a small part of the smallest rotifers and crustaceans was available. Only single larvae survived to the age 15 days. 6.2.3. Abudefduf sexfasciatus The eggs from ten females (n ~ 40 000) were obtained by the method of hormonal stimulation. The conditions of egg incubation and rearing of larvae were similar to those described above for D. trimaculatus. During ongrowing of larvae, the water exchange rate in the tanks 0.3 m3 in volume ranged from 0 to 100 ml/min, and average water temperature was 27.0°С. Initial stocking density of larvae was 10 exemplars/l. The start feeding of larvae began on the second day after hatching of embryos. Concentration of feeding organisms in the water ranged from 8 to 45 exemplars/ml in different experiments. Ovulated oocytes are 0.75–1.10 mm in diameter, more often, 0.90–1.00 mm. Embryonic development is described by Shadrin and Emel’yanova (2007). Several stages of embryonic development are illustrated in Color Plates, Fig. 20. Hatching of embryos is registered in 112–126 h after insemination of eggs. Newly hatched larva is on average 2.60 mm TL (Color Plates, Fig. 21a). Preanal length is 31.5% TL. The larva has 8 preanal and 21 postanal segments. The same number of segments is registered in A. sexfasciatus by Shadrin and Emel’yanova (2007). The yolk sac (0.22 x 0.27 mm) possesses a crimson coloration (similar to that in ovulated eggs), and the lipid droplet (0.11 mm in diameter) is yellow. Large melanophores are located behind the ventral part of the auditory capsule and in the dorsal part of the intestine, several large melanophores are distributed on the yolk sac, and two 117

melanophores are located on the anterior part of the head in front of the eyes. Small melanophores are seen in the preanal dorsal part of the body and along the ventral row of the postanal part. A transversal yellow band (formed by xanthophores) approximately 0.16 mm in width is seen behind the auditory capsule from the dorsal part of the body to the upper part of the intestine. The jaws are immobile. The free embryos are concentrated in the upper water layer and possess a weak positive phototaxis. At the age of 1 day from hatching, the yolk sac is almost totally resorbed, and movements of the jaws are registered. By the age 2 days, the average length of the larva reaches 2.63 mm (Color Plates, Figs. 21b, 21c). The length of the jaws is substantially increased. A pigmented stripe from comparatively large melanophores is formed behind the eyes along the anterior margins of the body cavity. The melanophores of the ventral row (n ~ 21) have become larger. The larvae are ready to the transition to exogenous feeding: rhythmic movements of branchial-jaw apparatus can be seen, the yolk sac is totally resorbed, and the size of the lipid droplet is substantially reduced. By the age 3 days, the swim bladder is filled by air, and the larvae are able to swim both near the water surface (as it is observed just after hatching) and in the water column. By the age 4–6 days, the lower jaw becomes longer (Color Plates, Figs. 21d–21g). The transversal yellow band is distributed up to the dorsal part of the fin fold. By the age 11 days, the relative length of the postanal part of the body becomes shorter: the length of the preanal part of the body reaches 34.5% TL (Color Plates, Fig. 21h). The anlages of the pelvic fins are seen in the ventral part of the body behind the dark pigmented stripe along the anterior margin of the body cavity. The number of melanophores in the ventral row of the postanal part of the body has become smaller reaching approximately 15 cells. Within the interval of development between the 11th and 15th days after hatching, an abrupt increase of the growth rate of the larva (from 2.93 to 4.38 mm TL) is registered. This increase is accompanied by substantial morphological changes. By the age 15 days, the postanal part of the body has become shorter relative to the body length, and the preanal part reaches 44.5% TL (Color Plates, Fig. 21i). At the same time, the numbers of preanal and postanal myomeres (8 + 21) is similar to the number of segments in the newly hatched larva. In the caudal peduncle, the notochord is straight, but onset of its upward flexion is registered in several larvae. The body has become substantially deeper. The anlages of the dorsal, anal, and caudal fins are separated from the fin fold, and the first rays are seen in the anterior part of the dorsal fin and in the caudal fin. The black pelvic fins have 118

become substantially longer protruding behind the anus. Pigmentation of the preanal part of the body by melanophores and xanthophores (from the anterior margin of the preoperculum to the posterior border of the body cavity) is very intensive. The walls of the body cavity possess an intensive dark coloration. However, the anterior part of the head and the postanal part of the body are almost free from pigment, with the exclusion of nine or ten melanophores in the ventral row. By the age 18 days at 4.75 mm TL, relative length of the preanal part of the body increases reaching 47.2% TL (Color Plates, Figs. 21j, 21k). The spined rays (10 from 13 in adult individuals) and all soft rays (12) are seen in the dorsal fin. All spined rays (2) and soft rays (12) are observed in the anal fin. The number of rays in the caudal fin is 19. In the caudal peduncle, the notochord upward flexion is completed. The black pelvic fins reach the spined rays of the anal fin. Pigmentation is almost exclusively restricted to the preanal part of the body behind the eyes. Several small melanophores are located on the head, and up to ten comparatively small melanophores remain in the ventral row of the postanal part of the body. A black pigmented stripe surrounded by orange pigment is seen along each of the first seven or eight spined rays of the dorsal fin. The larvae are still distributed in the water column. Thus, the coloration of the larvae almost reaching the definitive morphology substantially differs from that in adult exemplars. Almost total absence of pigment in the anterior part of the head and in the postanal part of the body of the larva, most likely, is used for camouflage on the background of the coral reef and for disorientation of predators. Similar morphological changes are described in the larvae of A. saxatilis raised in the laboratory (Alshuth et al., 1998). In this species, onset of notochord flexion is observed at 5.3 mm TL (vs. 4.4 mm TL in A. sexfasciatus). Following by the completion of the notochord flexion in A. saxatilis, an intensive development of pigmentation in the postanal part of the body is registered, and pigmentation of the juveniles becomes similar to that in adult individuals by 19 mm TL The proportion of hatched embryos of A. sexfasciatus reached on average 90% from the number of eggs with normal development. (This number ranged from 50 to 95% in the egg batches from different females.) Mass mortality of the larvae was registered at the age 4–8 days, and single larvae survived to the age 18 days. The mortality was connected mainly with the discrepancy between the gape size of the larva and the size of food organisms. The average gape size of the larva is 133 µm, and the width of its intestine at the age 7 days is approximately 70 µm (Color Plates, Fig. 21l). In the intestines of several larvae of this age, the food objects 40 µm in size 119

are seen. The size of food organisms used for start feeding of the larvae (rotifers 122–272 µm in length and crustaceans from zooplankton mainly more than 144 µm in length), most likely, is substantially larger than the optimal size of prey for consumption.

6.3. Conclusions The experiments on the raising of larvae of Zebrasoma scopas (Acanthuridae), Dascyllus trimaculatus, and Abudefduf sexfasciatus (Pomacentridae) showed that the main constraints were connected with unfavorable environmental conditions and none-adequate size of organisms used for start feeding. The former problem is especially important for the larvae of coral fish species, which should be reared at a high temperature and at large concentrations of microalgae and prey items. At the same time, both water exchange rate and intensity of aeration should not be large to prevent damage of fragile larvae. Thus, water quality (including oxygen content) can reduce rapidly during start feeding of larvae. This problem can be solved using large upwelling systems with recirculated water and proportional water flow. The water should be cleaned and oxygenated before inflowing to the tank. The latter problem is crucial for coral reef fish species with extremely small larvae. The mouth size in the larvae of three model species is represented in Fig. 38. The optimum size of prey items should be substantially lower than the mouth gape. The organisms of optimal size (40–60 µm for Z. scopas and 60–100 µm for D. trimaculatus and A. sexfasciatus) were almost totally absent in the experimental tanks that represented a main reason for a large larval mortality. Our attempts to start feed of the larvae on protozoans failed apparently due to a poor nutritional value of these organisms and their low concentration in the water of the tanks. Rotifers and Artemia nauplii are usually applied for start feeding of larvae of coral fishes of the genus Amphiprion reaching up to 6.7 mm TL at hatching (Durville et al., 2004). The organisms of smaller size are necessary for smaller larvae of other species. For example, natural zooplankton (63–100 µm) was used as a start food in the experiments on rearing of the larvae of Dascyllus albisella and D. aruanus (Danilowicz and Brown, 1992). In one of these experiments, survival of the young of the former species reached 41% in 10 weeks after hatching. In our experiments, large numbers of developing eggs of ornamental coral reef fishes were obtained for the first time using the method of hormonal therapy. The larvae of D. trimaculatus and A. sexfasciatus were raised until 120

the age of 15 and 18 days from hatching, respectively; and they reached a morphological condition similar to that in the juveniles. In addition, a possibility of natural spawning of D. trimaculatus after hormonal stimulation is shown. In natural habitats, first feeding marine larvae usually feed on a wide variety of microzooplankton with a very high content of HUFA (Riley and Holt, 1993). Thus, the collection of natural microzooplankton for start feeding of coral fish larvae is a possible way to solve the problem of their transition to exogenous feeding. Some other solutions are suggested recently. The attempts on the raising of larvae of marine ornamental fishes using small prey items are conducted, and protozoans (Tintinnoidea, Cilia-

Fig. 38. Mouth gape at the transition to exogenous feeding in (a) Zebrasoma scopas, (b) Dascyllus trimaculatus, and (c) Abudefduf sexfasciatus. Total lengths (TL) at hatching and at the transition to exogenous feeding: (a) 1.5 and 1.8 mm; (b) 2.1 and 2.3 mm; (c) 2.6 and 2.7 mm. Размер рта при переходе на экзогенное питание у (a) Zebrasoma scopas, (b) Dascyllus trimaculatus и (c) Abudefduf sexfasciatus. Общая длина (TL) при вылуплении и при переходе на экзогенное питание: (a) 1.5 и 1.8 мм; (b) 2.1 и 2.3 мм; (c) 2.6 и 2.7 мм.

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ta), dinoflagellates, and especially copepods eggs and nauplii are tested (Olivotto et al., 2005, 2006, 2008, 2011). For example, different strains of wild and cultured zooplankton were evaluated to find a suitable food for rearing the larvae of lemonpeel angelfish Centropyge flavissimus (Pomacanthidae) (Olivotto et al., 2006). Foods selected for evaluation were some of the plankton species that typically represent first food for fish larvae: copepod nauplii Acartia tonsa (70 x 90 µm), Parvocalanus spp. (45 × 70 µm), and dinoflagellate Oxyrrhis marina. Copepod stock cultures of A. tonsa and Parvocalanus spp. originally isolated from plankton collections were maintained in a temperature-controlled room and were fed daily with the microalgae Isochrysis galbana. O. marina was cultured in 25 l tanks on I. galbana. Ten percent of the larvae (2.3 mm TL at hatching) survived for 14 days only when fed wild plankton plus O. marina or Parvocalanus sp. nauplii. For the first feeding of cleaner goby Gobiosoma evelynae (Gobiidae), Euplotes sp. naked ciliates (average size 100 x 85 µm) were used together with two different rotifer species (Brachionus plicatilis and B. rotundiformis) and super small Artemia sp. nauplii (Olivotto et al., 2005). The ciliates were cultured on Nannochloropsis oculata applying the same techniques used for rotifers. A highest juveniles survival rate (50%) for 40 days after hatching was observed in larvae initially fed on Euplotes sp. ciliates and successively on B. rotundiformis rotifers showing the importance of naked ciliates as first food for larval rearing. The study on the culture of longsnout seahorse Hippocampus reidi (Syngnathidae) demonstrated the potential of harpacticoid copepod Tisbe spp. as a supplemental food to the traditional diet based on rotifers and Artemia (Olivotto et al., 2008, 2011). Thus, both size and species compositions of the feeding organisms are important for improving the larval survival of tropical fish. The larvae of certain molluscs are also attractive as a first feed for brackish water and marine fish larvae because of their availability and small size. The most commonly used are the trochophore larvae of oysters and mussels. Concluding remarks Aquaculture is considered as a potential alternative to capture of ornamental coral fishes from nature, and the production in intensive systems of some of the most heavily collected representatives of the species would certainly contribute to relieving the current fishing pressure on coral reefs. At present, natural spawning in captivity is almost exclusively applied to obtain offspring of marine ornamental fishes. For the beginning of natural reproduction, a prolonged adaptation of the fish to artificial conditions is 122

required using complex and expensive systems with water recirculation and control of environmental parameters. The spawning of different females, as a rule, is not synchronous that leads to obtaining of small egg batches in different time. It is another reason that constrains economically feasible production of offspring. Hormonal stimulation of maturation is an alternative method for obtaining mature sex products in ornamental fish species. Hormonal injections can be applied just after delivering of the wild-caught fish to the laboratory. Therefore the eggs can be inseminated and the larvae can be obtained within a few days without substantial expenses to the keeping of the spawners at controlled environment. The fish subjected to hormonal therapy can be returned to the sea after stripping of eggs and sperm or can be kept in artificial conditions until subsequent natural spawning or repeated hormonal stimulation. Large amounts of eggs and larvae suitable for rearing in industrial cultivation systems can be obtained all year round. Thus, the use of the method will make the process of artificial breeding of the fish feasible. In addition, many ornamental fish species (including the species which command extremely high prices) are poor adapted to artificial systems (Wabnitz et al., 2003), and hormonal therapy is the only way to obtain mature sex products. A possibility of the application of the methods of hormonal stimulation for the fishes with both continuous and discontinuous types of oocyte maturation is shown in this study. The schemes of hormonal therapy can be different for the fishes from various taxonomic groups characterized by specific patterns of oocyte development. Such characteristics, as the type of oogenesis, diameter of oocytes with maturational competence, period of reaching of this competence after the spawning, and duration of reproductive season should be determined. The assessment of oocyte condition both before and after the injection should be conducted for the development of the optimal scheme. The importance of this assessment is supported in the experiment with Siganus spinus: the third injection has been applied to obtain ovulated oocytes. The assessment of a possibility of the application of single injections instead of double injections represents an important task: single injections reduce handling and simplify the method. This possibility, most likely, is connected with the specificity of endocrinological sensibility and features of oogenesis. For example, Zebrasoma scopas can produce ovulated oocytes only after double injections, but single injections can be used for some representatives from the families Pomacentridae and Monacanthidae. The injections should be conducted during the first or second day after the capture of the fish: keeping of the fish without food for at least three days 123

leads to destruction of the largest oocytes. The interval between the injections I and II can be changed substantially that is demonstrated in Z. scopas. Ovulation can be obtained at suitable time by means of manipulation by this interval. However, the duration of the interval should be enough (e.g., not less than 12 h in Z. scopas) to obtain ovulated oocytes from the majority of females. The assessment of optimal doses of the preparations is also important. The decrease of the summing dose of surfagon to a certain value (up to 10 µg/kg in Z. scopas) has no effect on the incidence of ovulation and leads to increased feasibility of the method. The application of exclusively GnRHa (surfagon) without dopamine antagonists can be applied for many ornamental coral fish species that simplifies the work of the personnel. Based on the method of hormonal stimulation of oocyte maturation in ornamental coral reef fishes (in particular, in three model species from two families with different reproductive strategies), the new data on the features of their oogenesis, spawning, structure and physiology of gametes, and early ontogeny are obtained. The materials on the properties of sperm and ovulated oocytes and features of their activation after the in vivo and in vitro storage are valuable for obtaining good quality eggs in marine aquaculture. A general scheme for the assessment of egg quality in marine fishes, in particular, in coral reef fishes, can be suggested. The egg batches including oocytes substantially ranging in size, having unusual inclusions in the cytoplasm, exhibiting exocytosis of at least a part of cortical alveoli, or oocytes with opaque content should be discarded. Subsequent assessment of egg quality can be conducted in several minutes after insemination. Egg batches including large proportions of eggs with uncompleted cortical reaction (unfertilized with subsequent appearance of uncleaved blastodiscs or fertilized, but developed abnormally) and non-activated oocytes (with subsequent partial cortical reaction) should not be used for incubation. A presence of cortical alveoli in the developing eggs suggests their poor quality. If the proportion of eggs with completed cortical reaction is high, the percentage of (fertilized) eggs exhibiting cleavage will be also high. However, fertilization rate can not represent reliable information about egg quality, and subsequent assessment of egg quality should be conducted at the stages of cleavage or during later development of the embryo. Similar to the farming of fish for human consumption, an analysis of the feasibility of cultivation of a new species involves two questions (Engelsen et al., 2004): (1) Is there a market for the cultured fish? and (2) Is it possible to produce and sell the fish at a lower cost than the market price 124

for wild-caught exemplars of the same species? Answering these questions will lead to a new one: what will happen to prices and production cost with time when production volume increases? Based on the general opinion, the species chosen for cultivation should command a high market price. However, the technology of rearing can be extremely complex and production cost would be very high. Thus, a decision on the species perspective for culture is always a compromise between its market price and production cost. The latter parameter can depend much on the type of early ontogeny. In ornamental coral fishes, closing of life cycles in intensive culture systems is conducted mainly for the species of the family Pomacentridae, in particular, for several representatives of the genus Amphiprion (Arvedlund et al., 2000; Gordon and Hecht, 2002; Astakhov et al., 2007; Bengoa-Ruigomez and Urkiaga, 2007). It is partly connected with the features of their life cycle: comparatively large larvae of clown fishes can be raised on rotifers just after start feeding. The species with small pelagic eggs, most often, are characterized by the ontogeny with well-expressed metamorphosis, and the technology of their culture is especially complex. In the species with demersal eggs or fishes brooded the eggs (e.g., seahorses), the larvae are usually larger and morphologically more derived that makes the process of their transition to exogenous feeding easier decreasing a production cost. Technical difficulties in mass-producing of the organisms of microzooplankton are still a constraint to their routine use for start feeding of coral fish larvae. Improvements in inert microdiets will likely lead to a progressive substitution of live feeds (Conceição et al., 2010). However, complete substitution is probably impossible for most species, at least for the first days of feeding. We believe that in the nearest future, with better propagation techniques the culture of marine ornamental fish will supply an increasing component of the overall supply of marine fish to the international aquaria trade.

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Çàêëþ÷èòåëüíûå êîììåíòàðèè Аквакультура рассматривается как потенциальная альтернатива изъятию декоративных коралловых рыб из природы: разведение представителей некоторых наиболее интенсивно отлавливаемых видов в полноцикловых системах, несомненно, приведёт к снижению рыболовной нагрузки на коралловые рифы. В настоящее время для получения потомства декоративных коралловых рыб практически исключительно используют их естественное размножение в неволе. Для начала нереста требуется продолжительный период адаптации рыб к искусственным условиям с использованием дорогостоящих систем с рециркуляцией воды и контролем параметров среды. Размножение разных самок, как правило, не синхронное, что ведёт к получению небольших порций икры на протяжении продолжительного времени. Это является ещё одной причиной сдерживающей экономически оправданное производство молоди. Гормональная стимуляция созревания является альтернативным методом получения половых продуктов у видов декоративных коралловых рыб. Гормональные инъекции могут применяться непосредственно после отлова живых рыб в природе и их доставки в лабораторию. Поэтому осеменённая икра и личинки могут быть получены на протяжении нескольких дней без значительных затрат на содержание производителей в контролируемых условиях. После гормонального воздействия и получения половых продуктов рыбы могут быть возвращены в естественную среду или оставлены в искусственных условиях до естественного размножения или повторной гормональной стимуляции. С использованием метода появляется возможность получения больших количеств икры и личинок на протяжении всего года для культивирования в промышленных масштабах и, таким образом, процесс разведения может стать рентабельным. Следует отметить, что многие виды декоративных коралловых рыб (включая представителей, имеющих особенно высокие цены) плохо адаптируются к искусственным условиям (Wabnitz et al., 2003) и гормональная стимуляция является единственным методом получения зрелых половых продуктов. В книге продемонстрирована возможность применения методов гормональной стимуляции созревания и овуляции ооцитов для рыб с непрерывным и прерывистым типами оогенеза. Схемы гормональной терапии могут различаться у рыб из разных таксономических групп, характеризующихся специфическими особенностями развития ооцитов. Следует учитывать такие характеристики как размер ооцитов, приобретающих компетенцию к созреванию, время, за которое они дос126

тигают этих размеров и продолжительность сезона размножения. Для отработки оптимальной схемы необходимо проводить экспресс-оценку состояния ооцитов не только перед проведением инъекций, но и после них. Значение такой оценки наиболее наглядно прослежено на созревании ооцитов Siganus spinus: дополнительная (третья) инъекция использована для получения овулировавших ооцитов. Весьма актуальным является определение возможности проведения однократных инъекции вместо градуальных. Это не только упрощает работу, но и в меньшей степени травмирует рыб. Такая возможность, очевидно, связана со спецификой эндокринологической чувствительности и, в определенной мере, с особенностями оогенеза. Например, особи Zebrasoma scopas способны продуцировать овулировавшие ооциты только после двукратных инъекций, тогда как представители Pomacentridae и Monacanthidae – и после однократных. Инъекции следует проводить в течение первого или второго дня после отлова рыб: содержание особей без кормления более 3 суток приводит к повреждению наиболее крупных ооцитов. Интервал между инъекциями I и II может варьировать в весьма значительных пределах, что особенно отчетливо показано на Z. scopas. Изменяя этот интервал можно получить овуляцию икры в удобное время. В то же время, для получения овулировавших ооцитов от подавляющего большинства самок, продолжительность его должна быть выше определенного времени (не менее 12 ч для Z. scopas). Актуальной является отработка оптимальных доз вводимых препаратов. Возможность снижения суммарной дозы сурфагона до определенного уровня (до 10 мкг/кг у Z. scopas) не влияет на процесс овуляции и ведёт к экономии препарата. Применение в качестве гормонального препарата исключительно GnRHas (в нашем случае – сурфагона) без антагонистов дофамина ведёт к удешевлению метода и упрощает работу персонала. На основе метода гормональной стимуляции созревания ооцитов декоративных коралловых рыб (в особенности, представителей трёх модельных видов из двух семейств с разной репродуктивной стратегией), получены новые данные об особенностях оогенеза, размножения, структуре и физиологии гамет и раннем онтогенезе. Данные о свойствах спермы и овулировавших ооцитов, а также об активации гамет после выдерживания in vivo и in vitro, являются полезными для получения икры высокого качества в морской аквакультуре. Предложена схема оценки качества яиц морских рыб, в особенности, видов коралловых рыб. Порции, включающие ооциты значительно варьирующие по размеру, с необычными включениями в цитоплазме и 127

экзоцитозом по меньшей мере части кортикальных альвеол, а также ооциты с непрозрачным содержимым, должны быть отбракованы. Последующая оценка качества яиц может быть проведена через несколько минут после осеменения. Порции со значительной долей яиц с незавершённой кортикальной реакцией (неоплодотворённые с последующим появлением недробящегося плазменного диска или оплодотворённые, но впоследствии дробящиеся аномально) и неактивированные ооциты (с последующей частичной кортикальной реакцией) не должны использоваться для инкубации. Наличие кортикальных альвеол в развивающихся яйцах свидетельствует о низком качестве яиц. Если доля яиц с полной кортикальной реакцией высока, доля (оплодотворённых) дробящихся яиц будет высокой. Вместе с тем, процент оплодотворения не является надёжным показателем качества яиц, и детальная оценка качества должна быть проведена на стадиях дробления. Так же, как и при разведении рыб для пищевых целей, анализ рентабельности культивирования нового вида включает два вопроса (Engelsen et al., 2004): (1) имеется ли рынок сбыта для культивируемого объекта? и (2) возможно ли производить и продавать рыбу по цене меньшей, чем цена особей того же вида, отловленных в природе? Далее необходим анализ изменения цены и себестоимости продукции по мере увеличения её объёма. В соответствии с общепринятыми представлениями, вид, перспективный для разведения, должен обладать высокой рыночной ценой. Однако технология культивирования может быть чрезвычайно сложной, а себестоимость продукции чрезмерно высокой. Последний параметр может в значительной степени зависеть от типа раннего онтогенеза. Среди декоративных коралловых рыб, воспроизведение всего жизненного цикла осуществлено главным образом для видов семейства Pomacentridae, в частности, для представителей рода Amphiprion (Arvedlund et al., 2000; Gordon and Hecht, 2002; Astakhov et al., 2007; Bengoa-Ruigomez and Urkiaga, 2007). В значительной степени это связано с особенностями их онтогенеза: сравнительно крупные личинки с начала экзогенного питания могут быть переведены на потребление коловраток. Виды с мелкими пелагическими яйцами обычно характеризуются онтогенезом с ярко выраженным метаморфозом, а технология культивирования особенно сложна. Виды с донной икрой или рыбы вынашивающие потомство (например, морские коньки) обычно характеризуются сравнительно крупными и морфологически продвинутыми личинками; процесс их перевода на экзогенное питание менее сложен и, следовательно, себестоимость культивирования ниже. 128

Технические трудности, связанные с массовым разведением организмов микрозоопланктона ограничивают их использование в качестве стартового корма для личинок коралловых рыб. Современные достижения в области производства искусственных кормов, по-видимому, приведут к постепенной замене живых организмов (Conceição et al., 2010); но для большинства видов рыб полная замена живых кормов на искусственные, по меньшей мере на протяжении первых суток экзогенного питания, является невозможной. По нашему мнению, в ближайшем будущем в процессе совершенствования технологии, культивирование морских декоративных рыб будет всё больше соответствовать потребности в живых организмах, существующей в аквариумистике.

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COLOR PLATES

Color Plates, Fig. 1. Research facilities and methods of the study. (a) Coastal Department of Russian-Vietnamese Tropical Research and Technological Center (Nha Trang, Vietnam); (b) Mariculture Station of the Institute of Aquaculture no. 3 (Nha Trang, Vietnam); (c) laboratory for cultivation of ornamental fish species at the Mariculture Station; (d) hormonal injection to Abudefduf sexfasciatus; (e) probes for biopsy; (f) Natal’ya Emel’yanova and Vo Thi Ha in the laboratory. Íàó÷íûå ñòàíöèè è ìåòîäû èññëåäîâàíèÿ. (a) Ïðèìîðñêîå îòäåëåíèå ÐîññèéñêîÂüåòíàìñêîãî Òðîïè÷åñêîãî íàó÷íî-èññëåäîâàòåëüñêîãî è òåõíîëîãè÷åñêîãî öåíòðà (Íÿ÷àíã, Âüåòíàì); (b) Ñòàíöèÿ ìàðèêóëüòóðû Èíñòèòóòà Àêâàêóëüòóðû ¹ 3 (Íÿ÷àíã, Âüåòíàì); (c) ëàáîðàòîðèÿ äëÿ êóëüòèâèðîâàíèÿ äåêîðàòèâíûõ êîðàëëîâûõ ðûá íà Ñòàíöèè ìàðèêóëüòóðû; (d) ãîðìîíàëüíàÿ èíúåêöèÿ Abudefduf sexfasciatus; (e) ùóïû äëÿ áèîïñèè; (f) Íàòàëüÿ Åìåëüÿíîâà è Âî Òõè Õà â ëàáîðàòîðèè.

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Color Plates, Fig. 2. Three ornamental fish species used as the model objects: (a) Zebrasoma scopas, 138 mm TL; (b) Dascyllus trimaculatus, 116 mm TL; and (c) Abudefduf sexfasciatus, 138 mm TL. Òðè âèäà äåêîðàòèâíûõ êîðàëëîâûõ ðûá, èñïîëüçóåìûõ â êà÷åñòâå ìîäåëüíûõ îáúåêòîâ: (a) Zebrasoma scopas, 138 ìì TL; (b) Dascyllus trimaculatus, 116 ìì TL; (c) Abudefduf sexfasciatus, 138 ìì TL.

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Color Plates, Fig. 3. Urogenital region on the abdomen of Zebrasoma scopas: (a) male; (b) female with immature oocytes; (c) female with mature oocytes before ovulation or during ovulation. 1, anal pore; 2, urogenital pore; 3, genital pore; 4, urinal pore; 5, pellicle covering the genital pore before and during ovulation. Ìî÷åïîëîâàÿ îáëàñòü áðþøèíû Zebrasoma scopas: (a) ñàìåö; (b) ñàìêà ñ íåçðåëûìè îîöèòàìè; (c) ñàìêà ñ ñîçðåâøèìè îîöèòàìè ïåðåä îâóëÿöèåé èëè âî âðåìÿ îâóëÿöèè. 1, àíàëüíàÿ ïîðà; 2, ìî÷åïîëîâàÿ ïîðà; 3, ïîëîâàÿ ïîðà; 4, ìî÷åâàÿ ïîðà; 5, ìåìáðàíà, ïîêðûâàþùàÿ ïîëîâóþ ïîðó ïåðåä îâóëÿöèåé è â ïðîöåññå îâóëÿöèè.

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Color Plates, Fig. 4. Ovaries (left side) and testes (right side) of (a, b) Zebrasoma scopas, (c, d) Dascyllus trimaculatus, and (e, f) Abudefduf sexfasciatus. Scale bar 5 mm. ßè÷íèêè (ñëåâà) è ñåìåííèêè (ñïðàâà) (a, b) Zebrasoma scopas, (c, d) Dascyllus trimaculatus è (e, f) Abudefduf sexfasciatus. Ìàñøòàá 5 ìì.

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Color Plates, Fig. 5. Sex cells of sexually mature exemplars of Zebrasoma scopas: (a) before injection I; (b) in 6 h after injection II; (c) in 12 h after injection II; (d) after ovulation; (e) destruction of the largest oocytes in 6 days after the beginning of starvation of the fish; (f) a fragment of the testis. 1, nucleus; 2, small lipid droplets; 3, lipid droplet; 4, homogenized oocyte; 5, empty follicle; 6, resorbing oocyte; 7, spermatozoon. Ïîëîâûå êëåòêè ïîëîâîçðåëûõ ýêçåìïëÿðîâ Zebrasoma scopas: (a) ïåðåä èíúåêöèåé I; (b) ÷åðåç 6 ÷ ïîñëå èíúåêöèè II; (c) ÷åðåç 12 ÷ ïîñëå èíúåêöèè II; (d) ïîñëå îâóëÿöèè; (e) äåñòðóêöèÿ íàèáîëåå êðóïíûõ îîöèòîâ ÷åðåç 6 ñóò. ïîñëå íà÷àëà ãîëîäàíèÿ ðûá; (f) ôðàãìåíò ñåìåííèêà. 1, ÿäðî; 2, ìåëêèå æèðîâûå êàïëè; 3, æèðîâàÿ êàïëÿ; 4, ãîìîãåíèçèðîâàííûé îîöèò; 5, ïóñòîé ôîëëèêóë; 6, ðåçîðáèðóþùèéñÿ îîöèò; 7, ñïåðìàòîçîèä.

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Color Plates, Fig. 6. Sex cells in females and males of Dascyllus trimaculatus: (a) oocytes before injection I; (b) oocytes in several hours after injection II; (c) ovarian fragment in two days after ovulation; (d) testicular fragment. 1, nucleus; 2, small lipid droplets; 3, lipid droplet; 4, spermatozoa. Ïîëîâûå êëåòêè ñàìîê è ñàìöîâ Dascyllus trimaculatus: (a) îîöèòû ïåðåä èíúåêöèåé I; (b) îîöèòû ÷åðåç íåñêîëüêî ÷àñîâ ïîñëå èíúåêöèè II; (c) ôðàãìåíò ÿè÷íèêà ÷åðåç 2 ñóò. ïîñëå îâóëÿöèè; (d) ôðàãìåíò ñåìåííèêà. 1, ÿäðî; 2, ìåëêèå æèðîâûå êàïëè; 3, æèðîâàÿ êàïëÿ; 4, ñïåðìàòîçîèäû.

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Color Plates, Fig. 7. Sex cells in the females and males of Abudefduf sexfasciatus: (a) oocytes before injection I; (b) ovarian fragment after ovulation; (c) ovarian fragment with enlarged follicular cells; (d) testicular fragment; 1, empty follicle; 2, nucleus; 3, zona radiata; 4, follicular cells; 5, spermatozoa. Ïîëîâûå êëåòêè ñàìîê è ñàìöîâ Abudefduf sexfasciatus: (a) îîöèòû ïåðåä èíúåêöèåé I; (b) ôðàãìåíò ÿè÷íèêà ïîñëå îâóëÿöèè; (c) ôðàãìåíò ÿè÷íèêà ñ óâåëè÷åííûìè ôîëëèêóëÿðíûìè êëåòêàìè; (d) ôðàãìåíò ñåìåííèêà. 1, ïóñòîé ôîëëèêóë; 2, ÿäðî; 3, zona radiata; 4, ôîëëèêóëÿðíûå êëåòêè; 5, ñïåðìàòîçîèäû.

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Color Plates, Fig. 8. Dynamics of cortical reaction (number of cortical alveoli) in the oocytes of Zebrasoma scopas: the data for ten inseminated and fertilized eggs and one non-inseminated oocyte (for this oocyte, the number of cortical alveoli is designated as black triangles, and the line of a trend is given). The time is given from insemination; for the non-inseminated oocyte, from its placing into water. Äèíàìèêà êîðòèêàëüíîé ðåàêöèè (÷èñëî êîðòèêàëüíûõ àëüâåîë) â îîöèòàõ Zebrasoma scopas: äàííûå äëÿ 10 îñåìåí¸ííûõ è îïëîäîòâîðèâøèõñÿ ÿèö è äëÿ íåîñåìåí¸ííîãî îîöèòà (÷èñëî êîðòèêàëüíûõ àëüâåîë ýòîãî îîöèòà îáîçíà÷åíî ÷åðíûìè òðåóãîëüíèêàìè è ïðîâåäåíà ëèíèÿ òðåíäà). Âðåìÿ îòñ÷èòàíî îò îñåìåíåíèÿ, à äëÿ íåîñåìåí¸ííîãî îîöèòà – îò åãî ïîìåùåíèÿ â âîäó.

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Color Plates, Fig. 9. Quality of eggs (percentage of eggs with normal cleavage, abnormal cleavage, uncleaved blastodisc, and without blastodisc) of Zebrasoma scopas obtained from 15 females after in vivo storage of ovulated oocytes in the female's ovarian cavity. Same number of the stars means the absence of significant differences between the average percentages of normally cleaved eggs in different series. Êà÷åñòâî ÿèö 15 ñàìîê Zebrasoma scopas (ñîîòíîøåíèå ÿèö ñ íîðìàëüíûì è àíîìàëüíûì äðîáëåíèåì, íåäðîáÿùèìñÿ áëàñòîäèñêîì è áåç áëàñòîäèñêà) ïðè èõ çàäåðæêå â îâàðèàëüíîé ïîëîñòè ñàìêè (in vivo) ïîñëå îâóëÿöèè. Îäèíàêîâîå ÷èñëî çâåçäî÷åê îçíà÷àåò îòñó òñòâèå äîñòîâåðíûõ îòëè÷èé ìåæäó ñðåäíèìè çíà÷åíèÿìè ïîêàçàòåëÿ íîðìàëüíîãî äðîáëåíèÿ â ðàçíûõ ñåðèÿõ.

Color Plates, Fig. 10. Quality of eggs of Zebrasoma scopas obtained from seven females after in vitro storage of ovulated oocytes at 25 and 7°C. Same number of the stars means the absence of significant differences between the average percentages of normally cleaved eggs from different series. Êà÷åñòâî ÿèö 7 ñàìîê Zebrasoma scopas ïîñëå âûäåðæèâàíèÿ îîöèòîâ in vitro ïðè 25 è 7°C. Îäèíàêîâîå ÷èñëî çâåçäî÷åê îçíà÷àåò îòñóòñòâèå äîñòîâåðíûõ îòëè÷èé ìåæäó ñðåäíèìè çíà÷åíèÿìè ïîêàçàòåëÿ íîðìàëüíîãî äðîáëåíèÿ â ðàçíûõ ñåðèÿõ.

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Color Plates, Fig. 11. Egg quality in Zebrasoma scopas after storage of oocytes in the water before insemination: (a) oocytes from two females, maximum duration of the storage 20 min; (b) oocytes from five females, maximum duration of the storage 90 min; (c) overripened oocytes from five females, maximum duration of the storage 40 min. Êà÷åñòâî ÿèö Zebrasoma scopas ïîñëå âûäåðæèâàíèÿ îîöèòîâ â ìîðñêîé âîäå ïåðåä îñåìåíåíèåì: (a) îîöèòû îò 2 ñàìîê, ìàêñèìàëüíàÿ ïðîäîëæèòåëüíîñòü âûäåðæèâàíèÿ 20 ìèí; (b) îîöèòû îò 5 ñàìîê, ìàêñèìàëüíàÿ ïðîäîëæèòåëüíîñòü âûäåðæèâàíèÿ 90 ìèí; (c) ïåðåçðåâøèå îîöèòû îò 5 ñàìîê, ìàêñèìàëüíàÿ ïðîäîëæèòåëüíîñòü âûäåðæèâàíèÿ 40 ìèí.

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Color Plates, Fig. 12. Quality of eggs of Dascyllus trimaculatus obtained from two females after in vitro storage of ovulated oocytes at 25 and 5°C. Êà÷åñòâî ÿèö Dascyllus trimaculatus îò äâóõ ñàìîê ïîñëå âûäåðæèâàíèÿ îîöèòîâ in vitro ïðè 25 è 5°C.

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Color Plates, Fig. 13. Egg quality in Dascyllus trimaculatus after storage of oocytes from two females in the water before insemination. Êà÷åñòâî ÿèö Dascyllus trimaculatus ïîñëå âûäåðæèâàíèÿ îîöèòîâ îò äâóõ ñàìîê â ìîðñêîé âîäå ïåðåä îñåìåíåíèåì.

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Color Plates, Fig. 14. Quality of eggs of Abudefduf sexfasciatus obtained in two experiments from (a) five females and (b) seven females after in vivo storage of ovulated oocytes in the female’s ovary. The eggs from different females have different colors. Êà÷åñòâî ÿèö Abudefduf sexfasciatus, ïîëó÷åííûõ â äâóõ ýêñïåðèìåíòàõ îò (a) ïÿòè ñàìîê è (b) ñåìè ñàìîê ïîñëå çàäåðæêè îâóëèðîâàâøèõ îîöèòîâ (in vivo) â ÿè÷íèêå ñàìêè. ßèöà îò ðàçíûõ ñàìîê îáîçíà÷åíû ðàçíûì öâåòîì.

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Color Plates, Fig. 15. Quality of eggs of Abudefduf sexfasciatus obtained from two females after in vitro storage of ovulated oocytes at 25 and 5°C. Êà÷åñòâî ÿèö Abudefduf sexfasciatus îò äâóõ ñàìîê ïîñëå âûäåðæèâàíèÿ îâóëèðîâàâøèõ îîöèòîâ in vitro ïðè 25 è 5°C.

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Color Plates, Fig. 16. Egg quality in Abudefduf sexfasciatus after storage of oocytes from two females in the water before insemination. Êà÷åñòâî ÿèö Abudefduf sexfasciatus ïîñëå âûäåðæèâàíèÿ îîöèòîâ îò äâóõ ñàìîê â ìîðñêîé âîäå ïåðåä îñåìåíåíèåì.

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Color Plates, Fig. 17. Larval development of Zebrasoma scopas at 25 and 27°C. 25°C from insemination of the eggs: (a) stage of hatching, a real position of the free embryo, 1.5 mm TL, age 27 h from insemination; (b) resorption of the yolk 50% of its volume at hatching, large melanophores behind the eye, above the yolk sac, above the anus, and in the ventral middle area of the caudal part of the body, 1.7 mm TL, age 11 h from hatching; (c) resorption of the yolk 80% of its volume at hatching, 1.75 mm TL, age 21 h; (d) total resorption of the yolk and lipid droplet, three transversal pigment bands on the body, small melanophores in the ventral row of the caudal part, exogenous feeding, 1.8 mm TL, age 3 days 6 h. 27°C from hatching: (e) food objects in the intestine of the larva, age 3 days from hatching; (f) age 4 days, 1.85 mm TL; (g) food objects in the intestine of the larva, age 4 days. Ëè÷èíî÷íîå ðàçâèòèå Zebrasoma scopas ïðè 25 è 27°C. 25°C îò îñåìåíåíèÿ îîöèòîâ: (a) ñòàäèÿ âûëóïëåíèÿ, åñòåñòâåííàÿ îðèåíòàöèÿ ïðåäëè÷èíêè, 1.5 ìì TL, âîçðàñò 27 ÷ ïîñëå îñåìåíåíèÿ; (b) ðåçîðáöèÿ æåëòêà íà 50% åãî îáú¸ìà ïðè âûëóïëåíèè, êðóïíûå ìåëàíîôîðû çà ãëàçîì, íàä æåëòî÷ñíûì ìåøêîì, íàä àíóñîì è â âåíòðàëüíîé ñðåäíåé ÷àñòè õâîñòîâîãî îòäåëà, 1.7 ìì TL, âîçðàñò 11 ÷ ïîñëå âûëóïëåíèÿ; (c) ðåçîðáöèÿ æåëòêà íà 80% åãî îáú¸ìà ïðè âûëóïëåíèè, 1.75 ìì TL, âîçðàñò 21 ÷; (d) ïîëíàÿ ðåçîðáöèÿ æåëòêà è æèðîâîé êàïëè, òðè ïîïåðå÷íûõ ïèãìåíòíûõ ïîëîñêè íà òåëå, ìåëêèå ìåëàíîôîðû â âåíòðàëüíîì ðÿäó õâîñòîâîé ÷àñòè òåëà, ýêçîãåííîå ïèòàíèå, 1.8 ìì TL, âîçðàñò 3 ñóò. 6 ÷. 27°C ïîñëå âûëóïëåíèÿ: (e) ïèùåâûå îáúåêòû â êèøå÷íèêå ëè÷èíêè, âîçðàñò 3 ñóò. ïîñëå âûëóïëåíèÿ; (f) âîçðàñò 4 ñóò., 1.85 ìì TL; (g) ïèùåâûå îáúåêòû â êèøå÷íèêå ëè÷èíêè, âîçðàñò 4 ñóò.

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Color Plates, Fig. 18. Incubation of the eggs of Dascyllus trimaculatus attaching to the gauze. Èíêóáàöèÿ ÿèö Dascyllus trimaculatus ïðèêðåïë¸ííûõ ê ïîëîñêàì ãàçà.

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Color Plates, Fig. 19. Larval development of Dascyllus trimaculatus at 27°C. (a, b) Stages of hatching, yolk sac and lipid droplet, large melanophores in the dorsal and ventral rows, small melanophores around the eyes, number of muscular segments 8 before the anus and 24 after the anus, age from insemination of the eggs 3 days 4 h, 2.10 mm TL; (c) resorption of the yolk, lipid droplet remains, pigmented eyes, opening in the mouth, 1 day from hatching, 2.20 mm TL; (d) resorption of the lipid droplet, transition to exogenous feeding, 2 days, 2.24 mm TL; (e, f) 4 days, 2.27 mm TL; (g) 5 days, 2.28 mm TL; (h, i) 6 days, 2.40 mm TL; (j) 8 days, 2.57 mm TL; (k) relative shortening of the postanal part of the body, age 10 days, 2.82 mm TL; (l) first rays in the caudal fin, dark spots on the flanks, age15 days, 3.08 mm TL. Ëè÷èíî÷íîå ðàçâèòèå Dascyllus trimaculatus ïðè 27°C. (a, b) Ñòàäèè âûëóïëåíèÿ, æåëòî÷íûé ìåøîê è æèðîâàÿ êàïëÿ, êðóïíûå ìåëàíîôîðû â äîðñàëüíîì è âåíòðàëüíîì ðÿäàõ, ìåëêèå ìåëàíîôîðû âîêðóã ãëàç, ÷èñëî ìèîìåðîâ 8 äî àíóñà è 24 ïîñëå íåãî, âîçðàñò 3 ñóò. 4 ÷. ïîñëå îñåìåíåíèÿ, 2.10 ìì TL; (c) ðåçîðáöèÿ æåëòêà, ñîõðàíèëàñü æèðîâàÿ êàïëÿ, ïèãìåíò â ãëàçàõ, ðîò îòêðûò, 1 ñóò. ïîñëå âûëóïëåíèÿ, 2.20 ìì TL; (d) ðåçîðáöèÿ æèðîâàîé êàïëè, ïåðåõîä íà ýêçîãåííîå ïèòàíèå, âîçðàñò 2 ñóò., 2.24 ìì TL; (e, f) âîçðàñò 4 ñóò., 2.27 ìì TL; (g) âîçðàñò 5 ñóò., 2.28 ìì TL; (h, i) âîçðàñò 6 ñóò., 2.40 ìì TL; (j) âîçðàñò 8 ñóò., 2.57 ìì TL; (k) îòíî ñèòåëüíîå óê îðî÷åíèå ïîñòàíàëüíîé ÷àñòè òåëà, âîçðàñò 10 ñóò., 2.82 ìì TL; (l) ïåðâûå ëó÷è â õâîñòîâîì ïëàâíèêå, ò¸ìíûå ïÿòíà íà áîêàõ òåëà, âîçðàñò 15 ñóò., 3.08 ìì TL.

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Color Plates, Fig. 20. Embryonic development of Abudefduf sexfasciatus at 27°C. (a) Egg swelling, aggregation of the cytoplasm at the animal pole, age 23 min from insemination; (b) cytoplasmic disc at the animal pole, age 60 min; (c) 2 blastomeres, age 1 h 50 min; (d) 16 blastomeres, age 2 h 50 min; (e) middle-cell morula, age 6 h 30 min; (f) end of gastrulation before the yolk plug closure, age 17 h; (g) beginning of separation of the caudal part of the body from the yolk sac, age 1 day 14 h; (h) substantial resorption of the yolk, formation of large melanophores on the yolk sac and the ventral preanal part of the body, turnover of the embryo, age 1 day 22 h; (i) appearance of intensive black pigment in the eyes, 2 days, 22 h; (j) size of the yolk sac is less than the size of the eye, a stage before hatching, age 4 days 17 h. 1, cytoplasm; 2, yolk; 3, lipid droplet; 4, presumptive head of the embryo; 5, margin of the blastoderm. Ýìáðèîíàëüíîå ðàçâèòèå Abudefduf sexfasciatus ïðè 27°C. (a) Íàáóõàíèå ÿéöà, àãðåãàöèÿ öèòîïëàçìû íà àíèìàëüíîì ïîëþñå, âîçðàñò 23 ìèí ïîñëå îñåìåíåíèÿ; (b) öèòîïëàçìàòè÷åñêèé äèñê íà àíèìàëüíîì ïîëþñå, âîçðàñò 60 ìèí; (c) 2 áëàñòîìåðà, âîçðàñò 1 ÷ 50 ìèí; (d) 16 áëàñòîìåðîâ, âîçðàñò 2 ÷ 50 ìèí; (e) ìîðóëà ñðåäíåðàçìåðíûõ êëåòîê, âîçðàñò 6 ÷ 30 ìèí; (f) çàâåðøåíèå ãàñòðóëÿöèè, ïåðåä çàìûêàíèåì æåëòî÷íîé ïðîáêè, âîçðàñò 17 ÷; (g) íà÷àëî îòäåëåíèÿ õâîñòîâîé ÷àñòè ýìáðèîíà îò æåëòî÷íîãî ìåøêà, âîçðàñò 1 ñóò. 14 ÷; (h) çíà÷èòåëüíàÿ ðåçîðáöèÿ æåëòêà, ôîðìèðîâàíèå êðóïíûõ ìåëàíîôîðîâ íà æåëòî÷íîì ìåøêå è â âåíòðàëüíîé ïðåàíàëüíîé ÷àñòè òåëà, ïîâîðîò ýìáðèîíà, âîçðàñò 1 ñóò. 22 ÷; (i) ïîÿâëåíèå èíòåíñèâíîãî ÷¸ðíîãî ïèãìåíòà â ãëàçàõ, âîçðàñò 2 ñóò. 22 ÷; (j) ðàçìåð æåëòî÷íîãî ìåøêà ìåíüøå, ÷åì ðàçìåð ãëàçà, ñòàäèÿ ïåðåä âûëóïëåíèåì, âîçðàñò 4 ñóò. 17 ÷. 1, öèòîïëàçìà; 2, æåëòîê; 3, æèðîâàÿ êàïëÿ; 4, çà÷àòîê ãîëîâû ýìáðèîíà; 5, êðàé áëàñòîäåðìû.

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Color Plates, Fig. 21. Larval development of Abudefduf sexfasciatus at 27°C. (a) Stage of hatching, yolk sac and lipid droplet under the intestine, muscular segments: 7 before the anus and 22 after the anus, age from insemination of the eggs 4 days 20 h, 2.60 mm TL; (b, c) resorption of the yolk, enlarged melanophores in the ventral row, transition to exogenous feeding, age 2 days from hatching, 2.63 mm TL; (d) age 4 days, 2.71 mm TL; (e, f) yellow pigment on the dorsal part of the head, age 5 days, 2.72 mm TL; (g) age 6 days, 2.87 mm TL; (h) age 11 days, 2.93 mm TL; (i) abrupt shortening of the postanal part of the body, first rays in the dorsal and caudal fins, age 15 days, 4.38 mm TL; (j, k) appearance of rays in all fins, intensive pigmentation of the preanal part of the body, age 18 days, 4.75 mm TL; (l) digestive system at the age of 7 days: 1, intestine; 2, liver; 3, gall bladder; 4, swim bladder. Ëè÷èíî÷íîå ðàçâèòèå Abudefduf sexfasciatus ïðè 27°C. (a) Ñòàäèÿ âûëóïëåíèÿ, æåëòî÷íûé ìåøîê è æèðîâàÿ êàïëÿ, ÷èñëî ìèîìåðîâ 7 äî àíóñà è 22 ïîñëå íåãî, âîçðàñò îò îñåìåíåíèÿ ÿèö 4 ñóò. 20 ÷, 2.60 ìì TL; (b, c) ðåçîðáöèÿ æåëòêà, êðóïíûå ìåëàíîôîðû â âåíòðàëüíîì ðÿäó, ïåðåõîä íà ýêçîãåííîå ïèòàíèå, âîçðàñò 2 ñóò. ïîñëå âûëóïëåíèÿ, 2.63 ìì TL; (d) âîçðàñò 4 ñóò., 2.71 ìì TL; (e, f) æ¸ëòûé ïèãìåíò â äîðñàëüíîé ÷àñòè ãîëîâû, âîçðàñò 5 ñóò., 2.72 ìì TL; (g) âîçðàñò 6 ñóò., 2.87 ìì TL; (h) âîçðàñò 11 ñóò., 2.93 ìì TL; (i) ñóùåñòâåííîå óêîðî÷åíèå ïîñòàíàëüíîé ÷àñòè òåëà, ïåðâûå ëó÷è â ñïèííîì è õâîñòîâîì ïëàâíèêàõ, âîçðàñò 15 ñóò., 4.38 ìì TL; (j, k) ïîÿâëåíèå ëó÷åé âî âñåõ ïëàâíèêàõ, èíòåíñèâíàÿ ïèãìåíòàöèÿ ïðåàíàëüíîé ÷àñòè òåëà, âîçðàñò 18 ñóò., 4.75 ìì TL; (l) ïèùåâàðèòåëüíàÿ ñèñòåìà â âîçðàñòå 7 ñóò.: 1, êèøå÷íèê; 2, ïå÷åíü; 3, æåë÷íûé ïóçûðü; 4, ïëàâàòåëüíûé ïóçûðü.

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Acknowledgments This book was prepared according to initiative of the coordinator of our investigations professor Georgij G. Novikov (Department of Ichthyology, Biological Faculty of Moscow State University, Moscow, Russia) who passed away prematurely in 2007. We are grateful to V.K. Nezdoliy and N.L. Filichev, former and present directors of the Russian part of the Coastal Department of the Russian–Vietnamese Tropical Research and Technological Center (RVTRTC, Nha Trang, Vietnam) and Tran Thanh Quang, director of the Vietnamese part of the Coastal Department of the RVTRTC. This study would be impossible without help of our Vietnamese colleagues Luong Thi Bich Thuan and Vo Thi Ha. We thank all staff of the Coastal Department, and especially Nguyen Duy Toan and Dinh Thi Hai Yen participated in organization of our works, Nguyen Dang Thanh for technical help associated with different kinds of equipment, and Nguyen Van Hung (deputy director of Mariculture Station of the Institute of Aquaculture no. 3) for joint investigations at the Station. The study was carried out with the financial support from the RVTRTC and was supported in part by the joint projects between the RVTRTC and Institute of Aquaculture no. 3 and a grant for the leading scientific schools (no. NSh-719.2012.4;2012-2013). AUTHORS Natal’ya G. Emel’yanova Principal scientist Biological Faculty, Moscow State University, Vorob’evy gory, Moscow, 119899 Russia E-mail: [email protected] Dimitri A. Pavlov Principal scientist Biological Faculty, Moscow State University, Vorob’evy gory, Moscow, 119899 Russia E-mail: [email protected]

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