Section 2 Other Species

**91**

**1. Introduction**

**Chapter 6**

Species

*Mônica Cassel*

**Abstract**

A Review of the Macroscopic,

Ultramicroscopic Characteristics

Developmental Processes in Fish

Studies involving the reproductive biology of fish have several possibilities of approach, such as the understanding of gonadal development, oocyte development, and the reproductive cycle of the species. In addition, analyses of gonadal morphology can be made at macro-, micro-, and ultramicroscopic levels. This knowledge helps to define factors that determine the different stages of gonadal development, as well as the "triggers" that initiate the reproductive process. In females, the growth and maturation of the ovarian follicles depend on a carefully elaborated communication between the follicular cells and the oocyte and a precisely organized contractile system. Changes in these systems appear to be related to apoptotic cells. This extensive remodeling of gonadal tissue, due to cell proliferation and differentiation, promotes also changes in the extracellular matrix. With this in mind, we provide herein a complementary and in-depth information on cell-cell and cell-matrix interactions related to the process of oocyte development in fish species. This information, together with the existing structural and ultrastructural descriptions of ovaries of different species, will enable a better understanding of

**Keywords:** fish species, reproductive biology, gonadal and oocyte development,

The knowledge on the reproductive characteristics of fish is fundamental to understand the adaptations developed to maximize the reproductive success in a given environment, considering the life history aspects of each species [1]. Studies involving the reproductive biology of fish have several possibilities of approach, such as the understanding of gonadal development and the reproductive cycle of the species. Analyses of gonadal morphology are important for the understanding of the biology of the species and have been widely applied in

Microscopic, and

of Some Key Oocyte

the reproductive processes for the group of fish.

cell-cell interactions, cell-matrix interactions

#### **Chapter 6**

## A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key Oocyte Developmental Processes in Fish Species

*Mônica Cassel*

### **Abstract**

Studies involving the reproductive biology of fish have several possibilities of approach, such as the understanding of gonadal development, oocyte development, and the reproductive cycle of the species. In addition, analyses of gonadal morphology can be made at macro-, micro-, and ultramicroscopic levels. This knowledge helps to define factors that determine the different stages of gonadal development, as well as the "triggers" that initiate the reproductive process. In females, the growth and maturation of the ovarian follicles depend on a carefully elaborated communication between the follicular cells and the oocyte and a precisely organized contractile system. Changes in these systems appear to be related to apoptotic cells. This extensive remodeling of gonadal tissue, due to cell proliferation and differentiation, promotes also changes in the extracellular matrix. With this in mind, we provide herein a complementary and in-depth information on cell-cell and cell-matrix interactions related to the process of oocyte development in fish species. This information, together with the existing structural and ultrastructural descriptions of ovaries of different species, will enable a better understanding of the reproductive processes for the group of fish.

**Keywords:** fish species, reproductive biology, gonadal and oocyte development, cell-cell interactions, cell-matrix interactions

#### **1. Introduction**

The knowledge on the reproductive characteristics of fish is fundamental to understand the adaptations developed to maximize the reproductive success in a given environment, considering the life history aspects of each species [1]. Studies involving the reproductive biology of fish have several possibilities of approach, such as the understanding of gonadal development and the reproductive cycle of the species. Analyses of gonadal morphology are important for the understanding of the biology of the species and have been widely applied in Teleostei, as in recent studies on spermatogenesis [2–5], folliculogenesis [6–8], reproductive cycle [8–12], and fecundity [13].

Studies have been carried out to describe and classify the stages of gonadal development and reproductive stages of fish in the Neotropical region. One of the most classic and used bibliographic sources has been Vazzoler [14]. However, other proposals for description have already been made by Grier and Taylor [15], Grier [16], and Lo Nostro et al. [17], which detail the continuity and discontinuity of the germinal epithelium and the cell types present in the gonads. Recently, Brown-Peterson et al. [18] developed a simpler terminology to facilitate the communication and comparison of studies on the reproductive biology of fish. Still in order to make the nomenclature more comprehensive, the stages of oocyte development were simplified by Quagio-Grassiotto et al. [19], and the development of stages of atresia, which are characterized as involutive processes, follows according to Miranda et al. [20].

Gonadal development can be analyzed macroscopically, and changes in shape, size, color, and texture of the gonads have been used as parameters for the classification of maturation status in many studies of ecology and reproductive dynamics[14, 21]. However, the most used analysis has been of the microscopic characters, since it allows a more detailed and precise description of the transitions and morphological and structural transformations that happen during gonadal development [8, 22, 23]. Thus, regarding the microscopic aspects of the gonad, it is verified that [24]:


The oocyte development in a mature egg is a complex process modulated by numerous environmental and endocrine factors [25], and understanding the morphological characteristics of oocytes is important to interpret the dynamics of oogenesis [26]. Among the oocyte processes, folliculogenesis results in the removal of the primary oocyte from oogonium nests and consequent formation of ovarian follicles [27]. Descriptions for the germinal epithelium made by Grier [28] conceptualized "follicular complex" as the functional unit of the ovary. This complex is formed by two compartments separated by a basement membrane. One compartment is the follicle, which consists of the oocyte surrounded by follicular cells and originated from the germinal epithelium. The second compartment is the theca, made up of undifferentiated ovarian stromal cells.

In the previtellogenic oocyte phase, multiple nucleoli are observed, as described by Grier et al. [29]. These oocytes are also called perinucleolar oocytes, when the nucleoli migrate to the nuclear periphery. There is also the formation of the zona pellucida, a complex structure consisting generally of two layers crossed by pores or channels containing the oocyte microvilli and/or follicular cell extensions. The zona pellucida reflects adaptations to different ecological conditions in which the eggs develop [30], whose inner layer protects the egg from mechanical damage and whose outer layer protects it from microorganisms.

Another cell characteristic that is used to describe the stages of oocyte development is the presence of nüages, Balbiani corpuscles, and cortical alveoli. The nüages are originated by the transfer from the nucleus to the cytoplasm of large amounts of heterogeneous and ribosomal RNA synthesized [31] and associated with proteins. Balbiani corpuscles or yolk nuclei, described by Hubbard [32], were recognized

**93**

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key…*

as clusters of organelles located near the nucleus, which proliferate intensely and spread throughout the cytoplasm. And, the cortical alveoli, as observed by Grier et al. [29], are vesicles filled with glycoproteins, formed by depressions of the oocyte membrane that become progressively larger, marking the final stage of

The described changes are followed by an expressive growth of the oocyte during vitellogenesis, in which the oocyte accumulates the nutritive reserves necessary for the development of the embryo. The oocyte also accumulates RNA and completes the differentiation of its cellular and noncellular envelopes. During this time, the oocyte interrupts the meiosis at the end of the prophase and in the diplotene stage. Maturation processes are characterized by the reduction or halting of endocytosis, resumption of meiosis, breakdown of the germinal vesicle, formation of a monolayer of cortical alveoli under the plasma oocyte membrane, and dissolution

The understanding of cellular modifications is used to describe the reproductive cycle. This allows the recognition of the reproduction period and the gonadal morphological changes that occur. Descriptions of the reproductive cycle were initially elaborated by Yamamoto [33] and Agostinho et al. [34, 35], revalidated by Vazzoler [14], and later used by many authors. Next, Nuñez and Duponchelle [10] defined five stages of ovarian development with greater cellular detail and other four stages of testicular development based on macro- and microscopic characteristics. The last descriptions made by Lowerre-Barbieri et al. [24] and Quagio-Grassiotto et al. [19] on oocyte development, coupled with the stages of the reproductive cycle described by Brown-Peterson et al. [18], brought a proposal to homogenize the terms used and that has been applied in more recent studies. Research on the reproductive cycle of a given species helps to define determinant phases of gonadal development, as well as the "triggers" that initiate the process of cell proliferation and differentiation in the

**2. Important cellular morphological modifications during the oocyte** 

communication between the somatic cells of the follicle and the oocyte. This association between somatic cell and germ cell in the ovaries of various vertebrate and invertebrate species is established through intercellular junctions [39–43]. In vertebrate ovarian follicles, direct cytoplasmic connections between the oocyte and follicular cells of the granulosa layer associated with it are established early in the oocyte development. In fish, amphibians, and mammals, these cytoplasmic connections are established at the points of contact between the oocyte microvilli and follicular cells or between follicular cell microvilli and oocyte, via specialized

membrane junctions known as GAP junctions [44–47].

**2.1 Cellular junctions and your distribution throughout the oocyte development**

The growth and maturation of the ovarian follicles depend on carefully crafted

GAP junctions are intermembrane channel aggregates between adjacent cells composed by connexin proteins [48]. These junctions are considered homologous when they connect follicular cells to follicular cells and heterologous when they connect follicular cells to the oocyte [49]. Recent observations suggest that the functional coupling of GAP junctions, especially homologous ones, is necessary for the occurrence of the oocyte maturation process [50]. A possible role for the heterologous GAP junctions is the transfer of cAMP (PKA activator) from the follicular cells to the oocyte in order to induce the production or activity of membrane receptors

of yolk platelets; pelagic oocytes still undergo hydration [6].

*DOI: http://dx.doi.org/10.5772/intechopen.87967*

primary or previtellogenic growth.

formation of gametes [14, 36–38].

**development process**

#### *A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key… DOI: http://dx.doi.org/10.5772/intechopen.87967*

as clusters of organelles located near the nucleus, which proliferate intensely and spread throughout the cytoplasm. And, the cortical alveoli, as observed by Grier et al. [29], are vesicles filled with glycoproteins, formed by depressions of the oocyte membrane that become progressively larger, marking the final stage of primary or previtellogenic growth.

The described changes are followed by an expressive growth of the oocyte during vitellogenesis, in which the oocyte accumulates the nutritive reserves necessary for the development of the embryo. The oocyte also accumulates RNA and completes the differentiation of its cellular and noncellular envelopes. During this time, the oocyte interrupts the meiosis at the end of the prophase and in the diplotene stage. Maturation processes are characterized by the reduction or halting of endocytosis, resumption of meiosis, breakdown of the germinal vesicle, formation of a monolayer of cortical alveoli under the plasma oocyte membrane, and dissolution of yolk platelets; pelagic oocytes still undergo hydration [6].

The understanding of cellular modifications is used to describe the reproductive cycle. This allows the recognition of the reproduction period and the gonadal morphological changes that occur. Descriptions of the reproductive cycle were initially elaborated by Yamamoto [33] and Agostinho et al. [34, 35], revalidated by Vazzoler [14], and later used by many authors. Next, Nuñez and Duponchelle [10] defined five stages of ovarian development with greater cellular detail and other four stages of testicular development based on macro- and microscopic characteristics. The last descriptions made by Lowerre-Barbieri et al. [24] and Quagio-Grassiotto et al. [19] on oocyte development, coupled with the stages of the reproductive cycle described by Brown-Peterson et al. [18], brought a proposal to homogenize the terms used and that has been applied in more recent studies. Research on the reproductive cycle of a given species helps to define determinant phases of gonadal development, as well as the "triggers" that initiate the process of cell proliferation and differentiation in the formation of gametes [14, 36–38].

#### **2. Important cellular morphological modifications during the oocyte development process**

#### **2.1 Cellular junctions and your distribution throughout the oocyte development**

The growth and maturation of the ovarian follicles depend on carefully crafted communication between the somatic cells of the follicle and the oocyte. This association between somatic cell and germ cell in the ovaries of various vertebrate and invertebrate species is established through intercellular junctions [39–43]. In vertebrate ovarian follicles, direct cytoplasmic connections between the oocyte and follicular cells of the granulosa layer associated with it are established early in the oocyte development. In fish, amphibians, and mammals, these cytoplasmic connections are established at the points of contact between the oocyte microvilli and follicular cells or between follicular cell microvilli and oocyte, via specialized membrane junctions known as GAP junctions [44–47].

GAP junctions are intermembrane channel aggregates between adjacent cells composed by connexin proteins [48]. These junctions are considered homologous when they connect follicular cells to follicular cells and heterologous when they connect follicular cells to the oocyte [49]. Recent observations suggest that the functional coupling of GAP junctions, especially homologous ones, is necessary for the occurrence of the oocyte maturation process [50]. A possible role for the heterologous GAP junctions is the transfer of cAMP (PKA activator) from the follicular cells to the oocyte in order to induce the production or activity of membrane receptors

*Reproductive Biology and Technology in Animals*

reproductive cycle [8–12], and fecundity [13].

Teleostei, as in recent studies on spermatogenesis [2–5], folliculogenesis [6–8],

Studies have been carried out to describe and classify the stages of gonadal development and reproductive stages of fish in the Neotropical region. One of the most classic and used bibliographic sources has been Vazzoler [14]. However, other proposals for description have already been made by Grier and Taylor [15], Grier [16], and Lo Nostro et al. [17], which detail the continuity and discontinuity of the germinal epithelium and the cell types present in the gonads. Recently, Brown-Peterson et al. [18] developed a simpler terminology to facilitate the communication and comparison of studies on the reproductive biology of fish. Still in order to make the nomenclature more comprehensive, the stages of oocyte development were simplified by Quagio-Grassiotto et al. [19], and the development of stages of atresia, which are character-

Gonadal development can be analyzed macroscopically, and changes in shape, size,

color, and texture of the gonads have been used as parameters for the classification of maturation status in many studies of ecology and reproductive dynamics[14, 21]. However, the most used analysis has been of the microscopic characters, since it allows a more detailed and precise description of the transitions and morphological and structural transformations that happen during gonadal development [8, 22, 23]. Thus,

• Spermatogenesis shows stages of development that include spermatogonia,

• Oogenesis usually shows the following progression: oogonia, primary growth oocytes, a previtellogenic stage in which oocytes grow larger and often have cortical alveolar vesicles, an extensive vitellogenic phase, oocyte maturation,

The oocyte development in a mature egg is a complex process modulated by numerous environmental and endocrine factors [25], and understanding the morphological characteristics of oocytes is important to interpret the dynamics of oogenesis [26]. Among the oocyte processes, folliculogenesis results in the removal of the primary oocyte from oogonium nests and consequent formation of ovarian follicles [27]. Descriptions for the germinal epithelium made by Grier [28] conceptualized "follicular complex" as the functional unit of the ovary. This complex is formed by two compartments separated by a basement membrane. One compartment is the follicle, which consists of the oocyte surrounded by follicular cells and originated from the germinal epithelium. The second compartment is the theca,

In the previtellogenic oocyte phase, multiple nucleoli are observed, as described by Grier et al. [29]. These oocytes are also called perinucleolar oocytes, when the nucleoli migrate to the nuclear periphery. There is also the formation of the zona pellucida, a complex structure consisting generally of two layers crossed by pores or channels containing the oocyte microvilli and/or follicular cell extensions. The zona pellucida reflects adaptations to different ecological conditions in which the eggs develop [30], whose inner layer protects the egg from mechanical damage and

Another cell characteristic that is used to describe the stages of oocyte development is the presence of nüages, Balbiani corpuscles, and cortical alveoli. The nüages are originated by the transfer from the nucleus to the cytoplasm of large amounts of heterogeneous and ribosomal RNA synthesized [31] and associated with proteins. Balbiani corpuscles or yolk nuclei, described by Hubbard [32], were recognized

ized as involutive processes, follows according to Miranda et al. [20].

regarding the microscopic aspects of the gonad, it is verified that [24]:

spermatocytes, spermatids, and spermatozoa.

made up of undifferentiated ovarian stromal cells.

whose outer layer protects it from microorganisms.

and ovulation.

**92**

for the maturation-inducing hormone, or MIH [50], indirectly participating in the oocyte maturation process. GAP junctions may also be involved in specifying the pattern of polarity in the oocytes of various animal groups, so this junctional route can be used to pass intercellular signals from follicular cells to the oocyte to determine oocyte symmetry [51].

As previously reported, the fish oocyte is enveloped by the zona pellucida (microvillus area), by the follicular cells and by the basement membrane. Thus, from a morphological and functional point of view, it is important to know if there are any tight junctions between adjacent follicular cells, since these joints promote barriers for the passage of fluids through the extracellular space between adjacent cell membranes and maintain tissue and cell integrity [45, 52–54]. The main components of the intercellular junctions are the tight junctions [55, 56], which are composed of different transmembrane proteins that promote a homophilic interaction. The cytoplasmic domain of the transmembrane adhesion molecules connects the binding proteins which, in turn, anchor the cytoskeletal adhesion complex. Of these molecules, occludins and claudins are the most extensively studied. Although occludin is a highly conserved molecule, claudins comprise a family of more than 20 different proteins, some of which are expressed in a tissue-specific manner [57–59].

As claudins, cadherins are a transmembrane superfamily of proteins that contain several homologous members, exhibiting tissue diversity and distinct binding specificities [60–62], with a highly conserved cytoplasmic domain [63, 64]. These molecules mediate cell-cell contact at adhesion junctions also anchored in the cytoskeleton, thus playing an important role in the separation, positioning and control of cell movements, and in morphogenesis [65–67]. In a study with *Danio rerio*, E-cadherin homologous proteins were identified, and their synthesis and storage during oogenesis were verified [62]. Also, the establishment of heterotypic junctions linking the oocyte to follicular cells throughout folliculogenesis and cooperating in the determination of follicle architecture was observed [62]. When oocytes progress in vitellogenesis, the localization of adhesion proteins in the oocyte becomes restricted to a more specific pattern, which reflects the points of contact between the oocyte and the follicle cells and their adjustment to changes in the oocyte cytoskeleton throughout this phase [62].

#### **2.2 Distribution and structuration of the cytoskeleton throughout the oocyte development**

All intracytoplasmic and cortical events in oocytes involve a precisely organized and collaborative contractile system and a stable support matrix [68]. The cytoskeleton of the oocytes and embryos is implicated in key developmental events, such as creation and maintenance of axial polarity, cytoplasmic reorganization, cell division, change of surface architecture, morphogenetic motions, and internal arrangement of organelles [69]. It seems very likely that cytoskeletal structures are responsible for spatial distribution of yolk, cortical and pigment granules, lipid droplets, or mitochondria [68, 70]. Thus, the spatial organization of cytoskeletal filaments may be important for the preservation of oocyte viability [71].

Among the different proteins expressed in the cytoskeleton, the intermediate filament proteins are exceptionally complex [72, 73], especially in the class of cytokeratins. This is a class of proteins typical and specifically induced in cells compromised for epithelial differentiation [72, 74, 75], and their identification in oocytes and eggs presents an interesting contrast when comparing to other cytoskeletal proteins in germ cells. Cytokeratins are not synthesized in previtellogenic oocytes but are expressed and accumulated in the vitellogenic stage. These filament proteins are first detectable in the cortex of oocytes in later stages of previtellogenesis; at

**95**

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key…*

the beginning of vitellogenesis, they are distributed primarily in the region closest to the nucleus and appear to become cortical again in mature oocytes [76]. Intermediate filaments of cytokeratin contribute to the complex structure of the oocyte and egg cortex, which is also rich in other cytoskeletal filaments such as

The microtubule matrix seems to be a very important component in the immature oocyte cortex in fish. The function of the cortical matrix of microtubules in oocytes remains undetermined but may be related to the mechanical stiffness that has been attributed to the cortex [80]. Even the basic mechanism of germinal vesicle migration and its mechanical anchoring in the region of the animal pole occur from the depolarization of the microtubules, leading to a consequent change in the posi-

Evident changes occur in the distribution and localization of tubulin-containing structures in growing oocytes. In previtellogenic oocytes, a great amount of tubulin is concentrated in the Balbiani corpuscle [82–85]. During vitellogenesis, mitochondria are displaced from the Balbiani corpuscle to the surface of the cell, while others remain around the nucleus [82, 86], and this movement seems to be related to the reorganization of tubulin [87]. With the disintegration of the Balbiani corpuscle, due to the anterior displacement of membranous organelles, the released space is gradually filled with yolk, i.e., the yolk granules are in a tubulin-positive region. As vitellogenesis progresses, rearrangement of cell growth and its contents occurs with the movement of endosomes to transport yolk through the microtubules [87].

The proper organization and assembly of the cytoskeleton microtubule is an integral phenomenon, which is related to the expression of cellular asymmetry. Particularly in oocytes, the microtubules exhibit a unique paradigm as forming an eccentric meiotic spindle which, consequently, gives rise to asymmetric cytokinesis to form the first and second polar bodies. Its existence and function are dynamically regulated throughout the process of cell division, particularly during the S and M phases of the cell cycle [88]. Another element that contributes to the oocyte asymmetry is the actin cytoskeleton. In oocytes, the actin filaments are not randomly distributed within the cell [89]. In germ cells, as in many other cells, two types of actin are present: filamentous (F-actin) and non-filamentous (G-actin) actins [90]. Actin polymerization-depolymerization process is essential for the translocation of many organelles, as mitochondria [91], Golgi system [92], and cortical granules [93, 94], as well as for the regulation of ion channel activity [95]. In addition, a certain proportion of F-actin and G-actin is required for the normal course of meiotic and mitotic divisions [96]. In many cells, a significant part of these filaments is in the area of the cellular cortex, so it has been proposed that they take part in the transduction of transmembrane information signals, including hormonal signaling [97, 98]. Still in the oocyte cortex, the cortex-specific F-actin layer is peculiarly absent in the space between the germinal vesicle and the plasma membrane at the animal pole. In fact, it is through this "corridor" that the two polar bodies are extruded in the posterior phase of meiosis [99, 100]. The formation of actin bundles in the oocyte cortex is one of the first morphological markers of induction to maturation [99]. The role of actin microfilaments in oocyte maturation seems to be related to the translocation of the endoplasmic reticulum structures to the germinal vesicle area and to the coordination of the cortical granules in the plasma membrane zone [93, 101]. Even during follicular atresia, the actin cytoskeleton undergoes changes associated with the yolk degradation, while it remains preserved in follicular cells. Thus, maintenance of the actin cytoskeleton may be a sign of survival for follicular cells during early and/or advanced atresia processes [102]. Cytoskeleton changes have been extensively reported in apoptotic cells, among which changes in cell shape and anchorage are dependent on

the reorganization of actin filaments and focal adhesion contacts [103].

*DOI: http://dx.doi.org/10.5772/intechopen.87967*

actin filaments and microtubules [68, 77–79].

tion of the germinal vesicle [80, 81].

#### *A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key… DOI: http://dx.doi.org/10.5772/intechopen.87967*

the beginning of vitellogenesis, they are distributed primarily in the region closest to the nucleus and appear to become cortical again in mature oocytes [76]. Intermediate filaments of cytokeratin contribute to the complex structure of the oocyte and egg cortex, which is also rich in other cytoskeletal filaments such as actin filaments and microtubules [68, 77–79].

The microtubule matrix seems to be a very important component in the immature oocyte cortex in fish. The function of the cortical matrix of microtubules in oocytes remains undetermined but may be related to the mechanical stiffness that has been attributed to the cortex [80]. Even the basic mechanism of germinal vesicle migration and its mechanical anchoring in the region of the animal pole occur from the depolarization of the microtubules, leading to a consequent change in the position of the germinal vesicle [80, 81].

Evident changes occur in the distribution and localization of tubulin-containing structures in growing oocytes. In previtellogenic oocytes, a great amount of tubulin is concentrated in the Balbiani corpuscle [82–85]. During vitellogenesis, mitochondria are displaced from the Balbiani corpuscle to the surface of the cell, while others remain around the nucleus [82, 86], and this movement seems to be related to the reorganization of tubulin [87]. With the disintegration of the Balbiani corpuscle, due to the anterior displacement of membranous organelles, the released space is gradually filled with yolk, i.e., the yolk granules are in a tubulin-positive region. As vitellogenesis progresses, rearrangement of cell growth and its contents occurs with the movement of endosomes to transport yolk through the microtubules [87].

The proper organization and assembly of the cytoskeleton microtubule is an integral phenomenon, which is related to the expression of cellular asymmetry. Particularly in oocytes, the microtubules exhibit a unique paradigm as forming an eccentric meiotic spindle which, consequently, gives rise to asymmetric cytokinesis to form the first and second polar bodies. Its existence and function are dynamically regulated throughout the process of cell division, particularly during the S and M phases of the cell cycle [88].

Another element that contributes to the oocyte asymmetry is the actin cytoskeleton. In oocytes, the actin filaments are not randomly distributed within the cell [89]. In germ cells, as in many other cells, two types of actin are present: filamentous (F-actin) and non-filamentous (G-actin) actins [90]. Actin polymerization-depolymerization process is essential for the translocation of many organelles, as mitochondria [91], Golgi system [92], and cortical granules [93, 94], as well as for the regulation of ion channel activity [95]. In addition, a certain proportion of F-actin and G-actin is required for the normal course of meiotic and mitotic divisions [96].

In many cells, a significant part of these filaments is in the area of the cellular cortex, so it has been proposed that they take part in the transduction of transmembrane information signals, including hormonal signaling [97, 98]. Still in the oocyte cortex, the cortex-specific F-actin layer is peculiarly absent in the space between the germinal vesicle and the plasma membrane at the animal pole. In fact, it is through this "corridor" that the two polar bodies are extruded in the posterior phase of meiosis [99, 100].

The formation of actin bundles in the oocyte cortex is one of the first morphological markers of induction to maturation [99]. The role of actin microfilaments in oocyte maturation seems to be related to the translocation of the endoplasmic reticulum structures to the germinal vesicle area and to the coordination of the cortical granules in the plasma membrane zone [93, 101]. Even during follicular atresia, the actin cytoskeleton undergoes changes associated with the yolk degradation, while it remains preserved in follicular cells. Thus, maintenance of the actin cytoskeleton may be a sign of survival for follicular cells during early and/or advanced atresia processes [102]. Cytoskeleton changes have been extensively reported in apoptotic cells, among which changes in cell shape and anchorage are dependent on the reorganization of actin filaments and focal adhesion contacts [103].

*Reproductive Biology and Technology in Animals*

mine oocyte symmetry [51].

for the maturation-inducing hormone, or MIH [50], indirectly participating in the oocyte maturation process. GAP junctions may also be involved in specifying the pattern of polarity in the oocytes of various animal groups, so this junctional route can be used to pass intercellular signals from follicular cells to the oocyte to deter-

As previously reported, the fish oocyte is enveloped by the zona pellucida (microvillus area), by the follicular cells and by the basement membrane. Thus, from a morphological and functional point of view, it is important to know if there are any tight junctions between adjacent follicular cells, since these joints promote barriers for the passage of fluids through the extracellular space between adjacent cell membranes and maintain tissue and cell integrity [45, 52–54]. The main components of the intercellular junctions are the tight junctions [55, 56], which are composed of different transmembrane proteins that promote a homophilic interaction. The cytoplasmic domain of the transmembrane adhesion molecules connects the binding proteins which, in turn, anchor the cytoskeletal adhesion complex. Of these molecules, occludins and claudins are the most extensively studied. Although occludin is a highly conserved molecule, claudins comprise a family of more than 20 different proteins, some of which are expressed in a tissue-specific manner [57–59]. As claudins, cadherins are a transmembrane superfamily of proteins that contain

several homologous members, exhibiting tissue diversity and distinct binding specificities [60–62], with a highly conserved cytoplasmic domain [63, 64]. These molecules mediate cell-cell contact at adhesion junctions also anchored in the cytoskeleton, thus playing an important role in the separation, positioning and control of cell movements, and in morphogenesis [65–67]. In a study with *Danio rerio*, E-cadherin homologous proteins were identified, and their synthesis and storage during oogenesis were verified [62]. Also, the establishment of heterotypic junctions linking the oocyte to follicular cells throughout folliculogenesis and cooperating in the determination of follicle architecture was observed [62]. When oocytes progress in vitellogenesis, the localization of adhesion proteins in the oocyte becomes restricted to a more specific pattern, which reflects the points of contact between the oocyte and the follicle cells and their adjustment to changes in

**2.2 Distribution and structuration of the cytoskeleton throughout the oocyte** 

filaments may be important for the preservation of oocyte viability [71].

All intracytoplasmic and cortical events in oocytes involve a precisely organized and collaborative contractile system and a stable support matrix [68]. The cytoskeleton of the oocytes and embryos is implicated in key developmental events, such as creation and maintenance of axial polarity, cytoplasmic reorganization, cell division, change of surface architecture, morphogenetic motions, and internal arrangement of organelles [69]. It seems very likely that cytoskeletal structures are responsible for spatial distribution of yolk, cortical and pigment granules, lipid droplets, or mitochondria [68, 70]. Thus, the spatial organization of cytoskeletal

Among the different proteins expressed in the cytoskeleton, the intermediate filament proteins are exceptionally complex [72, 73], especially in the class of cytokeratins. This is a class of proteins typical and specifically induced in cells compromised for epithelial differentiation [72, 74, 75], and their identification in oocytes and eggs presents an interesting contrast when comparing to other cytoskeletal proteins in germ cells. Cytokeratins are not synthesized in previtellogenic oocytes but are expressed and accumulated in the vitellogenic stage. These filament proteins are first detectable in the cortex of oocytes in later stages of previtellogenesis; at

the oocyte cytoskeleton throughout this phase [62].

**development**

**94**

#### **3. Morphological characteristics related to ovarian reorganization**

#### **3.1 Processes of atresia and cellular proliferation**

Atresia is a degenerative process by which the ovarian follicles lose their integrity and are eliminated [104]. It is a common process in vertebrate ovaries under natural and/or experimental conditions [105] and can be induced by a series of exogenous and endogenous factors [106–109]. Oocyte degeneration, or follicular atresia, is a process that may occur before spawning, in oocytes that have not reached maturity and thereafter in oocytes that are no longer ovulated [110, 111]. In fish, atresia is involved in normal ovary growth [112, 113] and postovulatory regression [114–116], especially in females that are not able to perform maturation or ovulation after the vitellogenesis process [117].

Fish, in general, exhibits a reproductive periodicity, and, therefore, oocytes at various stages of development may be resorbed with the resultant formation of an atretic body. Considering the foregoing, Rajalakshmi [118] made a classification of the atretic processes taking into account the following stages: (1) "immature oocyte atresia" begins with the distortion of the cell shape, followed by loss of cytoplasmic homogeneity and reabsorption of the structure (in this type of atresia, the follicular cells do not exhibit any activity so the reabsorption of oocytes without yolk seems to be a relatively simple process); (2) "mature oocyte atresia" begins with the loss of the soft outline of the zona pellucida and dissociation of the follicular cells, which will then present phagocytic characteristic (i.e., enzymatic activity of acid phosphatase that will liquefy the yolk), followed by a slow dissolution of the zona pellucida and culminating in total resorption of the follicle; (3) "postovulatory complex atresia" begins with the distortion of the follicular cell shape, followed by loss of cell boundaries and formation of a syncytial structure, and finally the follicle shrink, with consequent degenerative changes.

The morphological characteristics of the atretic bodies and their stages of involution, independent of cellular development stage, were summarized in the study of Miranda et al. [20], as (1) initial atresia, with the disintegration of the oocyte nucleus, fragmentation of the zona pellucida, and follicular cell hypertrophy; (2) intermediate atresia, with follicular cells presenting phagocytic characteristics and ingesting the yolk; (3) advanced atresia, with numerous myelinic figures in the cytoplasm of follicular cells; and (4) final atresia, with the reduction in the number of follicular and theca cells and presence of granules of lipofuscin and granulocytes near the atretic follicle. With the current emergence of the theme of cell death pathways, studies about ovarian involutive processes in fish were brought to the spotlight again with new descriptions being made [102, 108, 116, 119–124] that add and/or corroborate those morphological characteristics already proposed by Miranda et al. [20].

In fish, mammals and, presumably, other vertebrates, the molecular mechanism responsible for ovarian follicular atresia is cell death by apoptosis [102, 124, 125]. Apoptosis, or programmed cell death, is a physiological process controlled by various hormones and growth factors. This is an evolutionarily conserved process, involved in remodeling, differentiation, and tissue degeneration in a variety of cell types [125]. It is characterized by biochemical and morphological changes such as chromatin condensation, DNA fragmentation, and the formation of apoptotic bodies [126]. The main effector proteins in apoptosis are the caspases, a family of highly conserved cysteine proteases [127, 128]. Among the caspases, caspase-3 is the major effector one, including in the ovarian tissue in which it is expressed in the follicular cells of atretic follicles of fish and mammals [102, 124, 129].

In addition to apoptosis, Thomé et al. [130] presented a new route to cell death the autophagy. This route differs from apoptosis by the purpose of the processes:

**97**

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key…*

The mechanism of cell proliferation is a highly regulated process that seems to be essential for the maintenance of ovarian homeostasis [137], and yet the hormonal mechanism controlling oocyte proliferation and recruitment of oocytes is not understood completely for any vertebrate [6]. In contrast to mammals, oogonia continue to proliferate in adult female fish [138], thus renewing stocks of young oocytes and follicles [139, 140]. The pre-follicular and follicular cells begin to proliferate when in the folliculogenesis phase, to support the oocyte growth [19]. However, ovarian mitosis in fish is usually observed at the end of each reproductive cycle [137], when ovarian tissues are reorganized [141, 142]. Throughout ovigerous lamellae in adult females, germ cell proliferation and the formation of germline cysts result in extensions of the germinal epithelium that are segregated from the connective tissue by a basement membrane [19]. These extensions of the germinal epithelium are known as oogonium nests [28, 143] and mark the beginning of the

**3.2 Extracellular matrix and its changes through the reproductive cycle**

During the reproductive cycle, ovarian tissue is constantly remodeled, with extensive cell proliferation and differentiation, as well as extracellular matrix changes from early follicular development to tissue involution after ovulation [144]. Among the processes and factors involved in tissue remodeling are apoptosis, changes in hormone levels, and degradation of the extracellular matrix in contact

The extracellular matrix is an insoluble network of several structural and functional macromolecules found in connective tissues and basement membranes [145]. It is both a barrier that separates the organism into tissue compartments and a substrate for cell adhesion [146]. In addition to these structural functions, the extracellular matrix is an essential regulator of cellular physiology, predominantly

A coordinated interaction of signals is necessary to regulate the proliferation, differentiation, adhesion, and migration of specific cell types for the development and organization of structural tissues [148]. During the normal development of an organ or in pathological modifications, the matrix undergoes intense changes in its composition. This process, called matrix remodeling, is involved in many

in cell survival, cell cycle, cell migration, and morphogenesis [147].

apoptosis is the programmed cell death, and autophagy is a stress adaptation to prevent cell death. The functional relationship between apoptosis and autophagy is complex. In some cases, autophagy is a form of adaptation to suppress apoptosis, whereas, in other cases, autophagy constitutes an alternative pathway of cellular elimination called autophagic or type II cell death [131–133]. It has been understood that apoptosis is the main mechanism involved in the involution of postovulatory follicles [116, 121], while autophagy contributes to the regression of atretic follicles [20, 130]. Even though the limits and interrelationships between these two processes have not yet been well established, recent studies have shown that there may be a crosstalk between autophagy and apoptosis pathways in the ovarian involution processes. A fine balance between the signs for survival and cell death appears to be essential for determining the fate of follicular cells, particularly in follicular atresia [102, 124]. During follicular development, a low rate of follicular cell apoptosis can be considered as a physiological event for the control of the appropriate number of cells and elimination of the undesirable ones [134]. However, high apoptosis values can be observed under unfavorable conditions, compromising follicular viability [135]. Thus, organic homeostasis is dependent on the balance between cell proliferation, differentiation, and death, so populations of rapidly proliferating cells usually have

*DOI: http://dx.doi.org/10.5772/intechopen.87967*

high rates of cell death by apoptosis [125, 136].

reproductive cycle again.

with cells [134].

#### *A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key… DOI: http://dx.doi.org/10.5772/intechopen.87967*

apoptosis is the programmed cell death, and autophagy is a stress adaptation to prevent cell death. The functional relationship between apoptosis and autophagy is complex. In some cases, autophagy is a form of adaptation to suppress apoptosis, whereas, in other cases, autophagy constitutes an alternative pathway of cellular elimination called autophagic or type II cell death [131–133]. It has been understood that apoptosis is the main mechanism involved in the involution of postovulatory follicles [116, 121], while autophagy contributes to the regression of atretic follicles [20, 130]. Even though the limits and interrelationships between these two processes have not yet been well established, recent studies have shown that there may be a crosstalk between autophagy and apoptosis pathways in the ovarian involution processes. A fine balance between the signs for survival and cell death appears to be essential for determining the fate of follicular cells, particularly in follicular atresia [102, 124].

During follicular development, a low rate of follicular cell apoptosis can be considered as a physiological event for the control of the appropriate number of cells and elimination of the undesirable ones [134]. However, high apoptosis values can be observed under unfavorable conditions, compromising follicular viability [135]. Thus, organic homeostasis is dependent on the balance between cell proliferation, differentiation, and death, so populations of rapidly proliferating cells usually have high rates of cell death by apoptosis [125, 136].

The mechanism of cell proliferation is a highly regulated process that seems to be essential for the maintenance of ovarian homeostasis [137], and yet the hormonal mechanism controlling oocyte proliferation and recruitment of oocytes is not understood completely for any vertebrate [6]. In contrast to mammals, oogonia continue to proliferate in adult female fish [138], thus renewing stocks of young oocytes and follicles [139, 140]. The pre-follicular and follicular cells begin to proliferate when in the folliculogenesis phase, to support the oocyte growth [19]. However, ovarian mitosis in fish is usually observed at the end of each reproductive cycle [137], when ovarian tissues are reorganized [141, 142]. Throughout ovigerous lamellae in adult females, germ cell proliferation and the formation of germline cysts result in extensions of the germinal epithelium that are segregated from the connective tissue by a basement membrane [19]. These extensions of the germinal epithelium are known as oogonium nests [28, 143] and mark the beginning of the reproductive cycle again.

#### **3.2 Extracellular matrix and its changes through the reproductive cycle**

During the reproductive cycle, ovarian tissue is constantly remodeled, with extensive cell proliferation and differentiation, as well as extracellular matrix changes from early follicular development to tissue involution after ovulation [144]. Among the processes and factors involved in tissue remodeling are apoptosis, changes in hormone levels, and degradation of the extracellular matrix in contact with cells [134].

The extracellular matrix is an insoluble network of several structural and functional macromolecules found in connective tissues and basement membranes [145]. It is both a barrier that separates the organism into tissue compartments and a substrate for cell adhesion [146]. In addition to these structural functions, the extracellular matrix is an essential regulator of cellular physiology, predominantly in cell survival, cell cycle, cell migration, and morphogenesis [147].

A coordinated interaction of signals is necessary to regulate the proliferation, differentiation, adhesion, and migration of specific cell types for the development and organization of structural tissues [148]. During the normal development of an organ or in pathological modifications, the matrix undergoes intense changes in its composition. This process, called matrix remodeling, is involved in many

*Reproductive Biology and Technology in Animals*

vitellogenesis process [117].

with consequent degenerative changes.

**3.1 Processes of atresia and cellular proliferation**

**3. Morphological characteristics related to ovarian reorganization**

Atresia is a degenerative process by which the ovarian follicles lose their integrity and are eliminated [104]. It is a common process in vertebrate ovaries under natural and/or experimental conditions [105] and can be induced by a series of exogenous and endogenous factors [106–109]. Oocyte degeneration, or follicular atresia, is a process that may occur before spawning, in oocytes that have not reached maturity and thereafter in oocytes that are no longer ovulated [110, 111]. In fish, atresia is involved in normal ovary growth [112, 113] and postovulatory regression [114–116], especially in females that are not able to perform maturation or ovulation after the

Fish, in general, exhibits a reproductive periodicity, and, therefore, oocytes at various stages of development may be resorbed with the resultant formation of an atretic body. Considering the foregoing, Rajalakshmi [118] made a classification of the atretic processes taking into account the following stages: (1) "immature oocyte atresia" begins with the distortion of the cell shape, followed by loss of cytoplasmic homogeneity and reabsorption of the structure (in this type of atresia, the follicular cells do not exhibit any activity so the reabsorption of oocytes without yolk seems to be a relatively simple process); (2) "mature oocyte atresia" begins with the loss of the soft outline of the zona pellucida and dissociation of the follicular cells, which will then present phagocytic characteristic (i.e., enzymatic activity of acid phosphatase that will liquefy the yolk), followed by a slow dissolution of the zona pellucida and culminating in total resorption of the follicle; (3) "postovulatory complex atresia" begins with the distortion of the follicular cell shape, followed by loss of cell boundaries and formation of a syncytial structure, and finally the follicle shrink,

The morphological characteristics of the atretic bodies and their stages of involution, independent of cellular development stage, were summarized in the study of Miranda et al. [20], as (1) initial atresia, with the disintegration of the oocyte nucleus, fragmentation of the zona pellucida, and follicular cell hypertrophy; (2) intermediate atresia, with follicular cells presenting phagocytic characteristics and ingesting the yolk; (3) advanced atresia, with numerous myelinic figures in the cytoplasm of follicular cells; and (4) final atresia, with the reduction in the number of follicular and theca cells and presence of granules of lipofuscin and granulocytes near the atretic follicle. With the current emergence of the theme of cell death pathways, studies about ovarian involutive processes in fish were brought to the spotlight again with new descriptions being made [102, 108, 116, 119–124] that add and/or corroborate those morphological characteristics already proposed by Miranda et al. [20].

In fish, mammals and, presumably, other vertebrates, the molecular mechanism responsible for ovarian follicular atresia is cell death by apoptosis [102, 124, 125]. Apoptosis, or programmed cell death, is a physiological process controlled by various hormones and growth factors. This is an evolutionarily conserved process, involved in remodeling, differentiation, and tissue degeneration in a variety of cell types [125]. It is characterized by biochemical and morphological changes such as chromatin condensation, DNA fragmentation, and the formation of apoptotic bodies [126]. The main effector proteins in apoptosis are the caspases, a family of highly conserved cysteine proteases [127, 128]. Among the caspases, caspase-3 is the major effector one, including in the ovarian tissue in which it is expressed in the follicular

In addition to apoptosis, Thomé et al. [130] presented a new route to cell death the autophagy. This route differs from apoptosis by the purpose of the processes:

cells of atretic follicles of fish and mammals [102, 124, 129].

**96**

physiological processes, such as activation of immune cells [149], wound healing [150, 151], embryogenesis [152, 153], or reproductive cycle [154].

The extracellular matrix-cell interactions influence gene regulation, cytoskeletal structure, differentiation, and many aspects of cell growth [155]. Changes in the expression of components that make up the extracellular matrix accompany follicular growth, ovulation, and involution of postovulatory follicles, which in its turn may influence follicular maturation, cell survival, and steroidogenesis [134, 156, 157]. Studies with mammals demonstrate that gonadal support cells synthesize a variety of components comprising the extracellular matrix and the basement membrane, such as collagen, laminin, keratin, fibronectin, lectin, and fibril chains [158, 159].

The balance between the degradation and regeneration of the extracellular matrix in ovarian tissues is maintained, in part, by the action of extracellular proteolytic enzymes that are secreted by the local cells. Most of these enzymes are matrix metalloproteinases (MMPs), which depend on the Ca+2 or Zn+2 binding to their activity [160]. During oogenesis, great changes in the extracellular environment of the ovary were largely attributed to the action of MMPs [144]. MMPs play an important role in the ovulation process in different groups of vertebrates, acting on follicular rupture, basement membrane fragmentation, and follicular connective fibers [144, 161, 162].

The integrity of the basement membrane is also evidenced by the continuous marking of laminin-β2 and type IV collagen, which allows the development of ovarian follicles [159, 163]. On the other hand, the discontinuous labeling of laminin-β2 and type IV collagen in the basal membrane of postovulatory follicles indicates that basement membrane degradation occurs due to the breakdown of these major components [134]. The loss of the basement membrane integrity may contribute to the increase of follicular cell apoptosis, suggesting its influence on the survival of postovulatory follicle cells [116].

Fibronectin and laminin have been shown to be extracellular matrix proteins synthesized by follicular cells [164, 165]. The presence of fibronectin on the surface of postovulatory follicle cells is due to the need of interaction between their domains with type IV collagen and cell surface integrin, and it is important for the maintenance of cell adhesion in the extracellular matrix [159]. According to Iwahashi et al. [166], the type IV collagen detected in the connective tissue among theca cells may be involved in the organization of extracellular fibronectin. This interaction between type IV collagen and fibronectin may act on cell migration that occurs during the late remodeling of postovulatory follicles [134].

Thus, the structure and composition of the extracellular matrix play an important role during follicular development and post-spawning involution in teleost fish. The basement membrane integrity is important for follicular cell survival, and the loss of integrity contributes to increased follicular apoptosis. In addition, MMP-9 may be involved in the final oocyte maturation and regression of postovulatory follicles [134]. Therefore, it follows that different combinations and proportions in the assembly of extracellular matrix components, together with the presentation of a large variety of proteoglycans at various times during the development and maturation of the gonads, can orchestrate distinct gene expression programs and culminate in more diverse tissue variations and adaptations [148].

#### **4. Conclusions**

Studies in gametogenesis help to understand the ecological, adaptive, and evolutionary relationships in the groups of species, especially when the oocyte structures are analyzed in an ultrastructural level. This is even more important when we

**99**

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key…*

consider that there are few fish species that present descriptions with adequate morphological and/or functional detail. Most of the studies do not evaluate the reproductive characteristics with the necessary histological and ultrastructural details, which can lead to incomplete interpretations of the reproductive characteristics of the species. Likewise, studies involving organelles and their distribution throughout the reproductive cycle and cellular development in fish species are punctual or restricted to a developmental stage. The understanding of these processes is then due to the sum of several studies at different stages of development, but they do not necessarily represent the same environmental, behavioral, and population pressures that are being addressed to the individuals of a given species. Thus, the continuous study of these variables throughout the reproductive cycle of key species may allow more real parameters on the dynamics of the intracellular structures in germ cells and follicular cells, as well as the extracellular matrix. All of the above is even more relevant when applied to such a diverse group, as fish, that have great ecological,

I would like to thank all those who did or are part of the laboratories where I conducted my undergraduate and graduate studies and who culminated in the improvement of the knowledge reproduced here. I thank Dr. Adelina Ferreira, Dr. Mahmoud Mehanna, and Dr. Débora Fabiane Neves da Silva, advisors and colleagues at the Morphology and Morphometry Laboratory of the Federal University of Mato Grosso (UFMT), for the first steps taken in the area of Animal Reproduction. I would also like to thank Dr. Maria Inês Borella, Dr. Chayrra Chehade Gomes, Dr. Gisele Cristiane de Melo Dias, Dr. Lázaro Wender Oliveira de Jesus, MSc Giovana de Souza Branco, MSc Marília de Paiva Camargo, and laboratory technician Cruz Alberto Mendoza Rigonatti of the Laboratory of Fish Endocrinology, University of São Paulo, for the support during the doctorate and for all the knowledge obtained in that period. Lastly, I would like to thank the FAPEMAT, CAPES, and FAPESP funding agencies for the financial support provided during my academic trajectory,

The author declares that there is no conflict of interest regarding the publication

*DOI: http://dx.doi.org/10.5772/intechopen.87967*

social, and economic importance.

which culminated in the formulation of this chapter.

**Acknowledgements**

**Conflict of interest**

of this chapter.

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key… DOI: http://dx.doi.org/10.5772/intechopen.87967*

consider that there are few fish species that present descriptions with adequate morphological and/or functional detail. Most of the studies do not evaluate the reproductive characteristics with the necessary histological and ultrastructural details, which can lead to incomplete interpretations of the reproductive characteristics of the species. Likewise, studies involving organelles and their distribution throughout the reproductive cycle and cellular development in fish species are punctual or restricted to a developmental stage. The understanding of these processes is then due to the sum of several studies at different stages of development, but they do not necessarily represent the same environmental, behavioral, and population pressures that are being addressed to the individuals of a given species. Thus, the continuous study of these variables throughout the reproductive cycle of key species may allow more real parameters on the dynamics of the intracellular structures in germ cells and follicular cells, as well as the extracellular matrix. All of the above is even more relevant when applied to such a diverse group, as fish, that have great ecological, social, and economic importance.

#### **Acknowledgements**

*Reproductive Biology and Technology in Animals*

fibers [144, 161, 162].

postovulatory follicle cells [116].

physiological processes, such as activation of immune cells [149], wound healing

as collagen, laminin, keratin, fibronectin, lectin, and fibril chains [158, 159]. The balance between the degradation and regeneration of the extracellular matrix in ovarian tissues is maintained, in part, by the action of extracellular proteolytic enzymes that are secreted by the local cells. Most of these enzymes are matrix metalloproteinases (MMPs), which depend on the Ca+2 or Zn+2 binding to their activity [160]. During oogenesis, great changes in the extracellular environment of the ovary were largely attributed to the action of MMPs [144]. MMPs play an important role in the ovulation process in different groups of vertebrates, acting on follicular rupture, basement membrane fragmentation, and follicular connective

The integrity of the basement membrane is also evidenced by the continuous marking of laminin-β2 and type IV collagen, which allows the development of ovarian follicles [159, 163]. On the other hand, the discontinuous labeling of laminin-β2 and type IV collagen in the basal membrane of postovulatory follicles indicates that basement membrane degradation occurs due to the breakdown of these major components [134]. The loss of the basement membrane integrity may contribute to the increase of follicular cell apoptosis, suggesting its influence on the survival of

Fibronectin and laminin have been shown to be extracellular matrix proteins synthesized by follicular cells [164, 165]. The presence of fibronectin on the surface of postovulatory follicle cells is due to the need of interaction between their domains with type IV collagen and cell surface integrin, and it is important for the maintenance of cell adhesion in the extracellular matrix [159]. According to Iwahashi et al. [166], the type IV collagen detected in the connective tissue among theca cells may be involved in the organization of extracellular fibronectin. This interaction between type IV collagen and fibronectin may act on cell migration that

Thus, the structure and composition of the extracellular matrix play an important role during follicular development and post-spawning involution in teleost fish. The basement membrane integrity is important for follicular cell survival, and the loss of integrity contributes to increased follicular apoptosis. In addition, MMP-9 may be involved in the final oocyte maturation and regression of postovulatory follicles [134]. Therefore, it follows that different combinations and proportions in the assembly of extracellular matrix components, together with the presentation of a large variety of proteoglycans at various times during the development and maturation of the gonads, can orchestrate distinct gene expression programs and

Studies in gametogenesis help to understand the ecological, adaptive, and evolutionary relationships in the groups of species, especially when the oocyte structures are analyzed in an ultrastructural level. This is even more important when we

occurs during the late remodeling of postovulatory follicles [134].

culminate in more diverse tissue variations and adaptations [148].

The extracellular matrix-cell interactions influence gene regulation, cytoskeletal structure, differentiation, and many aspects of cell growth [155]. Changes in the expression of components that make up the extracellular matrix accompany follicular growth, ovulation, and involution of postovulatory follicles, which in its turn may influence follicular maturation, cell survival, and steroidogenesis [134, 156, 157]. Studies with mammals demonstrate that gonadal support cells synthesize a variety of components comprising the extracellular matrix and the basement membrane, such

[150, 151], embryogenesis [152, 153], or reproductive cycle [154].

**98**

**4. Conclusions**

I would like to thank all those who did or are part of the laboratories where I conducted my undergraduate and graduate studies and who culminated in the improvement of the knowledge reproduced here. I thank Dr. Adelina Ferreira, Dr. Mahmoud Mehanna, and Dr. Débora Fabiane Neves da Silva, advisors and colleagues at the Morphology and Morphometry Laboratory of the Federal University of Mato Grosso (UFMT), for the first steps taken in the area of Animal Reproduction. I would also like to thank Dr. Maria Inês Borella, Dr. Chayrra Chehade Gomes, Dr. Gisele Cristiane de Melo Dias, Dr. Lázaro Wender Oliveira de Jesus, MSc Giovana de Souza Branco, MSc Marília de Paiva Camargo, and laboratory technician Cruz Alberto Mendoza Rigonatti of the Laboratory of Fish Endocrinology, University of São Paulo, for the support during the doctorate and for all the knowledge obtained in that period. Lastly, I would like to thank the FAPEMAT, CAPES, and FAPESP funding agencies for the financial support provided during my academic trajectory, which culminated in the formulation of this chapter.

#### **Conflict of interest**

The author declares that there is no conflict of interest regarding the publication of this chapter.

*Reproductive Biology and Technology in Animals*

#### **Author details**

Mônica Cassel Mato Grosso Federal Institute of Education, Science and Technology—Campus Alta Floresta, Alta Floresta, MT, Brazil

\*Address all correspondence to: cassel.mcp@gmail.com

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

**101**

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key…*

[8] Cassel M, Chehade C, Branco GS, Canepelle D, Romagosa E, Borella MI.

Ovarian development and the reproductive profile of *Astyanax altiparanae* (Teleostei, Characidae) over one year: Applications in fish farming. Theriogenology. 2017;**98**:1-15. DOI: 10.1016/j. theriogenology.2017.04.044

[9] Carvalho PA, Paschoalini AL, Santos GB, Rizzo E, Bazzoli N. Reproductive biology of *Astyanax fasciatus* (Pisces: Characiformes) in a reservoir in southeastern Brazil. Journal of Applied Ichthyology. 2009;**25**(3):306-313. DOI: 10.1111/j.1439-0426.2009.01238.x

[10] Nuñez K, Duponchelle F. Towards a universal scale to assess sexual maturation and related life history traits in oviparous teleost fishes. Fish Physiology and Biochemistry. 2009;**35**(1):167-180. DOI: 10.1007/

[11] Casali RCV, Vono V, Godinho HP, Luz RK, Bazzoli N. Passage and reproductive activity of fishes in the Igarapava fish ladder, Grande River, southeastern Brazil. River Research and Applications. 2010;**26**(2):157-165. DOI:

[12] Chehade C, Cassel M, Borella MI. Induced reproduction in a migratory

drawdown. Neotropical Ichthyology.

teleost species by water level

[13] Normando FT, Arantes FP, Luz RK, Thomé RG, Rizzo E, Sato Y, et al. Reproduction and fecundity of tucunaré, *Cichla kelberi* (Perciformes:

Cichlidae), an exotic species in Três Marias reservoir, south eastern Brazil. Journal of Applied Ichthyology. 2009;**25**(3):299-305. DOI: 10.1111/j.1439-0426.2008.01174.x

2015;**13**(1):205-212. DOI: 10.1590/1982-0224-20140028

s10695-008-9241-2

10.1002/rra.1242

*DOI: http://dx.doi.org/10.5772/intechopen.87967*

Caramaschi EP. Reproductive biology of *Astyanax janeiroensis* (Osteichthyes, Characidae) from the Ubatiba River, Maricá, RJ, Brazil. Brazilian Journal of Biology. 2005;**65**(4):643-649. DOI: 10.1590/S1519-69842005000400012

França LR. An overview of functional and stereological evaluation of spermatogenesis and germ cell

transplantation in fish. Fish Physiology and Biochemistry. 2008;**35**(1):197-206. DOI: 10.1007/s10695-008-9252-z

[3] Schulz RW, França LR, Lareyre JJ,

Nóbrega RH, et al. Spermatogenesis in fish. General and Comparative Endocrinology. 2010;**165**(3):390-411. DOI: 10.1016/j.ygcen.2009.02.013

[4] Costa FG, Adolfi MC, Gomes CC, Jesus LWO, Batlouni SR, Borella MI. Tests of *Astyanax altiparanae*:The Sertoli cell functions in a semicystic

spermatogenesis. Micron. 2014;**61**:20-27. DOI: 10.1016/j.micron.2014.02.004

[5] Camargo MP, Cassel M, Jesus LWO, Nóbrega RH, Borella MI. Characterization of undifferentiated spermatogonia and the spermatogonial niche in the lambari fish *Astyanax altiparanae*. Theriogenology. 2017;**96**:97-102. DOI: 10.1016/j.theriogenology.2017.03.027

[6] Lubzens E, Young G, Bobe J,

Cerdà J. Oogenesis in teleosts: How fish eggs are formed. General and Comparative Endocrinology. 2010;**165**(3):367-389. DOI: 10.1016/j.ygcen.2009.05.022

[7] Martins YS, Moura DF, Santos GB, Rizzo E, Bazzoli N. Comparative folliculogenesis and spermatogenesis of four teleost fish from a reservoir in south-eastern Brazil. Acta

Zoologica. 2010;**91**(4):466-473. DOI: 10.1111/j.1463-6395.2009.00437.x

Legac F, Chiarini-Garcia H,

[1] Mazzoni R, Mendonça RS,

[2] Nóbrega RH, Batlouni SR,

**References**

*A Review of the Macroscopic, Microscopic, and Ultramicroscopic Characteristics of Some Key… DOI: http://dx.doi.org/10.5772/intechopen.87967*

#### **References**

*Reproductive Biology and Technology in Animals*

**100**

**Author details**

Floresta, Alta Floresta, MT, Brazil

provided the original work is properly cited.

\*Address all correspondence to: cassel.mcp@gmail.com

Mato Grosso Federal Institute of Education, Science and Technology—Campus Alta

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

Mônica Cassel

[1] Mazzoni R, Mendonça RS, Caramaschi EP. Reproductive biology of *Astyanax janeiroensis* (Osteichthyes, Characidae) from the Ubatiba River, Maricá, RJ, Brazil. Brazilian Journal of Biology. 2005;**65**(4):643-649. DOI: 10.1590/S1519-69842005000400012

[2] Nóbrega RH, Batlouni SR, França LR. An overview of functional and stereological evaluation of spermatogenesis and germ cell transplantation in fish. Fish Physiology and Biochemistry. 2008;**35**(1):197-206. DOI: 10.1007/s10695-008-9252-z

[3] Schulz RW, França LR, Lareyre JJ, Legac F, Chiarini-Garcia H, Nóbrega RH, et al. Spermatogenesis in fish. General and Comparative Endocrinology. 2010;**165**(3):390-411. DOI: 10.1016/j.ygcen.2009.02.013

[4] Costa FG, Adolfi MC, Gomes CC, Jesus LWO, Batlouni SR, Borella MI. Tests of *Astyanax altiparanae*:The Sertoli cell functions in a semicystic spermatogenesis. Micron. 2014;**61**:20-27. DOI: 10.1016/j.micron.2014.02.004

[5] Camargo MP, Cassel M, Jesus LWO, Nóbrega RH, Borella MI. Characterization of undifferentiated spermatogonia and the spermatogonial niche in the lambari fish *Astyanax altiparanae*. Theriogenology. 2017;**96**:97-102. DOI: 10.1016/j.theriogenology.2017.03.027

[6] Lubzens E, Young G, Bobe J, Cerdà J. Oogenesis in teleosts: How fish eggs are formed. General and Comparative Endocrinology. 2010;**165**(3):367-389. DOI: 10.1016/j.ygcen.2009.05.022

[7] Martins YS, Moura DF, Santos GB, Rizzo E, Bazzoli N. Comparative folliculogenesis and spermatogenesis of four teleost fish from a reservoir in south-eastern Brazil. Acta Zoologica. 2010;**91**(4):466-473. DOI: 10.1111/j.1463-6395.2009.00437.x

[8] Cassel M, Chehade C, Branco GS, Canepelle D, Romagosa E, Borella MI. Ovarian development and the reproductive profile of *Astyanax altiparanae* (Teleostei, Characidae) over one year: Applications in fish farming. Theriogenology. 2017;**98**:1-15. DOI: 10.1016/j. theriogenology.2017.04.044

[9] Carvalho PA, Paschoalini AL, Santos GB, Rizzo E, Bazzoli N. Reproductive biology of *Astyanax fasciatus* (Pisces: Characiformes) in a reservoir in southeastern Brazil. Journal of Applied Ichthyology. 2009;**25**(3):306-313. DOI: 10.1111/j.1439-0426.2009.01238.x

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Press; 1990. pp. 1-11

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29243%3A3<265%3A%3AAID-

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1894;**33**(144):74-83

BF00220430

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10.1023/A:1022613404123

*undecimalis* (Teleostei: Centropomidae). Journal of Morphology. 2000;**243**(3):265-281. DOI: 10.1002/%28SICI%291097-

4687%28200003%

JMOR4>3.0.CO%3B2-I

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fertilization and egg activation in teleost fish. Reviews in Fish Biology and Fisheries. 2002;**12**(1):33-58. DOI: 10.1023/A:1022613404123

*Reproductive Biology and Technology in Animals*

reprodução de peixes teleósteos: teoria e prática. Maringá: Eduem; 1996. 169 p

ultrastructural study. Tissue and Cell. 1999;**31**(5):480-488. DOI: 10.1054/

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[22] Dias JF, Peres-Rios E, Chaves PTC, Rossi-Wongtschowski CLD. Análise macroscópica dos ovários de

teleósteos: problemas de classificação e recomendações de procedimentos. Revista Brasileira de Biologia.

[23] Honji RM, Vaz-dos-Santos AM,

CLDB. Identification of the stages of ovarian maturation of the argentine hake Merluccius hubbsi Marini, 1933 (Teleostei: Merlucciidae): Advantages and disadvantages of the use of the macroscopic and microscopic scales. Neotropical Ichthyology. 2006;**4**(3):329-337. DOI: 10.1590/ S1679-62252006000300004

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[25] Coward K, Bromage NR. Reproductive physiology of female tilapia broodstock. Reviews in Fish Biology and Fisheries. 2000;**10**(1):1-25.

DOI: 10.1023/A:1008942318272

BF00122584

[26] Tyler CR, Sumpter JP. Oocyte growth and development in teleosts. Reviews in Fish Biology and Fisheries. 1996;**6**(3):287-318. DOI: 10.1007/

[27] Coward K, Bromage NR, Hibbitt O, Parrington J. Gamete physiology,

tice.1999.0045

10.1093/icb/21.2.325

1998;**58**(1):55-69

Rossi-Wongtschowski

[16] Grier HJ. The germinal epithelium: Its dual role in establishing male

reproductive classes and understanding

the basis for indeterminate egg production in female fishes. In: Creswell RL, editor. Proceedings of the Fifty-Third Annual Gulf and Caribbean Fisheries Institute. Fort Pierce: Mississippi/Alabama Sea Grant

Consortium; 2002. pp. 537-552

10.1002/jmor.10105

[17] Lo Nostro F, Grier H, Andreone L, Guerrero GA. Involvement of the gonadal germinal epithelium during sex reversal and seasonal testicular cycling in the protogynous swamp eel, *Synbranchus marmoratus* Bloch 1795 (Teleostei, Synbranchidae). Journal of Morphology. 2003;**257**(1):107-126. DOI:

[18] Brown-Peterson NJ, Wyanski DM,

Saborido-Rey F, Macewicz BJ, Lowerre-Barbieri SK. A standardized terminology for describing reproductive development in fishes. Marine and Coastal Fisheries. 2011;**3**(1):52-70. DOI:

10.1080/19425120.2011.555724

[19] Quagio-Grassiotto I, Grier H, Mazzoni TS, Nóbrega RH, Amorim JPA.

Activity of the ovarian germinal epithelium in the freshwater catfish, *Pimelodus maculatus* (Teleostei: Ostariophysi: Siluriformes): Germline cysts, follicle formation and oocyte development. Journal of Morphology. 2011;**272**(11):1290-1306. DOI: 10.1002/

[20] Miranda ACL, Bazzoli N, Rizzo E, Sato Y. Ovarian follicular atresia in two teleost species: A histological and

[14] Vazzoler AEAM. Biologia da

[15] Grier HJ, Taylor RG. Testicular maturation and regression in the common snook. Journal of Fish Biology. 1998;**53**(3):521-542. DOI: 10.1111/j.

1095-8649.1998.tb00999.x

**102**

jmor.10981

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[29] Grier JH, Uribe-Aranzábal MC, Patiño R. The ovary, folliculogenesis and oogenesis in teleosts. In: Jamieson BJM, editor. Reproductive Biology and Phylogeny of Fishes (Agnathans and Bony Fishes) Phylogeny Reproductive System Viviparity Spermatozoa. Enfield: Science Publishers; 2009. pp. 25-84

[30] Stehr CM, Hawkes JW. The comparative ultrastructure of the egg membrane and associated pore structures in the starry flounder, *Platichthys stellatus* (Pallas), and pink salmon, *Oncorhynchus gorbuscha* (Walbaum). Cell and Tissue Research. 1979;**202**(3):347-356. DOI: 10.1007/ BF00220430

[31] Francolini M, Lora Lamia C, Bonsignorio C, Cotelli F. Oocyte development and egg envelope formation in *Oreochromis niloticus*, a mouth-brooding cichlid fish. Journal of Submicroscopic Cytology and Pathology. 2003;**35**(1):49-60

[32] Hubbard JW. The yolk nucleus in *Cynematogaster aggregatus* Gibbons. Proceedings of the American Philosophical Society. 1894;**33**(144):74-83

[33] Yamamoto K. Studies on the formation of fish eggs. Annual cycle in the development of the ovarian eggs in the flounder, *Lipsetta obscura*. Journal

of the Faculty of Science, Hokkaido University, Series IV—Zoology. 1956;**12**(3):362-373

[34] Agostinho AA, Suzuki HI, Sampaio AA, Borges JD. Índice de atividade reprodutiva: uma proposta para avaliação da atividade reprodutiva em peixes. In: Encontro Brasileiro De Ictiologia, Abstract. Maringá – PR; 1991. p. 53

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[36] Munro AD. General introduction. In: Munro AD, Scott AP, Lam TJ, editors. Reproductive Seasonality in Teleosts: Environmental Influences. Florida: CRC Press; 1990. pp. 1-11

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[38] Suzuki HI, Vazzoler AEAM, Marques EE, Lizama MAP, Inada P. Reproductive ecology of the fish assemblages. In: Thomaz SM, Agostinho AA, Hahn NS, editors. The Upper Paraná River and Its Floodplain: Physical Aspects, Ecology and Conservation. Leiden: Backhuys Publishers; 2004. pp. 271-291

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[52] Friend DS, Gilula NB. Variations in tight and GAP junctions in mammalian tissues. The Journal of Cell Biology. 1972;**53**(3):758-776. DOI: 10.1083/

[53] Staehelin LA. Structure and function of intercellular junctions. International Review of Cytology. 1974;**39**:191-283. DOI: 10.1016/ S0074-7696(08)60940-7

[54] Rodewald M, Herr D, Fraser HM, Hack G, Kreienberg R, Wulff C. Regulation of tight junction proteins occludin and claudin-5 in the primate ovary during the ovulatory cycle and after inhibition of vascular endothelial growth factor. Molecular Human Reproduction. 2007;**13**(11):781-789. DOI: 10.1093/molehr/gam066

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0000030565.74702.0a

jcb.53.3.758

dependent meiotic resumption in ovarian follicles of Atlantic croaker. General and Comparative Endocrinology. 2003;**131**(3):291-295. DOI: 10.1016/S0016-6480(03)00015-7

oogenesis and atresia in an insect. Journal of Ultrastructure Research. 1981;**74**(1):95-104. DOI: 10.1016/

[42] Biliński S, Klag J. GAP junctions between oocyte and follicle cells in Acerentomon sp. (Insecta, Protura). International Journal of Invertebrate Reproduction and

Development. 1982;**5**(6):331-335. DOI: 10.1080/01651269.1982.10553486

[43] Khan HR, Saleuddin ASM. Cell contacts between follicle cells and oocyte of Helisoma (Mollusca, Pulmonata). Journal of Morphology. 1983;**177**(3):319-328. DOI: 10.1002/

[44] Anderson E, Albertini DF. GAP junctions between the oocyte and companion follicle cells in the

mammalian ovary. The Journal of Cell Biology. 1976;**71**(2):680-686. DOI:

[45] Toshimori K, Yasuzumi F. Tight junctions between ovarian follicle cells in the teleost (*Plecoglossus altivelis*). Journal of Ultrastructure Research. 1979;**67**(1):73-78. DOI: 10.1016/

[46] Kobayashi W. Communications of oocyte-granulosa cells in the Chum salmon ovary detected by transmission electron microscopy. Development, Growth and

Differentiation. 1985;**27**(5):553-561. DOI: 10.1111/j.1440-169X.1985.00553.x

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10.1083/jcb.71.2.680

S0022-5320(79)80019-2

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The biochemistry of cancer dissemination. Critical Reviews in Biochemistry and Molecular Biology. 1997;**32**(3):175-253. DOI: 10.3109/10409239709082573

S0955-0674(96)80116-5

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S0952-7915(00)00217-X

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[149] Dustin ML, De Fougerolles AR. Reprogramming T cells: The role of extracellular matrix in coordination of T cell activation and migration. Current Opinion in Immunology. 2001;**13**(3):286-290. DOI: 10.1016/

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10.1002/jmor.1052180209

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*Reproductive Biology and Technology in Animals*

[136] Saito D, Morinaga C, Aoki Y, Nakamura S, Mitani H, Furutani-Seiki M, et al. Proliferation of germ cells during gonadal sex differentiation in medaka: Insights from germ cell-depleted mutant zenzai. Developmental Biology. 2007;**310**(2):280-290. DOI: 10.1016/j.

[137] Krysko DV, Diez-Fraile A, Criel G,

2008;**13**(9):1065-1087. DOI: 10.1007/

[138] Tokarz RR. Oogonial proliferation, oogenesis and folliculogenesis in

nonmammalian vertebrates. In: Jones RE, editor. The Vertebrate Ovary. Comparative Biology and Evolution. New York: Plenum

reproduction and oogenesis in teleost fish compared to mammals. Reproduction Nutrition Development. 2005;**45**(3): 261-279. DOI: 10.1051/rnd:2005019

Nishimura T, Tanaka M. Ovarian germline stem cells in the teleost fish, medaka (*Oryzias latipes*). International Journal of Biological Sciences. 2011;**7**(4): 403-409. DOI: 10.7150/ijbs.7.403

[141] Sriramulu V, Rajalakshmi M. Origin of a new crop of oocytes in *Gobius giuris* (Hamilton-Buchanan). Zeitschrift für Mikroskopisch-Anatomische Forschung.

[142] Billard R. The reproductive cycle of male and female brown trout (Salmo trutta fario): A quantitative study. Reproduction Nutrition Development. 1987;**27**(1A):29-44. DOI: 10.1051/

[143] Selman K, Wallace RA, Sarka QIX. Stages of oocyte development in the zebrafish, *Brachydanio rerio*. Journal of

Svistunov AA, Vandenabeele P, D'Herde K. Life and death of female gametes during oogenesis and folliculogenesis. Apoptosis.

ydbio.2007.07.039

s10495-008-0238-1

Press; 1978. pp. 145-179

1966;**75**(1):64-73

rnd:19870104

[139] Jalabert B. Particularities of

[140] Nakamura S, Kobayashi K,

[129] Boone DL, Tsang BK. Caspase-3 in the rat ovary: Localization and possible role in follicular atresia and luteal regression. Biology of Reproduction. 1998;**58**:1533-1539. DOI: 10.1095/

[130] Thomé RG, Santos HB, Arantes FP, Domingos FFT, Bazzoli N, Rizzo E. Dual roles for autophagy during follicular atresia in fish ovary. Autophagy. 2009;**5**(1):117-119. DOI: 10.4161/

[131] Maiuri MC, Zalckvar E, Kimchi A, Kroemer G. Self-eating and self-killing: Crosstalk between autophagy and apoptosis. Nature Reviews. Molecular Cell Biology. 2007;**8**(9):741-752. DOI:

[132] Mizushima N, Levine B, Cuervo AM, Klionsky DJ. Autophagy fights disease through cellular self-digestion. Nature. 2008;**451**(7182):1069-1075. DOI:

[133] Maiuri MC, Criollo A, Kroemer G. Crosstalk between apoptosis and autophagy within the Beclin 1 interactome. The EMBO Journal. 2010;**29**(3):515-516. DOI: 10.1038/

[134] Thomé R, Santos HB, Sato Y, Rizzo E, Bazzoli N. Distribution of laminin beta2, collagen type IV, fibronectin and MMP-9 in ovaries of the teleost fish. Journal of Molecular Histology. 2010;**41**(4-5):215-224. DOI:

[135] Drevinick PE, Sandheinrich MB, Oris JT. Increased ovarian follicular apoptosis in fathead minnows (*Pimephales promelas*) exposed to dietary methylmercury. Aquatic Toxicology. 2006;**79**(1):49-54. DOI: 10.1016/j.aquatox.2006.05.007

10.1007/s10735-010-9281-7

of vertebrate apoptosis. Apoptosis.

2009;**14**:1-21. DOI: 10.1007/

s10495-008-0281-y

biolreprod58.6.1533

auto.5.1.7302

10.1038/nrm2239

10.1038/nature06639

emboj.2009.377

**110**

[144] Curry TE, Osteen KG. The matrix metalloproteinase system: Changes, regulation, and impact throughout the ovarian and uterine reproductive cycle. Endocrine Reviews. 2003;**24**(4): 428-465. DOI: 10.1210/er.2002-0005

[145] Larreta-Garde V, Berry H. Modeling extracellular matrix degradation balance with proteinase/ transglutaminase cycle. Journal of Theoretical Biology. 2002;**217**(1): 105-124. DOI: 10.1006/jtbi.2002.3010

[146] Price JT, Bonovich MT, Kohn EC. The biochemistry of cancer dissemination. Critical Reviews in Biochemistry and Molecular Biology. 1997;**32**(3):175-253. DOI: 10.3109/10409239709082573

[147] Basbaum CB, Werb Z. Focalized proteolysis: Spatial and temporal regulation of extracellular matrix degradation at the cell surface. Current Opinion in Cell Biology. 1996;**8**(5):731-738. DOI: 10.1016/ S0955-0674(96)80116-5

[148] Schalburg KR, Cooper GA, Yazawa R, Davidson WS, Koop BF. Microarray analysis reveals differences in expression of cell surface and extracellular matrix components during development of the trout ovary and testis. Comparative Biochemistry and Physiology. 2008;**3**(1):78-90. DOI: 10.1016/j.cbd.2007.10.001

[149] Dustin ML, De Fougerolles AR. Reprogramming T cells: The role of extracellular matrix in coordination of T cell activation and migration. Current Opinion in Immunology. 2001;**13**(3):286-290. DOI: 10.1016/ S0952-7915(00)00217-X

[150] Witte MB, Barbul A. General principles of wound healing. The Surgical Clinics of North America. 1997;**77**(3):509-528. DOI: 10.1016/ S0039-6109(05)70566-1

[151] Davis GE, Bayless KJ, Davis MJ, Meininger GA. Regulation of tissue injury responses by the exposure of matricryptic sites within extracellular matrix molecules. The American Journal of Pathology. 2000;**156**(5):1489-1498. DOI: 10.1016/S0002-9440(10)65020-1

[152] Hay ED. Collagen and embryonic development. In: Hay ED, editor. Cell Biology of Extracellular Matrix. New York: PlenumPress; 1981. pp. 379-409

[153] Perris R, Perissinotto D. Role of the extracellular matrix during neural crest cell migration. Mechanisms of Development. 2000;**95**(1-2):3-21. DOI: 10.1016/S0925-4773(00)00365-8

[154] Hulboy DL, Rudolph LA, Matrisian LM. Matrix metalloproteinases as mediators of reproductive function. Molecular Human Reproduction. 1997;**3**(1):27-45

[155] Irving-Rodgers HF, Rodgers RJ. Extracellular matrix in ovarian follicular development and disease. Cell and Tissue Research. 2005;**322**(1):89-98. DOI: 10.1007/s00441-005-0042-y

[156] Oksjoki S, Sallinen S, Vuorio E, Anttila L. Cyclic expression of mRNA transcripts for connective tissue components in the mouse ovary. Molecular Human Reproduction. 1999;**5**(9):803-808. DOI: 10.1093/ molehr/5.9.803

[157] Rodgers RJ, Lavranos TC, Van Wezel IL, Irving-Rodgers HF. Development of the ovarian follicular epithelium. Molecular and Cellular Endocrinology. 1999;**151**(1-2):171-179. DOI: 10.1016/S0303-7207(99)00087-8

[158] Skinner MK, Tung PS, Fritz IB. Cooperativity between Sertoli cells and testicular peritubular cells in

the production and deposition of extracellular matrix components. The Journal of Cell Biology. 1985;**100**(6):1941-1947. DOI: 10.1083/ jcb.100.6.1941

[159] Rodgers RJ, Irving-Rodgers HF, Russell DL. Extracellular matrix of the developing ovarian follicle. Reproduction. 2003;**126**(4):415-424

[160] Sternilicht MD, Werb Z. How matrix metalloproteinase regulate cell behavior. Annual Review of Cell and Developmental Biology. 2001;**17**(1):463-516. DOI: 10.1146/ annurev.cellbio.17.1.463

[161] Smith MF, Ricke WA, Bakke LJ, Dow MPD, Smith GW. Ovarian tissue remodeling: Role of matrix metalloproteinases and their inhibitors. Molecular and Cellular Endocrinology. 2002;**191**(1):45-56. DOI: 10.1016/ S0303-7207(02)00054-0

[162] Ogiwara K, Takano N, Shinohara M, Murakami M, Takahashi T. Gelatinase A and membrane-type matrix metalloproteinases 1 and 2 are responsible for follicle rupture during ovulation in the medaka. Proceedings of the National Academy of Sciences of the United States of America. 2005;**102**(24):8442-8447. DOI: 10.1073/pnas.0502423102

[163] Berkholtz CB, Lai BE, Woodruff TK, Shea LD. Distribution of extracellular matrix proteins type I collagen, type IV collagen, fibronectin, and laminin in mouse folliculogenesis. Histochemistry and Cell Biology. 2006;**126**(5):583-592. DOI: 10.1007/s00418-006-0194-1

[164] Carnegie JA. Secretion of fibronectin by rat granulosa cells occurs primarily during early follicular development. Journal of Reproduction and Fertility. 1990;**89**(2):579-589. DOI: 10.1530/jrf.0.0890579

[165] Zhao Y, Luck MR. Gene expression and protein distribution of collagen,

fibronectin and laminin in bovine follicles and corpora lutea. Journal of Reproduction and Fertility. 1995;**104**(1):115-123. DOI: 10.1530/ jrf.0.1040115

[166] Iwahashi M, Muragaki Y, Ooshima A, Nakano R. Type VI collagen expression during growth of human ovarian follicles. Fertility and Sterility. 2000;**74**(2):343-347. DOI: 10.1016/ S0015-0282(00)00618-X

**113**

**Chapter 7**

**Abstract**

Reproductive Cycle of *Hexaplex* 

*José Luis Gómez-Márquez, Ana Bertha Villaseñor-Martínez,* 

During two annual periods, the reproductive cycle of the gastropod *Hexaplex princeps* from Puerto Ángel, Oaxaca, Mexico was studied through gonadal histology. Sex proportion for the collected individuals was not statistically different from parity although most of the time, the number of males was slightly larger than that of females, which only outnumbered males during the spawning season. The maturity stages established for females were: (1) initial oogenesis, (2) previtellogenic maturity, (3) vitellogenic maturity, (4) maturity, (5) spawning, and (6) resting; and for males: (1) initial spermatogenesis, (2) maturity, (3) spawning, (4) onset of the rest, and (5) resting. Monthly variations of maturation stages showed that H. princeps has an annual reproductive cycle with a long period of gonadal activity. The spawning season comprised from November (females) and December (males) to March, with activity peaks in January. From March to October (females) and from May to June (males), reproduction resting occurred. Spawning was related to high chlorophyll concentrations due to the upwelling processes resulting from the winds and to the cooler sea surface temperatures occurring from November to March. This study provides baseline information that may serve to establish measures for sustainable exploitation strategies and for future aquaculture implementation of this species.

*princeps* (Broderip, 1833)

*Ma. De Lourdes Jiménez-Badillo* 

*and Isaías Hazarmabeth Salgado-Ugarte*

*Verónica Mitsui Saito-Quezada, Esther Uría-Galicia,* 

**Keywords:** *Hexaplex princeps*, reproductive cycle, sexual proportion,

Marine mollusks constitute one of the more important world fisheries representing around 10% of the total value and quantity [1]. These invertebrates have been exploited since ancient times. Recently, it has been reported that omega fatty acids, including docosahexaenoic acid (DHA) are key to brain health and most likely helped to drive the evolution of the modern human brain, when hominin ancestors consumed rich DHA marine shellfish [2]. In the world, approximately 720 gastropod species are exploited [3, 4]. In Mexico, the gastropod catch in 2013 has the 19th

The gastropod *Hexaplex princeps* has a spiny shell, height from 7.6 to 15.2 cm and whirl count 6 or 9; edge of lip armed with long, hollow, and frond-like spines;

histological analysis, maturity scales

**1. Introduction**

place with 6011 ton [5].

#### **Chapter 7**

*Reproductive Biology and Technology in Animals*

fibronectin and laminin in bovine follicles and corpora lutea. Journal of Reproduction and Fertility. 1995;**104**(1):115-123. DOI: 10.1530/

[166] Iwahashi M, Muragaki Y,

S0015-0282(00)00618-X

Ooshima A, Nakano R. Type VI collagen expression during growth of human ovarian follicles. Fertility and Sterility. 2000;**74**(2):343-347. DOI: 10.1016/

jrf.0.1040115

the production and deposition of extracellular matrix components. The Journal of Cell Biology.

jcb.100.6.1941

1985;**100**(6):1941-1947. DOI: 10.1083/

[159] Rodgers RJ, Irving-Rodgers HF, Russell DL. Extracellular matrix of the developing ovarian follicle. Reproduction. 2003;**126**(4):415-424

[160] Sternilicht MD, Werb Z. How matrix metalloproteinase regulate cell behavior. Annual Review of Cell and Developmental Biology. 2001;**17**(1):463-516. DOI: 10.1146/

[161] Smith MF, Ricke WA, Bakke LJ, Dow MPD, Smith GW. Ovarian tissue

metalloproteinases and their inhibitors. Molecular and Cellular Endocrinology. 2002;**191**(1):45-56. DOI: 10.1016/

[162] Ogiwara K, Takano N, Shinohara M, Murakami M, Takahashi T. Gelatinase

metalloproteinases 1 and 2 are responsible for follicle rupture during ovulation in the medaka. Proceedings of the National Academy of Sciences of the United States of America. 2005;**102**(24):8442-8447. DOI: 10.1073/pnas.0502423102

[163] Berkholtz CB, Lai BE, Woodruff TK, Shea LD. Distribution of extracellular matrix proteins type I collagen, type IV collagen, fibronectin, and laminin in mouse folliculogenesis. Histochemistry and Cell Biology. 2006;**126**(5):583-592. DOI: 10.1007/s00418-006-0194-1

annurev.cellbio.17.1.463

remodeling: Role of matrix

S0303-7207(02)00054-0

A and membrane-type matrix

[164] Carnegie JA. Secretion of fibronectin by rat granulosa cells occurs primarily during early follicular development. Journal of Reproduction and Fertility. 1990;**89**(2):579-589. DOI:

[165] Zhao Y, Luck MR. Gene expression and protein distribution of collagen,

10.1530/jrf.0.0890579

**112**

## Reproductive Cycle of *Hexaplex princeps* (Broderip, 1833)

*Verónica Mitsui Saito-Quezada, Esther Uría-Galicia, José Luis Gómez-Márquez, Ana Bertha Villaseñor-Martínez, Ma. De Lourdes Jiménez-Badillo and Isaías Hazarmabeth Salgado-Ugarte*

#### **Abstract**

During two annual periods, the reproductive cycle of the gastropod *Hexaplex princeps* from Puerto Ángel, Oaxaca, Mexico was studied through gonadal histology. Sex proportion for the collected individuals was not statistically different from parity although most of the time, the number of males was slightly larger than that of females, which only outnumbered males during the spawning season. The maturity stages established for females were: (1) initial oogenesis, (2) previtellogenic maturity, (3) vitellogenic maturity, (4) maturity, (5) spawning, and (6) resting; and for males: (1) initial spermatogenesis, (2) maturity, (3) spawning, (4) onset of the rest, and (5) resting. Monthly variations of maturation stages showed that H. princeps has an annual reproductive cycle with a long period of gonadal activity. The spawning season comprised from November (females) and December (males) to March, with activity peaks in January. From March to October (females) and from May to June (males), reproduction resting occurred. Spawning was related to high chlorophyll concentrations due to the upwelling processes resulting from the winds and to the cooler sea surface temperatures occurring from November to March. This study provides baseline information that may serve to establish measures for sustainable exploitation strategies and for future aquaculture implementation of this species.

**Keywords:** *Hexaplex princeps*, reproductive cycle, sexual proportion, histological analysis, maturity scales

#### **1. Introduction**

Marine mollusks constitute one of the more important world fisheries representing around 10% of the total value and quantity [1]. These invertebrates have been exploited since ancient times. Recently, it has been reported that omega fatty acids, including docosahexaenoic acid (DHA) are key to brain health and most likely helped to drive the evolution of the modern human brain, when hominin ancestors consumed rich DHA marine shellfish [2]. In the world, approximately 720 gastropod species are exploited [3, 4]. In Mexico, the gastropod catch in 2013 has the 19th place with 6011 ton [5].

The gastropod *Hexaplex princeps* has a spiny shell, height from 7.6 to 15.2 cm and whirl count 6 or 9; edge of lip armed with long, hollow, and frond-like spines; is gonochoristic and the gonad with the digestive gland occupies the visceral coils [6]; distributes from the Gulf of California to Peru inhabiting moderately shallow waters [7, 8]. The presence of this species has been reported in Panama [9]. *Hexaplex princeps* (Broderip, 1833) is captured by several artisanal fisheries of Mexico: because of its size and taste, *H. princeps* is observed with less frequency in the subtidal zone at the Tenacatita (Jalisco) coral reef in search of other prey gastropods, although can be found in rocky substrates with heavy wave action [10]. At Acapulco (Guerrero), *H. princeps* is the second major exploited species after the oyster *Striostrea prismatica* [11]. Our observations along the study period suggest that at Puerto Ángel, Oaxaca, this species is an important fishery resource as it supports around 80% of the gastropod catch for local population and tourism consumption.

In the world, artisanal fisheries are in continuous expansion due to the growing demand and increasing value of appreciated species, and thus, the fisheries effort is augmenting [12]. Under this scenario, it is advisable (if not indispensable) to gather baseline biological information that may be used to propose management measures promoting long-term sustainable resource exploitation [4].

The study of the reproductive processes in marine organisms is a fundamental biological aspect, which permits to understand their population dynamics [13]. The reproductive season is a crucial life history trait and the proper timing of breeding may be important to provide the offspring with favorable environmental conditions and to influence parental fitness [14, 15]. The analysis of the reproductive cycle of organisms permits to know the adequate moment and intensity of the capture to avoid the population depletion.

The reproduction of Muricidae members (at which *H. princeps* belongs) has been studied in several instances: the gonad cycle of *Bolinus brandaris* at the South of Portugal [16] and at the South of Tunisia [4]; in the Gulf of California, Mexico, observations on the "Black Murex" *Hexaplex nigritus* [17] and *Hexaplex erythrostomus* [18] similar oviposition. In spite of the importance of *H. princeps* as a fishery resource, there is no information on its reproductive cycle. This knowledge gap makes necessary to carry out studies on the biological cycle of this species in order to take adequate management decisions leading toward its sustainable exploitation and even its aquaculture in the future.

Thus, this study is aimed to investigate the reproductive cycle of *Hexaplex princeps*, considering sexual proportion, gonad maturation, spawning periods, and maturity stages variation in relation with the surface temperature and chlorophyll concentration along two annual periods at Puerto Angel, Oaxaca, Mexico.

#### **2. Materials and methods**

The organisms were obtained from the artisanal fishery with (as possible) monthly periodicity during two annual periods from January 2014 to November 2015. The organisms were caught with the help of two local free divers and the captain of an 8 m length vessel with a 40 HP outboard motor at depths from 5 to 15 m in rocky coast localities at the vicinity of Puerto Ángel, Oaxaca, Mexico, between 9:00 and 12:00 h local time (**Table 1**, **Figure 1**). The collecting sites were determined each date according to the atmospheric and sea conditions as well as the diver's knowledge on the species availability in the zone. Our aim was to have a representative number of specimens from the region to gather the histological information from reproductive organs and tissues.

From the caught organisms, 10–15 individuals in the interval from 8 to 12 cm in length (interval that contained more than 90% of the lengths, we collected since

**115**

**Figure 1.**

**Table 1.**

*geopositions.*

2012) were separated and their shell broken to extract the soft parts, which were fixed in formalin 10% prepared with seawater [19]. Once fixed, the specimens were transported to the Biometry and Fisheries Biology Laboratory of the Facultad de Estudios Superiores Zaragoza, UNAM, where after 48 h were washed with tap water and preserved in 70% ethylic alcohol. As there are no external characters to

*de Aceite, LB: La Boquilla, Sr: Secretario, Tj: Tijera, Dm: Dominguillo, PT: Playita Tembo.*

*Geographical location of the study area, Puerto Ángel, Oaxaca, Mexico. The collection sites are indicated: PC: Punta Cometa, SA: San Agustinillo, Ar: Aragón, PP: Playa Panteón, ES: Estacahuite. LM: La Mina, BA: Bajos* 

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

**Site Latitude (N) Longitude (W)** Punta Cometa (PC) 15° 39′ 35.4″ 96° 33′ 16.5″ San Agustinillo (SA) 15° 39′ 48.6″ 96° 33′ 01.0″ Playa Panteón (PP) 15° 39′ 56.1″ 96° 29′ 27.1″ Aragón (Ar) 15° 39′ 38.2″ 96° 31′ 46.8″ Estacahuite (Es) 15° 40′ 04.7″ 96° 28′ 54.5″ Bajos de Aceite (BA) 15° 40′ 10.6″ 96° 28′ 29.6″ La Mina (LM) 15° 40′ 26.7″ 96° 28′ 35.7″ La Boquilla (LB) 15° 40′ 48.3″ 96° 27′ 58.4″ Secretario (Sr) 15° 41′ 02.3″ 96° 27′ 00.5″ Tijera (Tj) 15° 41′ 20.2″ 96° 26′ 26.3″ Dominguillo (Dm) 15° 41′ 35.0″ 96° 26′ 02.2″ Playita, Tembo (PT) 15° 41′ 36.1″ 96° 25′ 54.3″ Temperature (SST) and chlorophyll (CL) 15° 38′ 44.9″ 96° 28′ 45.0″

*Specimen collection and environmental variables (surface sea temperature and chlorophyll) measure site* 

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833) DOI: http://dx.doi.org/10.5772/intechopen.88074*


#### **Table 1.**

*Reproductive Biology and Technology in Animals*

consumption.

avoid the population depletion.

and even its aquaculture in the future.

information from reproductive organs and tissues.

**2. Materials and methods**

is gonochoristic and the gonad with the digestive gland occupies the visceral coils [6]; distributes from the Gulf of California to Peru inhabiting moderately shallow waters [7, 8]. The presence of this species has been reported in Panama [9]. *Hexaplex princeps* (Broderip, 1833) is captured by several artisanal fisheries of Mexico: because of its size and taste, *H. princeps* is observed with less frequency in the subtidal zone at the Tenacatita (Jalisco) coral reef in search of other prey gastropods, although can be found in rocky substrates with heavy wave action [10]. At Acapulco (Guerrero), *H. princeps* is the second major exploited species after the oyster *Striostrea prismatica* [11]. Our observations along the study period suggest that at Puerto Ángel, Oaxaca, this species is an important fishery resource as it supports around 80% of the gastropod catch for local population and tourism

In the world, artisanal fisheries are in continuous expansion due to the growing demand and increasing value of appreciated species, and thus, the fisheries effort is augmenting [12]. Under this scenario, it is advisable (if not indispensable) to gather baseline biological information that may be used to propose management measures

The study of the reproductive processes in marine organisms is a fundamental biological aspect, which permits to understand their population dynamics [13]. The reproductive season is a crucial life history trait and the proper timing of breeding may be important to provide the offspring with favorable environmental conditions and to influence parental fitness [14, 15]. The analysis of the reproductive cycle of organisms permits to know the adequate moment and intensity of the capture to

The reproduction of Muricidae members (at which *H. princeps* belongs) has been studied in several instances: the gonad cycle of *Bolinus brandaris* at the South of Portugal [16] and at the South of Tunisia [4]; in the Gulf of California, Mexico, observations on the "Black Murex" *Hexaplex nigritus* [17] and *Hexaplex erythrostomus* [18] similar oviposition. In spite of the importance of *H. princeps* as a fishery resource, there is no information on its reproductive cycle. This knowledge gap makes necessary to carry out studies on the biological cycle of this species in order to take adequate management decisions leading toward its sustainable exploitation

Thus, this study is aimed to investigate the reproductive cycle of *Hexaplex princeps*, considering sexual proportion, gonad maturation, spawning periods, and maturity stages variation in relation with the surface temperature and chlorophyll

The organisms were obtained from the artisanal fishery with (as possible) monthly periodicity during two annual periods from January 2014 to November 2015. The organisms were caught with the help of two local free divers and the captain of an 8 m length vessel with a 40 HP outboard motor at depths from 5 to 15 m in rocky coast localities at the vicinity of Puerto Ángel, Oaxaca, Mexico, between 9:00 and 12:00 h local time (**Table 1**, **Figure 1**). The collecting sites were determined each date according to the atmospheric and sea conditions as well as the diver's knowledge on the species availability in the zone. Our aim was to have a representative number of specimens from the region to gather the histological

From the caught organisms, 10–15 individuals in the interval from 8 to 12 cm in length (interval that contained more than 90% of the lengths, we collected since

concentration along two annual periods at Puerto Angel, Oaxaca, Mexico.

promoting long-term sustainable resource exploitation [4].

**114**

*Specimen collection and environmental variables (surface sea temperature and chlorophyll) measure site geopositions.*

#### **Figure 1.**

*Geographical location of the study area, Puerto Ángel, Oaxaca, Mexico. The collection sites are indicated: PC: Punta Cometa, SA: San Agustinillo, Ar: Aragón, PP: Playa Panteón, ES: Estacahuite. LM: La Mina, BA: Bajos de Aceite, LB: La Boquilla, Sr: Secretario, Tj: Tijera, Dm: Dominguillo, PT: Playita Tembo.*

2012) were separated and their shell broken to extract the soft parts, which were fixed in formalin 10% prepared with seawater [19]. Once fixed, the specimens were transported to the Biometry and Fisheries Biology Laboratory of the Facultad de Estudios Superiores Zaragoza, UNAM, where after 48 h were washed with tap water and preserved in 70% ethylic alcohol. As there are no external characters to

distinguish sex, the specimens were dissected to examine and search for the presence or absence of penis.

The sexual proportion was analyzed by means of the chi-squared goodness of fit test following the corrected Yates expression [20, 21].

The histological sections were carried out at the Histology Laboratory from the Morphology Department at the Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional. The alcohol-preserved specimens were dehydrated following the usual alcohol series (70–100%) and cleared in xylol before being included in paraplax and paraffin. The embedded tissues were sliced into sections of 5 μm thickness using a microtome and mounted over glass slides. The preparations were stained with the Hematoxylin-Eosin method [22] to facilitate the determination of the gonad development stages. The sections were fixed with Entalan and covered with glass slips. Finally, the preparations were observed and photographed by means of an optical microscope with attached camera.

The sea surface temperature (SST) and chlorophyll *a* of Puerto Ángel data were consulted from the GES DISC-NASA database [23, 24]. The monthly values were taken from a site in the vicinities of the Puerto Angel Bay (**Table 1**). To assess the statistical significance of the relationships between the maturity stages and the mean values of temperature and chlorophyll, two procedures were employed. In the first place, in order to clarify the pattern showed by the gonad stage percentages, a nonlinear resistant smoothing procedure was applied. The preferred smoother was the 4253eh,twice, which combines the smooth result of even span running median smoothers (4,2), the resistance of odd running medians (5,3) with end point adjustment (e), the "Hanning" weighted mean smoother (h) and the "re-roughing" (twice) step [25, 26, 27, 28, 29–32, 33].

The comparisons of the resulting time series data were performed by means of the cross-correlation analysis [34, 35, 33] between the percentages of maturity stages against the temperature and chlorophyll values. Additional cross-correlation analyses among maturity stages were made.

#### **3. Results**

#### **3.1 Reproductive cycle**

In total, 232 males and 214 females were captured. The sexual proportions throughout the study are included in **Table 2**. From the 446 individuals, 250 were analyzed for recognition and characterization of the gonad stages considering the degree of development besides the occurrence and abundance of gametes. The different gonad development stages were classified as follows: for females, six stages were established (**Table 3**, **Figure 2**). Stage 1 (S1) initial oogenesis, stage 2 (S2) previtellogenic maturity, stage 3 (S3) vitellogenic maturity, stage 4 (S4) maturity, stage 5 (S5) spawning, and finally, stage 6 (S6) resting. For males, five stages were recognized (**Table 4**, **Figure 3**). Stage 1 (S1) initial spermatogenesis, stage 2 (S2) maturity, stage 3 (S3) expulsion, stage 4 (S4) onset of rest, and stage 5 (S5) resting.

Spawning (S5) females of *H. princeps* (**Figure 4**) presented large percentages during January (60%), March (67%), and November (67%) of 2014. On the other hand, it is noted that 100% of the resting stage (S6) was registered in July. The months with larger spawning (S5) percentages during 2015 were January (60%), March (80%), and November (75%) and the months with resting (S6) larger frequency values were April (75%) and July (75%).

**117**

**Table 3.**

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

**Total Sexo Male:female** 

**Males Females** 24/Jan/14 (Es) 93 40 53 1.00:1.33 1.55 0.21 21/Feb/14 (Dm) 02 02 0 —:— 0.50 0.48 21/Mar/14 (Ar) 26 16 10 1.60:1.00 0.96 0.33 25/Apr/14 (PP) 60 33 27 1.22:1.00 0.42 0.52 23/May/14 (ML) 53 27 26 1.04:1.00 0.00 1.00 15/Aug/14 (PC) 42 25 17 1.47:1.00 1.17 0.28 24/Oct/14 (PC) 13 04 09 1.00:2.25 1.23 0.27 28/Nov/14 (PT) 57 32 25 1.28:1.00 0.63 0.43 21/Dec/14 (Es) 12 07 05 1.40:1.00 0.08 0.77 30/Jan/15 (PT) 08 03 05 1.00:1.67 0.13 0.72 27/Mar/15 (PP) 07 02 05 1.00:2.50 0.57 0.45 30/Apr/15 (PP) 08 05 03 1.67:1.00 0.13 0.72 15/May/15 (LM) 08 04 04 1.00:1.00 0.13 0.72 12/Jun/15 (SA) 11 06 05 1.20:1.00 0.00 1.00 15/Jul/15 (BA) 09 06 03 2.00:1.00 0.44 0.50 21/Aug/15 (PP) 09 05 04 1.25:1.00 0.00 1.00 25/Sep/15 (PP) 10 05 05 1.00:1.00 0.10 0.75 16/Oct/15 (PP) 08 04 04 1.00:1.00 0.13 0.72 27/Nov/15 (PP) 10 06 04 1.50:1.00 0.10 0.75 Total 446 232 214 1.08:1.00 0.65 0.42

*Number of individuals by collecting date and site (main); sexual proportion and its statistical significance are* 

Stage 1 initial oogenesis (S1) Occurrence of developing ovogonia and oocytes, thick follicle

Stage 2 previtellogenic maturity (S2) Oocytes full of yolk granules; in some oocytes, the nucleus

Stage 4 maturity (S4) Follicles full of yolk granules and platelets; thin follicle walls Stage 5 spawning (S5) Light in the follicles is observed; follicles partially empty; follicle

Stage 6 resting (S6) Some resting follicles besides cells or phagocytes in thick follicle

Stage 3 vitellogenic maturity (S3) Follicles with thin walls and developing oocytes; yolk granules are

observed and yolk platelets appear

walls thin with some remnant oocytes

walls; conspicuous conjunctive tissue

and nucleolus are observed; follicles completely mature full of

walls

oocytes

*Characterization of gonad development stages for females of Hexaplex princeps.*

**proportion**

*χ 2* **(Yates)**

*P*

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

**Collecting data and site** 

*Site abbreviations according to* **Table 1***.*

 *with yates correction).*

**Table 2.**

*included (χ 2*

**(main)**


#### *Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833) DOI: http://dx.doi.org/10.5772/intechopen.88074*

#### **Table 2.**

*Reproductive Biology and Technology in Animals*

test following the corrected Yates expression [20, 21].

means of an optical microscope with attached camera.

"re-roughing" (twice) step [25, 26, 27, 28, 29–32, 33].

frequency values were April (75%) and July (75%).

analyses among maturity stages were made.

ence or absence of penis.

distinguish sex, the specimens were dissected to examine and search for the pres-

The sexual proportion was analyzed by means of the chi-squared goodness of fit

The histological sections were carried out at the Histology Laboratory from the Morphology Department at the Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional. The alcohol-preserved specimens were dehydrated following the usual alcohol series (70–100%) and cleared in xylol before being included in paraplax and paraffin. The embedded tissues were sliced into sections of 5 μm thickness using a microtome and mounted over glass slides. The preparations were stained with the Hematoxylin-Eosin method [22] to facilitate the determination of the gonad development stages. The sections were fixed with Entalan and covered with glass slips. Finally, the preparations were observed and photographed by

The sea surface temperature (SST) and chlorophyll *a* of Puerto Ángel data were consulted from the GES DISC-NASA database [23, 24]. The monthly values were taken from a site in the vicinities of the Puerto Angel Bay (**Table 1**). To assess the statistical significance of the relationships between the maturity stages and the mean values of temperature and chlorophyll, two procedures were employed. In the first place, in order to clarify the pattern showed by the gonad stage percentages, a nonlinear resistant smoothing procedure was applied. The preferred smoother was the 4253eh,twice, which combines the smooth result of even span running median smoothers (4,2), the resistance of odd running medians (5,3) with end point adjustment (e), the "Hanning" weighted mean smoother (h) and the

The comparisons of the resulting time series data were performed by means of the cross-correlation analysis [34, 35, 33] between the percentages of maturity stages against the temperature and chlorophyll values. Additional cross-correlation

In total, 232 males and 214 females were captured. The sexual proportions throughout the study are included in **Table 2**. From the 446 individuals, 250 were analyzed for recognition and characterization of the gonad stages considering the degree of development besides the occurrence and abundance of gametes. The different gonad development stages were classified as follows: for females, six stages were established (**Table 3**, **Figure 2**). Stage 1 (S1) initial oogenesis, stage 2 (S2) previtellogenic maturity, stage 3 (S3) vitellogenic maturity, stage 4 (S4) maturity, stage 5 (S5) spawning, and finally, stage 6 (S6) resting. For males, five stages were recognized (**Table 4**, **Figure 3**). Stage 1 (S1) initial spermatogenesis, stage 2 (S2) maturity, stage 3 (S3) expulsion, stage 4 (S4) onset of rest, and stage

Spawning (S5) females of *H. princeps* (**Figure 4**) presented large percentages during January (60%), March (67%), and November (67%) of 2014. On the other hand, it is noted that 100% of the resting stage (S6) was registered in July. The months with larger spawning (S5) percentages during 2015 were January (60%), March (80%), and November (75%) and the months with resting (S6) larger

**116**

**3. Results**

5 (S5) resting.

**3.1 Reproductive cycle**

*Number of individuals by collecting date and site (main); sexual proportion and its statistical significance are included (χ 2 with yates correction).*


#### **Table 3.**

*Characterization of gonad development stages for females of Hexaplex princeps.*

#### **Figure 2.**

*Histological sections of Hexaplex princeps females showing ovary stages. a: initial oogenesis (20X), b: previtellogenic mature (10X), c: vitellogenic mature (20X), d: mature (10X), e: spawning (10X), f: resting (10X). Oo: oogonia, Oc: oocytes, DG: digestive gland, CT: connective tissue, PV: vitelline platelets, PvO: previtellogenic oocytes, VO: vitellogenic oocytes, FW: follicular wall, RO: residual oocytes, P: phagocytes.*

#### Males of *H. princeps* (**Figure 5**) presented spawning (S3) stage in January (100%) and March (50%); the reproductive resting stage (S5) occurred in


**119**

**Figure 4.**

**Figure 3.**

*phagocytes.*

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

*Histological sections of Hexaplex princeps males showing testis stages. a: initial spermatogenesis (50 μm), b: mature (100 μm), c: spawning (10x), d: onset of rest (100 μm), e: resting (50 μm). Eg: spermatogonia, Ec: spermatocytes, Ez: spermatozoa, RS: residual spermatozoa, DG: digestive gland, CT: connective tissue, P:* 

*Gonad stage frequency 2014–2015 for H. princeps females by sampling date. Chlorophyll concentration* 

*(mg m–3) and Surface Water Temperature (°C) values are showed.*

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

#### **Table 4.**

*Characterization of gonad development stage for males of Hexaplex princeps.*

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833) DOI: http://dx.doi.org/10.5772/intechopen.88074*

#### **Figure 3.**

*Reproductive Biology and Technology in Animals*

Males of *H. princeps* (**Figure 5**) presented spawning (S3) stage in January (100%) and March (50%); the reproductive resting stage (S5) occurred in

Stage 2 maturity (S2) Follicles utterly full with a greater quantity of spermatozoids, spermatogonia,

Stage 5 resting (S5) Empty follicle lumen; resting follicles due to the expulsion of spermatozoids are observed; conspicuous conjunctive tissue

spermatocytes, and spermatids Stage 3 spawning (S3) Mature spermatozoids in expulsion, ciliated cylindric epithelium with

Stage 4 onset of rest (S4) Some follicles in expulsion; empty and resting follicles are observed

Follicles active, developed with immature cells; small separated follicles with numerous immature cells (spermatogonia and spermatocytes), thick follicle

*Histological sections of Hexaplex princeps females showing ovary stages. a: initial oogenesis (20X), b: previtellogenic mature (10X), c: vitellogenic mature (20X), d: mature (10X), e: spawning (10X), f: resting (10X). Oo: oogonia, Oc: oocytes, DG: digestive gland, CT: connective tissue, PV: vitelline platelets, PvO: previtellogenic oocytes, VO: vitellogenic oocytes, FW: follicular wall, RO: residual oocytes, P: phagocytes.*

walls

foldings

*Characterization of gonad development stage for males of Hexaplex princeps.*

**118**

**Table 4.**

**Figure 2.**

Stage 1 initial spermatogenesis (S1) *Histological sections of Hexaplex princeps males showing testis stages. a: initial spermatogenesis (50 μm), b: mature (100 μm), c: spawning (10x), d: onset of rest (100 μm), e: resting (50 μm). Eg: spermatogonia, Ec: spermatocytes, Ez: spermatozoa, RS: residual spermatozoa, DG: digestive gland, CT: connective tissue, P: phagocytes.*

#### **Figure 4.**

*Gonad stage frequency 2014–2015 for H. princeps females by sampling date. Chlorophyll concentration (mg m–3) and Surface Water Temperature (°C) values are showed.*

#### **Figure 5.**

*Gonad stage frequency 2014–2015 for H. princeps males by sampling date. Chlorophyll concentration (mg m–3) and temperature (°C) are included.*

#### **Figure 6.**

*The smoothed frequency of the female reproductive stages along the study period and cross-correlation correlograms for mature-spawning (S4-S5) and spawning-resting (S5-S6) reproductive stages comparison.*

**121**

**Figure 7.**

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

**3.2 Chlorophyll** *a* **concentration and gonad cycle**

), February (4.00 mg m<sup>−</sup><sup>3</sup>

), February (1.23 mg m<sup>−</sup><sup>3</sup>

May (60%), June (75%), and July (67%) 2014. The months of 2015 with larger spawning (S3) percentages were January (100%) and March (50%); finally, the months with the larger frequency of resting individuals (S5) were May (60%)

In 2014, the highest chlorophyll *a* concentrations were observed in January

spawning stages (S5 females, S3 males) (**Figures 4** and **5**) was observed. For the same year, low chlorophyll concentrations were recorded in June (0.15 mg m<sup>−</sup><sup>3</sup>

females and males in the resting stages (S6 and S5, respectively) (**Figures 4** and **5**). In 2015, the high chlorophyll concentrations were observed in January

*The smoothed frequency of the male reproductive stages along the study period and cross-correlation correlograms for mature-spawning (S2-S3) and spawning-resting (S3-S5) reproductive stages comparison.*

); in the same months, the larger frequency of individuals in the

), March (2.05 mg m<sup>−</sup><sup>3</sup>

) months, which correspond with the larger frequency of

), March (1.08 mg m<sup>−</sup><sup>3</sup>

); except for April, in all other months, the stage with

), and December

), and (exception-

)

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

and June (75%).

(3.00 mg m<sup>−</sup><sup>3</sup>

(2.27 mg m<sup>−</sup><sup>3</sup>

(1.62 mg m<sup>−</sup><sup>3</sup>

and July (1.10 mg m<sup>−</sup><sup>3</sup>

ally) April (11.89 mg m<sup>−</sup><sup>3</sup>

*Reproductive Biology and Technology in Animals*

*Gonad stage frequency 2014–2015 for H. princeps males by sampling date. Chlorophyll concentration (mg m–3)* 

*The smoothed frequency of the female reproductive stages along the study period and cross-correlation correlograms for mature-spawning (S4-S5) and spawning-resting (S5-S6) reproductive stages comparison.*

**120**

**Figure 6.**

**Figure 5.**

*and temperature (°C) are included.*

May (60%), June (75%), and July (67%) 2014. The months of 2015 with larger spawning (S3) percentages were January (100%) and March (50%); finally, the months with the larger frequency of resting individuals (S5) were May (60%) and June (75%).

#### **3.2 Chlorophyll** *a* **concentration and gonad cycle**

In 2014, the highest chlorophyll *a* concentrations were observed in January (3.00 mg m<sup>−</sup><sup>3</sup> ), February (4.00 mg m<sup>−</sup><sup>3</sup> ), March (2.05 mg m<sup>−</sup><sup>3</sup> ), and December (2.27 mg m<sup>−</sup><sup>3</sup> ); in the same months, the larger frequency of individuals in the spawning stages (S5 females, S3 males) (**Figures 4** and **5**) was observed. For the same year, low chlorophyll concentrations were recorded in June (0.15 mg m<sup>−</sup><sup>3</sup> ) and July (1.10 mg m<sup>−</sup><sup>3</sup> ) months, which correspond with the larger frequency of females and males in the resting stages (S6 and S5, respectively) (**Figures 4** and **5**).

In 2015, the high chlorophyll concentrations were observed in January (1.62 mg m<sup>−</sup><sup>3</sup> ), February (1.23 mg m<sup>−</sup><sup>3</sup> ), March (1.08 mg m<sup>−</sup><sup>3</sup> ), and (exceptionally) April (11.89 mg m<sup>−</sup><sup>3</sup> ); except for April, in all other months, the stage with

#### **Figure 7.**

*The smoothed frequency of the male reproductive stages along the study period and cross-correlation correlograms for mature-spawning (S2-S3) and spawning-resting (S3-S5) reproductive stages comparison.*

**Figure 8.**

*The smoothed frequency of spawning females (S5), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S5-SST and S5-CL comparison.*

larger frequency was spawning (S5 and S3 for females and males, respectively) (**Figures 4** and **5**). In the same year, the lower chlorophyll concentrations occurred in July (0.18 mg m<sup>−</sup><sup>3</sup> ), October (0.18 mg m<sup>−</sup><sup>3</sup> ), and November (0.19 mg m<sup>−</sup><sup>3</sup> ). These chlorophyll concentration values were related to the resting stage of females and males (S6 and S5, respectively) (**Figures 4** and **5**).

It is possible that the April 2015, notably high (11.89 mg m<sup>−</sup><sup>3</sup> ), chlorophyll concentration originated a different pattern, in comparison with that from the same month of the previous year. The resting phase of females (S6) and males (S5) occurred with less frequency and the spawning gonad stage (S5 females and S3 males) extended to June, July, and August (**Figures 4** and **5**).

#### **3.3 Temperature and gonad cycle**

The lowest registered temperatures occurred in January (27.48°C), February (27.74°C), and December (27.96°C). In these months, it was observed that the spawning females (S6) and males (S3) were those with the highest frequency (**Figures 4** and **5**). The months with the highest temperatures were May (30.70°C),

**123**

**Figure 9.**

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

June (31.14°C), and August (30.83°C), which were related to the larger frequency of

*The smoothed frequency of spawning males (S3), surface sea temperature (SST) and chlorophyll concentration* 

*(CL) along the study period and cross-correlation correlograms for S3-SST and S3-CL comparison.*

The lowest temperatures for 2015 were registered in January (28.04°C), February (27.44°C), and March (28.11°C), which corresponded with the highest frequencies of female and male in the spawning stage (S5 and S3, respectively). The months with the larger temperature values were August (31.19°C) and September (31.24°C). In these months, female stage 4 (mature), 5 (spawning), and 6 (resting) were observed with 25, 25, and 50%, respectively; the male stages were 3 (spawning) and 4 (end of spawning, onset of the rest) (50 and 30%, respectively), and in September, 35% of stage 2 (maturity), 25% of phase 3 (spawning), and 40% of stage 5 (resting) were observed.

To describe in more detail the variation and relationships of the reproduction stages and the environmental variables, sea surface temperature (SST) and chlorophyll *a* (Cl) concentrations, along the study period, the smoothed frequency of the reproductive stages were analyzed by cross-correlation. It can be easily seen

the resting stages of females (S6) and males (S5) (**Figures 4** and **5**).

**3.4 Smoothing and cross-correlation**

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

#### **Figure 9.**

*Reproductive Biology and Technology in Animals*

larger frequency was spawning (S5 and S3 for females and males, respectively) (**Figures 4** and **5**). In the same year, the lower chlorophyll concentrations occurred

*The smoothed frequency of spawning females (S5), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S5-SST and S5-CL* 

chlorophyll concentration values were related to the resting stage of females and

concentration originated a different pattern, in comparison with that from the same month of the previous year. The resting phase of females (S6) and males (S5) occurred with less frequency and the spawning gonad stage (S5 females and S3

The lowest registered temperatures occurred in January (27.48°C), February (27.74°C), and December (27.96°C). In these months, it was observed that the spawning females (S6) and males (S3) were those with the highest frequency (**Figures 4** and **5**). The months with the highest temperatures were May (30.70°C),

), and November (0.19 mg m<sup>−</sup><sup>3</sup>

). These

), chlorophyll

), October (0.18 mg m<sup>−</sup><sup>3</sup>

It is possible that the April 2015, notably high (11.89 mg m<sup>−</sup><sup>3</sup>

males) extended to June, July, and August (**Figures 4** and **5**).

males (S6 and S5, respectively) (**Figures 4** and **5**).

**3.3 Temperature and gonad cycle**

**122**

in July (0.18 mg m<sup>−</sup><sup>3</sup>

**Figure 8.**

*comparison.*

*The smoothed frequency of spawning males (S3), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S3-SST and S3-CL comparison.*

June (31.14°C), and August (30.83°C), which were related to the larger frequency of the resting stages of females (S6) and males (S5) (**Figures 4** and **5**).

The lowest temperatures for 2015 were registered in January (28.04°C), February (27.44°C), and March (28.11°C), which corresponded with the highest frequencies of female and male in the spawning stage (S5 and S3, respectively). The months with the larger temperature values were August (31.19°C) and September (31.24°C). In these months, female stage 4 (mature), 5 (spawning), and 6 (resting) were observed with 25, 25, and 50%, respectively; the male stages were 3 (spawning) and 4 (end of spawning, onset of the rest) (50 and 30%, respectively), and in September, 35% of stage 2 (maturity), 25% of phase 3 (spawning), and 40% of stage 5 (resting) were observed.

#### **3.4 Smoothing and cross-correlation**

To describe in more detail the variation and relationships of the reproduction stages and the environmental variables, sea surface temperature (SST) and chlorophyll *a* (Cl) concentrations, along the study period, the smoothed frequency of the reproductive stages were analyzed by cross-correlation. It can be easily seen

#### **Figure 10.**

*The smoothed frequency of resting females (S6), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S6-SST and S6-CL comparison.*

(**Figure 6**) that during the cool months of the year (from October to February), the females mature (S4) and spawn (S5) from October to March. During 2015, the spawning period lasted longer than the previous year. Very clearly, the females are in reproductive rest (S6) during the warmer half of the year (from April to August). The cross-correlation of the series shows a significant positive trend between S4 and S5 stages synchronically and an inverse correlation with 4–5 months lag between spawning (S5) and rest (S6).

The males seem to mature (S2) early (July) but the trend is clear from September to January (**Figure 7**). The spawning males (S3) occur from October to February. Similarly than females, males attain the reproductive rest stage (S5) from April to August, during the warmer months.

The cross-correlograms show a direct relationship lagged 2 months between S2 and S3 (maturity, spawning stage) and a 6 month lagged high cross-correlation between spawning (S3) and resting (S5) males, corroborating significantly the above statements.

The spawning (S5) females showed a clear opposite (negative) correlation with the sea surface temperature values with a lag of 6 months, and concordant

**125**

**Table 5.**

*significance values.*

**Figure 11.**

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

*The smoothed frequency of resting males (S5), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S5-SST and S5-CL comparison.*

**Sex Sequence Lag Cross-correlation** *P***-value** Females S4-S5 0 0.7144 0.0001

Males S2-S3 2 0.4165 0.0182

Females S5-SST 6 0.6278 0.0001

Males S3-SST 7 0.5366 0.0013

Females S6-SST 2 0.7281 0.0000

Males S5-SST 2 0.7159 0.0000

*Cross-correlation analysis results resume: sex, sequences compared, time lag (month), cross-correlation, and* 

S5-S6 4 0.7525 0.0000

S3-S5 5 0.7192 0.0000

S5-CL 0 0.8442 0.0000

S3-CL 1 0.6835 0.0000

S6-CL −4 0.8384 0.0000

S5-CL −4 0.8467 0.0000

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

#### **Figure 11.**

*Reproductive Biology and Technology in Animals*

(**Figure 6**) that during the cool months of the year (from October to February), the females mature (S4) and spawn (S5) from October to March. During 2015, the spawning period lasted longer than the previous year. Very clearly, the females are in reproductive rest (S6) during the warmer half of the year (from April to August). The cross-correlation of the series shows a significant positive trend between S4 and S5 stages synchronically and an inverse correlation with 4–5 months lag

*The smoothed frequency of resting females (S6), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S6-SST and S6-CL comparison.*

The males seem to mature (S2) early (July) but the trend is clear from September to January (**Figure 7**). The spawning males (S3) occur from October to February. Similarly than females, males attain the reproductive rest stage (S5) from

The cross-correlograms show a direct relationship lagged 2 months between S2 and S3 (maturity, spawning stage) and a 6 month lagged high cross-correlation between spawning (S3) and resting (S5) males, corroborating significantly the

The spawning (S5) females showed a clear opposite (negative) correlation with the sea surface temperature values with a lag of 6 months, and concordant

**124**

above statements.

**Figure 10.**

between spawning (S5) and rest (S6).

April to August, during the warmer months.

*The smoothed frequency of resting males (S5), surface sea temperature (SST) and chlorophyll concentration (CL) along the study period and cross-correlation correlograms for S5-SST and S5-CL comparison.*


#### **Table 5.**

*Cross-correlation analysis results resume: sex, sequences compared, time lag (month), cross-correlation, and significance values.*

(positive) cross-correlation with chlorophyll *a* concentration with no lag (**Figure 8**). In **Figure 9**, it is possible to see that, as the females, spawning (S3) males had negative cross-correlation values with SST values (lagged around 7 months), and positive with chlorophyll (1-month lag). In contrast with the former trends, the resting females (S6) showed positive cross-correlations (lagged 2 months) with SST and negative relationship (with a lag of 7 months) with the chlorophyll concentrations (**Figure 10**). In a similar way as resting females, males in reproductive rest stage (S5) showed a direct trend with SST (lagged 1 or 2 months) and the opposite of the chlorophyll values (a lag of 7 months) (**Figure 11**). A resume of the cross-correlations significances is included in **Table 5**.

#### **4. Discussion**

The sex proportion of *H. princeps* from Puerto Ángel was found to be statistically balanced with a slight preponderance of males during the warmer months of the year. These findings correspond with the study of Vasconcelos *et al.* [36], who reported a balanced sex ratio and males dominating among smaller individuals of *Hexaplex* (*Trunculariopsis*) *trunculus* in the Ria Formosa Lagoon in Portugal. This is contrary to the unbalanced sex ratio reported by Elhasni *et al.* [4] for *Bolinus brandaris* (another Muricid) in Tunisia, where females surpassed males, mainly during the reproduction period. Although not significant, at Puerto Ángel in the cold months (January 2014 and January, March 2015), when the reproductive event occurs, the number of females was larger than that of males, which may be associated with the reproductive behavior of this species as the females tend to aggregate for oviposition.

The histological examination of the gonads of *Hexaplex princeps* at Puerto Ángel permitted to characterize six stages of maturity development in females (**Table 3**) and five for the males (**Table 4**). Although there are no previous reports on the histological maturity for *H. princeps*, the stages characterized in the present study for the females correspond closely to those suggested for *H. trunculus* from Portugal [36]. In the case of male maturity and based on our observations of the histological sections, we consider that only five stages are enough to describe the spermatogenic cycle.

Comparing both sexes, spawning and expulsion occurred in January, April, May, October, and November, and the larger frequency of resting individuals was registered in June and July. In this way, it is possible to recognize a period of spawning and expulsion from November to March with pikes in January and February. The resting period of females occurred from March to October, with peaks in July (2014) and April–July (2015), and the males presented high resting frequency values in June (2014) and May (2015). This does not corresponds to the reproduction times reported for *H. erythrostomus* from Bahía Concepción, Baja California Sur, where the reproductive events were annotated during the warmer months (May–July) [37]; though it has to be noted that the highest temperature of 28°C of the Bahía Concepción sea surface water temperature corresponds with the cooler temperatures of Puerto Ángel. The temperature is one of the most important external environmental factors that affects molluscan reproduction and in the case of *H. princeps*, for both sexes, the spawning and expulsion stages occurred at relatively low temperatures and the resting period at warmer temperatures of surface waters at the studied locality.

Chlorophyll concentrations have a direct relationship with the development of gonads as this reproductive process demands high energetic quantities that must be obtained from the eaten food extracted from the environment or from reserves previously accumulated or from both [6, 38]. *H. princeps* is a predator

**127**

**5. Conclusions**

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

gastropod that depends on the energy obtained from its preys. So, during 2014, for the females, it was noted that when the chlorophyll *a* concentrations were high, maturity and spawning stages presented a higher occurrence percentage and when the concentrations were low, the most frequent gonad stage is resting. For the males, when the chlorophyll concentrations were high, the most frequent gonad stage was the expulsion and when low concentrations occurred, the most frequent gonad stage was resting. In 2015, chlorophyll concentrations were very variable having high values from January to March and an increment in November corresponding with larger percentages of spawning females and expulsing males. However, an anomalous high peak of chlorophyll concentration occurred in April when, unlike the same month from 2014, could have caused the reduction of the resting stage and oogenesis and spermatogenesis occurred as indicated by presence of the spawning stage. Therefore, the periods with large chlorophyll availability coincide with the gonad development. *H. princeps* tends to reproduce when the phytoplankton population is blooming, so its offspring could have a higher probability of survival

In relation to temperature is worth to mention that along the period of study, the water temperature differences between surface-bottom lectures were not detected. The direct explanation for this finding is that the rocky coast localities where the individuals of *H. princeps* were collected are places very energetic under the effects of strong wave action, precisely the zones usually inhabited by this organism [7, 8]. In this study, the predominance of reproductive stages occurred during the winter months under relatively colder temperatures. As noted before, the reproduction of the related species *H. erythrostomus* in the Gulf of California happened when the water temperature was around 28°C [37], figure similar to the January surface

From October to April but mainly from November to February, the blowing winds, known as the "Tehuanos," originate upwelling and water vertical mixing causing an increase in chlorophyll concentrations [39, 40] by phytoplanktonic blooming. This water mixing process promotes spawning, breeding, and feeding of the aquatic species [41]. Thus, it is possible that the food availability is the main

On the other side, we would like to mention that the smoothing technique applied to the maturity stage frequencies allow distinguishing in a clearer way the subjacent pattern of the reproductive cycle. With the availability of long-term data records, it is possible to use time series analysis statistical techniques, making it possible to assess the significance of the observed behaviors. From this, it can be stated that *H. princeps* matures and spawns during the cold months of the year (October– March) and rest its reproduction during the warm part of the year (May–August). It observed a positive correlation with the quantity of food (indirectly indicated by the chlorophyll *a* concentrations) and in all the cases, the leads and lags of these

This is the first study on the reproductive cycle of *H. princeps* in the region and present baseline information for: (i) potential management measures, in particular, the knowledge of the timing of spawning season, (ii) assessment of aquaculture

The reproductive cycle of *Hexaplex princeps* was studied through monthly histological analysis of gonads during two annual periods (2014, 2015) from specimens obtained from the artisanal fishery at Puerto Ángel, Oaxaca, Mexico,

factor affecting the onset of reproduction of *H. princeps* in this region.

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

due to food abundance [14, 15].

temperatures of Puerto Ángel.

variables can be determined [35].

potential. More research on the subject is needed.

*Reproductive Biology and Technology in Animals*

in **Table 5**.

**4. Discussion**

(positive) cross-correlation with chlorophyll *a* concentration with no lag (**Figure 8**). In **Figure 9**, it is possible to see that, as the females, spawning (S3) males had negative cross-correlation values with SST values (lagged around 7 months), and positive with chlorophyll (1-month lag). In contrast with the former trends, the resting females (S6) showed positive cross-correlations (lagged 2 months) with SST and negative relationship (with a lag of 7 months) with the chlorophyll concentrations (**Figure 10**). In a similar way as resting females, males in reproductive rest stage (S5) showed a direct trend with SST (lagged 1 or 2 months) and the opposite of the chlorophyll values (a lag of 7 months) (**Figure 11**). A resume of the cross-correlations significances is included

The sex proportion of *H. princeps* from Puerto Ángel was found to be statistically balanced with a slight preponderance of males during the warmer months of the year. These findings correspond with the study of Vasconcelos *et al.* [36], who reported a balanced sex ratio and males dominating among smaller individuals of *Hexaplex* (*Trunculariopsis*) *trunculus* in the Ria Formosa Lagoon in Portugal. This is contrary to the unbalanced sex ratio reported by Elhasni *et al.* [4] for *Bolinus brandaris* (another Muricid) in Tunisia, where females surpassed males, mainly during the reproduction period. Although not significant, at Puerto Ángel in the cold months (January 2014 and January, March 2015), when the reproductive event occurs, the number of females was larger than that of males, which may be associated with the reproductive

The histological examination of the gonads of *Hexaplex princeps* at Puerto Ángel permitted to characterize six stages of maturity development in females (**Table 3**) and five for the males (**Table 4**). Although there are no previous reports on the histological maturity for *H. princeps*, the stages characterized in the present study for the females correspond closely to those suggested for *H. trunculus* from Portugal [36]. In the case of male maturity and based on our observations of the histological sections, we consider that only five stages are enough to describe the spermatogenic

Comparing both sexes, spawning and expulsion occurred in January, April, May, October, and November, and the larger frequency of resting individuals was registered in June and July. In this way, it is possible to recognize a period of spawning and expulsion from November to March with pikes in January and February. The resting period of females occurred from March to October, with peaks in July (2014) and April–July (2015), and the males presented high resting frequency values in June (2014) and May (2015). This does not corresponds to the reproduction times reported for *H. erythrostomus* from Bahía Concepción, Baja California Sur, where the reproductive events were annotated during the warmer months (May–July) [37]; though it has to be noted that the highest temperature of 28°C of the Bahía Concepción sea surface water temperature corresponds with the cooler temperatures of Puerto Ángel. The temperature is one of the most important external environmental factors that affects molluscan reproduction and in the case of *H. princeps*, for both sexes, the spawning and expulsion stages occurred at relatively low temperatures and the resting period at warmer temperatures of surface waters at the studied locality. Chlorophyll concentrations have a direct relationship with the development of gonads as this reproductive process demands high energetic quantities that must be obtained from the eaten food extracted from the environment or from reserves previously accumulated or from both [6, 38]. *H. princeps* is a predator

behavior of this species as the females tend to aggregate for oviposition.

**126**

cycle.

gastropod that depends on the energy obtained from its preys. So, during 2014, for the females, it was noted that when the chlorophyll *a* concentrations were high, maturity and spawning stages presented a higher occurrence percentage and when the concentrations were low, the most frequent gonad stage is resting. For the males, when the chlorophyll concentrations were high, the most frequent gonad stage was the expulsion and when low concentrations occurred, the most frequent gonad stage was resting. In 2015, chlorophyll concentrations were very variable having high values from January to March and an increment in November corresponding with larger percentages of spawning females and expulsing males. However, an anomalous high peak of chlorophyll concentration occurred in April when, unlike the same month from 2014, could have caused the reduction of the resting stage and oogenesis and spermatogenesis occurred as indicated by presence of the spawning stage. Therefore, the periods with large chlorophyll availability coincide with the gonad development. *H. princeps* tends to reproduce when the phytoplankton population is blooming, so its offspring could have a higher probability of survival due to food abundance [14, 15].

In relation to temperature is worth to mention that along the period of study, the water temperature differences between surface-bottom lectures were not detected. The direct explanation for this finding is that the rocky coast localities where the individuals of *H. princeps* were collected are places very energetic under the effects of strong wave action, precisely the zones usually inhabited by this organism [7, 8].

In this study, the predominance of reproductive stages occurred during the winter months under relatively colder temperatures. As noted before, the reproduction of the related species *H. erythrostomus* in the Gulf of California happened when the water temperature was around 28°C [37], figure similar to the January surface temperatures of Puerto Ángel.

From October to April but mainly from November to February, the blowing winds, known as the "Tehuanos," originate upwelling and water vertical mixing causing an increase in chlorophyll concentrations [39, 40] by phytoplanktonic blooming. This water mixing process promotes spawning, breeding, and feeding of the aquatic species [41]. Thus, it is possible that the food availability is the main factor affecting the onset of reproduction of *H. princeps* in this region.

On the other side, we would like to mention that the smoothing technique applied to the maturity stage frequencies allow distinguishing in a clearer way the subjacent pattern of the reproductive cycle. With the availability of long-term data records, it is possible to use time series analysis statistical techniques, making it possible to assess the significance of the observed behaviors. From this, it can be stated that *H. princeps* matures and spawns during the cold months of the year (October– March) and rest its reproduction during the warm part of the year (May–August). It observed a positive correlation with the quantity of food (indirectly indicated by the chlorophyll *a* concentrations) and in all the cases, the leads and lags of these variables can be determined [35].

This is the first study on the reproductive cycle of *H. princeps* in the region and present baseline information for: (i) potential management measures, in particular, the knowledge of the timing of spawning season, (ii) assessment of aquaculture potential. More research on the subject is needed.

#### **5. Conclusions**

The reproductive cycle of *Hexaplex princeps* was studied through monthly histological analysis of gonads during two annual periods (2014, 2015) from specimens obtained from the artisanal fishery at Puerto Ángel, Oaxaca, Mexico, considering sexual proportion, gonad maturation, spawning periods, and maturity stages variation in relation with environmental factors (sea surface temperature and chlorophyll concentration).

The sexual proportion was not statistically different from parity, although most of the times, the number of males was slightly larger than the number of females. Only during the spawning season, females were more frequent than males.

The histological analysis permitted to establish maturity stages.

Females (six stages): (1) initial oogenesis, (2) previtellogenic maturity, (3) vitellogenic maturity, (4) maturity, (5) spawning, and (6) resting.

Males (five stages): (1) initial spermatogenesis, (2) maturity, (3) spawning, (4) onset of the rest, and (5) resting.

Monthly variations of maturation stages showed that *H. princeps* has an annual reproductive cycle with a long period of gonadal activity. The spawning season comprised from November to March (females) and from December to March (males) with activity peaks in January. From March to October (females) and May to June (males) reproduction resting occurred.

Reproductive events were related to high chlorophyll *a* concentrations due to the upwelling processes resulting from the predominant winds and to the relatively cooler temperatures proper to the winter (November–March) season at Puerto Ángel, Oaxaca, Mexico.

#### **Acknowledgements**

We express our thankfulness to the Posgrado en Ciencias Biológicas, Universidad Nacional Autónoma de México (UNAM), and the Consejo Nacional de Ciencia y Tecnología (CONACyT) for their support. Without the help of the local divers Manolo Jarquín, Primitivo Herrera Ordóñez and Captain "Beto" (Abraham Reyes López), the collection of specimens would not have been possible. We are grateful to Dr. Erika Rosales Cruz for her help with the histological photographs and to the Central de Instrumentación de Microscopía, Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional. This research received support from Dirección del Personal Académico, UNAM under the programs PAPIME (PE206213, PE207417) and (partially) PAPIIT (IG201215). Besides, we received support from the Carrera de Biología, Facultad de Estudios Superiores (FES) Zaragoza, UNAM. Lastly, we would thank all the students and colleagues of the Laboratorio de Biometría y Biología Pesquera, FES Zaragoza, UNAM. This research report is part of first author's PhD thesis.

**129**

Mexico

**Author details**

Verónica Mitsui Saito-Quezada1

Ana Bertha Villaseñor-Martínez4

and Isaías Hazarmabeth Salgado-Ugarte1

provided the original work is properly cited.

, Esther Uría-Galicia2

\*

1 Laboratorio de Biometría y Biología Pesquera, FES Zaragoza, UNAM, Mexico

2 Laboratorio de Histología, Departamento de Morfología, ENCB, IPN, Mexico

5 Instituto de Ciencias Marinas y Pesquerías, Universidad Veracruzana, Veracruz,

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium,

4 Departamento de Paleontología, Instituto de Geología, UNAM, Mexico

3 Laboratorio de Limnología, FES Zaragoza, UNAM, Mexico

\*Address all correspondence to: ihsalgadougarte@gmail.com

, Ma. De Lourdes Jiménez-Badillo5

, José Luis Gómez-Márquez3

,

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833)*

*DOI: http://dx.doi.org/10.5772/intechopen.88074*

#### **Conflict of interest**

We state that there is no "conflict of interest."

*Reproductive Cycle of* Hexaplex princeps *(Broderip, 1833) DOI: http://dx.doi.org/10.5772/intechopen.88074*

*Reproductive Biology and Technology in Animals*

chlorophyll concentration).

(4) onset of the rest, and (5) resting.

Ángel, Oaxaca, Mexico.

**Acknowledgements**

of first author's PhD thesis.

We state that there is no "conflict of interest."

**Conflict of interest**

to June (males) reproduction resting occurred.

considering sexual proportion, gonad maturation, spawning periods, and maturity stages variation in relation with environmental factors (sea surface temperature and

The sexual proportion was not statistically different from parity, although most of the times, the number of males was slightly larger than the number of females. Only during the spawning season, females were more frequent than males. The histological analysis permitted to establish maturity stages.

Females (six stages): (1) initial oogenesis, (2) previtellogenic maturity, (3) vitel-

Males (five stages): (1) initial spermatogenesis, (2) maturity, (3) spawning,

Monthly variations of maturation stages showed that *H. princeps* has an annual reproductive cycle with a long period of gonadal activity. The spawning season comprised from November to March (females) and from December to March (males) with activity peaks in January. From March to October (females) and May

Reproductive events were related to high chlorophyll *a* concentrations due to the upwelling processes resulting from the predominant winds and to the relatively cooler temperatures proper to the winter (November–March) season at Puerto

We express our thankfulness to the Posgrado en Ciencias Biológicas, Universidad

Nacional Autónoma de México (UNAM), and the Consejo Nacional de Ciencia y Tecnología (CONACyT) for their support. Without the help of the local divers Manolo Jarquín, Primitivo Herrera Ordóñez and Captain "Beto" (Abraham Reyes López), the collection of specimens would not have been possible. We are grateful to Dr. Erika Rosales Cruz for her help with the histological photographs and to the Central de Instrumentación de Microscopía, Escuela Nacional de Ciencias Biológicas, Instituto Politécnico Nacional. This research received support from Dirección del Personal Académico, UNAM under the programs PAPIME (PE206213,

PE207417) and (partially) PAPIIT (IG201215). Besides, we received support from the Carrera de Biología, Facultad de Estudios Superiores (FES) Zaragoza, UNAM. Lastly, we would thank all the students and colleagues of the Laboratorio de Biometría y Biología Pesquera, FES Zaragoza, UNAM. This research report is part

logenic maturity, (4) maturity, (5) spawning, and (6) resting.

**128**

#### **Author details**

Verónica Mitsui Saito-Quezada1 , Esther Uría-Galicia2 , José Luis Gómez-Márquez3 , Ana Bertha Villaseñor-Martínez4 , Ma. De Lourdes Jiménez-Badillo5 and Isaías Hazarmabeth Salgado-Ugarte1 \*

1 Laboratorio de Biometría y Biología Pesquera, FES Zaragoza, UNAM, Mexico

2 Laboratorio de Histología, Departamento de Morfología, ENCB, IPN, Mexico

3 Laboratorio de Limnología, FES Zaragoza, UNAM, Mexico

4 Departamento de Paleontología, Instituto de Geología, UNAM, Mexico

5 Instituto de Ciencias Marinas y Pesquerías, Universidad Veracruzana, Veracruz, Mexico

\*Address all correspondence to: ihsalgadougarte@gmail.com

© 2019 The Author(s). Licensee IntechOpen. This chapter is distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/ by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

### **References**

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[3] Leiva GE, Castilla JC. A review of the world marine gastropod fishery: Evolution of the catches, management and the Chilean experience. Reviews in Fish Biology and Fisheries. 2002;**1**:283- 300. DOI: 10.1023/a:1021368216294

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[5] Secretaría de Agricultura, Ganadería, Desarrollo Rural, Pesca y Alimentación (SAGARPA). Anuario Estadístico de Acuacultura y Pesca 2013 de la Comisión Nacional de Acuacultura y Pesca; Mexico2015. p. 299

[6] Fretter V. Prosobranchs. In: Tompa AS, Verdonk HH, Van Der Biggelaar J, editors. The Mollusca: Reproduction. Vol. 7. London: Academic Press; 1984. pp. 1-45

[7] Keen AM. Sea Shells of Tropical West America (Marine Mollusks from Baja California to Peru). Stanford: Stanford University Press; 1971. p. 632

[8] Morris AP. A Field Guide to Pacific Coast Shells Including Shells of Hawaii and the Gulf of California. Boston: Houghton Mifflin Company; 1976. p. 176

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**132**

### *Edited by Juan Carlos Gardón Poggi and Katy Satué Ambrojo*

Reproductive success is a very important objective to ensure the evolution of animal species. In this sense, interesting research has been carried out to clarify various aspects of reproduction in different animal species. In this way, recent advances in the knowledge of reproductive biology and biotechnology developed for both males and females have been key to improving efficiency in different aspects. Thus, advances in the knowledge of sperm handling, oocyte characteristics, different genomic aspects related to somatic cell nuclear transfer, and the reproductive microarchitecture system in sheep, cows, pigs, and other invertebrates such as gastropods and fish are presented in this book. Additionally, we also present the most relevant topics of each area, making a detailed review of the knowledge reported to date.

Published in London, UK © 2020 IntechOpen © golfbress / iStock

Reproductive Biology and Technology in Animals

IntechOpen Book Series

Veterinary Medicine and Science, Volume 4

Reproductive Biology and

Technology in Animals

*Edited by Juan Carlos Gardón Poggi* 

*and Katy Satué Ambrojo*