Cell Death in Mammalian Ovary

Cell Death in Mammalian Ovary

Gerardo H. Vázquez-Nin  •  María L. Escobar Massimo De Felici  •  Olga M. Echeverría Francesca G. Klinger

Cell Death in Mammalian Ovary

Prof. Gerardo H. Vázquez-Nin Department of Cell Biology Faculty of Sciences National Autonomous University of Mexico (UNAM) Avenida Universidad 3000 04510 Mexico, D.F. Mexico [email protected] Dr. Massimo De Felici Department of Public Health and Cell Biology Section of Histology and Embryology University of Rome Tor Vergata Via Montpellier 1 00133 Rome Italy

Dr. María L. Escobar Department of Cell Biology Faculty of Sciences National Autonomous University of Mexico (UNAM) Avenida Universidad 3000 04510 Mexico, D.F. Mexico Dr. Olga M. Echeverría Department of Cell Biology Faculty of Sciences National Autonomous University of Mexico (UNAM) Avenida Universidad 3000 04510 Mexico, D.F. Mexico

Dr. Francesca G. Klinger Department of Public Health and Cell Biology Section of Histology and Embryology University of Rome Tor Vergata Via Montpellier 1 00133 Rome Italy

ISBN 978-94-007-1133-4 e-ISBN 978-94-007-1134-1 DOI 10.1007/978-94-007-1134-1 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2011929630 © Springer Science+Business Media B.V. 2011 No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Contents

Part I  Introduction   1 Brief Description of the Histological, Cytological and Functional Aspects of the Ovary..................................................... María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría   2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis............................................................................................... Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría   3 Development of the Ovary in the Embryo, Infancy, Childhood, Pre-puberty and Puberty.................................................... Gerardo H. Vázquez-Nin, María Luisa Escobar, Olga M. Echeverría, and Massimo De Felici

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Part II  General Aspects of Cell Death   4 Apoptosis................................................................................................... María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

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  5 Autophagy................................................................................................. María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

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  6 Oncosis...................................................................................................... 103 María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

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  7 Necrosis..................................................................................................... 111 Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría Part III  Process of Cell Death Embryonic Ovary   8 Programmed Cell Death in Fetal Oocytes............................................. 125 Francesca Gioia Klinger and Massimo De Felici   9 DNA Damage and Apoptosis in Fetal and Ovarian Reserve Oocytes....................................................................................... 143 Massimo De Felici and Francesca Gioia Klinger 10 Prefollicular Cells..................................................................................... 165 María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría Part IV  Process of Cell Death Prepubertal Ovary 11 Prepubertal Oocytes................................................................................ 173 Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría 12 Follicular Cells......................................................................................... 185 María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría Part V  Process of Cell Death Adult Ovary 13 Follicular Atresia in Adult Animals....................................................... 203 Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría 14 Luteolysis.................................................................................................. 221 Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría Index.................................................................................................................. 233

Part I

Introduction

Chapter 1

Brief Description of the Histological, Cytological and Functional Aspects of the Ovary María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

Abstract  The ovary is formed by three main compartments: superficial epithelium, cortex and medulla. The superficial epithelium is constituted by one layer of cubic cells. The cortex is a wide peripheral zone containing the follicles, the functional and structural unit of the ovary, and a stroma formed by compact connective tissue. Every follicle is formed by one oocyte surrounded by follicular cells, also called granulosa cells, and a basal lamina surrounding them. The medulla is the central region of the ovary formed by connective tissue with numerous blood vessels. As the follicles develop they change their size, morphology and physiology. Primordial follicles are formed by the oocyte surrounded by flat follicular cells. Primary ­follicles are characterized by the initiation of follicular growth. Secondary follicles are characterized by two or more layers of granulosa cells and no antrum. The early antral follicles are characterized by the formation and progressive growth of a cavity, due to the accumulation of a fluid. Once the antrum is formed the follicle goes through several stages: (a) basal growth, (b) selection and (c) dominance. The process of follicular growth is controlled by extra-ovarian and intra-ovarian factors and the importance of each of these factors depends on the stage of follicle development. Extra-ovarian factors regulate growth of antral and preovulatory follicles, while intra-ovarian factors regulate growth of preantral and early antral follicles. The ovary is not only involved in sexual reproduction, but also has great influence on the entire hormonal functioning during development of the organism. The ovary is the site of the highest synthesis and secretion of progesterone and estrogen in mammals and gives rise to cyclical fluctuations in the levels of these hormones in the blood. Before ovulation, granulosa cells mature to form the corpus luteum, which is responsible for the secretion of progesterone and estrogen.

M.L. Escobar (*), G.H. Vázquez-Nin, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico City, Mexico e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_1, © Springer Science+Business Media B.V. 2011

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List of Abbreviations LH FSH TGFb EGF

1.1

Luteinizing hormone Follicle stimulant hormone Transforming growth factor beta Epidermal growth factor

Morphological Characteristics of the Ovary

The primary reproductive organs of the female are the ovaries. Their main functions are the production of fertilizable oocytes and the secretion of steroid hormones (estrogen and progesterone), which are required for the correct function of the reproductive organs such as the Fallopian tubes, uterus and vagina. The ovary is formed by three main compartments: superficial epithelium, cortex and medulla (Fig.  1.1). The superficial epithelium is constituted by one layer of cubic cells, which are continuous with peritoneal epithelium at the periphery of the ovary. The cortex is a wide peripheral zone containing the follicles, the functional and structural unit of the ovary, and a stroma formed by compact connective tissue.

Fig. 1.1  Rat ovary 5 days old. The ovary is constituted by three regions: superficial epithelium, cortex and stroma. In the cortex zone diverse follicles are observed. Haematoxylin-eosin stain technique

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Fig. 1.2  Follicular classification according to McGee and Hsueh (2000). The classification of the authors includes six different development follicular phases

Every follicle is formed by one oocyte surrounded by follicular cells, also called granulosa cells, and a basal lamina surrounding them. The medulla is the central region of the ovary formed by connective tissue with numerous blood vessels. In some individuals there is a net of epithelial cords or tubules near the hilum, the rete ovarii. As the follicles develop they change their size, morphology and physiology. This development has been traditionally classified in several stages. However, there are differences in the name and the characterization of the stages. McGee and Hsueh (2000) described the following phases of development of the follicles as: ­primordial, primary, secondary, early antral, antral and pre-ovulatory stages (Fig. 1.2). These authors also characterized the hormonal factors involved in the survival and ­development of rodent follicles. Primordial follicles are formed by the oocyte surrounded by flat follicular cells (Fig. 1.3). The oocytes in these follicles are called quiescent because they remain unchanged for months in rodents or even years in ruminants and primates. There is a system of communication between the oocyte and the granulosa cells which is important in the coordination of the initial follicular development. Several studies carried out on various mammals demonstrate that primordial follicles may express leukemia inhibitory factor (LIF), which promotes follicular growth and

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Fig. 1.3  Primordial follicles. The oocytes are surrounded by a few flattened cells (arrows). Scale bar 50 mm. Haematoxylin-eosin stain technique

stimulates oocyte growth, proliferation of thecal cells and transition from ­primordial follicle to primary stage. LIF is mainly produced by granulosa and somatic cells (Nilsson et al. 2002). In this system of communication the Kit ligand (KitL) is also involved as an essential factor for the proliferation of the granulosa cells (Huang et al. 1993). KitL is segregated to intercellular space before reaching its receptor Kit. KitL stimulates oocyte growth via Kit (Montro and Bernstein 1993; Tisdall et al. 1999; Klinger and De Felici 2002). LIF interacts with KitL in the activation of primordial follicles (Nilsson et  al. 2002). Mouse oocytes express the tyrosine kinase Kit receptor in all stages of follicular growth, including primordial follicles (Manova et al. 1990). Natural mutations in KitL or in Kit induce alterations of follicular development causing infertility (Manova et al. 1990; Driancourt et al. 2000). The gene nobox (newborn ovary homeobox) is also expressed in primordial follicles and helps in the transition to primary stage (Rajkovic et al. 2004). The pool of primordial follicles is now known to be maintained in a dormant state by various forms of inhibitory machinery, which are provided by several inhibitory signals and molecules. Several recently reported mutant mouse models have shown that a synergistic and coordinated suppression of follicular activation provided by multiple inhibitory molecules is necessary to preserve the dormant follicular pool. Loss of function of any of the inhibitory molecules for follicular activation, including PTEN (phosphatase and tensin homolog deleted on chromosome 10), Foxo3a, p27, and Foxl2, leads to premature and irreversible activation of the primordial follicle pool. Such global activation of the primordial follicle pool leads to the exhaustion of the resting follicle reserve, resulting in premature ovarian failure (POF) in mice (for a review, see Adhikari and Liu 2009). Primary follicles are characterized by the initiation of follicular growth, by a change in the shape of granulosa cells, from flat cells they become cubic, the increase in the size of the oocyte and the formation of the zona pellucida (Rankin et al. 1996). The image of primary follicles is characterized by an oocyte surrounded by one layer of cubic granulosa cells and the basal lamina. Blood vessels are present only in the surrounding connective tissue and do not penetrate through the basal lamina (Fig. 1.4). Granulosa cells are related by gap junctions, allowing the passage of some molecules between cells and thus forming a metabolic syncytium, which compensate the

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Fig. 1.4  Primary follicles. Oocytes surrounded by a single line of granulosa cells (arrows). The granulosa cells have acquired the cubic shape. Scale bar 50  mm. Haematoxylin-eosin stain ­technique

remoteness of the blood vessels for themselves as well as for the oocyte. The gap junctions allow for communication among granulosa cells and also between ­granulosa cells and the oocyte. The gap junctions allow the transference of nutrients and metabolic precursors such as amino acids and nucleotides, hormones, neutropins and growth factors, as well as regulatory signals of meiosis. As such, there are interchanges that may promote the growth and differentiation of the oocyte. These gap junctions contain different connexins as: −32, −43, −45 (Rankin et al. 1996). Secondary follicles are characterized by two or more layers of granulosa cells and no antrum. This is a period of intense oocyte growth and rapid proliferation of granulosa cells. During this stage the differentiation of thecal cells takes place. Some of the cells of the connective tissue become arranged parallel to the basal lamina. The cells located closer to the basal lamina give rise to the internal theca and the remaining cells form the external theca (Fig.  1.5). This differentiation begins when the follicle has three layers of granulosa cells. During development, these cells acquire important functions such as synthesis of androgens and paracrine secretions for granulosa cells, fundamental for follicular development and maturation (Magoffin 2005). The secondary follicles are close to a net of anastomosed capillaries originating in one or two arterioles, thus they are much better irrigated than smaller follicles (Bassett 1943). The oocytes in secondary follicles increase their volume and their cytoplasmic organization becomes more complex, due to the synthesis of new RNAs, proteins, glycogen and lipids, as swell as increase in the number of ribosomes, mitochondria and other organelles (Picton et  al. 1998). One of the main changes during oocyte growth is the secretion of glycoproteins, mainly the proteins ZP1, ZP2 and ZP3, the

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Fig. 1.5  Secondary follicles. Oocyte (arrow head) surrounded by various lines of granulosa cells (arrows). The follicles in this phase is surrounded by flattened cells of the theca (dotted arrow). Scale bar 50 mm. Haematoxylin-eosin stain technique

Fig. 1.6  Secondary follicle stained with Periodic Acid Schiff (PAS) method for polysaccharides (red). The oocyte is surrounded by the PAS positive zona pellucida (arrow). Several granulosa cells (arrow head) are disposed around the oocyte. Basal lamina (B) is also PAS positive. The dotted arrow points to theca cells. N oocyte nucleus, C oocyte cytoplasm. Scale bar 10 mm

principal constituents of the zona pellucida (Fig.  1.6). The formation of the zona pellucida is essential for follicular development. The transcription factor germline alpha (Fig. a) is required for the expression of these three proteins (Soyal et  al. 2000). Fig. a is expressed specifically in oocytes and is required for the formation of

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primordial follicles (Soyal et al. 2000). In Fig. a null mice the formation of the gonad takes place but there are no primordial follicles. Fig. a is required for the expression of the gene coding for ZP proteins. The protein Dazla, a germ cell ­specific RNA binding protein is also essential for differentiation and ­development of germ cells (Rugglu et al. 1997). The germ cell nuclear factor (GCNFd) is another specific factor needed for the development of germinal cells (Katz et al. 1997). The cytoplasmic prolongations of the granulosa cells surrounding the oocyte go through the zona pellucida and contact the cell membrane of the oocyte, as classical electron microscopical studies have demonstrated (Sotelo and Porter 1959; Franchi 1960; Albertini et al. 2001; Motta et al. 1994). These contacts are numerous in the preantral stage and during rapid growth of the oocyte (Fig. 1.7). These contacts are vital for metabolic interchange between the oocyte and the granulosa cells.

Fig. 1.7  Oocytes showing the zona pellucida (ZP), the granulosa cells (GC) are in contact with the cytoplasm of the oocyte (C) via cytoplasmatic prolongations (arrow heads). (a) Periodic Acid Schiff (PAS) method. (b) Electron micrograph contrasted with uranyl acetate-lead citrate staining method. Scale bars: a 10 mm; b 2 mm

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Fig.  1.8  Early antral follicles. In this phase the follicle begins to form the cavity that will c­ onstitute the antrum (arrows). Periodic Acid Schiff (PAS). Scale bar 50 mm

In spite of the limited vasculature of the secondary follicles, they develop to antral follicles under the stimulation of gonadotropic hormones as Follicle Stimulating Hormone (FSH) (van den Hurk et al. 2000; McGee and Hsueh 2000). The previous action of the Luteinizing Hormone (LH) seems to be important for the action of FSH because it acts through the LH receptors of the thecal cells initiating androgen biosynthesis. The androgens stimulate the formation of FSH receptors in granulosa cells allowing the action of FSH in the development of secondary ­follicles (van den Hurk et al. 2000; van den Hurk et al. 1999). The early antral follicles are characterized by the formation and progressive growth of a cavity, due to the accumulation of a fluid (Fig. 1.8). This antral fluid contains different substances derived from the blood and secretion of follicular cells as regulatory proteins, gonadotropins, steroids, growth factors, proteoglycans, lipoproteins and numerous small molecules. During follicular development the size of the follicle rises rapidly due to an increment in the production of the antral fluid which is caused by an increase in the vascularization of the theca interna and in the permeability of the capillaries surrounding the follicle. In this stage there are two types of granulosa cells, those which form the wall of the follicle and those surrounding the oocyte, forming the cumulus oophorus. During this phase of follicle development, there is an increase in the size of the oocyte and an increase in the number of the granulosa cells forming the cumulus oophorus. Once the antrum is formed the follicle goes through several stages: (a) basal growth; (b) selection and (c) dominance (Fig. 1.9). (a) Basal growth. The thecal cells increase the expression of enzymes involved in the synthesis of steroids and granulosa cells shut down the expression of ­aromatases. These changes probably mean that progesterone and androgens are the main steroid hormones produced by the growing follicle. (b) Selection. The selection of growing follicles begins in the presence of low ­levels of FSH and high LH secretion. In this process a number of dominant ­follicles are selected to continue their development, according to the size of the litter. During the selection phase, the follicles become more dependent on FSH

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Fig.  1.9  Low magnification image of rat ovary stained with haematoxylin-eosin technique. Antral follicles are in different phases of development (arrows)

and the proportion of atretic oocytes decreases if FSH is experimentally increased (Gougeon 1984). The non-selected oocytes undergo a process of atresia. The mechanism of selection of the dominant follicles involves intraovarian factors and endocrine signals. The dominant follicles have a higher expression of Luteinizing Hormone Receptor mRNA and of 3 b–­hydroxysteroid dehydrogenase (3b HSD) than the follicles heading for atresia (Webb et  al. 1999). During selection the ovulation bound oocytes change androgen production into estrogen production, expressing aromatase activity induced by FSH. (c) Dominancy. The EGF-like growth factors induced by LH causes the expansion of the cumulus oophorus and the maturation of the oocyte. The EGF-like growth factors are also paracrine signals mediating the action of LH during ovulation (Park et al. 2004). The differentiation of granulosa cells to cumulus cells (Fig. 1.10) involves the acquisition of additional regulatory mechanisms such as higher sensibility to cumulus expansion factors (CEEFs), and increased levels of transcripts related to cell proliferation, caused by the activation of MAPK3/1 and MAPK14. One of these transcripts is the product of the Tnfaip 6 gene. During the pre-ovulatory peak of gonadotrophins, the cells of the cumulus oophorus of the antral follicles begin to expand. This process is not present in the granulosa cells of the preantral follicles due to the lack of active CEEFs secreted by the oocytes, thus there is practically no activation of MAPKs and there is no Tnfaip6 mRNA (Park et al. 2004).

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Fig.  1.10  Antral follicles the oocyte is surrounded by cumulus cells (arrows). Haematoxylineosin technique. Scale bar 50 mm

Another characteristic of dominant follicles is the presence of at least two f­ actors, the insulin-like growth factor I (IGF-I) and the vascular endothelial growth factor (VEGF). Factor VEGF is a potent promoter of angiogenesis derived from thecal cells whose production is stimulated by LH (Garrido et al. 1993). Increased blood supply to the dominant follicle maximizes the influx of LH and FSH to the follicle. In addition to gonadotropins and the above mentioned intra-­ follicular factors, other factors may influence cell proliferation in the granulosa and the theca, as well as the differentiation and fate of an early antral follicle. Some of these factors are growth hormone (GH), IGF-I, insulin, metabolic factors and local factors such as fibroblast growth factors (FGFs) and BMPs (Knight and Glister 2003). At this stage the oocyte actively leads the somatic cell function, as in the antral stage, since the oocytes continue to form the zona pellucida proteins and ­connexin-37, which promotes the proliferation and differentiation of granulosa cells through the secretion of paracrine factors which may differentially affect cumulus cells and the mural granulosa cells. The dominant follicles exert the effect of dominance on subordinate follicles, limiting the development of gonadotropin-dependent follicles suppressing the FSH and inducing their atresia (Campbell et  al. 1995). It has been shown that in the antral phase the viability of antral granulosa cells becomes a factor in the fate of the follicle, as are those which provide support to germ cells (Morita and Tilly 1999; Tilly 2001). At this stage of development apoptosis in a large number of granulosa cells is indicative of a follicle in the process of elimination. The pre-ovulatory follicles are characterized by high dynamics of growth and they develop a great capacity to respond to stimulation with FSH, they secrete large amounts of estradiol and inhibin and they are potentially capable of being ovulated. All follicle cells and especially pre-ovulatory granulosa cells begin to assume their endocrine function and to complete a highly regulated endocrine process that leads to a state of proliferation and differentiation. This process is the primary ­control of two pituitary hormones: FSH and LH.

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Fig. 1.11  Corpus luteum. Arrows show secondary follicles. Haematoxylin-eosin technique. Scale bar 50 mm

During the process of maturation of ovulatory follicles, the following changes take place at the same time: increase in follicular size; rapid proliferation of the granulosa cells and morphological changes in cells caused by marked esteroidogenic activity. The aromatase activity increases progressively during the preovulatory phase. Just before ovulation, granulosa cells are fully differentiated, proliferation stops and high levels of steroids are produced. The ability of follicles to respond to gonadotropins increases progressively according to the development of preantral follicle stage to preovulatory stage. The basic function of an ovulatory follicle is to produce an oocyte the can be fertilized, and function as endocrine gland during maturation, and after ovulation through the transformation of the ovulated follicle in a functional corpus luteum (Fig. 1.11). Once the luteal phase starts, after ovulation, the granulosa cells increase in size and initiate an accumulation of a yellow pigment called lutein (hence the name corpus luteum). Corpus luteum has the ability to produce estrogen and progesterone. The regulation of the secretion of these compounds is defined by the estral or menstrual cycle. When fertilization does not occur, the corpus luteum begins to degenerate and the levels of estrogen and progesterone diminish, stimulating the growth of new follicles. However, when fertilized, the corpus luteum increases progesterone production to inhibit the growth of new follicles. The continuity of the corpus luteum is maintained by human chorionic gonadotropin (hCG) at the start of pregnancy, then the placenta takes the function of producing the ­necessary estrogen and progesterone.

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In the early stages of follicular growth there is no significant increase in the size of the oocyte. Evidence in bovine oocytes indicates that this may be due to the association of growth with the relocation of cytoplasmic organelles and to the development of specific structures of the oocyte, such as the zona pellucida and cortical granules (Fair et al. 1997a). Fair et al. (1997b) showed that the nucleolar functions are activated gradually and that the transcription in the oocyte begins when the follicles reach secondary stage. On the other hand, the granulosa cells are in active proliferation when the oocyte is at a relatively low transcriptional activity. In mice, it was also observed that the synthetic activity is low in oocytes in primordial follicles and increases in the oocytes in primary follicles (Moore et al. 1974).

1.2

Molecular Factors Regulate the Function of the Ovary

The process of follicular growth is controlled by extra-ovarian and intra-ovarian factors and the importance of each of these factors depends on the stage of follicle development. Extra-ovarian factors regulate growth of antral and preovulatory follicles, while intra-ovarian factors regulate growth of preantral and early antral ­follicles (Hirshfield 1991). The transforming growth factor beta family (TGFb) has been implicated in ­various aspects of follicular development (Chang et al. 2002). This family consists of more than 35 members in vertebrates. The receptor consists of a TGFb receptor type I and a type II. TGFb receptor is coupled to a protein complex on the surface of the cell membrane (Massague et  al. 1994; Brand and Schneider 1996). Both receptors contain an amino terminal signal sequence, an extracellular domain rich in cysteine ligand with sites of N-linked glycosylation, a single hydrophobic ­transmembrane domain and a cytoplasmic kinase domain (Massague 1992). The route of the TGFb signaling is mediated by Smad proteins through a ­cascade of phosphorylation induced by the ligand (reviewed in Zimmerman and Padgett 2000). The ligand-specific type II receptor phosphorylates the type I receptor, which subsequently activates the downstream cascade of Smad signaling ­proteins (Attisano and Wrana 1998). The role of each of the TGFb is regulated by the Smad, and the Downstream Smad signaling cascade is defined by the type of effect of the factor (Fig. 1.12). The TGFb exerts different stimuli on cells either in growth, differentiation, mobility, organization or cell death, depending on the environment and/or cell type (Massague 1992; Massague et al. 1994). In rat ovary the expression of TGFb receptor II mRNA has been found in cultured porcine cells and the expression of TGFb receptor-I and II mRNAs was also observed (Goddard et al. 1995). The ovary expresses several members of this family: in oocytes GDF-9, BMP-6 and BMP-15 occur; in granulosa cells: inhibins, activins, TGFb1, TGFb2 and TGFb3 are formed; and the thecal cells synthesize BMP-4 and BMP-7 (Chang et al. 2002; Drummond et al. 2003) (Fig. 1.13).

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Fig. 1.12  The TGFb signaling pathway

Fig. 1.13  TGFb members family expressed in the ovary. Specific factors are expressed in each cell type

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There are two growth factors produced by the oocyte that can impact initial f­ ollicle growth: the growth differentiation factor 9 (GDF-9) and bone morphogenetic protein 15 (BMP-15 or GDF-9B). Both factors are expressed exclusively in the oocyte (McGrath et al. 1995; Dube et al. 1998). GDF-9 is expressed in several species, including humans and mice, in all follicular stages except in primordial follicles of newborn and adult mice (McGrath et  al. 1995). GDF-9 acts as a ­paracrine regulator in the proliferation and differentiation of granulosa cells in primordial follicles. GDF-9 and BMP15 promote proliferation of granulosa cells of early antral follicles (Hayashi et al. 1999; Otsuka et al. 2000; Vitt et al. 2000), so the GDF-9 allows primary follicles pass to the following stages of development. BMP15 is considered a luteinization inhibitor as it inhibits the FSH production stimulated by progesterone in granulosa cells of the rat (reviewed in Shimasaki et al. 2004). The Smad3 receptor of TGFb is highly expressed in the ovarian surface ­epithelium, in granulosa cells and in oocytes of various animal models (Symonds et al. 2003; Xu et al. 2002; Tomic et al. 2002). In Smad3-deficient mice, the genes that control cell cycle progression are affected, causing failure in folliculogenesis (Tomic et al. 2004). The inhibins are protein hormones that are produced primarily in the gonads. They down regulate the synthesis and secretion of FSH from the pituitary. The inhibins are composed of an a subunit and one or two b subunits (bA or bB), the b-dimers with a-bA and inhibin bB form the A and B respectively. Besides sharing the b subunits, the activins and inhibins are functional antagonists in many physiological contexts (Wang et  al. 1996). These proteins, in addition to their role in modulating the release of FSH, may also serve as local regulators of folliculogenesis (Findlay 1993). Type I receptors for activins (ACVR1 and ACVR2B) are expressed in granulosa cells, meanwhile the ligands are expressed in the oocyte and in somatic cells. Once the primordial follicles begin folliculogenesis to form preantrales and antral follicles they begin to be sensitive to gonadotropins and the GCs surrounding the oocyte begin to synthesize predominantly B and also inhibin A (Jaatinen et al. 1994). In the late luteal phase and in early follicular phase of the menstrual cycle, FSH levels rise. Under the stimulation of FSH, inhibin levels in serum increase until a negative feedback occurs and FSH levels fall before ovulation. The dominant follicle which is now responsive to LH produces more inhibin A than B. After the LH peak, the corpus luteum produces inhibin bA. This shows that there is differential expression of inhibins during the menstrual cycle. The activins are homo or heterodimers of the subunits bA or bB. They are ­regulators of FSH and bind to the receptor type II. When activin binds to its receptor, signalling events leading to a specific biological response to activins are ­initiated. The activin response is blocked when the inhibins bind to the same receptor as the activin or to betaglycans, generating a nonfunctional receptor complex. The inhibins bind to inhibin binding protein (InhBP/p120), which is also known as the product of immunoglobulin superfamily gene 1 [IGSF1], which is expressed in the pituitary and testis. This binding activates a transduction pathway of inhibins,

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which causes a specific response for inhibin ligands (Massague and Chen 2000). In undifferentiated GCs, the activin increases response to FSH, but in differentiated cells it is an inhibitor (Miro et al. 1995). The inhibin regulates FSH at high concentrations, antagonizes the activin and is a potent stimulator of FSH secretion. In the ovary, FSH combined with activin causes a dose-dependent increase of DNA synthesis, suggesting that FSH in the presence of activin is mitogenic. It is believed that inhibin is a hormone that plays a key role in the regulation of FSH and it has been shown that it can have an ­important physiological role for embryonic development and survival in mink. The neutralization of inhibin increases the ovulation rate in many species, and probably increases the concentration of serum FSH; however, it has been shown that the inmunoneutralization of inhibin suppresses the embryonic development in mink (Ireland et al. 1992). It appears that the activin is an intrafollicular protein that controls the activation of primordial follicles and their effect depend on factors such as age and availability of follistatin. The follistatin is synthesized by granulosa cells and can bind to activin, contributing to a complex pattern of regulation of secretion of progesterone. Follistatin is a single chain polypeptide that was initially identified by its activity to suppress FSH (Robertson et al. 1987), but later it was discovered that it binds to activin and inhibin through the common b subunit and neutralizes the bioactivity of the activin (Shimonaka et  al. 1991). Activin regulates progesterone production stimulated by FSH in granulosa cells of the rat (Miro et al. 1995). The connective tissue growth factor (CTGF) is a member of CTGF / cysteinerich over-express 61/nephoblastoma gene family that mediates the regulation of connective tissue synthesis induced by TGFb in various cell types. An abundant expression of CTGF mRNA was also detected in granulosa and theca cells of the ovaries of pigs (Wandji et al. 2000). CTGF gene expression in granulosa cells is inversely related to the state of granulosa cell differentiation, being directly inhibited by FSH signaling pathway mediated by AMP. The abundance of CTGF mRNA in undifferentiated granulosa cells in vitro is regulated by TGFb1, GDF-9 and activin, which may indicate paracrine functions of these growth and differentiation factors in the regulation of CTGF synthesis in ovaries of mammals (Harlow et al. 2002). The Müllerian inhibitory substance (MIS) is another member of the TGFb ­family of glycoprotein hormones that presents certain patterns of expression in follicles at various stages of development. This hormone was identified as a testicular product that induces regression of Müllerian ducts in males (Cate et al. 1986). Also known as anti-Müllerian Hormone (MIS), it is expressed in granulosa cells of small and early growing follicles (Ueno et al. 1989). Granulosa cells of pre-antral follicles and small antral express MIS type II receptor (Teixeira et al. 1996). The theca cells also have a marked expression of MIS in small antral and preantral follicles, but unlike the granulosa cells, theca cells continue to express the receptor in antral follicles and in early atresia (Ingraham et  al. 2000). It was also noted that MIS inhibits the growth of follicles, as it is expressed in the granulosa cells of primary and early antral follicles (Durlinger et al. 1999). The transcription of MIS genes is

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r­ egulated by nuclear transcription steroidogenic factor-1 (SF-1), originally identified as a factor that regulates steroid hydroxylase genes (Ingraham et  al. 2000). SF-1 also regulates LHB (Keri and Nilson 1996) and FSH receptor (Levallet et al. 2001). The transcription of MIS and SF-1 is also regulated by members of the family of GATA transcription factors. GATA-4 and GATA-6 are expressed in granulosa cells of developing follicles (Tremblay and Viger 2001). The previous results highlight the broad impact of the members of the TGFb superfamily in various functions and stages of ovarian development.

1.3

Hormonal Aspects of the Ovarian Functions

The ovary is not only involved in sexual reproduction, but also has great influence on the entire hormonal functioning during development of the organism. The ovary is the site of the highest synthesis and secretion of progesterone and estrogen in mammals and gives rise to cyclical fluctuations in the levels of these hormones in the blood (Norman and Litwack 1987). The follicles are responsible for the secretion of both hormones and the release of eggs during the normal cycle. According to the theory ‘two cell, two gonadotropin’ (Armstrong et al. 1979), the theca interna cells are stimulated by LH to produce androgens which are transported to the granulosa cells (Fig. 1.14). There they are converted to estrogen by

Fig.  1.14  Two cell-two gonadtrophin theory. LH induces androstenedione synthesis in theca cells. The FSH stimulus provokes granulose cells process androstenedione into estrone which is further converted into estradiol via 17hsd enzyme

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aromatizing enzymes, which are induced by FSH. The theca interna appears to be the major site of synthesis of androstenedione, whereas testosterone is produced in smaller quantities (Bergh et al. 1993). During the preovulatory maturation stage, acquisition of LH responsiveness transforms the ability of granulosa cells to respond to gonadotropins, because the aromatase can be stimulated by hCG (Dennefors et  al. 1983). After mid-cycle gonadotropin peak granulosa cells change and, instead of mainly producing ­estrogen, they become a tissue producing progestin (Bomsel et al. 1979). The formation of primordial follicles and the onset of follicular growth may be independent of pituitary gonadotropins. However, small growing follicles and the  preovulatory follicles are FSH dependent. The granulosa cells proliferation and the spatial separation of the cells of the cumulus from mural granulosa cells are also FSH dependent. This separation may require the regulation of mural granulosa cells and factors that are secreted by the oocyte-cumulus complex. As mentioned, the follicle in the early stages of development is not vascularized; it has been ­proposed that gradients of factors seem to settle in microenvironments within the follicle (reviewed in Richards 2001). If the follicle reaches the appropriate level of maturation when FSH is at high levels in the bloodstream, the follicle may continue and complete its growth. If the FSH is at basal levels, follicles cannot continue their growth. An unsuitable concentration of gonadotropins causes inhibition of follicle progress and no ovulation takes place. In this condition the number of follicles is reduced, though follicular growth is not completely eliminated, as some follicles may grow and mature to the selection phase, since the growth of follicles from the preantral stage to selectable follicle require low levels of gonadotropin. Before ovulation, granulosa cells mature to form the corpus luteum, which is responsible for the secretion of progesterone and estrogen at the end of the cycle. The release of progesterone from the corpus luteum is influenced mainly by LH, whose activity is mediated by intracellular cyclic adenosine 3’, 5’-monophosphate (cAMP) (Jordan et  al. 1978). FSH, prolactin, prostaglandins, and b-adrenergic agents also have a role in the control of progesterone secretion (Norman and Litwack 1987). The activin stimulated by FSH inhibits progesterone secretion by granulosa cells and follistatin synthesized by granulosa cells can bind to activin, contributing to a complex pattern of progesterone secretion regulation. The effects of progesterone are mediated by their nuclear receptors. The ­progesterone receptor (PR) is a member of a large family of ligand-activated nuclear transcription regulators, which includes receptors for steroids, retinoids, thyroid hormones, and vitamin D. PR receptor is induced in granulosa cells of preovulatory follicles after the LH surge (Natraj and Richards 1993). There are two isoforms of PR, PR-A and PR-B, which are differentially expressed when required in various target tissues. Uterus cells of mammals, luteinizing granulosa cells and the cells of the corpus luteum express the PR. It was observed that in PR null mice there is no ovulation, although mature ­preovulatory follicles occur, therefore PR is required for ovulation (Lydon et al. 1995; Conneely et al. 2003). PR was detected in the primate corpus luteum, despite the high local concentrations of progesterone (reviewed in Graham and Clarke 1997).

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Circulating levels of FSH are determined by a feedback loop between the ovary and pituitary. Maturing follicles send a negative signal to the brain, suppressing the secretion of FSH. When there are no mature follicles, FSH levels increase. In this way there is a strict control of the number of follicles that will grow and mature to ovulation. The system is so regulated that it can quickly offset the loss of follicles that have reached advanced stage of maturation. This is because there is always a large number of young follicles that accelerates the ripening process (Hirshfield 1991). LH and FSH synergistically promote follicular development, as has been ­mentioned, and this is required for the production and secretion of follicular ­hormones, for ovulation and the hormone secretion of the corpus luteum. During infancy, the amount of LH is low and increases during puberty when the follicles have reached the antral stage. The presence of estrogen receptors (ER) has been detected in the ovary, mainly in granulosa cells, when follicle growth has advanced to the luteal phase. Several studies have been done to relate the modulation of rFSH and rLH (reviewed in Richards 1980). The ovary produces three estrogens: estradiol, estrone and estriol. During the process of folliculogenesis and follicular development, oocytes remain arrested in the diplotene stage of meiotic prophase I. In vivo, the resumption of meiosis occurs when the preovulatory peak of LH rises and takes place only in those oocytes competent of meiotic dominant follicles. Shortly before the LH surge, junctions that connect the cells of the cumulus to the oocyte break. During the period between the LH surge and ovulation, the oocyte undergoes a series of changes not only in its nucleus but also in its cytoplasm, a process known as oocyte maturation. Nuclear maturation involves several steps, including two consecutive M-phases in the absence of DNA replication (S phase). The oocytes stop their ­division in the MII stage until fertilization, when activation of the stimuli generated by sperm penetration triggers the completion of the meiotic cycle and initiates embryonic development. The maturation of the cytoplasm is required to obtain the condition for blocking polyspermia if fertilization takes place, to decondense the nucleus of the spermatozoon, to form a pro-nucleus after fertilization, and to ­redistribute cytoplasmic organelles. The whole system of growth, maturation and follicular elimination is finely regulated by various morphological and molecular events from the primordial stage until ovulation and corpus luteum formation. However this system may have some flaws that produce various types of functional anomalies in the number of follicles that pass through each stage of development. These changes can produce alterations in the reproduction.

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Armstrong DT, Goff AK, Dorrington JH (1979) Regulation of follicular estrogen biosynthesis. In: Midgley AR Jr, Sadler WA Jr (eds) Ovarian Follicular Development and Function. Raven, New York Attisano L, Wrana JL (1998) Mads and Smads in TGF beta signalling. Curr Opin Cell Biol 10:188–194 Bassett DL (1943) The changes in the vascular pattern of the ovary of the albino rat during the estrous cycle. Am J Anat 73:252–292 Bergh C, Olsson JH, Selleskog U, Hillens ST (1993) Steroid production in cultured thecal cell obtained from human ovarian follicles. Hum Reprod 8:519–524 Bomsel H, Gougeon A, Thebault A et al (1979) Healthy and atretic human follicles in the preovulatory phase: differences in evolution of follicular morphology and steroid content of the follicular fluid. J Clin Endocrinol Metab 48:686–694 Brand T, Schneider MD (1996) Transforming growth factor-b signal transduction. Circ Res 78:173–179 Campbell BK, Scaramuzzi RJ, Webb R (1995) Control of follicle development and selection in sheep and cattle. J Reprod Fertil Suppl 49:335–350 Cate RL, Mattaliano RJ, Hession C et al (1986) Isolation of the bovine and human genes for Müllerian inhibiting substance and expression of the human gene in animal cells. Cell 45:685–698 Chang H, Brown CW, Matzuk MM (2002) Genetic analysis of the mammalian transforming growth factor-b superfamily. Endocr Rev 23:787–823 Conneely OM, Mulac-Jericevic B, Lydon JP (2003) Progesterone-dependent regulation of female reproductive activity by two distinct progesterone receptor isoforms. Steroids 68: 771–778 Dennefors BL, Hamberger L, Nilsson L (1983) Influence of human chorionic gonadotropin in vivo on steroid formation and gonadotropin responsiveness of isolated human preovulatory follicular cells. Fertil Sterll 39:56–61 Driancourt MA, Reynaud K, Cortvrindt R, Smitz J (2000) Roles of Kit and Kit Ligand in ovarian function. Rev Reprod 5:143–152 Drummond AE, Dyson M, Le Tan M et al (2003) Ovarian follicle populations of the rat express TGF-ß signaling pathways. Mol Cell Endocrinol 202:53–57 Dube JL, Wang P, Elvin J et al (1998) The bone morphogenic protein 15 gene is X-linked and expressed in oocytes. Mol Endocrinol 12:1809–1917 Durlinger A, Kramer P, Karels B et al (1999) Control of primordial follicle recruitment by antiMüllerian hormone in the mouse ovary. Endocrinol 140:5789–5796 Fair T, Hulshof SCJ, Hyttel P et  al (1997a) Oocyte ultrastructure in bovine primordial to early tertiary follicles. Anat Embryol 195:327–336 Fair T, Hulshof SCJ, Hyttel P et al (1997b) Nucleus ultrastructure and transcriptional activity of bovine oocytes in preantral and early antral follicles. Mol Reprod Dev 46:208–215 Findlay JK (1993) An updata on the roles of inhibin, activin, and follistatin as local regulators of folliculogenesis. Boi Reprod 48:15–23 Franchi LL (1960) Electron microscopy of oocyte-follicle relationships in the rat ovary. J Biophys Biochem Cytol 7:397–398 Garrido C, Saule S, Gospodarowicz D (1993) Transcriptional regulation of vascular endothelial growth factor gene expression in bovine granulosa cells. Growth Factors 8:109–117 Goddard I, Hendrick JC, Bnahmed M, Morera AM (1995) Transforming growth factor b receptor expression in cultured porcine granulosa cells. Mol Cell Endocrinol 115:207–213 Gougeon A (1984) Influence of cyclic variations in gonadotrophin and steroid hormones on ­follicular growth in the human ovary. In: de Brux I. Gautrav JP (eds) Clinical Pathology of the Endocrine Ovary. MTP Press, Lancaster Graham JD, Clarke CL (1997) Physiological action of progesterone in target tissues. Endocr Rev 18(4):502–19 Harlow ChR, Davidson L, Burns KH et al (2002) FSH and TGF-b supefamily members regulate granulosa cell connective tissue growth factor gene expression in  vitro and in  vivo. Endocrinology 143(9):3316–3325

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Hayashi M, McGee EA, Min G et al (1999) Recombinant growth differentiation factor-9 (GDF9) enhances growth and differentiation of cultured early ovarian follicles. Endocrinology 140: 1236–1244 Hirshfield AN (1991) Development of follicles in the mammalian ovary. Int Rev Cytol 124:43–101 Huang EJ, Manova K, Packer AI et al (1993) The murine Steel Panda mutation affects kit ligand expression and growth of early ovarian follicles. Develop Biol 157:100–109 Ingraham HA, Hirokawa Y, Roberts LM et al (2000) Autocrine and paracrine hormone signaling in reproduction. Recent Prog Horm Res 55:30–38 Ireland JJ, Martin TL, Ireland JLH, Aulerich RJ (1992) Immunoneutralization of inhibin suppress reproduction in female mink. Biol Reprod 47:746–750 Jaatinen TA, Penttila TL, Kaipia A et  al (1994) Expression of inhibin a, bA y bB Messenger ribonucleic acids in the normal ovary and in polycystic ovarian syndrome. J Endocrin 143: 127–137 Jordan AW III, Caffrey JL, Niswender GD (1978) Catecholamine induced stimulation of progesterone and adenosine 39, 59-monophosphate production by dispersed ovine luteal cells. Endocrinology 103:385–392 Katz D, Niederberger C, Slaughter GR, Cooney AJ (1997) Characterization of germ cell-specific expression of the orphan nuclear receptor, germ cell nuclear factor. Endocrinology 138:4364–4372 Keri RA, Nilson JH (1996) A steroidogenic factor-1 binding site is required for activity of the luteinizing hormone b subunit promoter in gonadotropes of transgenic mice. J Biol Chem 271:10782–10785 Klinger FG, De Felici M (2002) In vitro development of growing oocytes from fetal mouse oocytes: stage-specific regulation by stem cell factor and granulosa cells. Develop Biol 244:85–95 Knight PG, Glister C (2003) Local roles of TGFbeta superfamily members in the control of ovarian follicle development. Anim Reprod Sci 78:15–183 Levallet J, Koskimies P, Rahman N, Huhtaniemi I (2001) The promoter of murine follicle-stimulating hormone receptor: functional characterization and regulation by transcription factor steroidogenic factor 1. Mol Endocrinol 15:80–92 Lydon JP, DeMayo FJ, Funk CR et al (1995) Mice lacking progesterone receptor exhibit pleiotropic reproductive abnormalities. Genes Dev 9:2266–2278 Magoffin DA (2005) Ovarian theca cell. Int J Biochem Cel Biol 37:1344–1349 Manova K, Nocka K, Besmer P, Bachvarova RF (1990) Gonadal expression of c-kit encoded at the W locus of the mouse. Development 110:1057–1069 Massague J (1992) Receptors for the TGF-b family. Cell 69:1067–1070 Massague J, Chen YG (2000) Controling TGF-b signaling. Genes Dev 14:627–644 Massague J, Attisano L, Wrana JL (1994) The TGF-b family and its composite receptors. Trends Cell Biol 4:172–178 McGee EA, Hsueh AJW (2000) Initial and cyclic recruitment of ovarian follicles. Endocr Rev 21(2):200–214 McGrath SA, Esquela AF, Lee SJ (1995) Oocyte specific expression of growth/differentiation factor-9. Mol Endocrinol 9:131–136 Miro F, Smyth CD, Whitelaw PF et al (1995) Regulation of 3b-hydroxysteroid dehydrogenase5/4isomerase and cholesterol side-chain cleavage cytochrome P450 by activin in rat granulosa cells. Endocrinology 136:3247–3252 Montro B, Bernstein A (1993) Dynamic changes in ovarian c-kit and Steel expression during the estrous reproductive cycle. Dev Dyn 197:69–79 Moore GPM, Lintern-Moore S, Peters H, Faber M (1974) RNA synthesis in the mouse oocyte. J Cell Biol 60:416–422 Morita Y, Tilly JL (1999) Oocyte apoptosis: like sand through an hourglass. Dev Biol 213:1–17 Motta PM, Makabe S, Naguro T, Correr S (1994) Oocyte follicle cells association during development of human ovarian follicle. A study by high resolution scanning and transmission electron microscopy. Arch Histol Cytol 57:369–394

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Natraj U, Richards JS (1993) Hormonal regulation, localization, and functional activity of the progesterone receptor in granulosa cells of rat preovulatory follicles. Endocrinology 133: 761–769 Nilsson EE, Kezele P, Skinner MK (2002) Leukemia inhibitory factor (LIF) promotes the primordial to primary follicle transition in rat ovaries. Mol Cell Endocrinol 188:65–73 Norman A, Litwack G (1987) Hormones. Academy Press, Inc., London Otsuka F, Yao Z, Lee T et al (2000) Bone morphogenetic protein-15. Identification of target cells and biological functions. J Biol Chem 275:39523–39528 Park JY, Su YQ, Ariga M et al (2004) EGF-like growth factors as mediators of LH action in the ovulatory follicle. Science 303(5658):682–684 Picton H, Briggs D, Gosden R (1998) The molecular basis of oocyte growth and development. Mol Cell Endocrinol 145:27–37 Rajkovic A, Pangas SA, Ballow D et al (2004) Nobox deficiency disrupts early folliculogenesis and oocyte-specific gene expression. Science 305:1157–1159 Rankin T, Familari M, Lee E et al (1996) Mice homozygous for an insertional mutation in the Zp3 gene lack a zona pellucida and are infertile. Development 122:2903–2910 Richards JS (1980a) Maturation of ovarian follicles: actions and interactions of pituitary and ­ ovarian hormones on follicular cell differentiation. Physiol Rev 60(1):51–89 Richards JS (2001) Perspective: the ovarian follicle: a perspective in 2001. Endocrinology 142(6):2184–2193 Robertson DM, Klein R, deVos FL et al (1987) The isolation of polypeptides with FSH suppressing activity from bovine follicular fluid which are structurally different to inhibin. Biochem Biophys Res Commun 149:744–749 Rugglu M, Speed R, Taggart M et al (1997) The mouse Dazla gene encodes a cytoplasmic protein essential for gametogenesis. Nature 389:73–77 Shimasaki S, Moore RK, Otsuka F, Erickson GF (2004) The bone morphogenetic protein system in mammalian reproduction. Endocr Rev 25:72–101 Shimonaka M, Inouye S, Shimasaki S, Ling N (1991) Follistatin binds to both activin and inhibin through the common subunit. Endocrinology 128(6):3313–15 Sotelo JR, Porter K (1959) An electron microscope study of rat ovum. J Biophys Biochem Cytol 5:327–342 Soyal SM, Amleh A, Dean J (2000) Fig a a germ cell-specific transcription factor required for ovarian follicle development. Development 127:4645–4655 Symonds D, Tomic D, Borgeest C et al (2003) Smad3 regulates proliferation of the mouse ovarian surface epithelium. Anat Rec 273A:681–686 Teixeira J, He WW, Shah PC et  al (1996) Developmental expression of a candidate Müllerian inhibiting substance type II receptor. Endocrinology 137:160–165 Tilly JL (2001) Commuting the death sentence: how oocytes strive to survive. Nat Rev Mol Cell Biol 2:838–848 Tisdall DJ, Fidler AE, Smith P et al (1999) Stem cell factor and c-kit gene expression and protein localization in the sheep ovary during fetal development. J Reprod Fertil 116:277–291 Tomic D, Brodie SG, Deng C et al (2002a) Smad3 may regulate follicular growth in the mouse ovary. Biol Reprod 66:917–923 Tomic D, Miller KP, Kenny HA et al (2004a) Ovarian follicle development requires Smad3. Mol Endocrinol 18(9):2224–40 Tremblay JJ, Viger RS (2001) GATA factors differentially activate multiple gonadal promoters through conserved GATA regulatory elements. Endocrinology 142:977–986 Ueno S, Kuroda T, Maclaughlin DT et al (1989) Müllerian inhibiting substance in the adult rat ovary during various stages of the estrous cycle. Endocrinology 125:1060–1066 van den Hurk R, Bevers MM, Dieleman SJ (1999) Folliculogenesis and oocyte development in mammals (livestock animals). In: Joy KP, Krishna A, Haldar C (eds) Comparative endocrinology and reproduction. Narosa Publishing House, New Delhi van den Hurk R, Abir R, Telfer EE, Bevers MM (2000) Primate and bovine immature oocytes and follicles as sources of fertilizable oocytes. Hum Reprod 6:457–474

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Vitt UA, Hayashi M, Klein C, Hsueh AJ (2000) Growth differentiation factor-9 stimulates proliferation but suppresses the follicle stimulating hormone-induced differentiation of cultured granulosa cells from small antral and preovulatory rat follicles. Biol Reprod 62:370–377 Wandji SA, Gadsby JE, Barber JA, Hammond JM (2000) Messenger ribonucleic acids for MAC25 and connective tissue growth factor (CTGF) are inversely regulated during folliculogenesis and early luteogenesis. Endocrinology 141:2648–2657 Wang QF, Tilly KL, Tilly JL et  al (1996) Activin inhibits basal an androgen-stimulated ­proliferation and induces apoptosis in the human prostatic cancer cell line, LNCaP. Endocrinology 137:5476–5483 Webb R, Campbell BK, Garverick HA et al (1999) Molecular mechanisms regulating follicular recruitment and selection. J Reprod Fertil Suppl 54:33–48 Xu J, Oakley J, McGee EA (2002) Stage-specific expression of Smad2 and Smad3 during folliculogenesis. Biol Reprod 66:1571–1578 Zimmerman CM, Padgett RW (2000) Transforming growth factor-b signalling mediators and modulators. Gene 249:17–30

Chapter 2

Embryonic Development of the Ovary, Sexual Reproduction and Meiosis Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría

Abstract  Sex first evolved in the common ancestor of all eukaryotes some two billons years ago. Fertilization is the union of two haploid gametes of different sex; the resulting diploid cell is the zygote. In mammalian species gametes are derived from precursors termed primordial germ cells (PGCs). Specification of the germline occurs through: (1) repression of somatic differentiation; (2) reacquisition of ­potential pluripotency; (3) genome wide epigenetic reprogramming. In several mammals the maintenance of germ cell linage is not due to a preformed “germ plasma”, but is induced by environmental signals. In mammals PGCs originate in extraembryonic region and eventually move to gonadal ridges, located in the medial part of the urogenital ridge. Observations and experimental evidence demonstrate that PGCs move actively from the hindgut to the gonadal ridges. However, some observations suggest that most displacements of these cells may be explained by global growth movements. In most mammals sex determination is genetically controlled by the presence or absence of Y chromosome and the expression of gene Sry. However, the ovary differentiation requires in addition the action of a precise gene cascade. The main process that marks PCGs/oogonia sex differentiation within fetal ovary is the beginning of meiosis. During the progression of the first meiotic prophase many germ cells are eliminated, around 70% in mice, rats and humans. Prenatal development of the follicles is regulated by a complex interplay of hormones produced by the hypophysis (FSH, LH), and paracrine regulatory factors produced by the oocytes and the surrounding somatic cells. Epigenetic signals are covalent modifications of histones or DNA that convey a specific “meaning” to the stretch of chromatin on which they are found. Frequently they induce changes in chromatin conformation and gene expression. Pluripotent embryonic germ cells are capable of giving rise to all tissues, thus they must be able to reorganize their mechanisms of gene regulation to allow multiple lines of cell differentiation. Besides epigenetic modifications regulate X inactivation/reactivation in the female

G.H. Vázquez-Nin (*), M.L. Escobar, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico City, Mexico e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_2, © Springer Science+Business Media B.V. 2011

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embryo and mark the paternal and maternal genome of germ cells, making the expression of a group of genes monoallelic, crucial for embryo development termed for modification such as “imprinted” genes.

2.1 Evolutionary Advantages of Sexual Reproduction In all mammalian species, gametes are derived from precursors termed the ­primordial germ cells (PGCs) which migrate into and colonize the gonadal ridges during embryonic development. PGCs and the cells originated from them, the oogonia in the female and the spermatogonia in the male, are diploid cells, as any somatic cell of the body of eukaryotes. This means that they have a pair of each chromosome formed by one chromatide (monochromatide chromosome) during interphase and two chromatides (bichromatide chromosome) when the cell is ready for division. The mature gametes (secondary oocytes or eggs and spermatozoa) are instead haploid cells since they have only one monochromatide chromosome of each pair. Meiosis is the germ cell exclusive cell division producing haploid cells. Meiosis involves two cell divisions with only one DNA duplication and results in four ­haploid spermatids in the male or one mature egg or ovum and two small cells called polar bodies in the female. Fertilization is the union of two haploid gametes of different sex resulting in a diploid cell termed zygote. Meiosis and fertilization are the two complementary processes of the sexual reproduction cycle, the reproductive process most favored by the evolution of eukaryotes. Sex first evolved in the common ancestor of all eukaryotes some two billion years ago (Zimmer 2008). To explain the rarity of asexual eukaryotes, even if they can rapidly spread a beneficial mutation in a population, they can pass mutation only to their direct offspring. Thus the beneficial mutation will be always inserted in the same genome. If another organism undergoes a different beneficial mutation in a different gene, both mutations cannot be combined in the one genome. Sexual reproduction recombines genes, joining beneficial mutation and possibly improving a population faster than asexual reproduction. Some examples testify the superiority of sexual reproduction over asexual. Genetically engineered yeast that could only reproduce sexually and others that could only reproduce asexually were raised on a near-starvation diet. Sexual yeast was able to adapt faster and as they evolved, their growth rate increased 94%, while asexual strain increased only 80%. This difference in growth allowed the sexual yeast to take over the population (Zimmer 2008). Asexual organisms may pick up slightly deleterious mutations that natural selection fails to eliminate. In this way slight deleterious mutations may replace undamaged genes in a population and permanently compromise fitness. Sexual organisms due to meiotic recombination may interchange a defective gene for a working one and keep healthy genomes (Zimmer 2008). A long term study (30 years) analyzed the relationship between two snail populations, Potamopyrgus antipodarum, one asexual

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and other sexual, with a parasitic trematode capable of sterilizing them. At the ­beginning the most common strains of asexual snails were resistant to the trematode. Over the years, however, the snails became progressively vulnerable to an adapting strain of trematode. Ten years later, the initial strain of asexual snails had almost disappeared and the population of sexual snails remained relatively stable (Zimmer 2008). In a study of sexual and asexual lineages of the microcrustacean Daphnia pulex it was found that deleterious amino acid substitutions in mitochondrial ­proteins are four time faster in asexual lineages than in sexual ones. These results demonstrate that sexual reproduction reduces the deleterious mutational burden in the mitochondrial genome as well (Paland and Lynch 2006).

2.2 Strategies for PGC Formation PGCs form outside the gonads during early stages of embryonic development and enter or are subsequently enclosed into the developing gonads (gonadal ridges) where they differentiate into primary oocytes in the female (Fig.  2.1) and ­prospermatogonia or gonocytes in the male. The specification of PGC precursors takes place very early in development in Chordata, Nematoda and several other Phyla. Specification and determination of the germline occur through the integration of three key events: repression of the somatic

Fig.  2.1  Primordial germ cells reach the gonadal ridges to colonize the gonad. A close-up is shown upper the panel

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differentiation program, reacquisition of potential differentiation pluripotency and ensuing genome-wide epigenetic reprogramming (Yamaji et al. 2008). Early studies in the mouse embryo showed that germ cell lineage is induced in cells of the proximal epiblast by the extra-embryonic ectoderm (reviewed in Buehr 1997). Thanks to very elegant, recent studies, crucial events of such a fascinating process have been revealed. The bone morphogenetic proteins (BMPs) have been shown to be central in PGC formation in the mouse. These proteins are members of the transforming growth factor b (TGFb) superfamily that includes TGFßs and BMPs, GDFs (Growth Differentiation Factors), activins, inhibins, MIS (Müllerian Inhibiting Substance), Nodal and Lleftys (Chang et al. 2002). The first indications of the crucial role of BMPs in PGC formation came from the observation that in Bmp4 or in Bmp8b knockout mice, PGCs were reduced or absent. Mice double heterozygous for Bmp4 and Bmp2, a close relative of BMP-4 also have a severe reduction of PGCs (Ying et  al. 2001; Chang et  al. 2002). Further investigations demonstrated that the induction of mouse PGCs actually depends on the synergistic action of BMP4, BMP8b and BMP2 produced by extraembryonic tissues surrounding the epiblast (Fig. 2.2). Transplantation of epiblast cells at different sites, gave rise to PGCs only if they were positioned in close proximity to the extraembryonic ectoderm, indicating that factors produced in this tissue are required for the generation of the PGC precursors (Ying et  al. 2001). These factors were identified as BMP4 and BMP8b, while BMP2 was found to be secreted by the extraembryonic visceral endoderm (Ying and Zhao 2001; see also the reviews by Chang et al. 2002; De Felici et al. 2004; Van den Hurk and Zhao 2005). Although similar PGC specification events are likely to occur within the epiblast of the human embryo around the end of the second week of development before gastrulation, the different organization of the mouse and human embryo at this time

Fig.  2.2  Schematic representation of the hypothetic synergistic action of BMP4, BMP8b and BMP2 produced by extraembryonic tissues surrounding the epiblast to induce PGC precursors in the human embryo around the beginning of the gatrulation period

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makes difficult to identify the mouse equivalent extraembryonic tissues responsible of the inductive germline specification action in humans. We can speculate that the mouse extraembryonic ectoderm and visceral endoderm correspond to the roof of the proamniotic cavity and hypoblast (primitive endoderm) of the human embryo, respectively. As in the mouse, it is likely that from the epiblast, human PGC precursors around the beginning of the third week move together with the extraembryonic mesoderm cells through the forming primitive streak (Fig. 2.2) toward a region of the wall of the yolk sac adjacent to allantois where they are determined and have been identified around 20th day in very old studies (reviewed in Baker and Eastwood 1983). Since activation of intracellular signaling by BMPs occurs through SMAD ­proteins, it was not surprising to find that Smad5 and Smad1 knockout mice have defects in PGC development resembling that caused by the ablation of the BMP genes (Chang et al. 2002). These findings demonstrate that in mouse and other mammals, the maintenance of germ cell lineage is not due to the inheritance of a preformed “germ plasm”, but is induced by environmental signals acting on pluripotent somatic cells (Saitou et al. 2002). In other species, however, the germlineage is specified by a different strategy. For example, in the nematode Ascaris the germ cell linage is segregated during the first few cleavage divisions of the embryo in cells retaining their chromatin complement complete, while the somatic lineages suffered chromatin diminution for the loss of terminal regions of the chromosomes (see review by McLaren 2003). Similar mechanisms of intrinsic germline specification occur in Xenopus, Frog, Drosophila, Caenorhabditis elegans. In such species, a region of the cytoplasm of unfertilized egg near the vegetal pole contains contains aggregates of mitochondria, proteins and RNA. Cells that during the first mitotic division receive this special cytoplasm region, the “germ plasm” become PGCs. (see review by McLaren 2003).

2.3 Molecular Mechanisms of PGC Specification in Mouse To determine the molecular mechanisms of the PGC specification, Saitou and ­collaborators (2002) performed genetic screens between PGCs and their somatic neighbors that share a common ancestry. The expression of Fragilis, an interferoninducible transmembrane protein, marks the beginning of germ cell competence. Using single-cell gene expression profiles, these authors demonstrated that the cells, at 7.25 days post coitum (dpc), show the highest expression of Fragilis, subsequently expressed Stella. The function of Stella gene product is uncertain but it has domains characteristic of protein involved in RNA splicing. Actually, Stella represses homeobox genes in nascent germ cells, and as such maintains the pluripotency of PGCs during their migration to the gonadal ridge. The induction of Fragilis by extra-embryonic tissues is dependent on BMP4 (Saitou et  al. 2002). Wnt3 (a secreted signaling protein transmitted via beta-catenin), provides epiblast cells with the ability to respond to BMP signals (Ohinata et al. 2009). Going further

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back, at 6.25 dpc, about six cells within the Fragilis-positive cells, at the posterior side of the embryo, begin to show expression of Blimp1/Prdm1and shortly after of Prdm14. The restricted expression of these two transcription factors in the PGC precursors is likely due to the inhibition of BMP4-2 action in the anterior region by antagonists produced by the anterior visceral endoderm (AVE) (Ohinata et al. 2009). Experiments tracing genetic lineage demonstrate that all Blimp1 positive cells are the lineage restricted PGC precursors (Ohinata et  al. 2005; Vincent et  al. 2005). The complex Blimp1/Prmt5 arginine methyltransferase mediates a change in the methylation of arginine 3 on histone H2A and H4 tails between 8.5 and 10.5 dpc. This change, accompanied by the Prdm14 action, causes other epigenetic changes necessary for repression of somatic program and ensures that PGCs retain or reacquire a pluripotent character. The Blimp1-positive PGC precursors while proliferating and moving into the extraembryonic mesoderm (exM) re-express key pluripotency-associated genes such as Sox2, Nanog and Oct-4. This latter belongs to the class V of the POU domain transcription factor family and has been the first transcription factor demonstrated to be lineage restricted in the developing PGCs (for a review, see Pesce et  al. 1998). Oct-4, Sox2 and Nanog are transcription ­factors all essential for maintaining the pluripotent stem cell phenotype. Around 7. 25–7.5 dpc, the PGC precursors in the exM at the basis of allantois are determined in a cluster of about 40–50  PGCs by unknown factors, but likely still including BMP-4. The adhesion molecule, E-cadherin is involved in clustering such PGCs and appears to be crucial for these precursors to be definitively allocated to the germ cell lineage (Okamura et  al. 2003). Determined PGCs, besides the classical PGC markers tissue not specific alkaline phosphatase (Tnap) and stage-specificantigen-1 (Ssea-1), express Kit receptor and germline-specific genes such as Nanos3 and Dead endI. What activates germline genes in PGCs? Paradoxically, recent evidence from mouse and C. elegans suggests that the same factors responsible for transcriptional repression may also be involved in such activation. Single-cell transcriptome profiling has shown that Blimp1 is required, not only for virtually all transcriptional repression, but also for the activation of ~50% of the genes that are upregulated in mouse PGCs. Genes that require Blimp1 for activation include those that show the highest specificity for PGC expression (e.g. Nanos3) (Kurimoto et al. 2008). The conclusion is that Blimp1 activity is required, directly or indirectly, to initiate part of the germline-specific transcriptional program. Being a Blimp1-expressing PGC precursors is not sufficient, however, to undergo specification to PGCs. It is likely that a second signal is needed locally to trigger specification, with another Blimp1-independent pathway also functioning in parallel (Kurimoto et al. 2008). This second pathway appears to be controlled by Prdm14 (Yamaji et al. 2008). Prdm14 expression in PGCs initially occurs independently of Blimp1, but becomes dependent on Blimp1 by E7.5. In the absence of Prdm14, presumptive PGCs fail to downregulate GLP (G9a-related protein), the H3K9 methyltransferase, and ­maintain H3K9me2 and do not upregulate H3K27me3. Prdm14-deficient PGCs also do not activate the pluripotency-related gene Sox2. How Prdm14 and Blimp1 contribute to both the upregulation of certain genes and the downregulation of ­others is not known.

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2.4 PGC Migration Since the nineteenth century, it was established that in mammals PGCs originate in extraembryonic region, and eventually move to the gonadal ridges (Nussbaum 1880 quoted by Merchant and Alvarez 1986). During this pregonadic phase PGCs can be identified by morphological criteria and surface markers, such as TNAP and SSEA-1, and the expression of genes typical of pluripotent cells (e.g. Oct-4, Sox2, Nanog) (Oktem and Oktay 2008). From the site of their determination, the wall of the yolk sac adjacent to the allantois, PGCs reenter into the embryo proper following the morphogenic movements incorporating the proximal part of the yolk sac into the hind gut. It is likely that at this stage PGCs acquire a motility phenotype and consequently after invading the hind gut endoderm move throughout the dorsal mesentery towards the gonadal ridges formed in the celomic epithelium. Ultrastructural observations favored the idea of preexisting pathways from the hind gut basal lamina to the celomic epithelium. Such pathways are formed by elongated mesenchymal cells and oriented elements of the extracellular matrix as collagen fibrils and fribronectin (Merchant and Alvarez 1986). Studies in vivo and in vitro demonstrated that at this stages mouse and human PGCs are able to emit filopodia (Fig. 2.3) by which they attach to different structures and retract pulling

Fig. 2.3  Migrating primordial germ cells present filopodia. Migrating cells advance in gradients of increasing chemoattractant concentration leading to polarized cell protrusions originating the filopodia. The chemorepellent are avoided for the migrating cells

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the cell to the point of attachment (Albrecht-Buechler 1976; Gustafson and Wolpert 1963). Therefore, locomotion may be atypical ameboidal (Zamboni and Merchant 1973; Merchant and Alvarez 1986). Taking into account these data, Merchant and Alvarez (1986) proposed: (1) the presence of short pathways containing fibronectin between the hind-gut and the celomic epithelium; (2) that fibronectin is required for PGC locomotion and (3) that mammalian PGCs emit filopodia which establish contact with surrounding substrata. Contact guidance migration of PGC might be mediated by KIT receptor expressed on the surface of these cells and its ligand, Kit ligand (also called Stem cell factor), which is expressed in somatic cells along the migratory pathway (Fleischman 1993; van den Hurk and Zhao 2005). Kit ligand also stimulates PGC migration, survival and proliferation in vitro (Dolci et al. 1993). Molyneaux et al. (2001) studied systematically the displacement of the PGCs in the mouse embryo, concluding that during some stages of the migration between the hind gut and the gonadal ridges PGCs move actively and directional to their final destination. More recently, in a time-lapse comparative analysis of the PGC migration between Drosophila, Zebrafish and mouse, these authors defined six distinct stages of mouse PGC behavior: (1) invasion of the endoderm; (2) ­passive or active migration into hindgut; (3) random migration within the hindgut; (4) migration from the gut to the gonadal ridges; (5) clustering at the ridges; (6) cell death within midline structures (Molyneaux and Wylie 2004). At stage 3 the PGCs are located in the hindgut epithelium moving actively around the epithelial cells. The epithelial cells express the Ca++ dependent adhesion molecule E-cadherin; whereas PGC do not. The lack of strong adhesive interactions allow PGCs to move freely within the gut. PGCs upregulate E-cadherin expression upon leaving the gut (Molyneaux and Wylie 2004; Di Carlo and De Felici 2000). Then PGCs exit from the gut, divide in two streams and migrate to the developing gonadal ridges. In organ culture of transverse slices of mice embryos, the addition of BMP4 elevates the PGC number. The addition of Noggin, a BMP-antagonist, reduced the PGC number and slowed and randomize their movements, resulting in failure of PGCs to colonize gonadal ridges (Dudley et al. 2007). The action of BMPs is mediated by the BMP-specific Smads (Smad1, Smad5 and Smad8). Immunolocalization of Smads1/5/8 demonstrates that migratory PGCs do not express detectable levels of these proteins. Instead the somatic cells of the gonadal ridges exhibit an intense Smads1/5/8 immunolabeling, thus suggesting that BMPs act on these cells (Dudley et al. 2007). Several growth factors contribute to enlarge the PGCs population. The migratory PGCs of mouse embryos express subunits of interleukin-2 receptor. The addition of Interleukine-2 to PGS in culture increased their number by a mitogenic effect. These data are indicative of a paracrine effect of Interleukine-2 on PGC proliferation (Eguizabal et al. 2007). When mouse PGCs were co-cultured with gonadal ridges they increased in number and moved towards the gonadal ridges. These effects were present at long distances and were absent when PGCs were co-cultured with other embryonic organs. Gonadal ridge conditioned medium attracts PGCs and TGFß may mimic these effects (Godin et al. 1990;Godin and Wylie 1991). These results strongly suggest that the cells of gonadal ridges secrete chemo-attractants (van den Hurk and Zhao 2005). More recently, some evidence that KL itself exerts chemiotactic effect on mouse PGCs in vitro was reported (Farini et al. 2007).

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In spite of the numerous observations and experimental evidence demonstrating that PGCs move actively, some observations in mouse and human embryos suggest that most of the displacements of these cells can be explained by the global growth movements of the embryo and that possibly no active migration exists at all (Freeman 2003). The same author concludes that the displacement the PGCs in vivo are unrelated to the “artefactual” movements of explanted germ cells.

2.5 Formation of the Ovary The gonads develop in the medial part of the urogenital ridge. The urogenital ridges are longitudinal thickenings located at both sides of the dorsal mesentery. The gonadal ridge is unique among all organ primordia because of its bipotential nature. A testis or an ovary can develop from a gonadal primordium. This first appears as thickenings of the celomic epithelium. As the epithelium invades the connective tissue the genital ridges protrude to the celomic cavity forming the gonadal ridge. This proliferation of the epithelium results in an internal epithelial mass and the stratified superficial epithelium of the gonadal ridge. In most mammals sex determination is genetically controlled by the presence or absence of a Y chromosome. The expression of the Y linked gene Sry in mice starts the rapid differentiation of testis specific cell types. In the absence of Sry even an XY mouse develops ovaries (Brennan and Capel 2004). The formation of other sexual organs and tissues is largely a consequence of the sex differentiation of the gonads and their hormone production. On the other hand, the targets of the molecular mechanisms of sex differentiation are the somatic cells of the gonads and not the germ cells. PGCs are actually sexually bipotent and differentiate in oogonia/oocytes or prospermatogonia under the influence of the somatic environment of the ovary and testis, respectively, and independently by their sex chromosomes (Fig. 2.4). Recent results indicate, however, that ovary differentiation does not simply occur for default in the absence of Sry but requires the action of a precise gene cascade. Actually, the “canonical” ß-catenin signaling pathway represents a central piece of the mechanisms controlling gonad differentiation into ovary. This signaling is initiated by WNT ligand binding to Frizzled and LRP5/6 co-receptors, ­leading to the inhibition of the ß-catenin destruction complex, containing glycogen synthase kinase-3-beta (GSK-3-beta), the adenomatous polyposis coli protein (APC), and the scaffolding protein axin, among others. This complex phosphorylates ß-catenin and targets it for degradation by the proteasome. Its inhibition leads to ß-catenin stabilization and entry to the nucleus. Nuclear ß-catenin binding to T-cell factor (TCF)/ lymphoid enhancer factors (LEF) leads to the activation of target gene expression. In the absence of ß-catenin, TCF/LEF proteins act as a transcriptional repressor. This ability to switch between repressor and activator states, probably tighten control over gene expression (see review by Tevosian and Manuylov 2008). Several Wnt family members are expressed in the developing

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Fig.  2.4  Schematic diagram showing that the targets of the molecular mechanisms of sex ­differentiation are the somatic cells of the gonads and not the germ cells

mammalian gonads. Initially there is a low level of expression of Wnt4 in both XX and XY gonadal primordial. This expression is down regulated in differentiating male gonad while it is increased the female. In mice, Wnt4-null females are ­masculinized as by the absence of Müllerian ducts, a dramatic decrease in the number of developing oocytes, production of ectopic steroids as testosterone and formation of male specific blood vessels. Markers of early ovarian development as Follistatin and Bmp2 are not expressed in the absence of Wnt4. The effects of the increase of Wnt4 dosage during embryogenesis in mice is relatively mild, mostly confined to the abnormal development of celomic blood vessels in the male. In summary, the absence of Wnt4 expression is insufficient by itself to program the testis differentiation of somatic cells. However, Wnt4 controls several elements of sexual development (see review by Tevosian and Manuylov 2008). Rspo is another gene family with a role in female sexual development which also targets the ß-catenin transduction pathway. A recessive mutation of the gene encoding RSPO1 results in a complete female to male sex reversal in humans. The Rspo1 mutation (XX Rspo1-/-) in mice results in a phenotypical female. However, the gonads undergo incomplete sex reversal and the masculinization occurs postnatally. The conclusion seems to be that Rsp1 plays a positive role in mammalian ovary development, but the genetic mechanism of the late sex reversal remains to be determined. It is well known that activating Sry or Sox9 in the female will override the ovarian pathway and activate the male program in a XX gonad. A human XY patient carrying a chromosome duplication which included the WNT4 and RSPO1 genes exhibited sex reversion. This suggested that the up-regulation of both of these

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ligands in mice may be enough to antagonize male development. Because both WNT4 and RSPO1 signaling converge in activating canonical ß-catenin pathway, the researchers produced an undegradable form of ß-catenin in somatic cells of the developing ovary. This was sufficient to block the male pathway in the XY gonads. The stabilization of ß-catenin blocked male-specific Sox9 and Amh expression, while genes normally found in the ovarian somatic cells as Foxl2, Bmp2, Wnt4 and Fst were increased. Testis cords did not form in these XY mutants (see review by Tevosian and Manuylov 2008). In summary we can conclude that sexual differentiation of the gonads in ­mammals is driven by two opposing antagonistic activities of the transcriptional regulatory proteins: SOX9 in males and ß-catenin and T-cell factor (TCF)/lymphoid enhancer factor (LEF) in females (see review by Tevosian and Manuylov 2008).

2.6 Germ Cell Sex For the colonization and development of the PGCs into the gonadal ridges the expression of a large number of genes is needed (van den Hurk and Zhao 2005). The proliferation of somatic and germ cells leads to the enlargement of the gonadal ridge. In the developing ovary, PGCs (in some species now called oogonia) interact with epithelial somatic cells of the ovary and give rise to ovarian or ovigerous cords, which are continuous with the surface epithelium. These cords are more clearly defined in some species that in others (Pepling 2006). Numerous epithelial cells surrounding the PGCs in the ovigerous cords originate in the mesonephros. These epithelial structures are delimited by a basal lamina and surrounded by mesenchymal cells (Guigon and Magre 2006). Other studies support the nonmesonephric origin of gonadal somatic cells. They are mainly based in the demonstration that mesonephric agenesis does not block the formation of the gonad. The cells in the origin of the mesonephros, the adrenal gland and the gonad correspond to a multivalent urogenital mesenchyme (Merchant-Larios 1985). Groups of oogonia that are surrounded by a layer of epithelial cells (granulosa cells) form cysts. In human and other mammals oogonia are connected by cytoplasmic bridges giving rise to a clonal proliferation (Pepling 2006). Mitotic activity of oogonia determine the maximum number germ cells in the life span, that is six to seven million at 20 weeks of gestation in the human (Oktem and Oktay 2008). Due to a high rate of programmed cell death, a girl has one million oocytes at birth and three to four hundred thousand at puberty. Of these only 300–400 will ovulate until menopause (Oktem and Oktay 2008). The main process that marks PGCs/oogonia sex differentiation within the fetal ovary is the beginning of meiosis. PGCs/oogonia entering into meiosis are termed primary oocytes. These pass through the stages of prophase I (leptotene, zygotene, pachytene and diplotene), and around the time of birth become arrested at dictyate stage, which is a pseudo-interphase stage corresponding to the last G2 stage of meiotic process. In contrast, the spermatogonia are not engaged in meiotic process until puberty.

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The debate whether the PGCs/oogonia enter into meiotic S phase ­autonomously following internal signals (Zamboni and Upadhyay 1998; McLaren and Southee 1997) or induced by external signals is still undergoing (reviewed in Bullejos and Koopman 2004). In any case, as reported in the previous section, entering into meiosis does not depend on sex chromosomes. Recent studies demonstrate that retinoic acid (RA), a derivate of vitamin A, induces oogonia to enter in meiosis during fetal life in the mouse ovary and that RA degradation in the fetal testis ­prevents spermatogonia to do the same at the same time (see review by Bowles and Koopman 2007). Stimulated by retinoic acid, gene 8 (Stra8) is involved in the initiation of the meiotic process. For the decision, it is required to enter in the S phase preceding meiosis (Baltus et al. 2006). Germ cells do not initiate meiosis in Stra8-null mice, so RA must trigger meiosis inducing the cytoplasmic protein STRA8, which is never expressed in gonadal somatic cells, possibly due to differences in the epigenetic configuration of the regulatory regions of the gene. Retinoic acid ­produced in the mesonephros, causes germ cell in the ovary to enter meiosis (Bowles et al. 2006), and is also the key to sex-specific timing of meiotic process (Bowles and Koopman 2007). It is likely that besides RA availability entering into meiosis in female PGCs or meiotic prevention in male PGCs require additional molecular control that is being to be identified. For example, epigenetic modification of chromatin in female PGCs and the action of FGF-9 in male PGCs are involved in favoring or preventing meiosis, respectively (Wang and Tilly 2010; Barrios et al. 2010). As reported above, oocytes ring into the last stage of prophase I become arrested in dictyate. The molecular mechanisms of such arrest, necessary for allowing the progressive recruitment of group of oocytes into the growing phase after birth, are unknown in mammals but are likely to involve players of cell cycle control such as cyclins, CDKs and CDKI (Western et al. 2008; Spiller et al. 2009; Miles et al. 2010). When the meiosis is arrested in diakinesis at the end of the prophase of the first meiotic division, a prolonged pseudo-interphase takes place. Around and early after birth, ovigerous cords brake down in follicles, the oocytes are progressively ­surrounded by a monolayer of flat epithelial cells, now called granulosa or follicular cells and a basal lamina, giving rise to the primordial follicles. This process begins in the central region of the ovary and proceeds to the periphery. These follicles are surrounded by a microvascular network of capillaries juxtaposed to basal lamina (Sato et  al. 2006). The development of primordial follicles seems to be related to meiotic dyctiate stage block (Borum 1961). Experimental studies on the relation of a protein of the synaptonemal complex (SCP1), which is typical of pachytene stage and the formation of primordial follicles, demonstrate that the SCP1 deficient ovaries exhibited an increased number of newly formed follicles. These findings suggest that the completion of pachytene stage or probably an artificial finalization of meiotic prophase I endow the oocytes with the ability to orchestrate the follicular assembly (Paredes et al. 2005). Spontaneous primordial follicle development was completely inhibited by a Kit antibody that blocks ­kit-ligand/stem cell factor actions. Experiments in  vitro with rat ovaries demonstrated that kit-ligand is necessary and sufficient to induce primordial follicle

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d­ evelopment. On the other hand, gonadotropins (FSH and hCG) do not induce follicle ­development (Parrot and Skinner 1999). During the progression of the first meiotic prophase and follicle formation many germ cells are eliminated, around 70% in mice, rats and humans. The reasons for this constitutive germ cell death are not clearly established and will be discussed in other chapters of this book. It is important to note that germ cells are necessary for the correct formation of the ovary. However, if PGCs are eliminated by busulphan treatment, a drug that selectively destroys them, (Merchant 1975) or by the pleiotropic mutation W/Wv (Merchant-Larios and Centeno 1981), in the rat and in the mouse, somatic cells undergo proliferation and form an undifferentiated gonad in both sexes. That is, a gonad consisting in packed cells that gradually become organized into two different tissues separated by a basal lamina: epithelial layers and/or epithelial cords and the stromal tissue (Merchant 1975). Similar results have been obtained in Amphibia (Bounoure 1950; Padoa 1964) and in chicks (Willier 1950). Fragmentation of the cords to form follicles takes place only when germ cells act as differentiating centers (Merchant 1975). Some PGCs survive in the gonad of mutant W/Wv mice and are able to transform in normal oocytes and organize follicles capable of inducing steroidogenic tissue (Merchant-Larios and Centeno 1981).

2.7 Endocrine and Paracrine Regulation of Folliculogenesis In order to understand the prenatal endocrine relation of the developing ovary it is necessary take into account the developmental relationships between the ­hypothalamus, the hypophysis and the gonad. Anterior pituitary is formed by an evagination of the oral ectoderm and the posterior lobe is formed by a ventral prolongation of the diencephalon. Autocrine and paracrine signaling are involved in the initial development of the gland. This system differentiates and acquires capability for endocrine function very early. In humans, luteinizing hormone and follicle stimulating hormone are produced in the pituitary as early as the fifth week of gestation. Gonadotropin releasing hormone and other releasing hormones are present in the hypothalamus at about the same time. The ensemble can function as a whole after the differentiation of the portal vessels between the hypothalamus and pituitary (Cummings and Kavlock 2004). When the gonads begin to produce hormones, interactions take place between the gonads and the hypothalamic-pituitary axis. In the female human fetus estrogen is present by the 10th–14th week, but the hormone peaks at about the 20th week, following increments in hypothalamic gonadotropin releasing hormone and pituitary gonadotropin concentrations. Near birth, the hypothalami of the female acquire the capability to respond to rising estrogen levels and the triggering of the surge of LH that is a requisite for ovulation. However, these events will not occur until puberty (Cummings and Kavlock 2004). The development of the follicles is regulated by a complex interplay of hormones produced by the hypophysis as follicle-stimulant hormone (FSH) and luteinizing

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Fig. 2.5  Folliculogenesis process. The preantral follicles are independent of gonadotropin. The development of the follicles is regulated by FSH and LH, and paracrine regulatory factors ­produced by the oocytes and the surrounding somatic cells

hormone (LH), and paracrine regulatory factors produced by the oocytes and the surrounding somatic cells. The latter are especially crucial for the formation of the primordial follicle population around and early after birth and likely for the selection of follicles from these populations to be driven through the subsequent maturation stages. Hormones begin to play a role from the primary follicle stage throughout the secondary, preantral and antral follicle stages up to ovulation (Fig. 2.5). FSH up regulates a number of genes in granulosa cells, such as cyclin D2 involved in proliferation, aromatase enzyme required for estrogen production, and LH receptor (Anttonen 2005). FSH receptor expression is restricted to granulosa cells, whereas LH receptor is expressed both in granulosa and theca cells. LH stimulates androgen synthesis in theca cells and FSH induces estrogen conversion from androgen, by means of aromatase enzyme, in granulosa cells (Gómez et al. 2001). The defects in folliculogenesis differ between FSH receptor- and LH receptordeficient mice. Both have no mature, preovulatory follicles or luteal glands, but in FSH receptor-null mice follicular growth arrests at the preantral stage, whereas in LH receptor-null mice the follicles arrest at the antral stage. These findings demonstrate the need for FSH action at the preantral stage and beyond, and both FSH and LH actions for the follicular maturation and ovulation stages (Anttonen 2005). Oocytes and somatic cells produce several paracrine factors, modifying gonadotropin actions. Oocyte supplies specific morphogens that preserves phenotype and gene expression in cumulus cells. On the other hand, the somatic cells provide the oocytes with nutrients and growth factors during folliculogenesis (Anttonen 2005). Members of the TGF-ß superfamily have essential roles in these paracrine interplays. Growth and differentiating factor 9 (GDF9), bone marrow-morphogenic

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protein 15 (BMP-15), and BMP6 are secreted by the oocyte, while activins, ­inhibins and anti-Müllerian hormone (AMH) are expressed and secreted by somatic cells. AMH, in addition to its role in the differentiation of male reproductive tract, is a granulosa cell product capable of suppressing follicular growth induced by FSH and epidermal growth factor. AMH-null mice have an increased number of growing follicles, resulting in loss of the primordial follicles at a premature age (Anttonen 2005). For a general physiology text book consult Peterson (2007).

2.8 The Meiotic Division The meiosis is composed of two cell divisions preceded by the only S phase of this process (Fig. 2.6). The result is four cells with half the number of chromosomes of the species and only one DNA double helix per chromosome. The haploid cells are the mature gametes, the oocytes and the spermatocytes. The first division is composed of an S phase followed by G2 or preleptotene stage, a long prophase compose of several stages: leptotene, zygotene, pachytene, diplotene, and diakinesis (dictyate). Several processes specific to meiosis take place during this prophase

Fig.  2.6  Schematic drawing of meiosis process (kindly provided by Abrahan HernándezHernández)

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such as homologous recognition, alignment of homologous chromosomes, pairing of homologues by means of a special structure, the synaptonemal complex, and chiasm formation. The oocyte remains in diakinesis until ovulation when it ­proceeds to the metaphase in which the homologous chromosomes are still associated by the chiasmata, and then to the anaphase of the first division, in which each member of homologous chromosome pairs migrates to a different pole of the ­spindle. Thus the diploid number of chromosomes is divided between the two ­products of the first telophase. The meiotic spindle is located at the periphery of the cytoplasm, near the cell membrane and it specifies the position of the cleavage furrow, thus the two new cells formed by the telophase are very different in size. The larger ­daughter cell corresponds to the secondary oocyte and the smaller is the first polar body. Each has the haploid number of chromosomes formed by two DNA double helices. The second meiotic division is when the sister chromatids of each chromosome separate and migrate to a different cell like in a mitotic division. In this cytokinesis, as in the first one, the cleavage furrow separates a small polar body and a large oocyte. The second meiotic division takes place only after fertilization. The biological role of meiosis is the production of gametes (haploid cells), which are cells with only one member of each pair of chromosomes formed by one DNA double helix, frequently formed by paternal and maternal segments. During meiotic process, genetic recombination takes place by two different processes: random distribution of maternal and paternal homologues, in the first anaphase, and recombination. The maternal and paternal homologous forming each bivalent chromosome distribute at random at the first metaphase, so each secondary oocyte may incorporate a different mixture of paternal and maternal chromosomes. The recombination is the exchange of segments of chromatid between homologous, maternal and paternal chromosomes, when they are paired at the pachytene stage of the first meiotic prophase. Each exchange is visualized as a connection called a chiasma between the chromosomes. The chiasmata are easily seen under a light or electron microscope in post-pachytene stages of meiotic prophase I. The ultrastructural and molecular aspects of meiosis are better studied in ­spermatocytes than in oocytes, but most processes taking place in the prophase of the first meiotic division are similar in both sexes. The phase G2 of the last interphase of the spermatogonia is characterized by small lampbrush structures of extended chromatin formed by an axis from which irradiates the loops. These micro-lampbrush chromosomes are not found in interphase nuclei of somatic cells. They persist during leptotene and zygotene stage. Immunolocalization and in situ localization DNA and RNAs using electron microscope demonstrate that these lampbrush structures are formed by extended chromatin transcribed mainly or exclusively by RNA polymerase II and that the product of transcription is still associated to the chromatin; they contain introns and ribonucleoproteic particles associated to splicing are scarce. The results of quantitative autoradiography after [3H]uridine labeling demonstrate that transcription of the extended chromatin of the micro-lampbrush is intense but the export of RNA to the cytoplasm is very slow. Then the newly synthesized RNA is probably involved in a nuclear function taking place within these cells: the process of homology searching and recognition,

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p­ revious to pairing (Vázquez-Nin et  al. 2003). The searching of homologies is facilitated by a much conserved mechanism among eukaryotes, the bouquet. The bouquet involves the association of the telomeres of all chromosomes to the nuclear envelope and the subsequent displacement to a small zone of the envelope near the centrosome, a microtubule organizing center in the cytoplasm. This mechanism facilitates finding homologue regions of homologous chromosomes. The final pairing of homologous chromosomes takes place with the formation of another structure extremely conserved by evolution, the synaptonemal complex. This complex is formed by the parallel alignment of corresponding regions of homologous chromosomes from telomere to telomere, and is typical of the pachytene stage. The synaptonemal complex is formed by two lateral arms and a central space devoid of DNA except in the recombination nodules. The lateral arms are formed by DNA, RNA and proteins. Two filaments of chromatin formed by two DNA double helixes each are folded inside the lateral arms. These filaments are continuous with the long loops surrounding the complex. RNA of unknown origin is associated with the DNA filaments (Vázquez-Nin and Echeverría 1976; Ortiz et al. 2009). The complex is surrounded by long loops of extended chromatin associated to the lateral arms. Each lateral arm and the loops associated with it correspond to one of the paired homologous chromosomes. The DNA of the lateral arms of the synaptonemal complex is composed of non-coding repetitive sequences (Hernández-Hernández et al. 2008). For a recent review of epigenetic aspects of the synaptonemal complex and its formation (Hernández-Hernández et al. 2009). The alteration of the structural maturation of the central element of the ­synaptonemal complex abolishes the formation of crossovers, preventing recombination (Hamer et al. 2008). Many genes are involved in the process of recombination. Some of them, such as Spo11, are involved in double strand breaks and are conserved between yeast and mammal. In mammals, yeast and higher plants, recombination takes place more frequently in 1–2 kb long regions of the genome called hot spots (Parvanov et  al. 2009). The frequency of recombination is low outside of hotspots. Recombination hot spots in humans are not associated with DNase I or microccocal nuclease-hypersensitive sites (Fan and Petes 1996); they are more influenced by base composition (DNA motif), coding context and DNA repeats (Myers et  al. 2006). In human genome both repeat DNA and exonic sequences are negatively associated with recombination; indeed recombination rates reach a peak at distances of 10–50 kb from genes (Myers et al. 2006). On the contrary, regions of high GC content are strongly associated with hotspots, also the seven bases nucleotide motive CCTCCCT is five times enriched in the hot spots of the THE1A and THE1B retrotransposons. Mutation of one base of this motive strongly reduces the hotspot activity (Myers et al. 2006); protein-DNA complexes are essential for hotspot activity. The ade6-M26 hotspot (chromosomal region) of fission yeast is the most extensively characterized and best understood meiotic hotspot in eukaryotes (Gao et al. 2009). Atf1-Pcr1 protein heterodimer associates with ade6-M26 hotspot. The homologous recombination –activation domain resides exclusively in Atf1, which is a member of ATF/CREB family of transcription factors. The Atf1-Pcr1-M26 protein-DNA complex promotes locally the

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i­ nitiation of recombination from double strand breaks catalyzed by Rec12 (Spo11) recombinase, which is a conserved protein of the meiotic recombination machinery. The recombination-promoting domain resides in Atf1 protein and the hotspot activation requires the phosphorylation of ATF1 by Spc1, a protein kinase of the p38-family. The regulation of basal meiotic recombination is independent of this phosphorylation, whose function is to correct position of Atf-Pcr1 complex (Gao et al. 2009). In Saccharomyces cerevisiae double strand breaks occur within a few hundred base pairs of the transcriptional sites. However, only some hotspots are activated by transcription factors. The well characterized hotspot located within the promoter of the gene HIS4 is dependent on several transcription factors and ablation of the transcription factor binding sites abolishes recombination (Borts 2009). The study of a hot spot in the mouse genome, called Psmb9, showed that its activity is induced by a locus named Dsbc1, for double strand break control 1. This locus is located within a 6.7-Mb region on chromosome 17. Dsbc1 influences double strand breaks and crossovers, not only at Psmb9, but in several other regions of the chromosome 17. Dsbc1 influences the crossover distribution on chromosomes 15 and 18, and even the crossover activity in the Hlx1 hotspot in chromosome 1 (Grey et al. 2009). Recently it was found that the recombination activity of several hotspots on mouse chromosome 1 can be either activated or suppressed by the CAST allele(s) of distant trans-acting loci and that the activation of several hotspots is controlled by the locus Rcr1 that exert its action at the initiation of recombination (Parvanov et al. 2009). These interesting results on the positioning, the molecular mechanism and the regulation of meiotic recombination are probably just the beginning of a long line of future research. Failure to repair DNA damage caused by recombination triggers meiotic checkpoints and destruction of the germ cells by apoptosis can take place. However, there is an important sexual dimorphism; checkpoint requirements are much less stringent in females than in males. This provides an explanation for the ten-fold higher number of meiotic errors in females compared with males (Cohen et al. 2006).

2.9 Epigenetic Reprogramming of Germ Cells Epigenetic signals are covalent modifications of histones or DNA that convey a specific “meaning” to the stretch of chromatin on which they are found. Frequently they induce changes in chromatin conformation and gene expression. Pluripotent embryonic germ cells are capable of giving rise to all tissues derived from the three embryonic primary epithelial layers: ectoderm, mesoderm and endoderm. Thus they must be able to reorganize their mechanisms of gene regulation to allow multiple lines of cell differentiation. Besides determining genome plasticity, epigenetic modifications regulate X inactivation/reactivation in the female embryo and mark the paternal and maternal genome of germ cells during gametogenesis, making the expression of a group of

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genes monoallelic, crucial for embryo development termed for modification such as “imprinted” genes. As reported above, BLIMP1 (PRDM1), a transcriptional repressor, plays a ­critical role in the specification of PGCs in mice. BLIMP1 interacts with PRMT5, an arginine-specific methyltransferase, which mediates symmetrical dimethylation of arginine 3 in histone H2A and/or H4 tails. At E11.5, BLIMP1-PRMT5 translocates from the nucleus to the cytoplasm, resulting in the loss of H2a/H4 R3 methylation, thus determining an extensive epigenetic reprogramming of germ cells. At this stage, Dhx38, a putative target of the BLIMP1-PRMT5 complex, is also upregulated. The expression of Dhx38 is seen in pluripotent embryonic germ cells that are derived from PGCs when Blimp1 expression is lost. These observations demonstrate that in mice Blimp1 is involved in posttranscriptional regulatory actions mediated by methylation-demethylation of histone H2A and/or H4 arginine 3 (Ancelin et al. 2006). Postranscriptional modifications of the histones may be reprogrammed more easily than DNA methylation, which is a very stable modification that may be heritable. In mammals methylation of the DNA occurs almost exclusively on CpG dinucleotide, without preference for sequence context surrounding this target. About 60% of human genes are associated with CpG islands, of which the great majority are unmethylated at all stages of development, in all types of tissues (see review by Bird 2002). It is important to note that the DNA methylation does not silence active promoters, but affects genes already silenced, making the turn off permanently. DNA methylation is associated with a stable repression of transcription, frequently accompanied by conformational changes of the repressed loci. Examples of the modifications due to DNA methylation are monoallelic expression of some imprinted genes, repression of transcription of retrotransposons and inactivation of one X chromosome, and other examples quoted by Bird (2002). In all cases there is a compaction of the repressed chromatin and the modifications are for the lifespan (Lees-Murdock and Walsh 2008). DNA methylation is heritable through cell division by copying of the parental strand and can pass from one generation to the next via the gametes. The methylating enzyme DNMT1 methylates preferentially CpG whose partners on the parental strand are already methylated (see review by Bird 2002). It is interesting to note that also PRDM14, a PR domain-containing transcriptional regulator is expressed at the PGC specification. Its action is critical in two events: the reacquisition of pluripotency and successful genome wide epigenetic reprogramming. In mice Prdm14 mutants, the failure of these events manifests even in the presence of PRDM1 (BLIMP1), a key transcriptional regulator for PGC specification (Yamaji et  al. 2008). De novo methyltransferases DNMT3A and DNMT3B are highly expressed during epigenetic reprogramming (see review by Bird 2002). Differentially methylated regions of imprinted genes are conserved in migrating PGCs (Hajkova et al. 2002). In females only one X chromosome is active in specified PGCs. The reactivation of one X occurs during PGC migration and is ­completed in all PGCs after their arrival in the developing ovary and before their entering into

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meiosis. Reversion of imprinted X chromosome inactivation in the inner cell mass of the female blastocyst is initiated by the repression of Xist from the paternal X chromosome. Three of the key factors supporting pluripotency, Nanog, Oct3/4 and Sox2, bind to Xist intron 1 in undifferentiated embryonic stem cells. The drastic release of all three factors triggers a rapid ectopic accumulation of Xist RNA and X inactivation in the epiblast. These three factors cooperate to repress Xist and thus couple X inactivation reprogramming to control of pluripotency during embryogenesis in mammals (Navarro et al. 2008). Studies carried out on mice showed that the differentially methylated regions of maternally or paternally imprinted genes are progressively and synchronously ­demethylated in PGCs. The repetitive DNA elements studied so far are coordinately demethylated in the male and female mice genomes. Imprinted sequences are ­completely demethylated and repeat sequences only lose some methylation, retaining a substantial degree of methylation (see review by Lees-Murdock and Walsh 2008). In mice, the female germline demethylation continues past e15.5 as the female PGC, now known as primary oocytes, progress through prophase of meiosis (LeesMurdock et al. 2003). The mechanisms of demethylation have not been clarified. There are at least two different procedures: active or enzymatic demethylation and slower replication-coupled demethylation. The relationships of demethylase complex to MBD2b remain unclear. Some works demonstrate that the 5-methylcytosine was replaced by cytosine during demethylation reaction. Other enzymatic complexes of hydrolyses 5-methylcytosine to cytosine and methanol (Wolffe and Wade 1999). However, biochemical data suggest this reversal of the methylation is energetically unfavorable (see review by Lees-Murdock and Walsh 2008). Replication-coupled demethylation propose that DNA replication can also contribute to eliminating 5-methylcytosine from DNA. If some factor prevents the access of the methyltrasnferase enzyme to these sites, progressive demethylation of selected sites can take place (Wolffe and Wade 1999). The mechanism of demethylation in mammals has not been confirmed and no reproducible ­demethylation activity has been identified (see review by Lees-Murdock and Walsh 2008). Recent results obtained in mouse support a role of the Activation-Induced cytidine Deaminase (AID), a member of the cytidine deaminase APOBEC family, and of Base Excision Repair (BER) pathway in genome-wide DNA demethylatioin occurring in PGCs after gonadal ridge colonization (Popp et  al. 2010; Hajkova et al. 2010). Following erasure of imprints and partial erasure of methylation on repeats, these sequences must undergo de  novo methylation during subsequent germ cell development to achieve the methylation patterns of the mature gametes (see review by Lees-Murdock and Walsh 2008). While in the male germ line resetting of methylation occurs before meiosis, in the female germ line the imprint control regions are hypomethylated until after the pachytene stage of meiosis I, which occurs in the postnatal growing oocyte (Lees-Murdock et  al. 2005; see also the review by Lees-Murdock and Walsh 2008).

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Chapter 3

Development of the Ovary in the Embryo, Infancy, Childhood, Pre-puberty and Puberty Gerardo H. Vázquez-Nin, María Luisa Escobar, Olga M. Echeverría, and Massimo De Felici

Abstract  The hypothalamic-pituitary-gonadal system becomes activated in the human fetus. Negative feedback becomes operative towards term. During infancy gonadotropin concentration increases steroid secretion more in boys than in girls. The signification of this early hypothalamic-pituitary-gonadal activation is not known. During infancy serum concentration of gonadotropins appear to be ­sufficient to maintain growing follicles to small antral stage. In the absence of appropriated levels of LH, FSH or both, these follicles are unable to enter the preovulatory stage. In infantile rodents there is an accelerated loss of follicles from the resting pool during the initial waves of follicle growth. The presence of atretic processes during childhood was described in human and rat ovaries. Puberty involves the reactivation of GnRH pulse generator by removal of central system restrain. A large number of oocytes of rats from newborn to puberty are eliminated by means of different cell death processes: apoptosis, autophagy and a mixed process in which both routes participate in the same cell. List of Abbreviations ADP DAPI

Adenyl di-phosphate 4’, 6-Diamidino-2-phenylindole. Blue fluorescent dye that intercalates between DNA bases and stains it specifically FSH Follicle-stimulating hormone GnRH Gonadotropin releasing hormone LAMP-1 Lysosomal associated membrane protein 1

G.H. Vázquez-Nin (*), M.L. Escobar, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico City, Mexico e-mail: [email protected] M. De Felici Department of Public Health and Cell Biology, University of Rome, Tor Vergata, Italy G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_3, © Springer Science+Business Media B.V. 2011

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Luteinizing hormone Polymerase chain reaction Ribonucleic acid Real time PCR Terminal Deoxy-UTP nick end labeling. Procedure that detects DNA breaks typical of apoptosis.

3.1 Infancy It is interesting to note that the hypothalamic-pituitary-gonadal system becomes activated in the human fetus. The negative feedback of steroids becomes operative towards term and with the removal of placental steroids gonadotropin secretion is activated during the first 4 months. This high gonadotropin concentration increases steroid secretion more in boys than in girls. The significance of this early hypothalamic-pituitary-gonadal activation is not known. In early postnatal heifer calves there is a gonadotrophin driven increase in ­ovarian antral follicle growth. This initial endocrine activity is controlled by ­negative feedback mechanisms until puberty (Rawlings et al. 2003). A pulse pattern of LH and FSH secretion was found in females of all ages. Serum LH and FSH concentration increases to concentrations even exceeding adult values during the first 4 months (Neely et al. 1995). A rise in serum levels of inhibin B and estradiol was also found in the first 3 months of life in girls, declining to prepubertal concentrations thereafter (Bergadá et al. 2001). The high gonadotropin concentrations cause gonadal activation and increase in steroid secretion. After 4–6 months of age, gonadotropin and steroid concentrations progressively decrease to very low levels by 1–2 years of age (Apter 1997). It is important notice that the atresia process occurs in neonate, in prepubertal and in adult organisms. Most follicles appear to undergo atresia at an early stage of growth. Therefore, conditions other than increased androgens production may explain failure of these follicles to continue growth. Serum concentrations of gonadotropins appear to be important. Basal concentrations of gonadotropins appear to be sufficient to maintain growing follicles to the small antral stage (Fig.  3.1). In the absence of an appropriate sustained increment of LH, FSH or both, these follicles are unable to enter the preovulatory stage. In infantile rodents there is an accelerated loss of follicles from the resting pool during the initial waves of follicle growth (Hage et al. 1978). This accelerated rate of loss of follicles has been attributed to a lack of mature follicles that might exert a negative effect on initial recruitment and to qualitative differences in the first groups (Edwards et al. 1977), reviewed in McGee and Hsueh (2000). Initial recruitment is believed to be a continuous process that starts just after follicle formation, long before pubertal onset. Studies on the ovary atresia have demonstrated that in the neonate and prior to puberty most atresia occurs in preantral follicles (reviewed in Richards 1980).

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Fig. 3.1  Schematic representation of growing follicles in gonadotropin independent stages. The follicles reach an early antral phase in an independent gonadotropin manner. These follicles reach an early antral phase in spite of the absence of gonadotropin

The localization and quantitative changes of estradiol receptor of uterine e­ pithelial cells, muscle cells and fibroblasts were studied in the interchromatin space, compact chromatin, nucleolus, cytoplasm and background. In the three types of cells the estradiol receptor was mainly associated to interchromatin space, and to nucleolus to a lower extent. It is interesting to note that in the immature chick ovary estrogen receptor is expressed in primary oocytes and in somatic cells including typical steroidogenic. At ultrastructural level the estrogen receptor was immunolocalized in extended chromatin. These data support the hypothesis that estrogens are involved in the function of somatic and germ cells in the immature ovary (Méndez et  al. 1999). Using immunoelectron microscopic localization estradiol receptor was found in male and female reproductive organs and in cells of non-reproductive organs as hepatocytes, epithelial duodenal cells, striated muscle fibers, cells of the proximal convoluted tubules of the kidney, lymphocytes, neurons and adipose cells. In all of these cells a higher density of the estradiol receptor was found in the nucleus, especially in the space between the clumps of compact chromatin, where extended chromatin is abundant, as was previously found in epithelial endometrial cells (Méndez et al. 1999). It is interesting to note that in endometrial cells ovariectomy reduce the volume of the nucleolus to a third of its normal value, while the number of perichromatin granules (a nuclear RNP particle containing mRNA) increases. A single injection of 20 mg of estradiol produces a significant decrease of the number of perichromatin granules to a fourth of the value of castrate animals in 15–30 min and a significant increase of nucleolar volume in 2  h. The changes in nucleolar volume are due exclusively to the effects of estradiol on pre-rRNA transcription, while the variations of the number of perichromatin granules is the results of a combination of the effects on pre-mRNA synthesis and on its processing and transportation to the cytoplasm (Vázquez-Nin et al. 1978). Estradiol receptor was found not only in female reproductive organs but also in male reproductive organs and in non-reproductive organs as hepatocytes, epithelial duodenal cells, striated muscle

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fibers, cells of proximal convoluted tubules of the kidney, lymphocytes, neurons, and adipose cells (Echeverría et al. 1994). Immunolocalization of estrogen receptor in rat uterine epithelial cells, muscle cells and fibroblasts demonstrated that estradiol receptor is found in the nucleus mainly on extranucleolar ribonucleoprotein fibrils and probably on extended chromatin. In epithelial and muscle cells the nucleolus was labeled, but compact chromatin was not labeled. In epithelial cells was found a low but significant labeling of the cytoplasm. Ovariectomy did not change these distributions (Vázquez-Nin et al. 1991).

3.2 Childhood The lowest gonadotropin concentrations were found in 1–9 year old girls. The mean serum estradiol concentration in prepubertal girls was found to be 2.2 pmol/l. This finding implies that the ovaries secrete estradiol long before puberty (Apter 97 and references therein). Prior to puberty, in several mammals there is an initial recruitment of some ­primordial follicles to initiate growth, while most primordial follicles remain ­quiescent. Growing follicles are generally located in the inner part of the cortex, while non-growing primordial follicles lie in the external cortex. These follicles grow and develop an antrum; the oocytes remain in diakinesis of the first meiotic prophase and finally undergo atresia. In the ovary of 3-week-old mice, which is childhood for mice, there are simultaneously degenerating large follicles, together with medium-sized ones and some starting to grow (McGee and Hsueh 2000; Ortiz et al. 2006; Escobar et al. 2008). The presence of atretic processes in the ovaries during childhood was described in humans (Peters et al. 1976) and in rats (Ortiz et al. 2006; Escobar et al. 2008). Before puberty, blood concentrations of FSH are not sufficient to sustain ­development, thus all growing follicles become atretic. Inhibin B, produced by granulosa cells, is a heterodimeric glycoprotein that ­suppresses synthesis and the secretion of the follicle stimulating hormone (FSH) (Fig. 3.2). Prepubertal girls have irregular luteinizing hormone (LH) pulses of low ­amplitude during the day, while LH and FSH pulses increase amplitude within 1 h after the onset of sleep. Mechanisms underlying the atresia of follicles prior to puberty set the number of follicles available to the adult animal to ensure normal fertility and the hormones necessary to the overall health of the organism. The effects of estrogens on follicular development has been studied in aromatase knockout mice, in these organisms the follicles develop from primordial to antral, demonstrating that the estradiol deficiency is not critical for the follicular development. It is interesting that in knockout mice the number of primordial follicles is 40% less than in wild type organisms (Britt et  al. 2001). In the newborns of pregnant baboons treated with aromatase inhibitors, the primordial follicles number was reduced (Zachos et  al. 2002).

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Fig. 3.2  Hypothalamic-pituitary-gonadal system in chilhood. In absence of GnRH low levels of LH are present. The granulosa cells secrete inhibins which suppress the LH levels. The absence of LH and FSH source provokes the selectable antral follicles become atretic

The reduced number of primordial follicles demonstrates that the steroids play a role in early folliculogenesis. Primordial follicles are cleared from the ovary at an extremely high rate, but they were not positively associated with nuclear condensation, cell shrinkage, activation of caspase 3, cleavage of poly (ADP ribose) polymerase 1(PARP1), or fragmentation of DNA. These data are consistent with a non-apoptotic death of primordial follicles in prepubertal mice (Tingen et al. 2009). The transforming growth factor beta (TGFb) family of proteins consists of ­multifunctional cytokines that modulate a wide variety of cellular functions. Smad proteins mediate TGFb (and activin) signaling through a cascade of ligand-induced phosphorylation. Smad3 is a receptor highly expressed in the ovarian surface ­epithelium, granulosa cells, and oocytes in several animals (Tomic et  al. 2002). Smad3-/- mice have reduced fertility compared with wild type, does not affect the size of primordial follicles pool, but is important for the normal growth of primordial follicles to the antral stage (Tomic et al. 2002). Posterior studies demonstrate that Smad3 deficiency also causes atretic follicles, increased apoptosis, and ­degenerated oocytes (Tomic et al. 2004). Ultrastructural alterations of oocytes and granulosa cells were described in atretic follicles of prepubertal and young adult rats long time ago (Vázquez-Nin and Sotelo 1967; Ortiz et  al. 2006). Oocytes present lamellar alteration of the cytoplasm, probably due to changes in the cytoplasmic matrix and changes in the ­distribution of the organelles. Regions of the cytoplasm become devoid of vesicles,

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mitochondria, endoplasmic reticulum, Golgi cisternae, ribosomes and are occupied by fibrils and lamellae. When the process has fully developed the lamellae form groups of multiple parallel lamellae. The multivesicular bodies and other lysosomal vesicles containing cytoplasmic structures increase in number. Final stage of follicular atresia is characterized by loss of contact between follicular cells and the oocytes, corona cells become extensively vacuolated retract their prolongations to the oocytes and detach from the zona pellucida. Atretic anomalous segmentation of the oocytes frequently takes place (Vázquez-Nin and Sotelo 1967). The structural features of the process of rat oocytes death are similar in infancy, childhood and prepubertal periods. The structural features are mainly characterized by the presence of abundant clear vacuoles, autophagosomes, as well as by the absence of large clumps of compact chromatin associated to the nuclear envelope and apoptotic bodies. Cytochemical features as positive reaction to TUNEL method, active caspase-3 and lamp1 immunolocalizations are common to cell death of oocytes in all types of follicles in rats from 1 to 28 days (Ortiz et al. 2006). Recent studies of the alterations of dying oocytes in prepubertal rats employed TUNEL method, immunolocalization of active caspase 3, lamp1, localization of acid phosphatase, and DAPI staining (Fig. 3.3). All procedures were performed in adjacent sections of each oocyte. This procedure allowed demonstrating that in most dying

Fig. 3.3  Oocytes of different ages. In (a) and (a’) 1 day old oocytes are shown. In (b) and (b’) 25 days old oocytes. The oocytes are simultaneously positive to different markers as lamp1 and to the TUNEL technique. Scale bars 10 mm

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oocytes exist simultaneous features of apoptosis as active caspase 3 and DNA breaks characteristic of apoptosis and increased lamp1 and acid phosphatase characteristics of autophagy. Large clumps of compact chromatin and membrane blebbing were absent. Electron microscope observations demonstrated the presence of large number of small clear vesicles and autophagolysosomes. All these features indicate that a large number of oocytes are eliminated by a process sharing features of apoptosis and autophagy. In dying oocytes of newborn rats the markers of apopotosis predominate over those of autophagy. However, fragmentation and apoptotic bodies were not found. These features suggest that in different cytophysiological conditions the process of cell death may be differently modulated (Escobar et al. 2008). The processes of cell death were also studied in  vitro in populations of oocytes isolated from ­prepubertal rats. In order to identify apoptosis, the externalized phosphatidylserine was recognized with Annexin-V coupled to FITC and fragmentation of DNA was demonstrated by means of electrophoresis. Oocytes were tested for autophagy by means of incorporation of mondansylcadaverine and monitoring Lc3-I/Lc3-II by western blot. The expression of mRNA marker genes of autophagy and of apoptosis was studied by means of RT-PCR in pure populations of oocytes. Some oocytes expressed at least one of the following markers: caspase-3, lamp1 and Lc3. Some oocytes were positive to Annexin-V or to monodansylcadaverine. However, most of them were simultaneously positive to both markers. The relative frequency of oocytes simultaneously present to markers of apoptosis and autophagy did not change in rats of different ages studied. The mRNAs of caspase-3, lamp1 and Lc3 were present in all populations of oocytes. These results demonstrate that oocytes of rats from new born to prepubertal age are eliminated by means of different cell death processes: apoptosis, autophagy, and a mixed process in which both routes to cell death participate in the same cell (Escobar et al. 2010). The LH and FSH source allow the selectable antral follicles progress to ovulation. In prepubertal organisms antral follicles undergo atresia. In granulosa cell layer scattered pyknotic nuclei can be seen as well as cells detached of the basal ­membrane, fragmentation of the basal membrane and cell debris in the antral cavity. Frequently macrophages invade the stromal area adjacent to the atretic follicle.

3.2.1 Puberty Puberty involves the reactivation of the GnRH pulse generator by the removal of central nervous system restraint (Apter 1997). In this process the diminution of the sensibility of the hypothalamus and hypophysis to the inhibitory effects of ­estrogens is probably also involved (Villavicencio-Macías 1991). At the beginning of puberty an increase in LH and particularly in FSH concentrations takes place in girls, both while asleep and awake (Apter 1997). At the beginning of pubertal period, the percentage of pituitary cells producing estradiol receptor a increases significantly. Simultaneously the plasma concentrations of LH increases while GH concentration decreases. These changes reflect an escalating

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Fig.  3.4  Hypothalamic-pituitary-gonadal system during puberty. The GnRH pulse triggers an increase in LH and FSH concentrations. The LH and FSH source allow the selectable antral ­follicles progress to ovulation

regulatory role of estrogens in the physiology of pituitary cells at the end of the prepubertal period (Polkowska et al. 2008). During puberty a rise in serum levels of inhibin B, FSH, LH, and estradiol takes place (Chada et al. 2003). It is initiated independent of gonadal activity and can be activated by exogenous pulses of GnRH (Apter 97 and references therein). The mechanism activating puberty is still largely unknown. It seems to be a process initiated in the central nervous system which results in the production and liberation of gonadotropin releasing hormone (GnRH). This factor transported to the hypophysis by the hypothalamus – pituitary portal vessels stimulate gonadotropin secretion (Fig. 3.4). Experimentally the onset of puberty may be initiated independently of gonadal activity, by exogenous pulses of GnRH in infantile monkeys and girls with delayed puberty (Apter 1997) or by the administration of prolactin, which increases the sensibility of the ovary to the low concentrations of circulating gonadotropins ­existing in prepubertal rats. This treatment increases the number of receptors to LH in granulosa cells, producing an augment in the liberation of progesterone and estrogen. (Villavicencio-Macías 1991). The ovaries of immature cattle, sheep and pigs respond to the injection of exogenous gonadotropin producing fertilizable oocytes, showing that the ovary is prepared for its adult function long before puberty (González-Padilla 1978). Several studies report the birth of live offspring from bovine and ovine using in vitro fertilization of oocytes from prepubertal animals. However, the ­developmental

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competence of in vitro-matured oocytes from prepubertal animals is lower than that of oocytes derived from adult animals (Ledda et  al. 2001). Prepubertal swine oocytes were also found to be less competent than their adult counterparts, but are also able to develop to term (Marchal et  al. 2001). The lesser competence of ­prepubertal oocytes may be due to morphological differences, as fewer transzonal projections or a lower level of maturation-promoting factor, a universal cell cycle regulator of both mitosis and meiosis (Ledda et al. 2001). The innervation of the mammalian ovary has been described several times. Earlier studies showed that the extrinsic innervation of the ovary consists of sympathetic, parasympathetic and sensory fibers. The presence of ganglionar polygonal neurons was also described in several mammals (D’Albora et  al. 2002). Neurons identified by the expression of a neuron-specific nuclear protein (NeuN), were detected at all postnatal intervals in rat ovary. About 20% are catecholaminergic, as determined by their content in tyrosine hydroxylase and others showed neuro­peptide Y immunoreactivity (D’Albora et al. 2000; D’Albora et al. 2002). Neurons are also present in the human ovary (Anesetti et al. 2001). In the rat, the complexity of these neurons increases during prepubertal development. However, the number of neurons decreases after 60 days of postnatal age, and most of the remaining neurons present morphological anomalies. In rhesus monkey the total number of ovarian neurons decreases gradually between 2 months and 12 years of age. Conversely the number of tyrosine hydroxylase positive neurons increases significantly at puberty and declines with loss of reproductive capacity. (Les Dees et al. 2006). The ovarian neurons originate from the neural crest (Les Dees et al. 2006). In rat ovaries, catecholaminergic nerves are present in prefollicular stages in fetal (19 day) and newborn (15 h after birth) animals, as revealed by tyrosine hydroxylase immunodetection. As prefollicular ovaries are insensitive to gonadotropins; the developing ovary is probably subject to direct neurogenic influences before it acquires responsiveness to gonadotropins (Malamed et  al. 1992). On the same token, the absence of nerve growth factor, a protein of the neurotrophin family, required for survival and differentiation of neurons, is also necessary for the growth of ovarian primordial follicles, a process known to occur independently of gonadotropins in immature mice. The ovaries from homozygote nerve growth factor null (-/-) mutant animals, exhibited a reduced population of primary and secondary follicles in the presence of normal gonadotropin levels, and an increased number of oocytes that failed to be incorporated into a follicular structure (Dissen et al. 2009). Experiments ex-vivo using the celiac ganglion-superior ovarian nerve-ovary system of prepubertal rats showed that celiac ganglion participates on the regulation of the prepubertal rat ovary in one of two ways: either increasing nitric oxide, a gaseous neurotransmitter with cytostatic characteristics that causes immature follicles to remain dormant or by increasing the liberation of androstendione and estradiol, the steroids necessary to begin the first estral cycle (Delgado et al. 2006). The ensemble of these results shows that the prepubertal development of the ovary is mainly controlled by neural signals, while after puberty there is a gradual decrease in the number of intrinsic ovarian neurons, possibly due to programmed cell death (D’Albora et al. 2002).

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References Anesetti G, Lombide P, D’Albora H et al (2001) Intrinsic neurons in the human ovary. Cell Tissue Res 306:231–237 Apter D (1997) Development of the hypothalamic-pituitary-ovarian axis. Ann NY Acad Sci 816:9–21 Bergadá I, Bergadá C, Campo S (2001) Role of inhibins in childhood and puberty. J Pediatr Endocrinol Metab 14:343–353 Britt KL, Drummond AE, Dyson M et al (2001) The ovarian phenotype of the aromatase knockout (ArKO) mouse. J Steroid Biochem Mol Biol 79:181–185 Chada M, Průša R, Bronsky J et al (2003) Inhibin B, stimulating hormone, luteinizing hormone, and estradiol and their relationships to the regulation of follicle development in girls during childhood and puberty. Physiol Res 52:341–346 D’Albora H, Lombide P, Ojeda S (2000) Intrinsic neurons in the rat ovary: an immunohisto­ chemical study. Cell Tissue Res 300:47–56 D’Albora H, Anesetti G, Lombide P et  al (2002) Intrinsic neurons in the mammalian ovary. Microscop Res Tech 59:484–489 Delgado S, Casais M, Sosa Z, Rastrilla A (2006) Ganglionic adrenergic action modulates ovarian steroids and nitric oxide in prepubertal rat. Endocr J 53:547–554 Dissen G, Romero C, Hirshfield A, Ojeda S (2009) Nerve growth factor is required for early ­follicular development in the mammalian ovary. Endocrinology 142:2078–2086 Echeverría OM, González-Maciel A, Traish AM et al (1994) Immuno-electron microscopic localization of estradiol receptor in cells of male and female reproductive and non-reproductive organs. Biol Cell 81:257–265 Edwards RG, Fowler RE, Gore-Langton RE et al (1977) Normal and abnormal follicular growth in mouse, rat and human ovaries. J Reprod Fertil 51:237–263 Escobar ML, Echeverría O, Ortiz R et al (2008) Combined apoptosis and autophagy, the process that eliminates the oocytes of atretic follicles in immature rats. Apoptosis 13:1253–1266 Escobar ML, Echeverría OM, Sánchez-Sánchez L et  al (2010) Analysis of different cell death processes of prepubertal rat oocytes in vitro. Apoptosis 15:511–526 González-Padilla E (1978) La aparición de la pubertad en vaquillas. Cienc Vet 2:293–325 Hage AJ, Groen-Klevant AC, Welschen R (1978) Follicle growth in the immature rat ovary. Acta Endocrinol (Copnh) 88:375–382 Ledda S, Bogliolo L, Leoni G, Naitana S (2001) Cell coupling and maturation-promoting factor activity in in vitro-matured prepubertal and adult sheep oocytes. Biol Reprod 65:247–252 Les Dees JK, Hiney NH, McArthur GA et al (2006) Origin and ontogeny of mammalian ovarian neurons. Endocrinology 147:3789–3796 Malamed S, Gibney J, Ojeda S (1992) Ovarian innervation develops before initiation of folliculogenesis in the rat. Cell Tissue Res 270:87–93 Marchal R, Feugang J, Perrau C et al (2001) Meiotic and development competence of prepubertal and adult swine oocytes. Theriol Int J Anim Reprod 56:17–29 McGee E, Hsueh A (2000) Initial recruitment of ovarian follicles. Endocr Rev 21:200–214 Méndez MC, Chávez B, Echeverría OM et al (1999) Evidence for estrogen receptor expression in germ cell and somatic cell subpopulations in the ovary of the new hatched chicken. Cell Tissue Res 298:145–152 Neely E, Hintz R, Wilson D et al (1995) Normal ranges for immunochemiluminometric gonadotropin assays. J Pedriatr 127:40–46 Ortiz R, Echeverría OM, Salgado R et al (2006) Fine structural and cytochemical analysis of the processes of the cell death of oocytes in atretic follicles in new born and prepubertal rats. Apoptosis 11:25–37 Peters H, Himelstein-Braw R, Faber M (1976) The normal development of the ovary in childhood. Acta Endocrinol 82:617–630

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Polkowska J, Wójcik-Gładysz A, Wańkowska M et al (2008) Prepubertal changes in the synthesis, storage and release of growth hormone and in the immunoreactivity of oestrogen receptor-a in lamb pituitary cells. J Chem Neuroanat 36(1):53–58 Rawlings NC, Evans AC, Honaramooz A et  al (2003) Antral follicle growth and endocrine changes in prepubertal cattle, sheep and goats. Anim Reprod Sci 78(3–4):259–270 Richards JS (1980b) Maturation of ovarian follicles: actions and interactions of pituitary and ­ovarian hormones on follicular cell differentiation. Physiol Rev 60:51–89 Tingen CM, Bristol-Gould SK, Diesewetter SE (2009) Prepubertal primordial follicle loss in mice is not due to classical apoptotic pathways. Biol Reprod 81:16–25 Tomic D, Brodie SG, Deng C et al (2002b) Smad3 may regulate follicular growth in the mouse ovary. Biol Reprod 66:917–923 Tomic D, Miller KP, Kenny HA et al (2004b) Ovarian follicle development requires Smad3. Mol Endocrinol 18:2224–2240 Vázquez-Nin GH, Sotelo JR (1967) Electron microscope study of the atretic oocytes of the rat. Z Zellforsch 80:518–533 Vázquez-Nin GH, Echeverría OM, Molina E et  al (1978) Effects of ovariectomy and estradiol injection on nuclear structures of endometrial epithelial cells. Acta Anat 102:308–318 Vázquez-Nin GH, Echeverría OM, Fakan S et al (1991) Immunolocalization of estrogen receptor on pre-mRNA containing constituents of rat uterine cell nuclei. Exp Cell Res 192:396–404 Villavicencio-Macías M (1991) Un estudio de los mecanismos que regulan el crecimiento y la diferenciación folicular en la rata prepúber: el papel de la hormona estimulante del folículo y sus isohormonas. Thesis for Master degree. ENEP Zaragoza, Universidad Nacional Autónoma de México Zachos NC, Billiar RB, Albrecht ED et  al (2002) Developmental regulation of baboon fetal ­ovarian maturation by estrogen. Biol Reprod 67:1148–1156

Part II

General Aspects of Cell Death

Chapter 4

Apoptosis María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

Abstract  Cell death is a fundamental physiological process involved in controlling the balance between proliferation and differentiation during embryonic development, and in the renewal of cellular tissue throughout adulthood. It is presently known that some alterations in this process modify the cellular behavior and can lead to different pathologies, such as certain cancers and neurodegeneration processes. Apoptosis is the best studied cell death, and it is considered as programmed type I cell death. This process is characterized by several morphological cellular changes where ­cellular shrinkage, chromatin condensation, nuclear fragmentation are included. The apoptosis is executed by a group of proteases, denominated caspases, which lead to the cellular characteristics of the process. List of Abbreviations APAF1 Bak Bax BH3 Bid CARD ced-3 DED DISC FADD ICE Mcl-1 MOMP

Apoptotic protease activating factor 1 Bcl-2 antagonist/killer Bcl-2-associated X protein Bcl-2-homology domain-3 BH3-interacting-domain Caspases recruitment domain Caenorhabditis elegans cell-death abnormality-3 Death effector domain Death-inducing signaling complex Fas associated death domain-containing protein Interleukin-1 converting enzyme Myeloid cell leukemia-1 Mitochondrial outer-membrane permeabilization

M.L. Escobar (*), G.H. Vázquez-Nin, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico, USA e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_4, © Springer Science+Business Media B.V. 2011

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PIDD p53-Indicible death domain PUMA p53-Upregulated modulator of apoptosis Smac/DIABLO Second mitochondrial activator of caspase/direct IAP-binding protein with low pI tBID Truncated BID TNFR TNF receptor TNF Tumor necrosis factor TRADD TNFR1-associated death domain TRAIL TNF-related apoptosis-inducing ligand XIAP X-linked inhibitor of apoptosis protein

4.1 General Characteristics of the Apoptosis Cell death is essential for the elimination of defective or out of use cells. This ­process is necessary during development for the removal of unnecessary cells and to maintain the homeostasis in several organs during adulthood life. The term apoptosis was introduced by Kerr and cols. (1972), they described a series of morphological cellular characteristics defining a type of cell death that is different from necrosis. The morphological features of apoptosis include cytoplasmic shrinkage, membrane blebbing, chromatin condensation (pyknosis), nuclear fragmentation (karyorrhexis), intranucleosomal DNA fragmentation (endonucleolysis), and the formation of apoptotic bodies with cytoplasmic content. During apoptosis the organelles are preserved and basically there is no autophagic degradation. Diverse characteristics of apoptosis are easily visible in light microscopy, such as the cell shrinkage and pyknosis during the early process of apoptosis (Kerr et al. 1972). Pyknosis is the result of chromatin condensation whereas cell shrinkage makes the cells smaller, the organelles strongly packed and the cytoplasm denser. As a consequence of the plasma membrane blebbing, the cell is fragmented into pieces named apoptotic bodies; this process is called “budding.” The apoptotic bodies contain cytoplasm with tightly packed organelles, which may or may not have nuclear material surrounded by an intact cytoplasm. These apoptotic bodies are phagocytosed by macrophages or neighboring cells (Kerr et al. 1972). An important and fundamental aspect of apoptosis is that the plasma membrane is conserved until the final process, a characteristic that confers to apoptosis the attribute that no inflammatory event takes place during the cellular death process (Savill and Fadok 2000; Kurosaka et al. 2003) (Fig. 4.1). Recently, several studies have provided important information concerning the proteins participating in the apoptotic process of cell death. Biochemically, apoptosis is associated with the proteases named caspases, the apoptosis is known as a caspase-dependent process. Two important features of the biochemical alterations are the loss of the cytoplasmic membrane asymmetry, which provokes the phosphatidylserine externalization, and the permeabilization of mitochondria, which is an important organelle in the apoptosis execution.

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Fig. 4.1  Morphological characteristic of the apoptosis process. At the left a schematic representation of different cellular alterations during apoptosis. The micrography shows the chromatin of nuclei of cultured cells labeled with DAPI. The arrow heads point to highly compacted nuclei. The arrow shows apoptotic bodies. Scale bar 50 mm

4.2 Caspases Apoptosis is executed by a series of proteases called caspases which are cysteinedependant aspartic acid-specific proteases. Their function is to cleave vital cellular substrates after aspartic acid. These proteases were first identified in Caenorhabditis elegans (Ellis and Horvitz 1986), and were known as cell-death abnormality-3 (ced-3) gene product. The first CED identified was CED-3, which shares sequences with the mammalian protease inlerleukin-1 (IL-1)-converting enzyme. Later this enzyme was renamed as caspase-1 (Alnemri et al. 1996). Now 14 caspases are known, 11 of which are present in human (caspase-1 to -10, and caspase-14). Although the members of caspase family share some characteristics, such as amino acid sequence and molecular structure, they have different functions; some of them are involved in the inflammatory process (caspase-1, caspase-4, caspase-5, caspase-11, caspase-13 and caspase-12), and others are involved in apoptosis cell death (caspase-2, caspase-3, caspase-6, caspase-7, caspase-8, caspase-9, and caspase-10). Caspase-14 is involved in the differentiation of keratinocytes. The caspases can be found in inactive form in the cytoplasm. They are synthesized as inactive precursors (zymogens) that become active proteases after their cleavage at specific sites within the molecule (Stegh and Peter 2001) via internal proteolysis or by induced proximity. Initiator caspases are activated through their

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Fig. 4.2  Graphic representation of the caspases activation. The inactive precursors (procaspases) become active proteases after their cleavage or dimerization

oligomerization, mediated by adaptor molecules; the proximity of protein-protein interaction domains of both the adaptor and the caspase allows the process. This oligomerization leads to autoproteolytic cleavage of the initiator caspases (Muzio et  al. 1998; Salvesen and Dixit 1999). The initiator caspases are activated by dimerization, and the executioner caspases are activated by cleavage (Fig. 4.2). The molecular structure of the caspases is remarkably conserved; they contain a prodomain at the N-terminus, followed by a catalytic protease domain at the C-terminus, which consists of a prodomain p20 large (of 20 kDa), and two identical p17 small subunits (10 kDa) (Chang and Yang 2000; Prior and Salvesen 2004). The activation of the zymogen precursor is mediated by a series of cleavage events, first separating the large and small subunits, followed by the removal of the prodomain (Ramage et al. 1995; Yamin et al. 1996). The active form of the caspases comprises a heterotetramer of two units, each of which is composed by one large-a and smallb subunits (Shiozaki et al. 2003). All caspases have a catalytic site consisting of a Cys285 and His237 (Stennicke and Salvesen 1999). These enzymes recognize tetrapeptide motifs and cleave their substrates on the carboxyl side of an aspartate residue. To catalyze the hydrolysis of the peptide substrate the caspases utilize a cysteine thiolate residue. The nucleophilic attack by the cysteine thiolate initially forms a tetrahedral intermediate that is stabilized by the enzyme active site. This is followed by elimination of the C-terminal portion of the peptide, leaving a thioester in the active site. Subsequent hydrolysis of the thioester releases the N-terminal portion of the substrate peptide and regenerates an available active site to reenter the catalytic cycle (reviewed in Fricker 2010). The apoptotic caspases have been divided into two groups: initiator caspases with long prodomains of over 100 amino acids, and executer caspases with short prodomains of usually less than 30 amino acids. The initiator caspases are responsible for the activation of the executor caspases; this group is composed of caspase-8, caspase-10, and caspase-9. The executer caspases have diverse cellular targets which are cleaved during apoptosis, inducing the morphological changes

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Fig. 4.3  Graphic characteristics of caspases zymogens. The caspases have been classified according their function and structure

during this process (Earnshaw et  al. 1999; Stegh and Peter 2001), this group is constituted by caspase-7, caspase-6, and caspase-3. The caspase-8 and caspase-10 possess long prodomains containing the characteristic protein-protein interaction motif, the death effector domain (DED). On the other hand, caspase-1, caspase-2, caspase-4, caspase-5, caspase-9, caspase-11 and caspase-12, have the caspase activation and recruitment domain (CARD), which provides the source for interaction with upstream adaptor molecules mediating the homophilic interaction between procaspase and their adapters (Fig. 4.3).

4.3 Bcl2 Proteins The B-cell lymphoma/leukemia-2 (Bcl-2) family, are proteins regulators of the mitochondrial pathway of apoptosis. The Bcl2 family is constituted by more than 30 members and is divided into three groups on the basis of their functions and the composition of their Bcl-2 homology (BH) domains (Adams and Cory 1998). All the Bcl-2 family members share at least one of four relatively conserved BH domains. The proteins of Bcl-2 family have been classified as: pro-apoptotic, Bcl2- homology domain-3 (BH3)-only (BH3-only), and anti-apoptotic. The BH3-only belong to the pro-apoptotic type (Fig. 4.4).

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Fig. 4.4  Bcl-2 family proteins. The Bcl-2 family members share at least one of four BH domains. This family has the pro-apoptotic, BH3-only, and anti-apoptotic proteins

The pro-apoptotic members are Bcl-2-associated X protein (Bax), Bcl-2 antagonist/ killer (Bak), and Bcl-2-related ovarian killer (Bok). All of them share three domains (BH1, BH2, and BH3). They also have a similar 3D structure, namely a globular bundle of a-helices with a hydrophobic surface groove (Fesik 2005; Suzuki et al. 2000). The multi-domain proteins Bax and Bak promote the formation of pores in the mitochondrial external membrane, which enables the release of apoptogenic ­factors such as cytochrome-c. The BH3-only proteins are a subgroup of the Bcl-2 ­family, comprising at least 12 members: Bcl-xL/Bcl-2 associated death promoter (Bad), Bcl-2-interacting mediator of cell death (Bim), Bik/Nbk/Blk, BH3-interacting domain death agonist (Bid), Bcl2-modifying factor (Bmf), p53 upregulated modulator of apoptosis (Puma), Noxa, Spike, BNIP3, Nix, Hrk/DP5 and Beclin-1 (Giam et  al. 2008). These BH3-only proteins contain only one homology domain. The BH3-only promote activation of these multi-domain family members upon an apoptotic insult which is sufficient for their killing action. This amphipathic a-helix of 16 residues binds to the hydrophobic surface groove of anti-apoptotic members to inhibit apoptosis (Sattler et al. 1997; Liu et al. 2003). Both types of pro-apoptotic

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proteins are necessary to exert the cell death, because the BH3-only firstly antagonize the Bcl-2-like protein (Huang and Strasser 2000), and Bax or Bax is necessary for organelle permeation (Cheng et al. 2003; Wei et al. 2001). BH3-only proteins serve to sequester the antiapoptotic proteins or to bypass them and directly activate Bak/Bak proteins (Willis et al. 2007; Kroemer et al. 2007). The inactive form of Bax exists as a soluble protein in the cytosol, associated with cytosolic retention factors; Bak is a resident protein of the mitochondria outer membrane forming an inactive complex with voltage-dependent anion-selective channel protein 2 (VDAC2) (Lucken-Ardjomande and Martinou 2005). The proapoptotic Bcl-2 members Bax and Bak are activated when they are released of their sequestration by anti-apoptotic Bcl-2. These proteins have a direct interaction with a subset of BH3-only proteins Bim, truncated Bid (tBid) and Puma. During apoptosis, Bax undergoes a number of changes, including translocation to mitochondria, a conformational change exposing hidden epitopes within the N-terminus, and its assembly into high molecular weight complexes (Valentijn et al. 2003). The proapoptotic Bcl-2 proteins directly impact on mitochondrial outer-membrane permeabilization (MOMP), through the Bax and Bak conversion from inactive monomers to oligomeric transmembrane structures (Upton et  al. 2007); these proteins oligomerize through direct or indirect activation by the domain BH3-only members. The anti-apoptotic Bcl-2 family members, Bcl-2, BclxLong (Bcl-xL), Bcl-w, Bf1/ A1, and myeloid cell leukemia-1 (Mcl-1), contain up to four Bcl-2-homology (BH)domains and localize mainly to intracellular membranes such as endoplasmic reticulum and mitochondria; these proteins promote the cell survival (Cuconati and White 2002). These proteins are capable of preventing the release of apoptogenic factors, because they contain a hydrophobic cleft that enable them to bind to BH3only proteins and to the pro-apoptotic Bcl-2 family members Bad, Bak, and Bax to inhibit apoptosis (Liang and Fesik 1997).

4.4 Extrinsic Route of Caspases Activation In mammals, two different routes of apoptosis activation have been described: the extrinsic mediated, via a death receptor; and the intrinsic where the mitochondria are a key player. The extrinsic route implies the participation of death receptors (DR) located in the cytoplasm membrane and the caspase-8 and/or the caspase-10. The caspase-8 is the most upstream caspase in the death receptor signaling pathway and its activation is brought about as a result of the adaptor-mediated multimerization. The DRs are activated by extracellular ligands; the most common of these ligands are the tumor necrosis factor-a (TNF-a), Fas ligand (FasL or CD95L/Apo1), TNF-like WEAK inducer of apoptosis (TWEAK), and TNF-related apoptosis-inducing ligand (TRAIL). The DRs belong to the TNF receptor (TNF-R) super-family, as Fas/CD95, TNF-R1 (p55/CD120a), TRAMP/Apo-3/DR3/WSL-1/LARD/, the TRAIL receptors DR4/TRAIL-R1 and DR5/TRAIL-R2/TRICK2/KILLER (Ashkenazi and Dixit 1998).

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These receptors are localized in the lipid rafts (Nagata and Goldstein 1995; Nagata 1997; Kramer 2000) and contain death domains (DD) and death effector domains (DED) capable of homophilic protein-protein interactions. The death receptors share multiple cysteine-rich repeats in their extracellular ligand-binding domain, a type I (single pass) transmembrane domain, and a cytoplasmic death domain required for intracellular signaling (Schneider and Tschopp 2000). The DD acts as the recruitment site for adaptor proteins whose role is to initiate the various downstream signaling pathways governed by these receptors. The binding on the cell surface of the death receptor with their specific ligand recruits the adaptor proteins as TNF receptor-1-associated protein (TRADD) and Fas associated via death domain (FADD or MORT1). This complex is able to recruit caspase-8 and caspase-10 through their death effector domain (DED) forming a high molecular weight multiprotein complex known as the death-inducing signaling complex (DISC) (Muzio et  al. 1996; Sandu et al. 2006). DISC is a signaling complex that contains multiple adaptor molecules as well as FADD and TRADD, as DAXX, receptor interacting protein kinase (RIP), RIP associated protein with a DD (RAIDD) and FLICE-like inhibitory protein (FLIP). Receptor assembly into oligomers is triggered by ligand binding, resulting in the formation of the DISC through conformational alterations. DISC constitutes a subset of cell surface receptors of the TNF-R superfamily. DISC acts as a cellular controller and in presence of stimulus it begins the oligomerization of its constituents to form the active oligomeric platform, in which the ligand binding occurs. The entire process of recruitment and oligomerization of caspase-8 in the DISC results in the autocatalytic activation of caspase-8 and/or caspase-10, which subsequently activate downstream caspases directly (Fig. 4.5).

Fig. 4.5  Extrinsic route of activation of apoptosis. The cellular receptor of death is activated by its ligand in the cytoplasmic membrane. This event activates the intracellular upstream signalization of caspases conducing to the apoptosis

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4.5 Intrinsic Apoptosis Activation Intrinsic apoptotic signaling can be activated by several stimuli, such as DNA ­damage and cytotoxic drugs, provoking mitochondrial damage. Mitochondria are comprised of a matrix surrounded by the inner membrane, the intermembrane space and the outer membrane. The inner membrane contains molecules such as ATP synthase, electron transport chain, and adenine nucleotide translocator. Under normal physiological conditions these molecules allow the respiratory chain to create an electrochemical gradient and thus a membrane potential (reviewed in Rupinder et al. 2007). In the intrinsic apoptosis route, the mitochondria play an essential function. Several signals arrive to the mitochondria which respond by releasing diverse proteins. In this route the Bcl-2 family members have an important function. The pro-apoptotic Bcl-2 as Bax and Bak, promote the permeabilization of the mitochondrial outer membrane, and the subsequent release of apoptogenic factors, as cytochrome-c, Smac/DIABLO, high temperature requirement protein A2 (Omi/HtrA2), and apoptosis-inducing ­factor (AIF) into the cytosol (Gross et al. 1999) (Fig. 4.6). Cytochrome-c is released from the mitochondrial intermembrane space, causing apoptotic protease activating factor 1 (Apaf-1) oligomerization resulting in apoptosome formation. This complex, in turn, recruits and activates procaspase-9, which then activates executioner caspase-3 and caspase-7. When the cytochrome-c leaves the mitochondria its free form in the cytoplasm can interact with Apaf-1. Apaf-1 preexists in the cytosol as a monomer of 130-kDa consisting of a multidomain protein that is formed of three functional regions: an

Fig. 4.6  Intrinsic route of activation of apoptosis. When the cell recipes the pro-apoptotic insult, the mitochondria responds releasing cytochrome-c, to form the apoptosome complex

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N-terminal caspase-recruitment domain (CARD), a nucleotide-binding domain and an oligomerization domain (NOD/NB-ARC). The C-terminal domain is composed of 12–13 WD40 repeats (from different spliced forms), a motif that mediates ­protein-protein interactions (Ried et al. 2005). Apaf-1 CARD functions to recruit pro-caspase-9 via its CARD, and to oligomerize into a heptameric scaffold (Acehan et al. 2009). The WD40 repeats are organized into two domains and are responsible for the binding of cytochrome-c. Between the CARD and the WD40 repeats, there is a region constituted by three domains denominated NOD/NB-ARC domains, which is at the centre of oligomerization and thus, of the site of apoptosome formation (Ried et al. 2005). In the Apaf-1 monomeric form the WD40-repeat domains maintain to Apaf-1 in an autoinhibited state. Once free, the cytochrome-c binds to the WD40-repeat region of Apaf-1 and it induces a conformational change in Apaf1, provoking that the lock in the autoinhibited form of Apaf-1 is released, thus allowing the apoptosome formation. The entire process previously described results in the formation of the apoptosome, a heptamer comprised of seven Apaf-1 adaptor molecules, each bound to one molecule of cytochrome c and a dimer of the initiator caspase-9, with a size of nearly 1 megadalton. The apoptosome form a symmetrical wheel-like structure; in this complex Apaf-1 interacts with the adjacent Apaf-1 molecules via their N-terminal CARD domains to form a central hub region, and the C-terminal WD40 repeats are extended to form the outside ring. Unlike the others caspases, the caspase-9 is activated primarily by allosteric change and dimerization rather than cleavage (Rodriguez and Lazebnik 1999; Boatright et al. 2003). The formation of the apoptosome is a multistep process that requires the presence of either 2¢-deoxy ATP (dATP) or ATP and it is simultaneous with major conformational changes in Apaf-1 (reviewed in Ried and Salvesen 2007). Finally the apoptosome activates the executer caspases -3 and -7, which initiate their proteolytic activity. Caspase-2 also is activated sequentially. To active this caspase, the assembly of a large protein complex, which has 670 kDa molecular weight, is necessary (Tinel and Tschopp 2004; Read et al. 2002). In this complex an adaptor protein RAIDD, and the p43-induced protein with a death domain (PIDD) are present. Caspase-2activating complex is named PIDDosome (Tinel and Tschopp 2004). The pro-caspase-2 has a CARD that facilitates the dimerization of procaspase-2. The interaction of the protein RAIDD with CARD is an event that triggers the pro-caspase-2 activation. Caspase-2 is recruited into a protein complex similar to the Apaf-1/caspase-9 apoptosome (Tinel and Tschopp 2004).

4.6 Crosslinking Between Extrinsic and Intrinsic Activation The caspase-8 was originally called FADD-like Interleukin-1b-converting enzyme (FLICE) (Enari et al. 1995). It has the singular characteristic that once activated can initiate the mitochondrial signaling pathway, cleaving the cytosolic BH3interacting domain death agonist (Bid). Active caspase-8 cleaves Bid at Asp59, yielding a p15 C-terminal truncated fragment, generating a truncated Bid (tBid).

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Fig. 4.7  Crosslinking between extrinsic and intrinsic apoptosis activation processes

tBid is then able to directly activate pro-apoptotic multi-domain proteins to induce mitochondrial outer-membrane permeabilization (MOMP), inducing a conformational change of the multi-domain Bcl-2 family proteins Bax and Bak, which allows their insertion into the outer mitochondrial membrane. The permeabilization of the outer membrane provokes the release of apoptogenic factors, including cytochrome-c, Smac/DIABLO, OMI/HTR2A, and apoptosis-inducing factor (AIF) into the cytosol (Gross et al. 1999) (Fig. 4.7).

4.7 Apoptosis-Inducing Factor (AIF) The mitochondrial membrane permeabilization promotes the release of caspase independent death effectors like apoptosis-inducing factor (AIF) (Susin et  al. 1999), endonuclease G (EndoG), as well as HtrA2, which also possesses a serine protease activity (Ravagnan et  al. 2002), establishing the pathway “caspaseindependent death”. AIF is a mitochondrial flavoprotein, which in healthy cells is confined to the mitochondrial intermembrane space with its N-terminal region oriented towards the matrix and the C-terminal to the intermembrane space (Otera et al. 2005). It is a mitochondrial flavin adenine dinucleotide (FAD)-dependent oxidoreductase that plays roles in oxidative phosphorylation and redox control (Modjtahedi et al. 2006).

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AIF displays NAD(P)H oxidase as well as monodehydroascorbate reductase activities (Miramar et al. 2001). AIF is synthesized as a 613 amino acid (67 kDa) precursor; the mammalian AIF precursor contains an N-terminal mitochondrial localization sequence (MLS, residues 1–100) and a large C-terminal part (residues 121–610) (Susin et al. 1999). The precursor is imported into mitochondria via its N-terminal prodomain and then processed to a 62 kDa mature protein (Susin et al. 1999; Otera et al. 2005). The mature form of AIF (57 kDa) is generated by cleaving of the MLS, after import into the mitochondrial intermembrane space; this activation is made by calpains and/or cathepsins (Otera et al. 2005). Once AIF is cleaved by calpains and/or cathepsins the pro-apoptotic truncated AIF (tAIF) is formed, becoming soluble and then released. The AIF activation by cathepsins is a calcium dependent process; unlike the activation via calpains which is calcium independent. After an apoptotic insult, the mitochondrial outer membrane is permeabilized and AIF translocates to the cytosol and the nucleus, via its C-terminal domain nuclear localization sequence (NLS). In the nucleus AIF is associated to peripheral chromatin condensation, as well as high-molecular-weight (50 kbp) DNA fragmentation (reviewed in Candé et al. 2002) is a hallmark of caspase-independent apoptosis. AIF cytosolic/nuclear transit is regulated by Heat Shock Protein-70 and cyclophilin A (CypA) (Gurbuxani et al. 2003; Zhu et al. 2007). Cyp A is a peptidylprolyl cis-trans isomerase that has been implicated in the DNA degradation during apoptosis cell death (Montague et al. 1997). Five isoforms of AIF have been characterized: AIF, AIF2/AIF-exB, AIFsh (AIF short), AIFsh2 (AIF short 2), and AIFsh3 (AIF short 3). AIF2/AIF-exB has a regulation and sub cellular distribution similar to AIF (Loeffler et al. 2001). Other isoforms are restricted to different spaces: AIFsh1 is in the cytoplasm and AIFsh2 located to the mitochondria. During the cell death process only AIF and AIFsh participate in large-scale DNA fragmentation (Delettre et  al. 2006). AIF becomes activated in response to different insults that induce its release from the mitochondria. AIF activation provokes nuclear DNA degradation, as has been reported, under treatment with nitrosoureas, AIF interacts with the histone H2AX (Artus et al. 2010)

4.8 Inhibitor of Apoptosis Proteins (IAPs) The IAP family of proteins has the ability to protect cells from death. Highly conserved baculoviral IAP repeat (BIR) is a zinc-binding fold of approximately 70 amino acid residues that mediates protein-protein interactions (Sun et  al. 1999). BIR is found in 1–3 copies in every IAP. It is the domain that mediates apoptosis inhibition, through direct binding of caspases. In addition, mammalian IAPs as X-chromosome-linked inhibitor of apoptosis protein (XIAP), cIAP1 and cIAP2, possess a C-terminal RING domain. This structural motif confers E3-ubiquitin ligase activity to IAPs (Yang et al. 2000), allowing them to target numerous substrates, including IAPs themselves, caspases, IAP antagonists, and other cellular proteins, for poly-ubiquitylation. Moreover the previous characteristics, cIAP1 and

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Fig. 4.8  Graphic representation of IAP proteins family

cIAP2 contain a CARD. The BIR domains can be grouped into type I domains, with presence of a deep peptide-binding groove, and type II domains, without this groove (Lu et al. 2007) (Fig. 4.8). The XIAP structure is comprised of three baculoviral IAP repeat domains (BIR1-3), and a zinc-finger RING domain. This caspase inhibitor exerts its function at physiological concentrations. The active caspase-3 and caspase-7 are inhibited by XIAP via its adjacent BIR2 region (Vaux and Silke 2005). In contrast, the XIAP BIR3 domain is required for inhibiting caspase-9 (Vaux and Silke 2005). The XIAP BIR3 binds to the homodimerization surface of caspase-9; in this manner XIAP interferes with caspase-9 dimerization. Mitochondrial permeabilization, through pores formed by activated Bax or Bak, allows the release of factors that antagonizes the IAPs function. In mammals, the IAPs antagonists, as the second mitochondria derived activator of caspases (Smac)/ direct IAP binding protein with low PI (DIABLO), Omi/HTRA2, or GSPT1/eRF3, are constitutively expressed and are sequestered into their site of expression until they receive a death stimulus. Both Smac and Omi contain IAP-binding motifs (IBMs) through which they interact with IAPs, thereby sequestering them and ensuring that cells in fact undergo apoptosis (Eckelman et al. 2006). These antagonists are binding to the IAP BIR domains, blocking their access to caspases (reviewed in Gyrd-Hansen and Meier 2010). Smac/DIABLO is the principal antagonist of XIAP; this antagonist may occlude the binding sites of XIAP in the caspase-3, caspase-7 and caspase-9 (Scott et al. 2005). Survivin (16.5 kDa) is the smallest member of the IAP family, containing a single baculovirus IAP repeat motif. When the Survivin is activated, it is exported into the cytoplasm where it inhibits the caspases.

4.9 Execution Phase All of the proteolytic activation described above leads to the different morphological changes produced during apoptosis. The proteolytic cleavage of cellular substrates by effector caspases is responsible for the apoptotic hallmarks, such as chromatin condensation, plasma membrane asymmetry and cellular blebbing.

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The hallmark of apoptosis is DNA fragmentation into pieces with lengths c­ orresponding to multiple integers of approximately 180 base pairs (Wyllie 1980). This characteristic is generated by the activation of nucleases as caspase-activated DNase (CAD/DFF40). This nuclease is present in living cells, bound to its inhibitor of CAD (ICAD/DFF-45). CAD is activated by caspase-3 and caspase-7 via cleavage of ICAD, resulting in the release of CAD (Enari et al. 1998). Activated CAD translocates to the nucleus where it exerts its DNase function. The lamins and cytoskeletal proteins, such as fodrin and gelsolin, are cleaved by executer caspases, leading to morphological changes in apoptotic cell as nuclear shrinkage and fragmentation (Buendia et  al. 1999). The cytoskeleton cleavage causes the loss of overall cell shape; the cleavage of the components of the focal adhesion complex leads to detachment of apoptotic cells from their neighbors and from the basement membrane (Wen et al. 1997). Plasma membrane blebbing results from the caspase-mediated activation of gelsolin, an actin depolymerizing enzyme (Kothakota et al. 1997). Caspase mediated cleavage of PAK2, a member of the p21activated kinase family, participates in the formation of apoptotic bodies (Rudel and Bokoch 1997). Actin undergoes several modifications that lead to cell detachment (Levee et al. 1996; Celeste Morley et al. 2003). The cellular contractility has also been related to RHO-associated coiled-coil containing protein kinase (ROCK) (Tosello-Trampont et  al. 2003). The kinase activity of ROCK influences the status of myosin light chain (ML C) phosphorylation; this kinase activation is RHO-GTP dependent (Riento and Ridley 2003). The apoptotic cell death process is accompanied by a change in plasma membrane structure which leads to the exposure of phosphatidylserine (PS) on its external surface, while the membrane integrity remains unchallenged. Surface exposed PS can be detected by its affinity for Annexin-V. Annexin-V belongs to a family of phospholipid-binding proteins that include 12 members in vertebrates. The Annexin family has the ability to bind phospholipids in a reversible and calcium dependent manner. Annexin-V has a high affinity to bind to the negatively charged phospholipid PS. The PS is actively localized on the inner leaflet of the plasma membrane in healthy cells. The cell apoptosis initiation induces the rapid externalization of PS on the outer leaflet of the plasma membrane as a response to increased intracellular calcium. PS exposure on the outer leaflet of the plasma membrane can also be recognized by phagocytes as a signal for engulfment.

4.10 Other Routes of Apoptosis Activation The T-lymphocytes under cytotoxic environment activate a Fas mediated route to activate the apoptosis via Fas-FasL binding. This binding provokes the target cell membrane to become perforated by perforins, and subsequently the protease granzyme B is released in the cytoplasm. Granzyme B is an enzyme that splices proteins after the amino acid aspartate in the same way as caspases, resulting in a caspasecascade with the characteristic morphological changes (Martin and Green 1995).

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The endoplasmic reticulum stress pathway has been attributed to the activation of caspase-12 (Nakagawa et al. 2000), and in this sense it has been proposed that Bcl-2 functions on the endoplasmic reticulum as well as on mitochondria under cytotoxic stimuli, activating Bax and Bak to permeabilize the outer membrane of the mitochondria and the endoplasmic reticulum. As caspase 12 resides on the cytoplasmic face of the endoplasmic reticulum (Nakagawa and Yuan 2000; Nakagawa et al. 2000), it is possible that the activation of the caspase-12 is triggered at the level of the endoplasmic reticulum. Poly(ADP-ribose) polymerase (PARP) is an abundant nuclear protein that catalyses poly(ADP-ribose) ligation to acceptor proteins, including itself, in response to DNA strand breaks. The Poly(ADP-ribosylation) is a posttranslational modification that plays a relevant role in DNA repair. PARP-1 is a nuclear enzyme that rapidly binds to DNA strand breaks leading to the formation of long, branched poly(ADP-ribose) polymers using NAD+ as substrate. The resulting negatively charged PARP-1 is subsequently dissociated from DNA ends, facilitating the DNA repair process (Kim et al. 2005). PARP-1 is a caspase-3 and caspase-7 substrate that is converted from the 116-kDa form to fragments of 89 and 24 kDa (Kaufmann et al. 1993). PARP-1 is constituted by three functional domains: (a) the N-terminus a DNA binding domain (DBD, 46  kDa), which contains a bipartite nuclear localization sequence (NLS) and two zinc fingers (DBD is responsible for DNA strand breaks recognition); (b) the central region of the protein is the auto modification domain (22 kDa), which mediates the PARP-1 autoribosylation; (c) the C-terminus, contains the NAD-binding domain (54 kDa), which is essential for the conversion of NAD1 into ADP-ribose. Caspases 3 and 7 cleave PARP-1 in Asp-Glu-Val-Aspaldehyde (DEVD) site within the DBD, thus splitting the NLS. Apoptosis cell death is a highly regulated process and its morphological and biochemical characteristics are strongly related, since the pro-apoptotic stimulus lead the molecular machinery activation to eliminate damaged or altered cells.

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Chapter 5

Autophagy María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

Abstract  Autophagy is a physiological process conserved by evolution for the recycling of cell constituents, as organelles or fragments of cytoplasm. Autophagy does not cause loss of cell chemical components as the cell reutilizes them. The lysosomes are acidic vesicles delimited by a single membrane formed in the Golgi apparatus. They contain acid hydrolases for degradation of proteins; lipids, nucleic acids and glucids, and they fuse with diverse types of phagocytic vesicles. Autophagosomes contain undegraded cytoplasmic ground substance and recognizable organelles. The transformation of an autophagosome in an autolysosome occurs by the acquisition of lysosomal membrane proteins and the delivery of acid hydrolases. The autophagy mechanism is considered as a protective system; organisms in starving conditions, autophagy is a procedure to reduce metabolic demand by the cell and thus avoid death. On the other hand, autophagic cell death is a form of programmed cell death morphological and metabolic different from apoptosis. Autophagic cell death is accompanied by cytoplasmic vacuolization and activation of lysosomal enzymes. Due to the absence of a rupture of cell membrane and leak of cytoplasmic content, this cell death program is characterized by the lack of a tissue inflammatory response.

List of Abbreviations TOR TSC TOR TSC RER Atg

Target of rapamycin Tuberous sclerosis complex Target of rapamycin Tuberous sclerosis complex Endoplasmic reticulum AuTophaGy-related

M.L. Escobar (*), G.H. Vázquez-Nin, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico, USA e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_5, © Springer Science+Business Media B.V. 2011

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Phosphatidylinositol 3-OH kinase Vacuolar protein sorting 34 Phosphatidylethanolamine Light chain 3 microtubule-associated protein 1 Mitochondrial permeability transition AMP-activated protein kinase Preautophagosomal structure (or phagophore assembly site) Phosphatidylinositol-4phosphate Phosphatidylinositol-4, 5-bisphosphate Phosphoinositide-dependent kinase-1

5.1 Autophagy Process Autophagy is a physiological process conserved by evolution for the recycling of cell constituents, as organelles or other cell constituents or fragments of cytoplasm. Actually, autophagy is the main route for degradation of cell constituents by proteasomes. This process is especially significant during development and under certain conditions of stress (Klionsky and Emr 2000). Autophagy does not cause loss of cell chemical components as they are reutilized by the cell. For instance, the breakdown of proteins by autophagy produces amino acids and other elements needed for intermediary metabolism and for biosynthetic pathways. In this way eukaryotic cells degrade parts of their cytoplasm and organelles (Dunn 1994). Defective proteins are also eliminated in this way. Moreover, autophagy is the primary intracellular catabolic mechanism for degrading and recycling long-lived proteins and organelles. Autophagy is involved in different events of physiological processes such as: tissue remodeling during development and differentiation, production of amino acids when nutrients fall short and elimination of unwanted and damaged organelles and molecules. These processes are likely to be an important function during the span of adult life (Meijer and Codogno 2004). In mammals, autophagy has been proposed as an anti-ageing mechanism for eliminating organelles damaged by the age-dependent peroxidation of molecules (Bergamini et  al. 2003). As autophagy and most of the lysosome-mediated degrading processes decline during ageing, keeping autophagy active would be important to delay ageing (Cuervo and Dice 2000). The deregulation of autophagy has been proposed to play a role in certain diseases, including cancer, cardiomyopathy, muscular diseases, and neurodegenerative disorders (reviewed in Levine and Klionsky 2004). These pathological processes are mainly due to the accumulation of autophagic vacuoles in the cytoplasm of the cells because of autophagic processes lacking the fusion of autophagosomes with lysosomes. In cancer, autophagy may function as an adaptive mechanism allowing cancer cells to overcome limitation in nutrient supply. In this way the process of autophagy may function as a promotor of tumorigenesis. However, the stimulation of autophagy in cancer cells was observed in response to anti-cancer

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Fig. 5.1  Dual roles for autophagy in cancer disease. The function of autophagy depends on the stimulus, environment and cellular response

treatments (Bursch et al. 2000; Inbal et al. 2002). It remains to be firmly established whether autophagy is a protective mechanism or if it is directly involved in cell death mechanisms to substitute the apoptotic programme that is frequently altered in ­cancer cells (Fig. 5.1). In insects one of the principal functions of autophagy is its participation in the development of the insect removing larval muscles and salivary glands during the metamorphosis process (reviewed in Lockshin and Zakeri 2004). The morphological traits of autophagy were described for the first time in mammals. Cells in the early stages of autophagy contain several autophagic vacuoli, and both the nucleoplasm and the cytoplasm appear slightly darkened, although nuclear structure still appears normal. Mitochondria and the endoplasmic reticulum are sometimes dilated, and the Golgi apparatus is often enlarged. The plasma membrane loses specializations such as microvilli and junctional complexes. In several cases an intense endocytosis is observed. During late stages, both the number and size of vacuoli increase, and many of them contain myelin figures or are filled with lipids, appearing as pale gray inclusions in the cytoplasm (Clarke 1990) (Fig. 5.2). The nucleus of a cell undergoing autophagic cell death can become pyknotic, identifiable by light microscopy either in early or in late stages of the degenerative process. This nuclear condensation is neither as common nor as remarkable as that of apoptosis. The late autophagic cell debris are frequently removed by heterophagy, but this tends to occur in very late stages and seems to be less conspicuous than the clearance of apoptotic bodies (Clarke 1990). A reorganization of intracellular membranes and high lysosomal activity was also observed. Like other lysosomal compartments, autophagic vacuoles possess an acidic pH which is generated by a V-type H+-ATPase (Al-Awqati 1986). Autophagy process includes at least four steps: the induction and the formation of the autophagosome, fusion of the autophagosome with the lysosome or vacuole, and autophagic vacuole breakdown (Klionsky and Emr 2000) (Fig. 5.3). The process of autophagy involves the participation of lysosomes to degrade the content of autophagy vesicles. In most eukaryotic cells the lysosomes and in yeast the vacuoles are the main degrading organelle. The lysosomes are vesicles formed in the Golgi apparatus. They

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Fig. 5.2  Ultrastructural features of autophagic cell death of an oocyte of a 28 days old rat. The cytoplasm contents numerous autophagic vacuoles with cytoplasmic contents in different degree of degeneration

Fig. 5.3  The autophagosomal pathway

contain acid hydrolases and fuse with diverse types of phagocytic vesicles in meeting points where different intracellular currents converge (González-Noriega 2003). The hydrolases are formed in the rough endoplasmic reticulum (RER) and they pass through the Golgi apparatus. Lysosomes are acidic organelles delimited by a single membrane, containing a characteristic set of acid hydrolases for degradation of proteins, lipids, nucleic acids and glucids arriving through endocytosis, autophagy and phagocytosis (Novikoff 1961). During endocytosis, molecules are incorporated in early endosomes and some of these molecules are recycled to the cell membrane, while others are incorporated in late endosomes. The lumen of these endosomes is at pH 6 and contains lysosomal hydrolases originated in the Golgi complex. They receive lysosome enzymes from the biosynthetic pathway and become lysosomes. The optimum pH for the catalysis of these enzymes is about 4.6. The low pH is maintained by a proton transporter (a H+-ATPase) located in the membrane of the organelle. Lysosomes are responsible for the degradation of internalized material from the endocytic and autophagic pathways.

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The lysosomal membrane has several functions: the acidification of the interior of the lysosome, the uptake of lysosome enzymes (Mellmann et al. 1986), the transport of degradation products from the lumen to the cytosol and the fusion between lysosomes and between lysosomes and other organelles (Fukuda 1991; Peters and von Figura 1994). The lysosomal membrane contains several N-glycosylated proteins, among which the better characterized are lamp1 and lamp2, which are structurally related and evolved from a common ancestral gene (Granger et al. 1990). They are the major components, constituting nearly 50% of the proteins of lysosomal membrane, according to estimations based on immuno-purification (Chen et  al. 1985; Marsh et al. 1987). These are transmembrane proteins type I, with a large luminal domain, a transmembrane domain and a cytoplasmic domain containing the carboxy terminus. The cytosolic polypeptide chain conserved in both proteins is composed by 11 amino acids and contains the information for the cytosolic location of the lysosome (Hunziker et al. 1996) (Fig. 5.4). In spite of the homology, 37% of aminoacids lamp1 and lamp2 are distinct proteins that diverged early in the evolution as ­suggested by the localization of their genes in different chromosomes (Fukuda 1991). Autophagocytosis begins with formation of autophagosomes, which then mature into autolysosomes by acquiring degradative activity. The process of autophagy has different routes for the elimination of cytoplasmic components: macroautophagy, microautofagia and ubiquitin-proteosomal system (Mizushima 2007). These ­processes differ in the mechanisms for transportation of substrates into the lysosome, in their regulation and in their selectivity. The macroautophagy, shortly autophagy, and the microautophagy are processes conserved from yeast to mammals, while the ubiquitin-proteosomal system is present only in mammals.

Fig. 5.4  Immunolocalization of lamp1 in adult rat oocytes. Note the abundant signal in the cytoplasmic region of the oocyte (arrows)

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5.2 Macroautophagy The sequestration of organelles and cytosol for subsequent degradation within the lysosomal vacuolar system occurs by macroautophagy. Degradation of bulky cell constituents by the lysosome/vacuole system occurs via macroautophagy; in the initial step of macroautophagy, a membrane structure known as the ‘isolation membrane’ is formed. This structure grows to form a double-membraned vesicle containing various cytosolic proteins, as well as cytoplasmic organelles in the lumen of double-membrane autophagosomes. These autophagosomes then mature, perhaps fusing with endosomes, before fusing with lysosomes. Autophagosomes then fuse with endosomes or lysosomes to become mature, single-membraned autolysosomes (Fig.  5.5a). Acidification of the lumen and acquisition of lysosomal hydrolytic enzymes enable this specialized membrane system to degrade sequestered cytoplasmic components (Ueno et  al. 1999). The acidic hydrolases of the lysosome then effect degradation of the contents of the autophagosome, resulting in amino acids and fatty acids that can then be either metabolized for energy production or ­recycled into biosynthetic pathways (Eskelinen 2005).

Fig. 5.5  Schematic drawing of different autophagic pathways. (a) macroautophagy, (b) microautophagy, (c) chaperone-mediated autophagy system

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5.3 Microautophagy In microautophagy relatively small portions of cytoplasm (e.g. glycogen and ­ribosomes) are sequestered by an invagination of the lysosome membrane, by the wrapping of a flap-like protrusion, and/or septation of the lysosomal limiting membrane. This process results in membrane bound intralysosomal vesicles whose contents are degraded by lysosomal hidrolases (Fig.  5.5b), and is regulated by ­environmental factors (reviewed in Dunn 1994).

5.4 Chaperone-Mediated Autophagy System Autophagy mediated by chaperones is activated by different agents causing stress, such as starvation, exposition to toxins and oxidative stress. In chaperone-mediated autophagy, proteins with a specific sequence signal are transported from the cytoplasm through the lysosomal membrane to the lysosomal lumen (Cuervo and Dice 1996). These proteins are selectively transported after interacting with chaperone hsc70, which is a 70 kDa member of the thermal shock proteins family (Fig. 5.5c). The interaction of hsc70 with the substrate takes place by means of pentapeptide KFERQ, which is present in all substrates that take this route (Dice 1990). The complex substrate-chaperone binds a lysosome membrane receptor, lamp2A or a similar protein associated to the lysosomal membrane. After unfolding, the substrate goes through the lysosomal membrane assisted by a ­chaperone lying in the lumen of the lysosome (Cuervo and Dice 1996).

5.5 Atg Genes The molecular components of autophagy were initially described in yeast. Different approaches led to the identification of the autophagy genes and the nomenclature of these genes has been unified, beening termed Atg genes (AuTophaGy-related). The Atg genes were identified first in the yeasts Saccharomyces cerevisiae, Hansenula polymorpha, and Pichia pastoris and many of these genes have their candidate orthologs in higher eukaryotes. Among these Atg genes, one subset is required for autophagosome formation in all subtypes. Atg genes were discovered using yeast mutants defective in autophagy. Atg genes are required for the activation of the ­signalization complex that initiates the formation of the autophagosome. The first mammalian autophagy genes Atg5 and Atg12 were identified by Mizushima et al. (1998a, b). Another critical step for autophagy analysis in higher eukaryotes was the identification of the mammalian Atg8 homologue MAP1LC3 (LC3), and the subsequent development of LC3-based assays for monitoring autophagy in mammals and other higher eukaryotic systems.

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Beclin 1 is the mammalian orthologue of yeast Atg6. Beclin 1 localizes to the trans-Golgi network, it belongs to the class III phosphatidylinositol 3-kinase complex and participates in the early stages of autophagosome formation, promoting the nucleation of autophagic vesicles (Kihara et al. 2001). This complex plays a key role in increasing the size of pre-autophagosomal membranes and in their biogenesis by recruiting proteins from the cytosol. Beclin 1 was originally discovered as a Bcl-2-interacting protein (Liang et  al. 1998) and was the first human protein shown to be indispensable for autophagy (Liang et al. 1999). The overall structure of Atg6/Beclin 1, as well as its essential role in autophagosome formation, is evolutionarily conserved throughout all eukaryotic phyla. The autophagy function of the Beclin 1-class III PI3K complex is activated by UVRAG, Ambra1, and Bif-1, and inhibited by Bcl-2 and Bcl-xL (Levine and Kroemer 2008). The core machinery of autophagy is composed of three major functional groups: –– Atg9 and its cycling system, which includes Atg9, the Atg1 kinase complex (Atg1 and Atg13), Atg2 and Atg18. –– The phosphatidylinositol 3-OH kinase (PI3K) associated with the membrane anchored p150 adapter, which tethers the enzyme to cytoplasmic membranes, forming a complex with the vacuolar protein sorting 34 (Vps34). The resulting complex is Vps34 (PI3K)/Vps15 (p150). –– The ubiquitin-like protein system includes different proteins with specific functions. The ubiquitination process involves three sequential reactions catalyzed by the E1 (ubiquitin activating), E2 (ubiquitin conjugating/carrier), and E3 (ubiquitin ligase) enzymes of the ubiquitin system (Hershko and Ciechanover 1998). In autophagy the ubiquitin system is composed of two ubiquitin-like proteins (Atg8 and Atg12); one activating enzyme (Atg7); two analogues of ubiquitin-conjugating enzymes (Atg10 and Atg3); one Atg8 modifying protease (Atg4); the Atg5 one protein target of Atg12 attachment; and the Atg16. In this system the Atg7 and the Atg10 proteins function as the E1 and E2 enzymes in the ubiquitin pathway.

5.6 Atg9 Cycling System Atg9 is an integral membrane protein located in the preautophagosomal membrane. It has a role prior to the closure of the double-bilayer vesicle but is absent in mature autophagosomes. The interaction of Atg2 with Atg9 is indispensable for autophagosome formation. Atg1 and Atg18 are required for the correct localization of Atg2 in the pre-autophagosomal membrane.

5.7 Class-III PI3K Class-III PI3K (and its adaptor p150) was also found to be necessary for autophagy in yeast. These organisms contain Vps34, a homolog of the class-III PI3K, which is part of a complex which includes Vps15 (the homolog of p150 in mammalian cells), Vps30,

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and Atg14 (Hutchins et al. 1999). This complex is required to recruit the Atg12–Atg5 conjugate to the preautophagosomal structure (Suzuki et  al. 2001). The autophagy effector Beclin 1, the mammalian ortholog of yeast Atg6/Vps30, is part of a Class III PI3K complex which is involved in both vacuolar protein sorting and autophagy.

5.7.1 Ubiquitin-Like Protein System Analyses of the Atg proteins allowed the identification of two ubiquitination-like conjugation systems required for autophagosome formation (Ohsumi 2001) and one of these systems mediates the conjugation of Atg12–Atg5 (Mizushima et al. 1998a, b). This Atg12–Atg5 conjugation system is conserved from yeast to human. The Atg12–Atg5 system associates non-covalently with the coiled-coil protein Atg16. The other system mediates a covalent linkage between Atg8 and phosphatidylethanolamine (PE) (Ichimura et al. 2000). Atg8 was the first molecule found to localize to the intermediate structures of the autophagosome, necessary for autophagosome formation (Kirisako et al. 1999; Huang et al. 2000). Atg12–Atg5 conjugate is present on the outer side of the isolation membrane and is required for elongation of the isolation membrane (Mizushima et al. 2001). The conversion of Atg8 to Atg8-PE requires Atg7. The latter is an enzyme which has a function similar to E1 enzyme in the ubiquitin pathway in the Atg12 conjugation system (Mizushima et  al. 1998a, b; Tanida et  al. 1999; Kim et  al. 1999). Atg8 conjugates with Atg3 through a thioester bond between the C-terminal glycine of Atg8 and Cys234 of Atg3. This system is dependent upon the activity of Atg3 which functions as an E2 enzyme (Ichimura et al. 2000). Light chain 3 (LC3) of neuronal microtubule-associated protein 1A/B, is a mammalian homolog of yeast Atg8 that is required for autophagosome formation (Kabeya et al. 2000). Immediately after the synthesis of LC3, the protein is converted to the cytoplasmic form, LC3-I, by cleavage at the C-terminal region (Kabeya et  al. 2000). LC3-I is further converted into LC-II a membrane-bound form, by Atg7 and Atg3, when autophagy is induced (Ohsumi 2001). LC3-II is localized mainly in the membranes of autophagosomes and, to a lesser extent, within autophagolysosomes (Kabeya et al. 2000). A PE-conjugated form of LC3 localizes on the isolation membrane and the autophagosome membrane (Kabeya et  al. 2000; Kabeya et  al. 2004). LC3 is required in the formation process of autophagomes, and colocalizes to the nascent and early autophagic vacuole ­membranes; LC3 is thus the first specific marker protein for autophagic vacuoles.

5.8 Autophagosomes Formation Autophagosomes are usually bound by two or more membranes and contain ­undegraded cytoplasmic ground substance and recognizable cellular organelles. Autophagosomes sequester peroxisomes, mitochondria, ER, ribosomes and cytosolic ground substance for degradation (Dunn 1990). The autophagy starts when a flat

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Fig.  5.6  Schematic representation of autophagosomes formation. The conjugation system Atg12–Atg15 associates non-covalently to Atg16. The autophagosome formation besides requires the participation of the conjugation system LC3-PE. LC3-II form is released to the cytosol

membrane cistern wraps around a portion of cytosol and/or organelles, forming a closed double-membrane bound vacuole that contains cytoplasm and cytoplasmic components called autophagosome. The autophagosomes can be classified as early or initial autophagic vacuoles, when they contain cytosol or organelles, and as late or degradative autophagic vacuoles, when they contain partially degraded cytoplasmic material (Dunn 1994; Dunn 1990). The isolation membrane, which is thought to be formed as a result of the activity of the Beclin 1–Vps34 complex, is elongated with the help of an ubiquitin-like conjugation system. Atg12 is first activated by Atg7, then transferred to Atg10 and finally covalently attached to Atg5: this process requires ATP. The Atg12–Atg5 conjugate is localizated in autophagosome precursors and is dissociated around the completion of autophagic vacuole formation (Mizushima et al. 2001; Mizushima et al. 2003). An ubiquitin-like modification of LC3 protein is required for completion of autophagosome formation. The cytosolic precursor of LC3 is cleaved at its C terminus by Atg4 to form LC3-I (Kirisako et al. 2000). The LC3-I is covalently conjugated to phosphatidylethanolamine to form LC3-II (as previously mentionated). LC3-II is specifically targeted to the complex Atg12–Atg5 autophagosome precursors and remains associated with autophagosomes even after fusion with lysosomes, subsequent to which LC3-II is delipidated and recycled (Kirisako et al. 1999) (Fig. 5.6).

5.9 Maturation of Autophagosomes to Autolysosomes The maturation of a newly formed (early) autophagosome into an autolysosome appears to occur in two steps: the formation of a late autophagosome upon acquisition of lysosomal membrane proteins including the acidifying H+-ATPase, and the

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delivery of acid hydrolases by either transfer from endosomes mediated by the mannose-6-phosphate receptor or fusion with pre-existing lysosomes, thereby transforming the late autophagosome into an autolysosome (Dunn 1994). The autophagosome fuses with endosomal and/or lysosomal vesicles, and the content is delivered to the endo/lysosomal lumen. Autophagosomes fused with endosomes are called amphisomes (Berg et al. 1998), an autophagosome or amphisome that has fused with a lysosome is called autolysosome, at this stage, autophagosomes acquire lysosome-associated membrane protein 1 (Lamp1) and Lamp2 (Eskelinen et  al. 2003). In S.  cerevisiae the autophagosomes fuse with the vacuole. The mechanism of fusion has similarities with homotypic vacuole fusion, and the SNAREs Vam3 and Vti1 (mammalian orthologues are syntaxin-7 and VTI1B, respectively) have both been implicated in the fusion of autophagosomes with the yeast vacuole (Darsow et al. 1997). Members of the homotypic fusion and vacuole protein sorting complex (HOPS complex), have also been shown to be important in autophagosome/vacuole fusion in S. cerevisiae and autophagosome–lysosome fusion in Drosophila melanogaster (Rieder and Emr 1997). In mammalian cells, RAB7 has been implicated in the fusion of autophagosomes with lysosomes (Jäger et al. 2004).

5.10 Autophagy as a Process of Cell Death Autophagy is generally accepted as a normal catabolic pathway for lysosomal ­ degradation of intracellular membranes, cytosolic organelles and secretory products. The autophagy mechanism is considered as a protective system, attempting to reduce metabolic demand by the cell to avoid death; many organisms in starving conditions will consume first labile and then less labile muscle proteins. Autophagy is also involved in removing damaged mitochondria and other organelles, in degrading intracellular pathogens, and protein aggregates too large to be removed by the ubiquitin-proteasomal system. These functions of autophagy could promote cellular survival during aging, infectious diseases, and neurodegenerative processes. In addition to a cell-autonomous role for autophagy in promoting survival, autophagy is involved in programmed cell death during physiologic processes in vivo (reviewed in Levine and Yuan 2005). The function of autophagy as a death process is debated; however diverse reports indicate that the autophagic cell death occurs primarily when the developmental program (e.g., insect metamorphosis) or homeostatic processes in adulthood require massive cell elimination. Despite its predominant role as a survival pathway, progressive autophagy can result in cell death if allowed to proceed to completion under persistent stress and during development. In Drosophila, autophagic cell death plays an important role during salivary gland development (Baehrecke 2000). Diverse studies have also described autophagic cell death in diseased mammalian tissues and in tumour cell lines treated with chemotherapeutic agents (Levine and Yuan 2005). In most situations, such as starvation, autophagy can be increased and

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provoke an oxidative stress or mitochondrial dysfunction (Aubert et  al. 1996). Under these circumstances the autophagy probably functions initially as a cytoprotective mechanism, but if cellular damage is too extensive, or if apoptosis is compromised, excessive autophagy may be used to kill the cell (Gorski et  al. 2003; Kissova et al. 2004) (Fig. 5.7). The term “autophagic cell death” describes a form of programmed cell death morphologically distinct from apoptosis and presumed to result from excessive levels of cellular autophagy (Schweichel and Merker 1973). Autophagic cell death is not only accompanied by the accumulation of autophagic vacuoles, but also involves an increase in cytoplasmic degradation that contributes to cell death. Autophagic degeneration is always accompanied by cytoplasmic vacuolization and activation of lysosomal enzymes (Afford and Randhawa 2000). This cell death program is characterized by the lack of a tissue inflammatory response due to the absence of a rupture of the cell membrane and the leak of cytoplasmic content to the extracellular space. Autophagy is characterized by the accumulation of autophagic vesicles (autophagosomes and autophagolysosomes), which occurs in the absence of chromatin condensation but is accompanied by large-scale autophagic vacuolization of the cytoplasm. Autophagic cell death is often observed when massive cell elimination is needed or when phagocytes do not have easy access to the dying cells, as in some forms of developmental programmed cell death (e.g., mammalian embryogenesis, insect metamorphosis), in which the availability of macrophagic cells may be insufficient for clearance of dead cells. In such cases, dying cells may activate

Fig. 5.7  Graphic representation of different pathway of autophagy. An excessive ­vacuolization lead to different molecular mechanisms that lead to autophagic cell death

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autophagy to target its contents for degradation by its own lysosomes (reviewed in Levine and Yuan 2005). During autophagic cell death, the cytoskeleton is largely preserved, even beyond the stage of nuclear destruction; presumably the cytoskeletal network is required for the initiation and progression of autophagocytosis, serving to eliminate cytoplasmic constituents such as endoplasmic reticulum, Golgi apparatus etc. (Bursch et al. 2000). During the sequestration of cytoplasmic structures, the initial step in formation of autophagosomes depends on intermediate filaments such as cytokeratin and vimentin (Aplin et al. 1992), whereas their subsequent fusion with lysosomes depends on microtubule (reviewed in Blommaart et al. 1997). All steps including the degradation of cytoplasmic material in autophagic vesicles are ATP-dependent (Luiken et al. 1996). Studies indicate that autophagy selectively degrades specific proteins (Onodera and Ohsumi 2004) lending support to the idea that the autophagy may kill cells through selective degradation of regulatory molecules or organelles that are essential for cell survival (Yu et al. 2004a, b), such as mitochondria. It has been suggested that autophagy may be initiated by the mitochondria depolarisation and permeability transition (MPT) (Lemasters et al. 1998). It has been suggested that the overall autophagic activity in a cell doomed to die is far more extensive than that associated with the normal cytoplasmic and organelle turnover occurring in healthy cells. The physiological and pathological role of autophagic cell death is emphasized with various evidences giving a therapeutic value to autophagic cell death. Autophagic cell death was proposed as the action mechanism of some anticancer agents. When the human mammary carcinoma is treated with the anti-estrogen tamoxifen, MCF-7 cells have a high accumulation of autophagic vacuoles (Gozuacik and Kimchi 2004). Treatment of L929 fibroblasts with the caspase inhibitor zVAD causes a form of cell death that is characterized by autophagosome formation. RNA interference of Atg7 or beclin-1 attenuates this reduction in cell number, indicating that autophagy is a key mediator of cell death in response to zVAD (Yu et al. 2004a, b). Autophagic cell death is also extensively observed in steroid-triggered cell death during development of Drosophila (Lee et al. 2003). In Drosophila, a marked increase in autophagy is observed at the end of the larval stage (Baehrecke 2005). This developmental programmed autophagy is hormonally controlled by ecdysone and is responsible for the elimination of organs, such as the fat body, during metamorphosis. The increase in autophagy of the fat body is caused by the inhibition of the class-I PI3K pathway by ecdysone (Rusten et  al. 2004). This inhibitory effect is observed when the ecdysone receptor is expressed. Also, beclin 1 and Atg7 were reported to be associated with autophagic cell death (Ogier-Denis and Codogno 2003; Yu et al. 2004a, b). Danon disease, cardiomyopathies, and skeletal myopathies with different genetic origins are characterized by the accumulation of autophagic vacuoles (Nishino et al. 2000; Tanaka et al. 2000). In Alzheimer disease, Huntington disease, and Parkinson disease, affected neurons have an increase in the number of autophagic vacuoles (Kegel et al. 2000; Nixon et al. 2000; Anglade et al. 1997).

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5.11 Autophagy Induction In yeast, flies, plants, and worms, autophagy is primarily an adaptive survival response to food deprivation. Humans, as other mammals, have molecular mechanism to maintain homeostasis during cellular metabolic stress. Autophagy is induced by metabolic energetic stresses and by catabolic hormones (Wang and Klionsky 2003). One of the more frequently used models to induce autophagy is starvation, which induces kinases as Gcn2 and its downstream target eIF2a. Starvation also activates the Gcn4 a transcriptional transactivator (Talloczy et  al. 2002), as well as Tor, which is a negative regulator of autophagy. Tor acts in a signal transduction cascade through various downstream effectors to control both translation and transcription. Tor is a good candidate as a keeper of the autophagic pathway, because it is a sensor for amino acids and ATP (Marygold and Leevers 2002). Tor appears to cause the hyperphosphorylation of Atg13, which binds to and activates Atg1. The Atg1–Atg13 association is required for autophagy; when Atg13 is hyper-phosphorylated it has a lower affinity for the Atg1 kinase and autophagy is inhibited (Kamada et al. 2000). The initial sensor of cellular bioenergetics crisis is AMPK. Its downstream ­target, TOR integrates signals from nutrients as well as growth factors to control cell growth (Wullschleger et al. 2006; Inoki et al. 2003). Thus, TOR acts as a metabolic rheostat controlling protein synthesis during cellular stress. Autophagy mediating catabolism of intracellular contents can maintain cellular bioenergetics during metabolic stress (Xu et al. 2007). Under nutrient-rich conditions, TOR blocks the initiation step of autophagy by facilitating dissociation of Atg13–Atg1 complex, an essential factor required for the formation of an autophagic vesicle ­(autophagosome) in budding yeast (Kamada et al. 2000).

5.12 Autophagy and Cancer Several studies established that autophagy may have different effects on cancer. At the early stage of tumor development, autophagy functions as a tumor suppressor. Autophagy may protect against cancer by sequestering damaged organelles, permitting cellular differentiation, increasing protein catabolism, and/or promoting autophagic death. Expression of Beclin 1 reduces tumorigenic capacity through induction of autophagy. Mice Beclin 1+/– display a remarkable increase in the incidence of lung cancer, hepatocellular carcinoma, and lymphoma. At advanced stages of tumor development, autophagy promotes tumor progression. The tumor cells that are located in the central area of the tumor mass undergo autophagy to survive low-oxygen and low-nutrient conditions and the autophagy may contribute to cancer by promoting the survival of nutrient starved cells. Autophagy protects some cancer cells against anticancer treatments by blocking the apoptotic pathway (‘protective autophagy’). By contrast, other cancer cells undergo autophagic cell death

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after cancer therapies. Autophagy is induced mainly through the phosphatidylinositol 3-phosphate kinase (PI3K)–AKT–mTOR (mammalian target of rapamycin) signalling pathway (reviewed in Kondo and Kanzawa 2005).

5.13 Inhibition of Autophagy Autophagy may be inhibited at various stages of the process (Fig. 5.8). There are several physiological inhibitors found in experimental models, such as media rich in serum, aminoacids, which cooperate at different levels in the suppression of autophagy. The pharmacologic inhibitor of autophagy 3-methyladenine (3-MA), a nucleotide derivative that can inhibit class III PI3K kinases (Xue et al. 1999), delays or partially inhibits death in different cells. Okadaic acid and related toxins suppress autophagy. The effects of these toxins are mediated by the over phosphorylation of the AMP-activated-protein kinase (AMPK); these properties are shared by the AMPK activator AICAR (Yue et al. 2003). They inhibit autophagy producing extensive fragmentation of the cellular keratin and plectin cytoskeleton networks (Stefanis et al. 2001), which are required for the initiation and progression of autophagocytosis.

Fig. 5.8  The autophagy process may be induced or inhibited by different pathways

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Rapamycin is the most specific kinase inhibitor known and its target, mTOR, controls many processes independent of autophagy (Wullschleger et al. 2006). TOR (mTOR in mammals) is an important protein in the signaling pathways of the autophagy. TOR is a serine/threonine kinase which inhibits autophagy through two general mechanisms: by controlling both translation and transcription, and by modifying (directly or indirectly) Atg proteins, thus interfering with the formation of autophagosomes. Under nutrient-rich conditions, active TOR causes hyperphosphorylation of Atg13, preventing its association with Atg1. The inhibition of TOR signaling (which causes Atg13 dephosphorylation) is therefore required to stimulate Atg1–Atg13 association and Atg1 kinase activity, needed for the formation of the autophagosomal membrane. The inhibition of TOR is also required to enhance the expression of specific genes, such as Atg8 and Atg14, involved in the autophagosome formation (reviewed in Ferraro and Cecconi 2007). AKT is a serine-threonine kinase, located downstream of class I PI3K. It activates the kinase mTOR (Schmelzle and Hall 2000; Gingras et al. 2001), leading to suppression of autophagy. A role for this signalling pathway in the negative regulation of autophagy has been observed in non-mammalian cells (Wang and Klionsky 2003). Class I and class III PI3K regulate autophagy differently: the class I PI3KAKT-mTOR signal, which is activated in cancer cells through the growth factor receptor, inhibits autophagy and, by contrast, class III PI3K promotes the sequestration of cytoplasmic material that occurs during autophagy. Ras is a protein that belongs to a large super-family of proteins known as G-proteins. The oncogenic forms of Ras are also implicated in the negative control of autophagy, through activation of class I PI3K. The RNA interference (RNAi) technique for blocking protein expression in mammalian cells has allowed efficient suppression of autophagy in cultured mammalian cells by targeting Atg8/LC3; Atg6/Beclin 1, and other Atg genes, proving successful results. There is a flow from the preautophagosomal structure (PAS or phagophore assembly site) to the autophagosome. The PAS seems to play a crucial role just before or during formation of the autophagosome (Noda et al. 2002). Class I PI3K enzymes phosphorylate phosphatidylinositol-4phosphate (PtdIns4P) and phosphatidylinositol-4, 5-bisphosphate (PtdIns(4,5)P2) to produce PtdIns (3,4) P2 and PtdIns(3,4,5)P3, which, via Pleckstrin Homology (PH) domains bind to protein kinase B (Akt/PKB) and its activator phosphoinositide-dependent kinase-1 (PDK1) (Brazil and Hemmings 2001; Katso et  al. 2001). PDK1 phosphorylates other kinases, including p70S6 kinase (Vanhaesebroeck and Alessi 2000). Activation of this pathway by challenging receptors that recruit class I PI3K or by expressing a constitutive active form of PKB has an inhibitory effect on autophagy (Arico et al. 2001; Petiot et al. 2000). eIF2 kinases (GCN2 and PKR), which regulate stress-induced translation arrest by phosphorylating the translation factor eIF2, positively controlled autophagic sequestration in yeast and mammalian cells in response to nutrient deprivation (Talloczy et al. 2002). It is therefore possible that functions of Beclin 1 independent of autophagy may be involved in its tumor ­suppressor activity.

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5.14 Autophagy Activation Autophagy initiation is stimulated by many different intracellular or extracellular stress stimuli (Jin and White 2007) (Fig. 5.8). The classical autophagy inducer is represented by amino-acid deprivation and this stimulus can increase the activity and phosphorylation of mTOR a protein kinase central to nutrient sensing signal transduction, regulation of translation and cell cycle progression control (Lum et al. 2005) which leads to inhibition of mTOR. Autophagy can also be induced by a mTOR-independent route by lowering myo-inositol-1,4,5-triphosphate (IP3) levels (Rubinsztein et al. 2007). The antiestrogen tamoxifen induces autophagy and cell death in MCF-7 cells (Bursch et  al. 1996). Tamoxifen stimulates autophagy by increasing the intracellular level of ceramide and abolishing the inhibitory effect of the class-I PI3K pathway on autophagy. In a rat cardiomyocyte-derived cell line, the activation of class-I PI3K during glucose deprivation induces the accumulation of autophagic vacuoles and causes cell death (Aki et al. 2003). During tamoxifeninduced autophagic cell death of MCF-7 cells, intermediate and microfilaments are redistributed, but largely preserved even beyond the stage of nuclear collapse (Bursch et al. 2000). These data support the concept that autophagic cell death is a separate form of programmed cell death that is distinctly different from apoptosis. The regulation of the phosphorylation state of some Atg proteins is one of the mechanisms through which autophagy is regulated. Autophagy regulation is a highly complex process involving many signaling complexes and pathways. Under nutrient starvation, Atg13 which is a regulatory protein that forms a complex with Atg1 (serine/threonine-kinase), and becomes partially dephosphorylated leading to an Atg1– Atg13 interaction and subsequent generation of autophagosomes (Yang et al. 2005).

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Nishino I, Fu J, Tanji K, Yamada T, Shimojo S, Koori T, Mora M, Riggs JE, Oh SJ, Koga Y, Sue CM, Yamamoto A, Murakami N, Shanske S, Byrne E, Bonilla E, Nonaka I, DiMauro S, Hirano M (2000) Primary LAMP-2 deficiency causes X-linked vacuolar cardiomyopathy and myopathy (Danon disease). Nature 406:906–910 Nixon RA, Cataldo AM, Mathews PM (2000) The endosomal-lysosomal system of neurons in Alzheimer’s disease pathogenesis: a review. Neurochem Res 9–10:1161–1172 Noda T, Suzuki K, Ohsumi Y (2002) Yeast autophagosomes: de novo formation of a membrane structure. Trends Cell Biol 12:231–235 Novikoff AB (1961) Lysosomes and related particles. In: Brachet J, Mirsky AE (eds) The Cell, vol II, Cells and their component parts. Academic Press, New York, pp 423–488 Ogier-Denis E, Codogno P (2003) Autophagy: a barrier or an adaptive response to cancer. Biochim Biophys Acta 1603:113–128 Ohsumi Y (2001) Molecular dissection of autophagy: Two ubiquitin-like systems. Nat Rev Mol Cel Biol 2:211–216 Onodera J, Ohsumi Y (2004) Ald6p is a preferred target for autophagy in yeast, Saccharomyces cerevisiae. J Biol Chem 279:16071–16076 Peters C, von Figura K (1994) Biogenesis of lysosomal membranes. FEBS Lett 346:108–114 Petiot A, Ogier-Denis E, Blommaart EF et al (2000) Distinct classes of phosphatidylinositol 3’-kinases are involved in signaling pathways that control macroautophagy in HT-29 cells. J Biol Chem 275(2):992–998 Rieder SE, Emr SD (1997) A novel RING finger protein complex essential for a late step in protein transport to the yeast vacuole. Mol Biol Cell 8:2307–2327 Rubinsztein DC, Gestwicki JE, Murphy LO, Klionsky DJ (2007) Potential therapeutic applications of autophagy. Nat Rev Drug Discov 6:304–312 Rusten TE, Lindmo K, Juhasz G, Sass M, Seglen PO, Brech A, Stenmark H (2004) Programmed autophagy in the Drosophila fat body is induced by ecdysone through regulation of the PI3K pathway. Dev Cell 7:179–192 Schmelzle T, Hall MN (2000) TOR, a central controller of cell growth. Cell 103:253–262 Schweichel JU, Merker HJ (1973) The morphology of various types of cell death in prenatal tissues. Teratology 7:253–266 Stefanis L, Larsen K, Rideout HJ, Sulzer D, Greene LA (2001) Expression of A53T mutant but not wild-type a-synuclein in PC12 cells induces alterations of the ubiquitin-dependent degradation, loss of dopamine release, and autophagic cell death. J Neurosci 21:9549–9560 Susin SA, Zamzami N, Kroemer G (1998) Mitochondira and regulators of apoptosis: Doubt no more. Biochim Biophys Acta 1366:151–165 Suzuki K, Kirisako T, Kamada Y, Mizushima N, Noda T, Ohsumi Y (2001) The pre-autophagosomal structure organized by concerted functions of APG genes is essential for autophagosome formation. EMBO J 20:5971–5981 Takeda M, Shirato I, Kobayashi M, Endou H (1999) Hydrogen peroxide induces necrosis, apoptosis, oncosis and apoptotic oncosis of mouse terminal proximal straight tubule cells. Nephron 81:234–238 Talloczy Z, Jiang W, Virgin HW, Leib DA, Sheuner D, Kaufman RJ, Eskelinen EL, Levine B (2002) Regulation of starvation -and virus- induced autophagy by the elF2a kinase signalling pathway. Proc Natl Acad Sci USA 99:190–195 Tanaka Y, Guhde G, Suter A, Eskelinen EL, Hartmann D, Lullmann-Rauch R, Janssen PM, Blanz J, Figura K, Saftig P (2000) Accumulation of autophagic vacuoles and cardiomyopathy in LAMP-2-deficient mice. Nature 406(6798):902–906 Tanida I, Mizushima N, Kiyooka M, Ohsumi M, Ueno T, Ohsumi Y, Kominami E (1999) Apg7p/ Cvt2p: A novel protein-activating enzyme essential for autophagy. Mol Biol Cell 10:1367–1379 Ueno T, Ishidoh K, Mineki R, Tanida I, Murayama K, Kadowaki M, Kominami E (1999) Autolysosomal membrane-associated betaine homocysteine methyltransferase. Limited degradation fragment of a sequestered cytosolic enzyme monitoring autophagy. J Biol Chem 274(21):15222–15229 Vanhaesebroeck B, Alessi DR (2000) The PI3K-PDK1 connection: more than just a road to PKB. Biochem J 3:561–576

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Wang CW, Klionsky DJ (2003) The molecular mechanism of autophagy. Mol Med 9:65–76 Wullschleger S, Loewith R, Hall MN (2006) TOR signaling in growth and metabolism. Cell 124:471–484 Xu ZX, Liang J, Haridas V, Gaikwad A, Connolly FP, Mills GB, Gutterman JU (2007) A plant triterpenoid, avicin D, induces autophagy by activation of AMP-activated protein kinase. Cell Death Differ 14:1948–1957 Xue L, Fletcher GC, Tolkovsky M (1999) Autophagy is activated by apoptotic signalling in sympathetic neurons: an alternative mechanism of death execution. Mol Cell Neurosci 14:180–198 Yang YP, Liang ZQ, Gu ZL, Qin ZH (2005) Molecular mechanism and regulation of autophagy. Acta Pharmacol Sin 26(12):1421–1434 Yu L, Lenardo MJ, Baehrecke EH (2004a) Autophagy and caspases. A new cell death program. Cell Cycle 3(9):1124–1126 Yu L, Alva A, Su H, Dutt P, von Freundt E, Welsh S, Baehrecke EH, Lenardo MJ (2004b) Regulation of an ATG7-beclin 1 program of autophagic cell death by caspase-8. Science 304:1500–1502 Yue Z, Jin S, Yang C, Levine AJ, Heintz N (2003) Beclin 1, an autophagy gene essential for early embryonic development, is a haploinsufficient tumor suppressor. Proc Natl Acad Sci USA 100:15077–15082

Chapter 6

Oncosis

María Luisa Escobar, Gerardo H. Vázquez-Nin, and Olga M. Echeverría

Abstract  Morphological features of oncosis are characterized by cell and ­organelle swelling and disruption; nucleus is initially preserved and later undergoes karyolysis and caspase-independent DNA fragmentation. The mitochondrial generation of ATP requires the presence of an electrochemical gradient; a reduction in mitochondrial membrane potential frequently precedes the morphological changes seen in oncotic cells. Oncotic cell death is characterized by early plasma membrane rupture and disruption of organelles including mitochondria. The plasma membrane breakdown allows the outflow of cell components, which affects near or distant organs. List of Abbreviations ATP Adenosine triphosphate DNA Deoxyribonucleic acid ROS Reactive oxygen species

6.1 Molecular Mechanisms of Oncosis Process In addition to the processes of programmed cell death type I and type II, previously described in this book, there is another type of cell death called oncosis. This process is different to necrosis, in which has sometimes been included as initial necrosis. For several authors necrosis is a process involving postmortem biochemical and morphologic changes. The term oncosis is derived from the Greek word onkos, meaning swelling. It was introduced one century ago by von Recklinghausen (1910) to describe the

M.L. Escobar (*), G.H. Vázquez-Nin, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico, USA e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_6, © Springer Science+Business Media B.V. 2011

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Fig. 6.1  Morphological characteristics of the oncotic cell death

death process of osteocytes in patients with rickets and osteomalacia and human hypertrophic chondrocytes during the growth of cartilage. Majno and Joris (1995) proposed using the term oncosis to designate any cell death characterized by a marked cell swelling. The morphological features of oncosis include cell and organelle swelling, membrane blebbing, increased membrane permeability, nuclear chromatin clumping in the absence of evident dense chromatin bodies, cytoplasmic vacuolation, dilation of the endoplasmic reticulum and Golgi, lysosomal disruption and leakage, as well as mitochondrial swelling and disruption. Oncotic cell death is characterized by early plasma membrane rupture and disruption of cellular organelles, including mitochondria (Fig. 6.1). During oncotic cell death the nucleus is initially preserved, and later undergoes karyolysis rather than karyorhexis and caspaseindependent DNA fragmentation, which does not involve internucleosomal fragmentation. In oncotic myocytes, DNA fragmentation was reported to be caused by the activation of a Ca++ and Mg++-dependent nuclease, which is indistinguishable from DNase I (Giannakis et al. 1991; Peitsch et al. 1993). In an oncotic cell, the chromatin is initially condensed and then dispersed and ruptured into pieces. This process is different from lytic necrosis, in which the cells are dissolved without chronic swelling. Oncotic death may culminate in the release of intracellular contents, which can produce inflammation, especially if the cellular corpses are not efficiently phagocytised. The process of oncosis ultimately leads to depletion of cellular energy stores and failure of the ionic pumps in the plasma membrane (Fig. 6.2). The cytoplasmic swelling observed in oncosis has been attributed to primary changes in the plasma membrane, resulting in altered ionic fluxes and loss of cell volume regulation. An increased intracellular calcium concentration initiates the translocation of cytosolic phospholipase A2s to cellular membranes, where the hydrolysis of membrane phospholipids decreases membrane integrity (Sapirstein et al. 1996; Cummings et al. 2000). At the biochemical level, oncosis is a form of cell death induced by energy depletion, which produces dramatic reductions in mitochondrial respiration and ATP synthesis (Majno and Joris 1995). Diverse data suggest a central role for mitochondria in regulating oncotic cell death. Changes in mitochondrial membrane

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Fig.  6.2  The oncotic cell death is a mechanism based in the failure of the ionic pumps of the plasma membrane; this provokes an increased cytosolic Ca2+ and a drastic decrease of the ATP levels

trigger several events, such as alterations in the ATP production. The mitochondrial generation of ATP requires the presence of an electrochemical gradient; a reduction in mitochondrial membrane potential frequently precedes the morphologic changes seen in oncotic cell death (Petit et al. 1996). The increased entry of Ca2+ into the mitochondria can activate specific phospholipases and proteases which disrupt mitochondrial permeability, resulting in arrest of ATP production, and activation of the generation of reactive oxygen species, which provoke an oxidative stress that consequently leads to open the mitochondrial permeability transition pores, which also contributes to the collapse of mitochondrial membrane potential and loss of mitochondrial function (Nayler and Elz 1986). The sequence of ischemia reperfusion, through increased oxygen free radicals, causes deletions in several areas of the mitochondrial genome. This cumulative mitochondrial DNA damage is associated with induction of nuclear oxidative phosphorylation of mRNA (Ferrari 1996). The process of oncosis often leads to nonspecific DNA fragmentation with impaired ATP generation, early mitochondrial damage, ionic pump dysfunction resulting in an escape of lysosomal enzymes to the cytoplasm, which increase plasma membrane permeability and leakage of intracellular structural proteins. Oncosis occurs when ATP generation is attenuated or when cellular energy consumption becomes unregulated (Majno and Joris 1995). Poly(ADP-ribose) polymerase (PARP) is a nuclear enzyme that is activated by DNA strand breaks to

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catalyze the addition of poly(ADP-ribose) to a variety of nuclear proteins. The inhibition of PARP in oxidant-stressed endothelial cells attenuates oncosis and induces caspase activation, shifting the oncotic cell death program to apoptosis (Walisser and Thies 1999). During the repair process of massive DNA destruction, the cellular energy depletion by PARP results in oncosis (Fink and Cookson 2005), because the massive DNA destruction causes an excessive PARP activity which depletes its ­substrate nicotinamide adenine dinucleotide (NAD), and the resynthesis of NAD depletes ATP. The eventual loss of energy stores leads to oncotic cell death (Walisser and Thies 1999). In this way, energy depletion and oncosis occur as a regulated response to severe DNA injury (Berger 1985). In constrast, during apoptosis caspases cleave and inactivate PARP preserving cellular ATP, despite significant DNA damage (Pieper et al. 1999). The changes accompanying oncosis result from active enzyme-catalyzed biochemical processes. Calpains, a family of calcium-activated neutral cysteine proteases, have been shown to play a role in oncotic cell death (Liu et al. 2004). The involvement of this particular cysteine protease family suggests that oncosis is carefully regulated. On the basis of their tissue expression patterns, calpains are classified as either ubiquitous or tissue specific (Sorimachi et al. 1997). Two ubiquitous isoforms have been identified: (calpain I) and m-calpain (calpain II). Physiologically, calpains play critical roles in embryogenesis, cell cycle progression, cell proliferation, differentiation, and migration (Liu et al. 2004). Mitochondrial release of Ca++, together with the inhibition of the Ca++-Mg++-ATPase in the plasma membrane (Tsokos-Kuhn et al. 1988), may lead to an increase of cytosolic Ca++ levels sufficient to activate Ca++-dependent intracellular calpains. In renal proximal tubules, extensive ATP depletion through endoplasmic reticulum Ca++ release result in a sustained increase in Ca++ concentration and calpain activation, which leads to cell swelling and loss of plasma membrane integrity (Harriman et al. 2002). It has been shown that the rise in the concentration of intracellular free Ca++ promotes the activation calpains via TNF (Chang et al. 2004) (Fig. 6.3). Increased calpain activity provokes a lack of caspase activity, the calpain cleaves Bid and other molecules regulating mitochondrial potential (Chen et al. 2001). Oncosis may result from toxic agents that interfere with ATP generation or ­processes that cause uncontrolled cellular energy consumption (Majno and Joris 1995). The cytotoxic agent 8-Iso-Prostaglandin F2 and its mediator TXA2 provoke oncotic cell death via an increase in intracellular calcium concentration (Brault et  al. 2003). Maitotoxin is a cytolytic agent isolated from the dinoflagellate Gambierdiscus toxicus. The maitotoxin initiates the oncotic cell death in a variety of cell types, causing a graded increase in cytosolic free Ca++ concentration, which is followed by the opening of cytolytic/oncotic pores that allow the exchange of organic molecules of molecular weight less than ~800 Daltons across the plasma membrane (Estacion and Schilling 2002). Marine palytoxin is a non-protein toxin first isolated from the soft coral of the genus Palythoa. This protein can join with its receptor in the plasmalemmal Na+-K+-ATPase pump and this conjunction converts the pump into a monovalent cation channel that exhibits a slight permeability

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Fig. 6.3  Calpains activation during oncotic cell death

to Ca++, causing a small but significant increase in intracellular Ca++ in vascular endothelial cells. This Ca++ increase leads to the activation of large dye-permeable pores, causing rapid oncotic cell death via a Ca++ overload mechanism (Schilling et al. 2006). The oncotic cell death has been associated to the histone H3 phosphorylation by ERK and p38MAPK (both MAPK family members). This phosphorylation leads to a premature chromatin condensation (Jia et al. 2004). Assays in vitro have shown that in renal cells induced by 2,3,5-Tris-(glutathion-S-yl)hydroquinone (TGHQ), a reactive metabolite of the nephrotoxic hydroquinone, the ROS-dependent activation of MAPKs is activated, followed by histone H3 phosphorylation and oncotic cell death in renal proximal tubule epithelial cells (LLC-PK1) (Dong et al. 2004).

6.2 Oncosis in Human Pathology Oncosis has typically been regarded as passive, and in many physiological and pathological processes, oncosis might play an important role. Oncosis is usually induced by pathological stimuli such as ischemia or ischemia and reperfusion, the same as toxic agents. Oncotic cell death is known to follow tissue ischemia and it is often observed in bone and cartilage that have little or no blood supply (Majno and Joris 1995). Cell death by oncosis has been recognized, and reported in diverse pathologies, such as acute lung injury, liver failure, stroke, smooth muscle cells in atherosclerotic lesions, myocytes in ischemic heart disease in humans and rabbits, human macrophages infected with Shigella flexneri, and in the rat pancreatitis model. Mura et al. (2007) have evidenced signs of oncosis in different cells including the lung airway epithelial cells, type II pneumocytes, microvascular endothelial cells, and heart and kidney cells in a study with animals which were subjected to

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­ echanical ventilation with oxygen and to the ischemia-reperfusion process. m Oncosis induced by pathogen infection has been identified in a number of experimental models, such as infection by Salmonella. This pathogen has developed a strategy to survive and escape from macrophages. Salmonella infection triggers swelling of macrophages; the oncotic macrophages are packed with motile Salmonella bacilli and later the flagellated Salmonella bacilli escape from oncotic macrophages (Sano et al. 2007). A local oncotic process may cause important consequences in other organs or systems. The outflow of cell contents occurring during an oncotic process may recruit and activate inflammatory cells, such as macrophages, inducing them to secrete excessive inflammatory factors into the blood, which may cause damage in near or distant organs. In acute pancreatitis disease, the oncosis is the primordial cell death process. The pancreatic acinar cells contain a variety of digestive enzymes, and when the oncotic cell death occurs the leaking of these enzymes may directly damage adjacent cells and even lead to pancreas autodigestion (Castejón and Arismendi 2006). The plasma membrane breakdown of the oncotic cells, allows the outflow of cell components as cytokines, which affect neighboring cells. The initiation of cytokine network in patients with diverse pathologies promotes the occurrence of systemic inflammatory response syndrome and multiple organ dysfunction syndromes. Due to the complexity of the cytokine network it is impossible to block all the pathways. A possible way to avoid the activation of the cytokine chain reaction may be the induction of apoptosis, which reduces the process of oncosis and the release of endocellular enzyme. The development of clinical strategies has been focused on the design of a route to avoid the progression of ischemic cellular damage, to make the oncotic process reversible and promote the apoptotic process before the cytoplasmic rupture. One of the strategies designed to avoid oncosis cell death is the use of the 11-Deoxy-16,16-dimethyl PGE2 (DDM-PGE2), which selectively induces a group of proteins, including RBP, cytoskeletal proteins, and molecular chaperones such as Grp78 and Hsp27 in pkNEO cells. These proteins may act in concert to mediated cytoprotection against oncotic/necrotic cell death, but not against apoptosis, when the toxic 2,3,5-tris(glutathion-S-yl) hydroquinone (TGHQ), a potent nephrotoxic and nephrocarcinogenic metabolite of hydroquinone, is employed (Jia et al. 2004). Another strategy utilized to reduce oncosis on pancreatic acinar cells has been inducing apoptosis. This induction brings about a decreased infiltration and activation of macrophages and a decrease of the generation of inflammatory cytokines (Zhao et al. 2007b). Myocytes become oncotic without reperfusion when the period of ischemia is prolonged. When the mechanical stress is weak enough to maintain sarcolemmal integrity during reperfusion, the myocytes undergo an apoptotic process of death; whereas the introduction of mechanical stress at the time of reperfusion induces myocyte oncosis by disrupting the sarcolemma. This treatment overrules mitochondrial function and inhibits the apoptotic process during the subsequent period of reperfusion, giving rise only to oncotic myocyte death (Otani et  al. 2006). In a simulated ischemic injury in cultured myocytes the cardioprotective protein AKT

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is activated, protecting myocytes from oncotic and apoptotic death. The activation of AKT suggests that myocardial stress activates a cytoskeleton-based survival pathway that may play an important role in protection against acute ischemia-­ reperfusion injury in ventricular myocardium (Wei and Vander Heide 2008). A deeper understanding of this important biological process will provide the framework for new therapies directed at protecting tissues and organs from severe metabolic and ischemic insults, preventing the increase in the number of cells undergoing oncosis which may result in disorders in the function of tissues and organs. Several authors have differentiated oncosis from necrosis. Oncosis and apoptosis are active processes of cell death, whereas necrosis refers to those processes of cellular degradation that follow cell death (Majno and Joris 1995). Oncosis and apoptosis represent different outcomes of the same pathway induced by MPT pore opening so that in the absence of ATP oncosis prevails, whereas the presence of ATP favors and promotes apoptosis. It has been suggested that in human oesophageal squamous cell carcinoma with sufficient blood supply cells die by apoptosis, while in those with poor blood supply most cells undergo oncosis. These results suggest that the selection of an oncotic or an apoptotic pattern of cell death depends on the energy supply (Zhao et al. 2007a). It has been proposed that in the absence of Fas pathway, activated T cells may die by at least by two different pathways: a TNF-independent oncotic pathway or an apoptotic pathway mediated by TNF that takes several days (Davidson et al. 2002). When the activity of caspases is inhibited, the process of cell death is mainly oncosis, leading some authors to name this process of cell death as primary necrosis.

References Berger NA (1985) Poly(ADP-ribose) in the cellular response to DNA damage. Radiat Res 101:4–15 Brault S et al (2003) Selective Neuromicrovascular Endothelial Cell Death by 8-Iso-Prostaglandin F2a. Stroke 34:776–782 Castejón OJ, Arismendi GJ (2006) Nerve cell death types in the edematous human cerebral cortex. J Submicrosc Cytol Pathol 38:21–36 Chang I et  al (2004) Role of calcium in pancreatic islet cell death by IFN-gamma/TNFalpha. J Immunol 172:7008–7014 Chen M et al (2001) Bid is cleaved by calpain to an active fragment in vitro and during myocardial ischemia/reperfusion. J Biol Chem 276:30724–30728 Cummings BS, McHowat J, Schnellmann RG (2000) Phospholipase A2s in cell injury and death. J Pharmacol Exp Ther 294:793–799 Davidson WF, Haudenschild Ch, Kwon J, Williams MS (2002) T Cell Receptor Ligation Triggers Novel Nonapoptotic Cell Death Pathways That Are Fas-Independent or Fas-Dependent. J Immunol 169:6218–6230 Dong J et al (2004) EGFR-independent activation of p38 MAPK and EGFR-dependent activation of ERK1/2 are required for ROS-induced renal cell death. Am J Physiol Ren Physiol 287:F1049–F1058 Estacion M, Schilling WP (2002) Blockade of maitotoxin-induced oncotic cell death reveals ­zeiosis. BMC Physiol 2:2–14

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Ferrari R (1996) The role of mitochondria in ischemic heart disease. J Cardiovasc Pharmacol 28(Suppl 1):S1–S10 Fink SL, Cookson BT (2005a) Apoptosis, pyroptosis, and necrosis: mechanistic description of dead and dying eukaryotic cells. Infect Immun 73:1907–1916 Giannakis C, Forbes IJ, Zalewski PD (1991) Ca21/Mg(21)-dependent nuclease: tissue distribution, relationship to inter-nucleosomal DNA fragmentation and inhibition by Zn21. Biochem Biophys Res Commun 181:915–920 Harriman JF et al (2002) Endoplasmic reticulum Ca2+ signaling and calpains mediate renal cell death. Cell Death Differ 9:734–741 Jia Z et al (2004) Grp78 is essential for 11-deoxy-16, 16-dimethyl PGE2-mediated cytoprotection in renal epithelial cells. Am J Physiol Ren Physiol 287:F1113–F1122 Liu X, Van Vleet T, Schnellmann RG (2004) The role of calpain in oncotic cell death. Annu Rev Pharmacol Toxicol 44:349–370 Majno G, Joris I (1995) Apoptosis, oncosis, necrosis An overview of cell death. Am J Pathol 146:3–15 Mura M, Andrade CF, Han B et al (2007) Intestinal ischemia-reperfusion induced acute lung injury and oncotic cell death in multiple organs. Shock 28:227–238 Nayler WG, Elz JS (1986) Reperfusion injury: Laboratory artifact or clinical dilemma? Circulation 74:215–221 Otani H, Matsuhisa S, Akita Y et al (2006) Role of Mechanical Stress in the Form of Cardiomyocyte Death During the Early Phase of Reperfusion. Circ J 70:1344–1355 Peitsch MC, Polzar B, Stephan H et al (1993) Characterization of the endogenous deoxyribonuclease involved in nuclear DNA degradation during apoptosis (programmed cell death). EMBO J 12:371–377 Petit PX, Susin SA, Zamazami N et al (1996) Mitochondria and programmed cell death: back to the future. FEBS Lett 396(1):7–13 Pieper AA, Verma A, Zhang J, Snyder SH (1999) Poly (ADPribose) polymerase, nitric oxide and cell death. Trends Pharmacol Sci 20:171–181 Sano G, Takada Y, Goto S et al (2007) Flagella Facilitate Escape of Salmonella from Oncotic Macrophages. J Bacteriol 189(22):8224–32 Sapirstein A, Spech RA, Witzgall R, Bonventre JV (1996) Cytosolic phospholipase A2 (PLA2), but not secretory PLA2, potentiates hydrogen peroxide cytotoxicity in kidney epithelial cells. J Biol Chem 271:21505–21513 Schilling WP, Snyder D, Sinkins WG, Estacion M (2006) Palytoxin induced cell death cascade bovine aortic endothelial cells. Am J Physiol Cell Physiol 291:C657–C667 Sorimachi H, Ishiura S, Suzuki K (1997) Structure and physiological function of calpains. Biochem J 328:721–732 Tsokos-Kuhn JO, Hughes H, Smith CV, Mitchell JR (1988) Alkylation of the liver plasma membrane and inhibition of the Ca2+ ATPase by acetaminophen. Biochem Pharmacol 37:2125–2131 Walisser JA, Thies RL (1999a) Poly(ADP-ribose) polymerase inhibition in oxidant-stressed endothelial cells prevents oncosis and permits caspase activation and apoptosis. Exp Cell Res 251:401–413 Wei H, Vander Heide RS (2008) Heat stress activates AKT via focal adhesion kinase-mediated pathway in neonatal rat ventricular myocytes. Am J Physiol Heart Circ Physiol 295:H561–H568 Zhao GF, Seng JJ, Zhao S et al (2007a) Oncosis in human esophageal squamous cell carcinoma and its relationship with apoptosis and microvessel density. Chin Med J 120(22):1999–2001 Zhao M, Xue DB, Zheng B et al (2007b) Induction of apoptosis by artemisinin relieving the severity of inflammation in caerulein-induced acute pancreatitis. World J Gastroenterol 13(42):5612–5617

Chapter 7

Necrosis

Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría

Abstract  The term necrosis was used for accidental, not programmed death of cells or tissues with uncontrolled release of cellular content resulting in inflammation. Necrosis is a term used by pathologists to designate the sum of changes that occurred in cell and tissues after they have died. Different experiments carried out in different cell types, species and cell death inducers show several traits that delineate a sequence of intracellular events specific to necrotic cell death. Morphological necrosis is characterized by cellular swelling including dilation of the endoplasmic reticulum, cellular lysis and subsequent inflammation. Core events of necrosis are bioenergetic failure and rapid loss of plasma membrane integrity. These failures results from defined molecular events including increased mitochondrial reactive oxygen species production, channel-mediated calcium uptake, activation of non-apoptotic proteases, swelling of mitochondria, perinuclear clustering of organelles and enzymatic destruction of cofactors required for ATP production, cell lysis and inflammation. The default occurrence of necrosis and its unmasking by inhibition of autophagy and/or apoptosis might reflect its early emergency in evolution.

List of Abbreviations NAD ATP ADP

Nicotinamide adenine dinucleotide Adenosine triphosphate Adenosine diphosphate

G.H. Vázquez-Nin (*), M.L. Escobar, and O.M. Echeverría Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences, National University of Mexico (UNAM), Mexico, USA e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_7, © Springer Science+Business Media B.V. 2011

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7.1 Processes Leading to Cell Death Various processes of cell death are characterized as active or programmed mechanisms of cell death such as apoptosis, autophagy, oncosis, pyroptosis and a form of necrosis different from accidental death and post-mortem alterations, frequently also called necrosis (Fig. 7.1). The best studied programmed cell death is apoptosis, a term proposed by Kerr and colleagues (1972) to describe a process of cell death with specific morphological pattern (Fig.  7.2). This process was characterized as an active, programmed process of autonomous dismantling that avoids inflammation. Various pathogens are known to cause host cell death with features of apoptosis (Fink and Cookson 2005). Numerous studies revealed that caspases, a group of cystein-dependent aspartate specific proteases, are the main actors in apoptotic process. All members of the caspase family are similar in their amino acids sequence, but they differ in their roles. Those related to apoptosis are divided into two subgroups: initiator caspases (caspase-2, -8, -9, and -10) and effector caspases (caspase-3, -6, and -7). Activated effector caspases selectively cleave a restricted set of target proteins (Fink and Cookson 2005). Morphological the cell shrinks and becomes denser, the

Fig.  7.1  Schematic representation of diverse processes of cell death, in this case the necrosis death is considered as a post-mortem condition

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Fig.  7.2  Different processes cell death. Morphological characteristics are indicators of diverse cell death events

chromatin becomes pyknotic and packed into masses applied against the nuclear membrane, the nucleus breaks and the cell emits processes that often contain pyknotic nuclear fragments. These processes break off and become apoptotic bodies which may be phagocytized by macrophages, neighboring cells or remain free (Majno and Joris 1995). Autophagy is an evolutionarily conserved mechanism for cell survival in starving cells which also functions in cell death (Baehrecke 2005). Autophagy involves the sequestration of cytosol and cytoplasmic organelles within vesicles limited by double membranes, thus creating autophagosomes or autophagic vacuoles (Fig. 7.2). Autophagosomes subsequently fuse with lysosomes, thereby forming autophagolysosomes also called autolysosomes. The limiting membranes of lysosomes and of autophagolysosomes are enriched in proteins called lysosomal associated proteins (lamps). These proteins are the most abundant lysosomal membrane glycoproteins, they may play a role in creating a barrier to lysosomal hydrolases protecting the membranes from hydrolytic destruction. The content of these vesicles is degraded by lysosomal hydrolases (Escobar et al. 2008). In spite of the extensive cytoplasmic destruction, the cell membrane does not break and the remains of the cell may be phagocytized by macrophages or neighboring cells. This process of cell death does not trigger inflammation. The oncosis is a process involving cellular and organelle swelling that may result from toxic agents that interfere with ATP generation or processes that cause

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uncontrolled energy consumption (Majno and Joris 1995) (Fig. 7.2). Massive DNA destruction promotes poly(ADP-ribose) polymerase activity that depletes its substrate (NAD) and the resynthesis of NAD depletes ATP. The loss of energetic stores leads to oncotic cell death. It is interesting to note that the inhibition of poly(ADPribose) polymerase prevents oncosis and permits caspase activation and apoptosis (Walisser and Thies 1999). Altered intracellular calcium levels may also regulate oncotic cell death. Pyroptosis is a form of cell death proinflammatory, dependent on caspase-1, frequently induced by infection with Salmonella and Shigella species. A function of Caspase-1, a cysteine protease, not involved in apoptotic cell death, is to process the proforms of the inflammatory cytokines IL-1b and IL-18 to their active forms in macrophages infected with those parasites resulting in cell membrane breakdown and release of the inflammatory cellular content (Fink and Cookson 2005).

7.2 Accidental Cell Death and Necrosis According to Majno and Joris (1995) and Fink and Cookson (2005), cell death and necrosis are two very different things. Cell death is a process that leads to a point of no return after which the cell dies and necrosis is the post-mortal decay changes of cells. However, these post mortem processes depend on the previous alterations of the leaving cell, thus there are several types of post-mortal processes frequently called apoptotic necrosis, ischemic necrosis or massive necrosis when the mechanism is not known (Fink and Cookson 2005). They use the term necrosis for accidental, not programmed death of cell or tissues with uncontrolled release of cellular contents resulting in inflammation. By the same token Goldstein and Kroemer (2006) also call necrotic process to the cell death in harsh conditions, such as detergent stress or freeze-thawing, resulting in a non-regulated, poorly defined process. Necrosis is a term used by pathologists to designate the sum of changes that occurred in cells or tissue after they have died. Used in this way several types of necroses are described according to the catabolism and post-mortem morphological traits. For instance the term apoptotic necrosis describes dead cells that have reached this state via the apoptotic program (Majno and Joris 1995; Fink and Cookson 2005).

7.3 Does Programmed Necrotic Cell Death Exist? Goldstein and Kroemer (2006) propose two types of necrotic processes, one nonregulated, poorly defined, taking place in harsh conditions such as detergent stress and other programmed processes in terms of both its course and its occurrence. Different experiments carried out by various authors in heterogeneous systems in terms of cell types, species (mammalian, nematode and slime mold) and death

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Fig. 7.3  Electron microscopy images showing necrotic cell death in mammal testis. In A normal cells, B-F altered cells in different phases of necrosis. Arrows show the altered cells. In E several neighboring cells are damaged, a classical morphological characteristic from necrosis. In F cellular remains are evidenced. General stained uranyl acetate and lead citrate. Scale bars 200 mm

inducers (ischemia, ligands of cell-surface receptors and so on) show several traits that delineate a sequence of intracellular events specific to necrotic cell death. Morphological necrosis is characterized by cellular swelling including dilation of the endoplasmic reticulum, Golgi apparatus, with or without chromatin condensation, leading to nuclear and cellular lysis and subsequent inflammation (Yuan et al 2003) (Fig. 7.3). Several studies demonstrate various resemblances between apoptosis, typical programmed cell death, and necrosis, suggesting that both processes are related: (1) diverse pathological stress such as heat, ionizing radiation, pathogens, cytokines cause both forms of cell death in the same population; (2) anti-apoptotic mechanisms, as Bcl-2, can protect cells from both necrotic and apoptotic destruction; (3) biochemical interventions (e.g. inhibitors of poly-(ADP-ribose)-polymerase) into the signal and executive mechanisms of programmed cell death can change the choice of the cell death form; (4) during both necrosis and epigenetic programs of apoptotic cell death that need no macromolecular synthesis (e.g. CD95-dependent death), the nucleus plays a passive role. These similarities allow the conclusion that necrosis, as apoptosis, is a form of programmed cell death (Proskuryakov et  al. 2002). The distinction between the two types of cell death is obvious in some ­systems, while in other cells necrotic and apoptotic features coexist.

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The resemblance between the process of apoptotic and necrotic processes of cell death and the coexistence of features of both processes in the same cell strongly suggest that necrosis is in fact, a programmed, active process of cell death. The extensive relationships between both processes strengthen this idea.

7.4 Relationships of Apoptosis and Necrosis Treatments with a pro-oxidant (2,3-dimethoxy-1,4-naphtoquinone) produce three different effects: stimulated growth of pancreatic cells in culture, triggered apoptosis or caused necrosis, depending on the dose and duration of exposure. The higher dose, which produced necrosis, caused a rapid intracellular Ca++ overload, ATP, NAD+, and glutathione depletion, and extensive DNA single strand breakage, which result in necrotic death (Dypbukt et al. 1994). The critical factor deciding the type of death is probably the energy level of the cell. Cells depleted of ATP and with a marked ion imbalance swell and rapidly lyses, whereas apoptotic features such as cell shrinkage and chromatin compaction would require preserved energy metabolism (Dypbukt et al. 1994). A similar result was obtained by exposing cultured cerebellar granule cells to different glutamate concentrations (1 mM to 3 mM) for 30  min and subsequently reincubating in culture medium lacking glutamate. The changes in mitochondrial membrane potential were monitored in individual neurons by determining the shift in wavelength and intensity of the fluorescence emission of the dye. By measuring ATP, ADP and AMP levels in the neuron cultures, the levels of cell energy were also detected. The cerebellar granule cells exposed to neurotoxic concentrations of glutamate undergo two distinct fates: a subpopulation succumbs to acute necrosis during and immediately after the exposure. In these cells, mitochondrial membrane potential collapses and their ability to metabolize methyl trazolium to formazan decreases, nuclei swell and intracellular debris are scattered in the incubation medium. These neurons undergo a typical necrotic cell death. Neurons surviving this early necrotic phase recovered mitochondrial potential and energy levels. Later these granular neurons undergo apoptosis. These results suggest that mitochondrial functions determine the mode of neuronal death. During the initial phase, when mitochondria were altered and energy scarce, cerebellar granule cells underwent a necrotic cell death and when energy levels were at least partially recovered the neurons underwent apoptosis. Relatively intact, mitochondrial activity and energy levels of neurons appears to be necessary for the apoptosis program to proceed (Ankarcrona et al. 1995). Cell death by hypoxia has been believed to be represented as necrosis based in ultrastructural findings. However, biochemical observations demonstrate that hypoxia also induce apoptosis. So once again, both necrosis and apoptosis are induced by the same agent (Shimizu et al. 1996). Furthermore, in cultures of cortical neurons exposed to different doses of neurotoxic N-Methyl-D-aspartate, it could also be observed that low doses mainly induce apoptosis and higher doses produce necrotic cell damage, characterized by swelling and lysis (Bonfoco et  al. 1995). Recent studies have

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demonstrated that in response to a given stimulus, there is a continuum of apoptosis and necrosis. Many insults induce apoptosis at lower doses and necrosis at higher doses. Even in response to a certain dose of a death-inducing agent, features of both apoptosis and necrosis may coexist in the same cell (Zong and Thompson 2006). All these observations taken together leave no doubt that necrosis is a programmed and active process of cell death initiated in precise metabolic conditions in which apoptosis can not take place and that mitochondrial events control the selection of these processes (Zamzami et al. 1997). It is interesting to note that, in spite of the at least partial conservation of energy production in cells undergoing apoptotic process, the caspase dependent breakdown of translation initiation factors as elF4G produce a drastic drop of translation (Saelens et al. 2005).

7.5 Relationships Between Autophagy and Necrosis In Dictyostelum autophagic or necrotic cell death can be triggered by distinct motifs of the same differentiation factor DIF-1. Distinct motifs of DIF-1 are required to trigger autophagy or necrosis. These motifs are separately recognized at the onset of autophagic or necrotic cell death. The metabolic state of the cell seems to guide the choice of the signaling pathway to cell death, which in turn imposes the cell death type and the recognition pattern of the differentiation factor (Luciani et al. 2009). Evolution may have modified a primitive dedicated programmed cell death mechanism, such as the one found in C. elegans, into multiple more flexible and adaptive cell death pathways (Yuan 2006). Under oxidative stress, poly(ADP-ribose)polymerase-1 (PARP-1) is activated and contributes to necrotic cell death through ATP depletion. On the other hand, oxidative stress is known to stimulate autophagy, and autophagy may act as either a cell death or a cell survival mechanism in cells under oxidative stress. It was demonstrated that autophagy plays a cytoprotective role in H2O2-induced cell death. Recently a novel autophagy signaling mechanism was identified that links PARP-1 to the serine/threonine protein kinase LKB1-AMP-activated protein kinase (AMPK)-mammalian target of rapamycin (mTOR) pathway, leading to stimulation of autophagy. (Huang et al. 2009). These authors also demonstrated that autophagy plays a cytoprotective role in H2O2-induced cell death, and that PARP-1 may function by promoting autophagy through LKB1-AMPK-mTOR pathway to enhance cell survival in cells under oxidative stress (Fig.  7.4). BNIP3 and NIX are two proteins involved in inducing an atypical process of cell death and also autophagy. The process of cell death is independent of apaf-1, ­caspase-9, caspase-3, and without cytochrome c release. The mechanism of cell death is similar but not identical in different cell types. They share certain features with the BH3-only protein subgroup of the BCL2 family, such as sequence homology in the BH3 domain, residence in the mitochondrial outer membrane, and the ability to interact with BCL2 and BCL-XL. However, both proteins are weak inducers of cell death that cause loss of mitochondrial Dym, opening of the mitochondrial

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Fig. 7.4  Relationship between autophagy and necrosis

permeability transition pores, and variable release of cytochrome c. In addition, the transmembrane domains, but not the BH3 domains of BNIP3 and NIX, play a major role in the induction of cell death mediated by mitochondrial depolarization, which may also serve as the event that initiates autophagy. BNIP3 or NIX induces autophagy under different conditions, which suggests that the ability to induce autophagy is an intrinsic property of these proteins. They are not homologous of any of the canonical autophagy proteins identified in yeast, but may recruit components of autophagy machinery to mitochondria. NIX induces autophagocytosis of mitochondria, mitophagy. Autophagy generates membranes which serve to compartmentalize destructive enzymes and perform a protective function. The relationships of these two functions are not clearly established (Zhang and Ney 2009).

7.6 Molecular Mechanisms of Necrosis Necrosis can be defined morphological by electron-lucent cytoplasm, swelling of cellular organelles, and loss of plasma membrane integrity. These events can be reproduced experimentally by impairing the ATP production of the cell because the  core events of necrosis are bioenergetic failure and rapid loss of plasma ­membrane integrity. These failures results from defined molecular events that occur in the dying cell, including increased mitochondrial reactive oxygen species (ROS)  production, channel-mediated calcium uptake, activation of non-apoptotic

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Fig. 7.5  Molecular events carried out during necrosis process

proteases in particular calpains and cathepsins, swelling of mitochondria, perinuclear clustering of organelles and/or enzymatic destruction of cofactors required for ATP production leading to ATP depletion (and ultimately plasma membrane rupture), cell lysis, and inflammatory response that may facilitate wound healing (Zong and Thompson 2006; Goldstein and Kroemer 2006; Degenhardt et al. 2006) (Fig. 7.5). The most studied systems are heterogeneous in terms of cell types and species: several mammalian cell types, the nematode Caenorhabditis elegans and the slime mold Dictyostelium discoideum. Various death conditions were used in experimental induction of necrosis; some of them were ischemia, cyanides, ligands of cell surface receptors, suppression of glucose, depletion of ATP, exposure to glutamate, etc. Most of these systems are similar in that they manifest early plasma membrane rupture, but no sign of apoptosis or autophagy. The chronological and molecular order of the events remains elusive. Taken alone each of these events is not specific to necrosis, and indeed some are shared with apoptosis; it is the accumulation of these events in an organized, programmed cascade of self-destruction that might define necrosis (Goldstein and Kroemer 2006). The concept of both programmed course and programmed occurrence is ­supported by several data. First, necrosis normally occurs in precise places and

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times during development and in adult tissue homeostasis, e.g. intestinal epithelial cells. Second, necrosis can be triggered by the occupation of specific plasma membrane receptors through their physiological ligands. This observation implies that specific signal transduction pathways are connected to the induction of necrosis rather than to the induction of other types of cell death. Third, susceptibility to necrotic cell death can be regulated by genetic and epigenetic factors. Fourth, the inhibition of some enzymes and processes can prevent necrosis, meaning that these factors have an active, decisive role in the lethal process. Fifth, inhibition of caspases can change the morphological appearance of cell death from apoptosis to autophagy or to necrosis. Thus, the same upstream signal can produce different types of cell death as a function of the activation or inhibition of catabolic enzymes in the cell (Goldstein and Kroemer 2006). Epithelial tumor cells incubated in hypoxic conditions undergo apoptosis cell death as the main process of death. BAX-/-, BAK-/- cells in the presence of BCL-2 go through autophagy. Autophagy may sustain viability in the short term and lead to cell death without inflammation in the long term. The combination of the inactivation of apoptosis and inhibition of autophagy by AKT activation and the reduction of the expression of the essential autophagy gene beclin1 (beclin1+/-), results in a necrotic cell death process (Degenhardt et al. 2006). In other models the depletion of ATP or the inhibition of caspases induce necrotic cell death (Goldstein and Kroemer 2006; Zong and Thompson 2006). Another process of inhibition of apoptosis and autophagy using zVAD also demonstrated that this double inhibition is responsible for induction of necrotic cell death (White 2008). Disruption of endoplasmic reticulum (ER) function leads to the initiation of a stress response known as the unfolded protein response, which goal is to resolve the ensuing stress; however, when this goal cannot be attained it induces cell death, which is primarily apoptosis. As in the previously described model, when normal cells or cells defective in apoptosis were used autophagy is induced. Prolonged ER stress on cells defective in apoptosis still results in cell death in a fashion resembling necrosis. This necrosis-like cell death is associated with autophagy. In response to ER stress, while autophagy serves as a survival response to delay apoptosis, it promotes cell death by necrosis in cells with impaired apoptosis (Ullman et al. 2008). These results strongly sustain the view that necrosis, in this context, is a programmed process of cell death. The default occurrence of necrotic cell death and its unmasking by inhibition of autophagy and/or apoptosis might reflect its early emergence in evolution, perhaps as the primordial eukaryotic cell death pathway (Goldstein and Kroemer 2006).

References Ankarcrona M, Dybukt JM, Bonfoco M et al (1995) Glutamate-induced neuronal death: a succession of necrosis or apoptosis depending on mitochondrial function. Neuron 15:961–973 Baehrecke EH (2005b) Autophagy: dual roles in life and death? Nat Rev Mol Cell Biol 6:505–510

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Bonfoco E, Krainc D, Ankarcrona M et  al (1995) Apoptosis and necrosis: two distinct events induced, respectively, by mild and intense insults with N-methyl-D-aspartate or nitric oxide/ superoxide in cortical cell cultures. Proc Nat Acad Sci USA 92:7162–7166 Degenhardt K, Mathew R, Beaudoin B et al (2006) Autophagy promotes tumor cell survival and restricts necrosis, inflammation, and tumorigenesis. Cancer Cell 10:51–64 Dypbukt JM, Ankarcrona M, Burkitt M et al (1994) Different prooxidant levels stimulate growth, trigger apoptosis, or produce necrosis of insulin-secreting RINm5F cells. J Biol Chem 269:30553–30560 Escobar ML, Echeverría OM, Ortiz R, Vázquez-Nin GH (2008) Combined apoptosis and autophagy, the process that eliminates the oocyte of atretic follicles in immature rats. Apoptosis 13:1253–1266 Fink SL, Cookson BT (2005b) Apoptosis, pyroptosis, and necrosis: mechanistic description of dead and dying eukaryotic cells. Infect Immun 73:1907–1916 Goldstein P, Kroemer G (2006) Cell death by necrosis: towards a molecular definition. Trends Biochem Sci 32:37–43 Huang Q, Wu YT, Tan HL et al (2009) A novel function of poly(ADP-ribose) polymerase-1 in modulation of autophagy and necrosis under oxidative stress. Cell Death Different 16:264–277 Kerr JFR, Wyllie AH, Currie AR (1972) Apoptosis: a basic phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer 26:239–257 Luciani MF, Kubohara Y, Kikuchi H et al (2009) Autophagic or necrotic cell death triggered by distinct motifs of the differentiation factor DIF-1. Cell Death Differ 16:564–570 Majno G, Joris I (1995) Apoptosis, oncosis, and necrosis. An overview of cell death. Am J Pathol 146:3–15 Proskuryakov SY, Gabai AG, Konoplyannikov AG (2002) Necrosis is an active and controlled form of programmed cell death. Biochem Mosc 67:387–408 Saelens X, Festjens N, Parthoens E et  al (2005) Protein synthesis persists during necrotic cell death. J Cell Biol 168:545–551 Shimizu S, Eguchi Y, Kamiike W et  al (1996) Induction of apoptosis as well as necrosis by hypoxia and predominant prevention of apoptosis by Bcl-2 and Bcl-XL. Cancer Res 56:2161–2166 Ullman E, Fan Y, Stawowczyk M et al (2008) Autophagy promotes necrosis in apoptosis-deficient cells in response to ER stress. Cell Death Diff 15:422–425 Walisser JA, Thies RL (1999) Poly(ADP-ribose) polymerase inhibition in oxidant-stressed endothelial cells prevents oncosis and permits caspase activation and apoptosis. Exp Cell Res 251:401–413 White E (2008) Autophagic cell death unraveled: pharmacological inhibition of apoptosis and autophagy enables necrosis. Autophagy 16:399–401 Yuan J (2006) Divergence from a dedicated cellular suicide mechanism: exploring the evolution of cell death. Mol Cell 23:1–12 Yuan J, Lipinski M, Degterev A (2003) Diversity in the mechanisms of neuronal cell death. Neuron 40:401–413 Zamzami N, Hirsch T, Dallaporta B et  al (1997) Mitochondrial implication in accidental and programmed cell death: apoptosis and necrosis. J Bionerg Biomembr 29:185–193 Zhang J, Ney PA (2009) Role of BNIP3 and NIX in cell death, autophagy, and mithophagy. Cell Death Diff 16:939–946 Zong WX, Thompson CB (2006) Necrotic death as a cell fate. Cell Develop 20:1–15

Part III

Process of Cell Death Embryonic Ovary

Chapter 8

Programmed Cell Death in Fetal Oocytes Francesca Gioia Klinger and Massimo De Felici

Abstract  In all mammalian species studied, a marked oocyte loss occurs during the fetal life before their enclosure within the primordial follicle. The concept that female mammals are born with all of the oocytes they will ever posses and the consequent notion that the age-related ovarian failure and menopause occur when the oocyte ovarian reserve is exhausted, render particular relevant to understand the reasons and the mechanisms of oocyte demise in the fetal ovary. Over the years, three main hypotheses to explain the cause of fetal oocyte death have been proposed: (1) the number of oocytes formed in the ovary is in excess respect to the supporting cell, (2) the process of cross over central in prophase I, requires critical molecular processes subjected to frequent errors leading to oocyte death and (3) most oocytes could sacrify themselves donating their cytoplasm content to a subset of surviving oocytes. Mainly during the last three decades, researchers have reported evidence favoring each of these hypotheses; thus suggesting that the survival or death of fetal oocytes depends on several conditions and mechanisms. The concept that cell death is a carefully controlled process both at genomic and molecular level termed apoptosis or programmed cell death (PCD) affirming at the beginning of 80 years, stimulated researchers to analyze in more details the morphological and molecular characteristics as well as the kinetics of the fetal oocyte elimination. In the next sections of the present chapter, we will critically review the principal studies carried out, mainly in the mouse, in our and other laboratories that are slowly disclosing the complexity of the fetal oocyte elimination.

8.1 Introduction The formation of germ cells in the female is a long complicated process called oogenesis, that begins with the formation of the germ cell precursors, the ­primordial germ cells (PGCs). Following migration into the gonadal ridges, PGCs differentiate F.G. Klinger (*) and M. De Felici Department of Public Health and Cell Biology, University of Rome, Tor Vergata, Italy e-mail: [email protected] G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary, DOI 10.1007/978-94-007-1134-1_8, © Springer Science+Business Media B.V. 2011

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into oogonia which, after a period of intense proliferation, enter into meiosis becoming primary oocytes. Entering meiosis does not occur synchronously and oogonia and oocytes may coexist within the fetal ovary for a certain period depending on the species. Prophase of meiosis I (MPI) can be divided into four distinct sub stages, leptotene, zygotene, pachytene and diplotene, defined by the chromosomal events occurring during each stage as well as the assembly of the synaptonemal complex (SC) needed for cross over. In the mouse, the entire population of PGCs/ oogonia enter meiosis between 13.5 and 14.5 days post coitum (dpc). During the subsequent 5–6 days, the oocytes progress through the MPI stages and around birth they progressively undergo meiotic arrest at a stage known as dyakinesis or dictyate. At this time, oocytes are individually enclosed by a specialized lineage of ovarian somatic cells, the pre-granulosa cells, to form primordial follicles. Within the primordial follicle, the oocyte remains arrested, resuming and completing meiosis only after ovulation and fertilization, respectively, at some point in adult life. The apparent disappearance in the fetal ovaries of PGCs/oogonia able to mitotic proliferation results in a fixed number of oocytes that if lost cannot be renewed. In mammals, actually, the oocyte number undergo a continuous physiological decline so that the oocyte stockpile (ovarian reserve) is exhausted about half to two-thirds of the way through the lifespan of the animal which leads to ovarian failure or, in other words, to menopause. In all mammalian species studied, a marked oocyte loss occurs during the fetal life before their enclosure within the primordial follicle. In fact, it has been reported that over two-thirds of the female germ cell pool is lost by the time of birth in rats (Beaumont and Mandl 1961), mice (Borum 1961; Baker 1963; Bakken and McClanahan 1978) monkeys (Nichols et al. 2005) and humans (Baker 1963; Forabosco et al. 1991; Faddy et al. 1992; Hansen et al. 2008). It has been estimated that in mice, the number of oocytes decreases from around 20,000 at 13.5 dpc to about 6,000–10,000 after 6  days, at birth (Burgoyne and Baker 1985; Tam and Snow 1981); in humans, 7 millions of germ cells in the fetal ovaries at around week 20 of gestation are decimated to 1–2 millions oocytes in early neonatal life (Gondos 1978). The concept that female mammals are born with all of the oocytes they will ever posses and the consequent notion that the age-related ovarian failure and menopause occur when the oocyte ovarian reserve is exhausted, render particular relevant to understand the reasons and the mechanisms of oocyte demise in the fetal ovary. Over the years, three main hypotheses to explain the cause of fetal oocyte elimination have been proposed: (1) the number of oocytes formed in the ovary is in excess respect to the supporting ovarian cells; surplus oocytes dye for shortness of nutrients/growth factors so that others have the resources to develop properly, (2) many fetal oocytes are defective for incomplete DNA repair or errors in chromosome synapses occurring during homologous recombination and (3) most oocytes sacrifice themselves donating their cytoplasm content to a subset of surviving oocytes. In all these circumstances, oocyte death could occur by either extracellular or intracellular inducers.

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8.2 Defining the Form/s of Fetal Oocyte Death The first observation in studying the degeneration of fetal oocytes is that it occurs with the characteristics of PCD: it takes place in a predictable place (fetal ovary) and time (during MPI) and results in rapid elimination of a large number of cells (oocytes) without any inflammatory reaction. As largely described in other chapters of this book, at morphological level, two types of PCD have been described: apoptotic cell death, which includes the morphological changes of cell shrinkage, membrane blebbing and extensive chromatin condensation followed by nucleus fragmentation and autophagic cell death, characterized by the formation of autophagic vacuoles in the cytoplasm of dying cells. Early studies carried out by light and electron microscope (TEM) in several mammalian species in vivo (reviewed in Gondos 1978) and more recent in vivo and in vitro researches on mouse (Coucouvanis et al. 1993; Pepling and Spradling 2001; Lobascio et al. 2007a, b), report morphological description of degenerating fetal oocytes compatible with both forms of PCD. It is to be pointed out, however, that in such studies, condensation rather than margination of chromatin and absence of nuclear fragmentation typical of classical apoptosis were reported. Since on the basis of biochemical aspects various types of PCD and mixed forms of apoptosis/autophagy can be distinguished (Kroemer et al. 2009), several methods for detecting fetal oocyte death have been employed in attempting to more precisely define the type and the molecular mechanisms of cell death in such cells. In this regard, distinction between in vivo and in vitro studies must be performed because under diverse conditions stimuli and mechanisms of PCD can be different.

8.2.1 In Vivo Studies Cleavage of DNA into oligonucleosomal size fragments is an integral part of typical apoptosis. This DNA fragmentation is often analyzed using agarose gel electrophoresis to demonstrate a “ladder” pattern at about 200-base pairs (bp) intervals. Atypical apoptotic DNA ladders were actually detectable by autoradiographic analysis of DNA extract form 15.5 dpc to day 0 mouse ovary likely due to fragmented DNA in the nucleus of degenerating oocytes (Ratts et al. 1995; Greenfeld et al. 2007). Fluorescent activated cell sorter (FACS) analyses confirmed the presence of degradated DNA in a subset of small size oocytes showing reduced DNA content (hypodiploid cells) obtained from mouse ovaries at 13.5 dpc and increased on 15.5 and 17.5 dpc (Coucouvanis et  al. 1993; our unpublished observations). Moreover, several authors using the deoxynucleotidyltransferase-mediated dUTP nick-end labelling (TUNEL) histochemistry to detect exposed 3-’OH end of DNA fragments in single cells were able to identify fetal oocytes positive to this staining scattered throughout ovary tissue sections (Pesce et al. 1997; Pepling and Spradling 2001; Greenfeld et al. 2007; Rodrigues et al. 2009; Fig. 8.1a–c) or in cytospreads (Ghafari et  al. 2007). TUNEL staining has been also applied coupled with other apoptosis markers such as cleaved poly(ADP ribose) polymerase (PARP) and active

Fig. 8.1  Detection of oocyte apoptosis by morphology and TUNEL assay in ovary tissue ­sections. Top panels: (a) Paraffin section of a 16.5 dpc ovary stained by TUNEL histochemistry. Note several TUNEL positive oocytes (arrowheads). Bar 150 mm; (b) and (c) subsequent paraffin sections of 16.5–17.5 dpc ovary showing: (b) TUNEL-positive oocytes and (c) Hematoxilin stained oocytes. Note that most of TUNEL-positive oocytes show apoptotic morphologies (arrowheads) and that some oocytes with degenerating features are TUNEL-negative (arrows). Bar = 50  mm. Bottom: Percentage of oocyte showing TUNEL staining in tissue sections of fetal ovaries of ­different developmental ages; (d) The percentage of apoptotic germ cells in tissue sections are plotted from 13.5 dpc to 7 dpp. All apoptotic cells in the ovaries were detected by using the TUNEL method

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caspases 3 and in association with fetal oocyte markers (GCNA1, VASA or SCP3) for more precise immunoidentification of degenerating oocytes (Pepling and Spradling 2001; Ghafari et al. 2007; Rodrigues et al. 2009). A quantitative evaluation of the number of TUNEL positive oocytes in ovary tissue sections showed small percent of positive cells at midgestation ages ranging from 0.5% to 2% and increasing (8–10%) around or early post birth (Pesce et  al. 1997; Pepling and Spradling 2001; Fig. 8.1d); most of these oocytes resulted also positive to cleaved PARP (Pepling and Spradling 2001). Notably, in these studies, fragmented oocytes and TUNEL-positive somatic cells were never observed. On the other hand in some instances, we observed oocytes in advanced stages of degeneration, showing necrotic (low cytosolic density and chromatin residues) rather than apoptotic features which resulted generally TUNEL-negative (Pesce et al. 1997). TUNEL on tissue sections was used successfully to correlate cell death in human fetal oocytes by Modi et al. (2003), who found that 3–7% of oocytes were TUNEL positive between weeks 13 and 23 in normal ovaries, rising to more than 50% in Turner’s syndrome ovaries, where extensive prenatal loss of oocytes occurs. Like Modi, Albamonte et  al. (2008) observed low levels (less than 10%) of TUNEL-positive oocytes throughout the second trimester, but they found a higher incidence (about 20%) at 18–20 weeks. It is to be pointed out that a small percentage of cells found in apoptosis at a single time may signify a large accumulated effect of apoptosis over time. In fact, it has been calculated that apoptotic cells are cleared within 1–2 h by neighbouring phagocytic cells, making the total amount of apoptosis in a tissue difficult to assess. Assuming a constant rate of apoptosis and a window of detection of dying cells of 3 h, a finding of 1% apoptosis at a single time would equate to a 56% loss of cells over 7 days (the standard period of MPI in the mouse). It is to be also noted that in cytospreads the number of TUNEL positive oocytes may result much higher, up to 30–40%, than in tissue sections (mouse: Ghafari et al. 2007; human: Hartshorne et al. 2009). This finding, that not necessarily concerns all cytospread preparations (our unpublished observations, see below), is likely due to increased accessibility of reagents to DNA and TUNEL staining of physiological DNA double strand breaks (DSBs) generated by SPO11 in germ cells entering into meiosis (see the review by Keeney and Neale 2006 and the companion Chap. 9), and complicates the interpretation of certain results. Taken together, these observations identify apoptosis as the likely mechanism of PCD in fetal oocytes. However, such process seems, perhaps for the distinctive meiotic status of the oocyte, to differ from classical apoptosis for several features (i.e. absence of chromatin margination and nuclear fragmentation, atypical DNA ladder, in some case absence of TUNEL staining). A marked amplification of the lysosome compartment recently described by Rodrigues et  al. (2009) in mouse oocytes around birth, supports the notion that mechanisms distinct from classical apoptosis are involved in oocyte loss at least at the perinatal period. In the aim to further characterize this process and identify causes and molecular pathways involved, we devised an in vitro culture system of isolated oocytes that, with the limits of the culture methods, allowed to obtain some useful information and rose new questions.

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8.2.2 In Vitro Studies Under our culture system, we observed that 15.5 dpc mouse oocytes were able to progress into MPI stages and underwent a peak of TUNEL positivity ranging from 50% to 70% after 3–4 days of culture. Since a similar pattern for Annexin V staining and pancaspase activity was observed, this confirmed apoptosis as the major form of cell death in fetal oocytes at least under our culture conditions (Lobascio et al. 2007a, b). This was also supported by the finding that DNA ladders and cleaved PARP were detectable in cultured oocyte at later times of culture and that caspase 2, a major effector of cell death in oocytes (Bergeron et al. 1998), was expressed in its activated form in cultured oocytes (Lobascio et al. 2007b). As for in vivo studies, however, some results of the in vitro oocyte culture were not entirely consistent with the oocyte apoptosis paradigm. For example, TEM observations of oocytes cultured for 3–4 days revealed recurrent atypical apoptotic morphologies characterized by the absence of chromatin margination and nuclear fragmentation; oocytes with autophagic and necrotic features were often observed. Moreover, pancaspase and caspase 2 inhibitors were able to slow down, but not to abolish, oocyte cell death. This prompted us to investigate the possibility that fetal oocytes undergo forms of cell death alternative or parallel to apoptosis eventually converging in DNA fragmentation revealed by TUNEL. The significant reduction of the number of TUNEL positive oocytes by the calpain inhibitor I, evidenced after 4 days of culture and the increased number of such oocytes both at day 3 and 4 caused by incubation in rapamycin (mTOR inhibitor) supported activation of a caspase-independent and autophagic pathways in the oocyte cell death. Furthermore, we observed in cultured oocytes that a progressive reduction of the expression and cytoplasmic staining of the apoptosis inducing factor (AIF), a caspase-independent mitochondrial death effector responsible for partial chromatinolysis (Daugas et  al. 2000), occasionally parallel to AIF and TUNEL nuclear staining. Finally, cytoplasmic staining for Beclin1, a protein part of the PI(3) kinase class III (PI(3)KC3) lipid-kinase complex that induces autophagy, was markedly increased from the beginning to day 3 of the culture (De Felici et al. 2008). Other in vitro culture results reported in the recent paper by Rodrigues et al. (2009) also support a role for autophagy in oocyte degeneration around the perinatal period. These authors found that starvation of newborn mouse ovaries in culture medium without serum caused a marked increase of the number of oocytes stained for acidified lysosomes, a feature typical of autophagy. Moreover, inhibitors of autophagy, but not of apoptosis, reduced oocyte acidification induced by serum starvation and expression of protein mediators of both autophagy (LC3B and beclin1) and apoptosis (caspase3 and cathepsinD) in the cultured ovaries.

8.3 Growth Factor-Regulated Oocyte Death PGCs/oogonia are known to be dependent for their survival and proliferation on several growth factors (for a review see De Felici 2000). The notion that also the survival/death of fetal oocytes is dependent on growth factors came from recent

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in vitro culture studies of whole ovary or isolated oocytes showing that SCF (stem cell factor) also known as Kit Ligand (KL), LIF (leukemia inhibitory factor) or IGF-1 (insulin growth factor-1), were able to prevent mouse oocyte degeneration and favours their meiotic progression (Pesce et  al. 1993; Morita et  al. 1999; Lyrakou et al. 2002; Lobascio et al. 2007b). In line with these results, it has been found that fetal oocytes express during late stages of MPI the SCF receptor KIT and constant levels of the IGF-1 receptor-1 (IGFR-1) throughout MPI (Manova et al. 1990; Klinger and De Felici 2002; Doneda et al. 2002). Although the oocyte pro survival growth factors are likely to be produced by the ovarian somatic cells, Doneda et al. (2002) have also found that just before birth oocytes express either SCF and KIT suggesting also a paracrine role for this system. According to Morita et al. (2001), fetal oocytes express also type 1 interleukin receptor-1 (ILR-1) and the addition of IL-1a/b to the culture medium significantly increases the oocyte survival in fetal ovaries cultured in  vitro. An unexpected role for neurotrophins, growth factors originally discovered in the nervous system, namely neurotrophin 4/5 (NT4/5) and brain-derived neurotrophic factor (BDNF), in favouring the survival of fetal mouse and human oocytes, has been also reported (Anderson et al. 2002; Spears et al. 2003; Paredes et al. 2004). NT4/5 and BDNF are present in the fetal ovary and oogonia and oocytes express their favourite receptor TrkB, preferentially a truncated isoform. Ablation of such receptor result in severe depletion of primordial follicle at birth (Spears et al. 2003; Paredes et al. 2004). All together these results gain strong support to the view that limited amount of growth factors in the microenvironment is one of the cause of the fetal oocyte dismiss and consequently that growth factors withdrawal can trigger apoptosis in such cells. Unfortunately proves of such process in vivo are still lacking. Some results support an opposite extrinsic way through which fetal oocyte could be eliminated: growth factors or compounds produced by ovarian somatic cells could exert a proapoptotic action on oocytes. Sakata et al. (2003) showed that the absence of Kit signalling activates Fas cell death pathways within oocytes of adult mice. Fas has been also implicated in the loss of postnatal oocytes in MRL+/+ mice, a strain that exhibits remarkable regenerative abilities (Guo et  al. 2002). Moreover, rat and mouse oocytes of primordial follicles at birth can be induced to undergo apoptosis by TNFa, likely produced by the granulosa cells (Morrison and Marcinkiewicz 2002; Greenfeld et al. 2007). Liu et al. (2010) on the basis of the depletion of fetal oocytes observed as a consequence of ablation of Wnt4 and b-catenin genes, proposed that the balance between somatic cell signalling (Wnt4/ b-Catenin) and activin B is critical for the maintenance of female germ cells during embryonic stage and that fetal oocyte elimination results from a shifted balance toward the proapoptotic action of activin B on the oocytes. It has been recently shown that in the Drosophila ovaries, somatic granulosa-like cells can be induced by rapamycin simulating cell starvation to destroy intact oocytes without a requirement for earlier oocyte compromise. The response to rapamycin was found to be conserved in mammals, as mouse ovarian follicles cultured in vitro in the presence of this drug showed a rapid destruction of the oocyte by adjacent granulosa cells (Thomson and Johnson 2010).

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It is unknown, however, how such mechanisms of active oocyte elimination might be normally activated during MPI and on which basis oocyte are selectively eliminated. This way of cell elimination could imply some sort of recognition of oocytes to be eliminated by the somatic cells as it occurs in the immune system and be part of the oocyte quality control process discussed below.

8.3.1 Apoptotic and Autophagic Pathways It has been shown by many studies that withdrawal of a variety of growth factors in several cell systems results in mitochondrial-dependent activation of apoptosis involving cytochrome c release from mitochondria, the formation of the apoptosome and activation of effector caspase. It has been demonstrated that the binding of a growth factors to its receptor results in phosphorylation events involving many different kinase pathways, including the PI3K/AKT, MAPK and GSK-3 pathways that eventually converge on Bcl-2 family members and other players of the machinery of the intrinsic apoptotic pathway (Kennedy et al. 1997; Songyang et al. 1997; Maurer et al. 2006) (Fig. 8.2).

Fig. 8.2  A schematic drawing of the possible molecular pathways involved in the apoptotic and autophagic forms of mouse fetal oocyte programmed cell death supported by the in  vivo and in vitro studies reviewed in the present chapter. For details, see text. GC granulosa cell

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Little is currently known about the intracellular effectors utilized by the ligand-activated receptors providing anti apoptotic signals in fetal oocytes. In particular nothing is still known about mitochondria and apoptosome involvement in fetal oocyte apoptosis. In oocytes of primordial follicles of rat and mouse, it has been found that SCF promotes survival by upregulating Bcl-2 and Bcl-xl and downregulating Bax proteins through the PI3K-AKT signaling pathway (Jin et al. 2005; Reddy et al. 2005). The only information available in fetal oocytes is that in cultured mouse ovaries, pharmacological inhibition of PI3K abolish the antiapoptotic actions exerted by SCF, LIF and IGF-1 on these cells (Morita et al. 1999). In this regard, preliminary results obtained in our laboratory have shown that stimulation of enriched population of mouse fetal oocytes by SCF and IGF-1 resulted in a robust increase of AKT and S6 but not of ERK 1-2 kinase phosphorylation (Klinger 2002). It is reasonable to think that a limited amount of growth factors can be sensed by the cells by the same players involved in detecting nutrient shortness. In this regard, autophagy is a major mechanism by which a starving cell reallocates nutrients from unnecessary processes to more-essential processes. Interestingly, mTOR (mammalian target of rapamycin) kinase, a key player of autophagy appears to be a downstream target for the cell prosurvival growth factor action. Under normal conditions, in the presence of growth factors and nutrients, mTOR is constitutively activated. This activation is generally partly maintained through the insulin receptor or insulin-like growth factor receptor pathways, via a cascade that involves the activation of phosphatidylinositide-3kinase (PI3K), then phosphatidyl inositol-3,4,5 phosphate-mediated activation of AKT/PKB-mediated phosphorylation of mTOR. In turn, mTOR phosphorylates S6-kinase and 4E-BP1 to enhance the translation of selected mRNA transcripts involved in multiple metabolic pathways. Another target of mTOR implicated in the regulation of protein translation is PP2A which dephosphorylates the mTOR substrates S6-kinase and 4E-BP1(reviewed in Castedo et  al. 2002). When mTOR is reduced or inhibited, autophagy and/or apoptosis is activated. It is likely that autophagy is first activated as a survival strategy to provide nutrients to the cell and then evolves in cell death if nutrients and growth factors depletion continue. Growing evidence supports the notion that apoptotic pathways are downstream mTOR activation. As a possibility, its downstream target S6-kinase, which can bind to mitochondrial membranes, can phosphorylate the pro-apoptotic molecule Bad on serine 136, a reaction which disrupts Bad binding to the mitochondrial death inhibitors Bcl-xl and Bcl-2 and thus inactivates Bad (Harada et al. 2001). In our culture system, we found that rapamycin was able to increase oocyte apoptosis evaluated by the TUNEL staining. Interestingly, the SCF anti-apoptotic action on oocytes did not appeared dependent on the downstream activation of mTOR since it was not affected by the presence of rapamycin in the culture medium. On the other hand, SCF was able to abolish the proapoptotic effect of rapamycin suggesting cross-talk between these pathways (Lobascio et al. 2007b).

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8.3.2 Bcl-x Family Members and Caspases In the companion chap. 9, we discuss the involvement of Bcl-x family genes and caspases in the possible DNA-damage depended cell death of the fetal oocytes; in the present section, we report evidence about their role in the growth factor-­ regulated oocyte death. The demonstration that Bcl-2 is able to prevent apoptosis following growth ­factor withdrawal was one of the seminal experiments elucidating the anti apoptotic function of Bcl-2 (Vaux et al. 1988). Yet there has been an incomplete understanding of the signalling events linking growth factor withdrawal to Bcl-2 family members and other players of the machinery of the intrinsic apoptotic pathway. In this regard, in the previous section we reported that Bcl-2 family members are also involved in authopagy since the mTOR-S6-kinase-Bad pathways can trigger autophagic cell death. As also reported in the previous section, in oocytes of primordial follicles of rat and mouse, SCF promotes survival by upregulating Bcl-2 and Bcl-xl and downregulating Bax through the PI3K-Akt signaling pathway (Jin et  al. 2005; Reddy et al. 2005). Studies of the functions of members of the Bcl-2 gene family suggested that apoptosis was controlled by a rheostat in which anti-apoptotic proteins like Bcl-2 bound and sequestered proapoptotic proteins like Bax. Fetal oocyte express Bcl-X (mostly Bcl-xl) and Bax (De Felici et al. 1999; Rucker et al. 2000; Klinger 2002). Bax expression has been also detected in oocytes of the human fetal ovary from the 14th week until term (Vaskivuo et al. 2001). Although mouse fetal oocytes seem less sensitive than male germ cells to the haplo-sufficiency of Bcl-x (Kasai et  al. 2003), in some instance the rheostat Bcl-x/Bax appears to regulate fetal oocyte apoptosis. Actually, Rucker et al. (2000) showed that in bcl-x hypomorph mice a marked demise of fetal oocytes occurred around 15.5 dpc and the ablation of bax restored the oocyte number. We found, however, that SCF and IGF-1 do not exert their oocyte prosurvival action on the Bcl-xl/Bax rheostat. In fact, both factors were unable to consistently influence the expression of Bcl-xl in cultured oocytes and, although the oocytes with apoptotic features accumulated Bax protein, neither SCF nor IGF-1 influenced this occurrence (De Felici et al. 1999; Klinger 2002). The existence of Bax-independent mechanisms of fetal oocyte elimination is supported by two papers showing that between 14.5 dpc and birth oocytes are eliminated at the same rate either in wild type and bax-/- ovaries (Alton and Taketo 2007; Greenfeld et al. 2007). These papers, in contrast to Perez et al. (1999), also observed that at birth the number of the primordial follicles was significantly higher in Bax null than in wild type ovaries. In accord with our previous results (De Felici et al. 1999), the authors concluded that the regulatory activity of Bax occurs during PGC development. Other studies in mouse fetal oocytes indicate that Bcl-2 protein was barely immunodetectable in oocytes entering into meiosis and its expression did not change during the stage of MPI (De Felici et al. 1999). However, others reported that in Bcl-2-/- females the ovarian reserve oocytes was reduced (Ratts et al. 1995) and over expression of Bcl-2 enhanced follicular endowment (Flaws et al. 2001),

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leaving open the possibility that Bcl-2 can play perhaps an indirect role in oocyte dismiss likely by sustaining granulosa cell survival. As reported in other chapters of this book, the caspase family of cysteine ­protease comprises a group of apoptosis-inducing enzymes that are sub typed by differences in their pro-domain, substrate specificity and their position in the caspases cascade. In this regard the caspases can be divided in initiator (caspases 2, 8, 9 or 10) and effector (caspases 3, 6 or 7) caspases. The only caspase whose activity appears to be involved in the growth factor-regulated oocyte death is caspase 2. Caspase 2 has a unique position in the apoptotic pathways, it has the molecular characteristics of initiator caspases and at the same time it can be activated by effector caspases (Slee et al. 1999). Higher numbers of primordial follicle are endowed in the ovaries of neonatal caspase 2 deficient female mice and caspase 2 deficiency prevents fetal oocyte degeneration resulting from cytokines, namely Interleukin-1a and 1b, insufficiency (Bergeron et  al. 1998; Morita et  al. 2001). Moreover, the active forms of caspases 2 is detectable in mouse oocyte undergoing apoptosis in culture (Lobascio et al. 2007b). On the other hand, the dismiss of fetal oocytes does not appear to require caspase 3 (Matikainen et  al. 2001). It is to be noted that if growth factor-dependent oocyte degeneration occurs or involves at some points autophagy, caspase activation can be dispensable in such process. This explains the absence of detectable caspase activity in a subset of cultured TUNEL positive oocytes and the partial and temporary protective effects of caspase inhibitors on the oocyte degeneration in culture (Lobascio et al. 2007b).

8.4 Quality Control-Dependent Oocyte Death Two models could explain how germ cell death maintain oocyte quality. In the first, apoptosis eliminates defective oocytes from the germ line. In the second, apoptosis modulates the number of developing oocytes to help allocate resources properly.

8.4.1 Crossover Defects as Possible Cause of Oocyte Elimination The complexity of the processes occurring during crossover has led to the idea that fetal oocytes could undergo apoptosis as a consequence of unrepaired DNA DSBs or defects in synapses (asynapses) detected by meiotic checkpoints. Alternatively, as reported in a previous section, the quality of oocytes could depend on other factors such as the number of mitochondria or the capability to express adequate number of receptors for growth factors or paracrine/autocrine factors. While the existence of these last two quality oocyte mark is only hypothetical, evidence supporting the notion of genomic defects as cause of oocyte death comes from studies of impaired germline development either in male and female mice with genetic

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mutations causing abnormalities in cross over. These studies and possible molecular pathways involved are discussed at length in the companion chap. 9. Although there is little doubt that in some instances cross over defects are able to trigger meiotic arrest and elimination of fetal oocytes, the efficiency of these checkpoints appears quite lower in female than in male meiotic cells and little evidence exists that such mechanisms are actually responsible for the physiological prenatal massive oocyte death. Moreover, the form of PCD elicited by the meiotic checkpoints remained to be clearly determined. In the present section, we limit the discussion to a few studies that have tried to directly correlate the presence of unrepaired DNA and asynapses in fetal oocytes in  vivo and in  vitro to expression of apoptotic markers. In one of these studies, mouse oocyte synaptonemal axial elements were labeled in cytospreads by SCP3 antibodies and immunocytochemistry for TUNEL and/or cleaved PARP was used as apoptotic markers (Ghafari et al. 2007). These authors found that while at the beginning of every MPI stages (mainly at leptotene and zygotene), TUNEL was able to stain a subset of oocytes freshly collected from the fetal ovary with physiological DNA DSBs or in apoptosis, later the labeling was progressively restricted to apoptotic oocytes. They concluded that PARP or double PARP-TUNEL positivity, ranging from 20% to 40% of the scored oocytes, identified early and late apoptotic oocytes, respectively. However, fragmentation of axial elements indicative of SC assembly defects was not consistently associated to PARP-TUNEL positivity. Similar results as regard TUNEL, were obtained by the same group in human oocytes (Hartshorne et al. 2009), which show high frequency of SC fragmentation, synaptic anomalies or both (zygotene: up to 70%; pachytene around 18–30%) (Tease et  al. 2006). We also investigated the association of TUNEL staining with the presence of DNA DSBs and asynapses evaluated by immunocytochemical labeling of the phosphorylated histone H2AX (gH2AX) in mouse fetal oocytes (unpublished observations). Previous studies either in mouse and humans have shown that gH2AX labeling appears at leptotene in large, cloudlike chromatin regions corresponding to DNA DSBs throughout the oocyte nucleus. Subsequently, the repair of DSBs results in the disappearance of most of the gH2AX signal at the pachytene stage. However, a second and independent wave of gH2AX staining appears in late zygotene and pachytene oocytes; this second labelling appears in spots distinct from that large cloud-like staining of leptotene/­ zygotene and is associated with the asynapsed axial core of meiotic chromosomes (de Vries et al. 1999; Turner et al. 2005; Garcia-Cruz et al. 2009). In line with these observations, we observed that virtually all leptotene and zygotene oocytes freshly isolated from mouse fetal ovaries showed gH2AX cloud-like staining due to DNA DSBs. At pachytene and diplotene, around 40% of the oocytes were gH2AX positive, mostly (>90%) with spot labelling (asynapsis) and a few with cloud-like DSB labelling. At all stages examined, only a few oocytes (0.5–3%) showed TUNEL positivity that appeared unrelated to any distinguishable MPI stages and gH2AX labelling pattern. Since we knew that the most part of isolated leptotene/zygotene oocytes cultured for 3–5 days are able to reach pachytene and diplotene stages or undergo apoptosis as evaluated by increased frequency of TUNEL positive oocytes

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(Lobascio et al. 2007a, b), we reasoned that a correlation between TUNEL staining with MPI stages and gH2AX labelling pattern should be more easily achieved in oocytes in culture where apoptotic cells are more easily captured. Although it was often difficult to determine the MPI stage of the TUNEL positive oocytes by SC labelling, in most cases they appeared in pachytene and diplotene. Moreover, TUNEL positivity was coupled mostly to gH2AX diffuse throughout the nucleus with variable number of patches underlying indicative of widespread DNA fragmentation (Pelzel et  al. 2010) and rarely (