Human Reproduction Update, Vol.8, No.3 pp. 243±254, 2002
Follicle culture in reproductive toxicology: a tool for in-vitro testing of ovarian function? R.G.Cortvrindt1 and J.E.J.Smitz Centre for Reproductive Medicine, Follicle Biology Unit, University Hospital and Medical School, Dutch-speaking Brussels Free University, Brussels, Belgium 1
To whom correspondence should be addressed at: Centre for Reproductive Medicine, Follicle Biology Unit, University Hospital and Medical School, Dutch speaking Brussels Free University, Laarbeeklaan 101, B-1090 Brussels, Belgium. E-mail:
[email protected]
Public opinion and of®cial bodies place great emphasis on the reduction, re®nement and replacement of the use of laboratory animals in testing protocols. In the ®eld of toxicology, major efforts are being made to commit to this goal. The testing of reproductive function is currently still performed by in-vivo tests, mainly in rodents. In the past, follicle culture models were developed for the in-vitro production of mature oocytes and used to study the process of folliculogenesis and oogenesis in vitro. These culture systems might be able to acquire a place in fertility testing, replacing in-vivo studies for ovarian function and female gamete quality testing. The pro®ciency data from a wellcharacterized follicle culture system suggest that this bioassay might be of potential use for in-vitro screening of xenobiotic substances affecting ovarian function and fertility. Keywords: follicle/in-vitro/oocyte/ovary/reprotoxicity
TABLE OF CONTENTS Introduction Female reproductive toxicity Female reproductive function testing Follicle culture systems In-vitro bioassay for testing ovarian function and female gamete quality Conclusion Acknowledgements References
Introduction New chemical entities (NCE) are produced in their thousands each year by the chemical and pharmaceutical industries. The pharmaceutical industry identi®es yearly, hundreds of potential new drugs. Before their acceptance for testing in humans (or animals), these compounds (IND = investigational new drugs) need extensive chemical and physical characterization besides testing for pharmacological activity. During this phase and throughout the clinical test phases I to III, toxicological effects are documented in safety studies. The effects on reproductive function are evaluated relatively late in this drug development process, being performed during the clinical phase II trials (Figure 1). The testing of toxic effects on reproductive function is complex, since reproduction is a continuous cycle that takes Ó European Society of Human Reproduction and Embryology
place in two individuals (Mattison et al., 1990). In reproductive toxicity, two sub®elds should be considered: (i) toxic effects on fertility; and (ii) toxic effects on development. Developmental toxicology covers toxic effects during pregnancy in females, including prenatal and post-natal effects on the development of the offspring until adulthood (Chapin, 1998). Fertility testing comprises analysis of adverse effects on libido, sexual behaviour, spermatogenesis, oogenesis, fertilization and implantation. Guidelines for reproductive toxicology testing have been issued by The International Conference on Harmonisation of Technical Requirements for Registration of Pharmaceuticals for Human use (ICH) and by the Organisation for Economic Co-operation and Development) (OECD) (MuÈller et al., 1999). Both organizations aim to standardize test procedures with the aim of saving resources, bringing treatments more rapidly to patients (ICH), and safeguarding the environment and its population from toxic insults (OECD). Although basic study designs are similar for pharmaceutical and environmental agents, somewhat different approaches are used, in part because of the different parameters of exposure to potential toxicants in these two ®elds. The therapeutic levels of exposure of potential drugs are known and exposures are reasonably well controlled. Exposures to environmental agents are dif®cult to predict, are usually involuntary and ill-de®ned, and also can last a lifetime. Currently, the guidelines produced by ICH and OECD for reprotoxicity testing are based mainly on in-vivo
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Figure 1. Pipeline drug development. The scheme shows the different phases of drug development on a time scale. Safety testing is a continuous process starting in the pre-clinical phase and ending with continued post-marketing surveillance. Reproductive toxicology testing is positioned during clinical test phase II. FDA = Food and Drug Administration; INA = Investigational New Drug; NCE = New Chemical Entities; NDA = New Drug Application; RT = Reprotoxicity testing.
animal studies. Nevertheless, animal welfare is stressed and great concern exists about the lack of alternative test systems. Efforts are being made to identify and develop alternative methodologies that can replace or reduce laboratory animal use and promote animal welfare (Davila et al., 1998; Jackson, 1998). As yet, no adequate in-vitro models have been validated to pinpoint potentially hazardous products with regard to the reproductive system. Research into the development of alternative tests has concentrated mainly on teratogenicity, which is only one of the manifestations of adverse effects on development and does not include fertility (Spielmann, 1998). For the class of endocrine disrupters, cell and cell-free assays are available for the screening of potential hazards (Andersen et al., 1999). However, these tests have no signi®cance for the in-vivo effect or mechanism of action and can be used only in preliminary investigations (pre-screening or priority selection) or in secondary screening protocols (Zacharewski, 1998).
Female reproductive toxicity Female reproductive toxicity is, in many unique and challenging ways, different from toxicities to other systems such as the renal, cardiac or pulmonary systems. Whereas toxicity in the latter typically involves the development of disease in an exposed individual, exposures of the reproductive system can be toxic but remain unnoticed over an extended period of time (Mattison et al., 1990). In humans, the effect will not be detected where steps are taken to avoid fertility. Effects on the genetic constitution of the gametes will be discovered only in the following generation(s).
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Within the frame of reproductive toxicology in females, there is an urgent need for toxicity-testing systems which are predictive at the early stage, provide a better insight into the mechanisms leading to reproductive failure, and detect in a quantitative and qualitative way the toxic damage to the gametes produced (Ecobichon, 2001). Toxicity to the reproductive system in the post-natal female
Toxicity can harm the adult female reproductive system at two distinct levels: ®rst by disrupting the reproductive function of the exposed female; and second by altering the viability and morbidity of the offspring. Disruption of reproductive function in the adult female
In females, normal reproductive function involves the appropriate interaction of the central nervous system, anterior pituitary, oviducts, uterus, cervix and ovaries. During the reproductive years, the ovary is the central organ in this axis. The functional unit within the ovary is the follicle, which is also a morphological unit composed of three basic cell types: theca cells, granulosa cells, and the oocyte. The somatic compartment synthesizes and secretes hormones (steroids and growth factors) necessary for the orchestration of the inter-relationship between the other parts of the reproductive tract and the central nervous system (Mattison, 1993; Hoyer, 1997). At birth, the pool of primordial resting follicles (oocytes) is de®nitively set; any destruction at this level is de®nitive and cannot be restored (for reviews, see Morita and Tilly, 1999; Reynaud and Driancourt, 2000). The follicle is at the basis of reproduction control through its principal products: oocytes,
Follicle culture in reproductive toxicology steroids and speci®c protein hormones. These are produced during the process of folliculogenesis, which involves recruitment of primordial follicles out of the resting into the growing pool, progressing through the several developmental stages [the preantral stage (primary, secondary) and the antral stage (tertiary) onto the Graa®an stage (pre-ovulatory)]. The development of the pre-antral follicle involves oocyte growth, granulosa-cell proliferation and recruitment of the endocrine theca cells from interstitial tissue. With the acquisition of a large ¯uid-®lled antrum, the granulosa cells differentiate into two subpopulations with distinct endocrine characteristics. Along with these structural changes, the different follicular cells acquire the intracellular machinery required to synthesize and secrete steroids and protein hormones and express stage-speci®c receptors to respond to endocrine stimuli. During the process of follicular growth, the enclosed immature oocyte is nursed so as to grow and develop into a mature fertilizable oocyte. Each stage of follicular development exhibits unique patterns of gonadotrophin sensitivity, steroid production, and feedback pathways to keep the hypothalamic±pituitary±gonadal axis in concert and aligned with the rest of the reproductive tract (Richards, 2001). The regulation of every component of the follicular unit (theca cell, granulosa cell, oocyte) can be affected at each speci®c level and as such disturb the reproductive system at different levels. This creates a situation in which the pattern of infertility induced by a particular agent is dependent on the types of follicle(s) which are affected. The following examples clarify this. Toxicity to primordial follicles: this will not bring about immediate signs of infertility, but will ultimately shorten the reproductive lifespan, resulting in precocious irreversible infertility. After a certain period of time, the exact mechanism of destruction will be impossible to trace. Only several years after exposure, will the female become subfertile and ®nally infertile because of early exhaustion of the primordial follicular pool (te Velde, 1998). Treatment with chemotherapeutics (mostly the alkylating drugs) and ionizing radiation (Springer et al., 1996) are the best-known destructive agents of the primordial pool. Toxicity to the pre-antral follicular pool (primary, secondary stage): this will cause cycle disturbances, which become evident only after a few months. Toxicity to antral or pre-ovulatory follicles: this will result in an immediate loss of reproductive function, but this can be restored once the agent is retracted from the system and a new wave of follicles is allowed to develop. The study of potentially hazardous compounds on the female reproductive system will therefore require the assessment of their speci®c interaction with each follicular component at each stage of development. Making use of in-vivo screening for this purpose is complicated, since the active adult ovary is made up of a variety of follicle populations at different stages of development, obscuring differential follicle toxicity. The information that can be generated from in-vivo studies is scanty because of the lack of reliable non-invasive tests to identify alterations in the exposed system immediately. It is fair to conclude that the origin and mechanisms of damage by a given compound cannot be reliably investigated using in-vivo experiments. In most cases, the remains of a degenerative process can only be contemplated (Hirsh®eld, 1997).
Affection of viability and morbidity of offspring
The ovary is responsible for the development of the female gamete. During fetal, life the germ cell becomes isolated within the follicular unit, creating a microenvironment for the oocyte in which it is nourished by the granulosa cells and allowed to grow and mature until ovulation. It is obvious that the fate of the oocyte will be largely determined by the health of the follicle cells. Damage to the follicle cells will affect oocyte quantity and quality. When follicle atresia is induced, the oocyte will also die (for reviews, see Hsueh et al., 1994; McGee and Hsueh, 2000; MarkstroÈm et al., 2002). Besides complete destruction of the follicle/oocyte (lethal) at any stage of follicular development (by direct or indirect mechanisms), toxicity can also act in a more subtle manner and alter follicular hormone biosynthesis and somatic±germ cell interactions in viable follicles, with the more subtle effects perhaps impairing oocyte quality. This occult defect will be recognized only after ovulation when the exposed oocyte is fertilized, and will in turn affect the viability and morbidity of the offspring (Hoyer and Sipes, 1996). The `quality' of an oocyte is determined by its cytoplasmic and nuclear (chromosomal) constitution. While the nuclear (chromosomal or genetic) constitution determines the viability and morbidity of the late-stage fetus and offspring, the cytoplasmic constitution of the oocyte determines fertilization and the early developmental capacity of the embryo (Moor et al., 1998). Cytoplasmic maturation
During oogenesis, the ooplasma is built up and organized from the moment of the recruitment of the primordial follicle until ovulation. Meanwhile, the oocyte increases its volume by a factor of 300 to 400. Gross abnormalities in cytoplasm formation during oogenesis will result in interruption of the meiotic cell cycle or failure of the fertilization process. More subtle imperfections will affect the late preimplantation cleavage stages (Moor and Trounson, 1977; Leibfried-Rutledge et al., 1987; Moor and Gandol®, 1987; Moor et al., 1998). The production of a developmentally competent oocyte is, ®rstly, dependent on whether the oocyte, with support from the (interconnected) granulosa cells and an optimal endocrine environment, can build its essential store of organelles, proteins and mRNA (pre-antral stage) (Nayudu et al., 1987; Haines and Emes, 1991). Secondly, these organelles and biomolecules need to be correctly mobilized and redistributed prior to, and at the time of, nuclear maturation (antral stage) (Maro et al., 1986; LeÂvesque and Sirard, 1996; Albertini and Carabatsos, 1998). For its growth and development, the oocyte is dependent on the follicular structure. All factors impairing follicular regulation and metabolism will also de facto affect oocyte developmental potential. Nuclear maturation
Oocytes are most vulnerable to genetic damage during the periods of active nuclear events. Two separate vulnerable periods can be distinguished during oogenesis. The ®rst period is during early fetal life, when primordial germ cells proliferate during migration from their site of origin and colonize the gonads. There, they enter the ®rst reduction division which spontaneously arrests at the prophase of meiosis I (germinal vesicle stage) (Henderson and
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Recently, it has become increasingly clear that the process called imprinting can alter the functionality of the de®ned chromosomal constitution. At the genomic level, patterns of methylated cytosine residues determine allele-speci®c expression of imprinted genes, which are essential for normal development. Methylation patterns are at ®rst largely erased in primordial germ cells and re-established in a sex-speci®c pattern in the mature sperm cell and the oocyte. After fertilization there occurs a second wave of demethylation during preimplantation development. Around the implantation time, the de-novo methylation of the DNA becomes established on the basis of the primary signals that had been established during gametogenesis. The so-called imprinted genes retain their patterns at all stages such as they were established during gametogenesis (Monk et al., 1987; Walsh and Bestor, 1999). The imprinted gene expression data show that the primary imprinting during oocyte growth plays a de®nitive role in parent-speci®c gene expression during embryogenesis (Kono et al., 1996; Kono, 1998). Environmental insults and xenobiotic substances to which gametes and embryos are exposed during development could have signi®cant implications in congenital growth disorders and carcinogenesis (Moore, 2001).
The segment I study is designed to evaluate fertility and general reproductive function and to assess possible effects on the development of the offspring. It involves the treatment of males and females prior to mating for a suf®cient time to cover all stages of spermatogenesis and folliculogenesis, as well as during pregnancy and lactation. The offspring are evaluated only up to the age of sexual maturity. Segment II is designed to detect embryotoxicity and teratogenicity, with pregnant animals being exposed during the period of embryogenesis. The fetuses are delivered just before term and are examined for morphological and structural malformations of the viscera and skeleton. Segment III testing provides an assessment of peri- and post-natal toxicity. Pregnant animals are exposed during the last third of the pregnancy, through parturition and lactation (Sullivan et al., 1993; Christian, 1997). The segment I test evaluates the integrated reproductive function of the female and male. It is recognized that the endpoints in these tests do not allow adequate evaluation of the complete reproductive function in the female and that they are inadequate in pinpointing the site or mechanism of toxic action or more subtle effects that might result in subfertility (Kimmel, 1993). To evaluate the female reproductive function by in-vivo animal studies it is necessary to collect from treated females extensive additional information. Parameters to be recorded include: vaginal opening, vaginal cytology, oocyte toxicity (destruction of the primary oocyte population leading to cessation of ovarian function), time to mating, gestation length, and reproductive organ weight base on stage of oestrous cycle at necropsy (Kimmel, 1993). Extensive histological evaluation (differential follicle counts) of the ovarian tissue of the exposed animals within these studies would allow quanti®cation of the ovarian damage, and in some cases would provide a more sensitive indicator of female reproductive toxicity than fertility (Smith et al., 1991; Heindel and Chapin, 1993; Plowchalk et al., 1993; Bolon et al., 1997; Bucci et al., 1997). Full evaluation of the oocyte quality is, in the in-vivo studies, impossible to achieve. End-points such as time to pregnancy, implantation and number of live offspring are all secondary parameters which do not allow the adverse effect to be pinpointed. In conclusion, evaluation of the female reproductive function by in-vivo studies is currently carried out on the basis of parameters related to fertility outcome. These parameters do not allow pinpointing of the site of action or elucidation of the mechanism of toxic damage to the ovary. For a complete evaluation of ovarian function, additional animals have to treated and analysed with ovary-speci®c evaluation parameters. Furthermore, it must be borne in mind that drugs might affect essential metabolic pathways that can in themselves in¯uence the reproductive system secondarily.
Female reproductive function testing
Follicle culture systems
Reproductive function is now tested by a tiered in-vivo testing system, with the aim being to explore the potential (adverse) effects at least through one complete life cycle from conception in one generation through to conception in the following generation. Three consecutive segments are distinguished.
In-vitro follicle culture systems have been developed with the aim of growing immature oocytes from early follicle stages to fertilizable oocytes. They are also often used as a tool to study the process of folliculogenesis and oogenesis in vitro. Several research groups have described the conditions for in-vitro culture
Edwards, 1968). The second vulnerable period is at the time of ovulation, at the midcycle LH surge with the induction of resumption of the ®rst meiotic process up to the second metaphase, where it will be blocked again until fertilization (LeMaire-Adkins et al., 1997). In the germinal vesicle stage, nuclear activity is limited to processes such as gene transcription and repair mechanisms which are thought to be less sensitive to drugs interfering with meiotic activities (DNA synthesis, spindle formation and chromosome segregation). However, drugs which intercalate into the DNA may induce chromosomal breakage and dominant lethal mutations. On the resumption of meiosis, intracellular dynamic processes are activated by a set of key cell-cycle molecules, which ensure that the oocytes undergo reduction division (Chesnel and Eppig, 1995; Heikinheimo et al., 1995). Oocytes do not have pairs of centrioles at their spindle poles, but instead contain microtubule organizing centres (MTOC) that are essential for organization of the symmetrical bi-polar spindle. The formation of MTOC requires components such as gamma-tubulin, centrin, cell cycleregulating enzymes (kinases) and other proteins before transition to the M-phase can occur. Dynamic interactions between the chromosomes, the cytoskeleton and the membrane are required for the formation of a spindle, its attachment to the cell periphery and the ful®lment of cytokinesis (Verlhac et al., 1996). At this moment of development, when the oocytes resume meiosis, they are very sensitive to induction of aneuploidy (Eichenlaub-Ritter and Sobek-Klocke, 1993; Kubiac et al., 1993). Genomic imprinting
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Follicle culture in reproductive toxicology
Figure 2. The follicle culture bioassay: morphology. Explanatory scheme showing morphological features of the follicle bioassay. Pictures show the morphological development and the follicle remodelling on different days of medium replenishment (days 1, 4, 8 and 12) and after `in-vitro' ovulation (day 13). EGF = epidermal growth factor.
of ovarian follicles, especially in rodents (Gosden et al., 1993; Hartshorne, 1997; van den Hurk et al., 1997; Cortvrindt and Smitz, 2000; Smitz and Cortvrindt, 2002). Few such culture systems were consolidated with the birth of live young. In the mouse model, such limited data have been provided only rarely (Eppig and Schroeder, 1989; Spears et al., 1994; Eppig and O'Brien, 1996; Cortvrindt et al., 1998a). The reported in-vitro culture systems differ in terms of the species used, the follicle stage at which the culture is started, the type of culture vessel, the composition of the medium, and the end-points evaluated. Selection of the culture set-up should be dictated by the aims of the study. If an interaction with the ovulation process is to be studied, then the protocol used should promote intact follicle culture (Rose et al., 1999), but if the aim is to study interactions with the oocyte during gametogenesis, then oocyte granulosa cell complexes could be isolated enzymatically and cultured in groups (Eppig and Schroeder, 1989). Another possibility is to isolate intact follicles mechanically to obtain oocyte±granulosa±theca cell cultures (Cortvrindt et al., 1996). As each system has its own characteristics and bene®ts, there is an obvious need for meticulous characterization of the in-vitro process and its comparison with the in-vivo process. The mouse follicle culture system (Cortvrindt et al., 1996) allows the growth and development of intact early pre-antral follicles up to the ovulatory stage (Figure 2). Mature metaphase II oocytes can be harvested at the end of the 13-day culture period
after induction of meiosis by an ovulatory stimulus. From these oocytes healthy offspring were repeatedly obtained by IVF and transfer of the embryos into pseudopregnant-foster mothers (Figure 3) (Cortvrindt et al., 1998b and unpublished observations). In-depth study of the culture system demonstrates that the whole in-vitro process mimics normal (in-vivo) physiology from the point of view of morphogenesis, differentiation, hormone production and hormone responsiveness (Cortvrindt and Smitz, 1998). The culture system is based on the use of a homogeneous narrow class of intact early pre-antral follicles of 100±130 mm diameter (3b follicles according to the classi®cation by Pedersen and Peters, 1968). Approximately 40 follicles of this class can be collected per ovary by mechanical means from a 14-day-old mouse. The selection of such a narrow class of follicles and exposure to a de®ned medium with gonadotrophins allows synchronous growth during culture with a strict developmental pattern (Smitz and Cortvrindt, 2002) (Figure 2). First, the surrounding theca cells attach to the bottom of the dish and start to proliferate. By days 4±6, granulosa cell proliferation is clearly visible, noticeable by the enlargement of follicular diameter and subsequent perforation of the basal membrane. Proliferating granulosa cells grow on top of the theca cell monolayer. During the second half of the culture period, granulosa cells differentiate into two subpopulations: steroid-producing mural granulosa cells separated by an antral-like cavity from the cumulus cells tightly
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Figure 3. The follicle culture bioassay. Evaluation parameters: oogenesis. Illustration of end-point parameters characterizing oocyte quality and developmental competence. COC = cumulus±oocyte complex.
enclosing the oocyte. The growing follicles can be easily observed microscopically, and their cell-speci®c secretion pattern (steroid and protein hormones) can be measured in the spent medium. Androgens are produced by the theca cells and converted to estrogens by the granulosa cells. Estrogen concentrations increase with follicle development and differentiation. Progesterone production remains low and rises modestly towards the end of the culture period (days 10±12). Inhibin A and B and activin production also follows the physiological pattern (Smitz and Cortvrindt, 1998; Smitz et al., 1998a). During in-vitro folliculogenesis the oocyte grows (as in vivo) mainly in the pre-antral phase and remains blocked at the germinal vesicle stage as long as it is kept within the follicle structure (Cortvrindt et al., 1998b,c). The follicles in culture are FSH-dependent for their growth and development and become LH-responsive by the end of culture (Cortvrindt et al., 1997, 1998c). The administration of an ovulatory stimulus (HCG or LH) on day 12 of culture provokes detachment of the cumulus±oocyte complex (COC), muci®cation of the cumulus cells and resumption of meiosis in the oocyte (Smitz et al., 1998b). At the same time, a steep rise of progesterone output can be measured in the spent medium (Figure 4). These in-vitro-produced oocytes reveal normal spindle morphology with well-aligned chromosomes comparable with oocytes grown and matured in vivo (Figure 3) (Hu et al., 2001).
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In-vitro bioassay for testing ovarian function and female gamete quality On the basis of the follicle culture system described above [early pre-antral mouse follicles (100±130 mm)], a bioassay was designed for testing ovarian function and gamete quality in vitro. The step-wise adaptation of the culture set-up to achieve this goal is shown in Table I. Column A represents the original culture system, in micro-drops under oil in Petri dishes. In order to avoid cross-talk between the follicles through the oil overlay by the lipophilic steroids, new conditions were set in 96well plates represented in column B. In this setting the culture medium was protected by an oil overlay to prevent ¯uctuations in temperature and pH during follicle handling, yet still limiting testing of compounds to those with non-lipophilic properties. Finally, the oil overlay was removed as depicted in column C. In this design, follicles are vulnerable to external in¯uences. Hence, handling must be carried out under strict conditions and minimizing the external physical stress (temperature, pH, evaporation) that is known to in¯uence folliculogenesis and oogenesis. The process of folliculogenesis and oogenesis in this new setting was compared with the original system (column A) (Table II, Figure 2). The morphological observation showed that the pre-antral follicles develop in both cultures in an identical manner; this
Follicle culture in reproductive toxicology
Figure 4. The follicle culture bioassay. Evaluation parameters: folliculogenesis. Scheme of the parameters characterizing folliculogenesis monitored during the 14day culture period. E2 = estradiol; P = progesterone; T = testosterone. Table I. Design and properties of three different culture systems
Culture design follicles medium volume oil volume Medium/oil Handling Evaluation Possible applications No. of follicles/ experiment Outcome parameters Folliculogenesis Morphology Conditioned medium-analysis Oogenesis
A Petri dish
B 96-well plates
C 96-well plates
n = 20 20 3 (20 ml) droplets 5 ml 1/12.5 Easy Individual (~ group) Large-scale testing Avoid lipophilic compounds 200±300 (10±15 plates)
n = 3312 50 ml 30 ml 5/3 Experienced Individual Individual testing Avoid lipophilic compounds 120±150 (10±12 plates)
n = 12 75 ml No No oil Experienced Individual Individual testing All compounds
Individual (~ group)
Individual
Individual
Pooled/plate Individual (~ group)
Pooled/row or individual Individual
Pooled/row or individual Individual
results in similar survival rates (de®ned below) and both respond on the ovulatory stimulus (= muci®cation rates). The oocyte grows equally well in both systems and matures to metaphase II at the same rate. Fertilization experiments showed that there were no differences at the level of the oocyte quality. The steroid concentrations in collected spent medium in both culture set-ups cannot be compared as these values are in¯uenced by the presence/absence of the steroid-extracting oil overlay. Each system must refer to its own reference range (see Figure 5).
120 (10 plates)
De®nition of parameters for evaluation of the process of folliculogenesis (Figure 4)
For evaluation of the process of folliculogenesis, the following end-point parameters were selected: 1. Follicle survival at day 12. Survival of a follicle is de®ned as those follicles which retain their oocyte within the granulosa-cell mass. Oocyte release is associated with cytotoxic effects on the granulosa cells and disassembly of the follicular structure. This
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R.G.Cortvrindt and J.E.J.Smitz Table II. Comparison of follicle and oocyte developmental characteristics and preimplantation development of two follicle culture systems in Petri dish with oil and 96-well plate without oil Parameter
Folliculogenesis Differentiation (%) (antrum formation) Survival rate (%) Muci®cation (%) Oogenesis Oocyte diameter (mm) Polar body rate (%) Preimplantation development 2 cell rate (%) Blastocyst/2 cell (%)
De®nition of hormone secretion pattern
Culture system Petri dish + oil Mean 6 SD
96-well + oil Mean 6 SD
86 94 94
86 96 97
6 17 6 7.3 6 9.3
6 4.8 6 4.3 6 4.3
71.5 6 0.84 87 6 13.9
71.3 6 1.20 88 6 12.8
73 72
74 54
69 6 16
69 6 10
Data for folliculogenesis and oogenesis were obtained from eight randomselected cultures and expressed as % of follicles reaching the evaluation parameter over the total number of follicles plated/plate (e.g. Petri dish = 20; 96-well plate = 12). Results for preimplantation development were obtained from four independent experiments. In each experiment, oocytes for fertilization were pooled per condition (dish/96-well plate).
parameter re¯ects the cytotoxic effect of the compound under study. 2. Differentiation stage on day 12. By the end of the culture almost all follicles need to have progressed to their ®nal differentiation stage being either antral (A) or antral-diffuse (A/ D) if follicles reside in the diffuse stage; this indicates that the process of follicle differentiation is impaired. If follicles are blocked in the follicular stage (F) or follicular-diffuse stage (F/D), this means that the compound is not cytotoxic but that cell proliferation is prohibited, or at least seriously reduced 3. Cumulus cell muci®cation on day 13. This is an indicator for normal differentiation and LH-receptor expression on the muralgranulosa cells. It also re¯ects the viability of the oocytes within the follicular structure. De®nition of parameters for evaluation of oogenesis (Figure 4)
1. Oocyte diameter re¯ects the oocyte growth process. 2. Oocyte nuclear maturation stage [polar body (PB) extrusion; germinal vesicle breakdown (GVBD), germinal vesicle (GV) stage] gives a ®rst indication of quality of the oocyte. Different options can be taken for more in-depth study of the oocytes' quality of the GVBD and PB extruded oocytes: d Spindle analysis by ¯uorescent probing: spindle morphology and chromosome alignment will predict genetic constitution of the embryo (aneuploidy). d Chromosome spreading: PB-extruded oocytes: indication of aneuploidy; GVBD oocytes: indication type of meiotic problem (delayed/cytokinesis). Eventually ¯uorescence in-situ hybridization (FISH) analysis of the chromosome preparations will even re®ne the analysis. d IVF: two cell rate and blastocyst formation rate gives an indication of the cytoplasmic quality of the oocyte.
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Eventually, embryos can be transferred to pseudopregnant foster mothers for evaluation of implantation capacity and normality of the offspring. Hormones produced by the follicle were measured routinely by validated immunoassays on the conditioned media sampled on refreshment days. In the bioassay setting where the oil overlay is omitted, the measurements for testosterone, estradiol and progesterone re¯ect the total amount produced by the individual follicles. The bioassay provided the expected pro®les for the following hormones: estradiol, progesterone, testosterone and total inhibin. Figure 5 illustrates data on steroid concentrations measured in Petri dishes with oil or wells without oil overlay. Pro®ciency testing of the in-vitro follicle bioassay
Prerequisites for using this culture system as a bioassay in a routine setting are that results are reproducible from assay to assay. Obvious measures such as precise animal age selection (12±14 days), strict control over temperature and pH during all handling (37°C) and proper maintenance of incubators (gas makeup, humidity) are essential for reproducible results. The composition of the medium should be kept constant over time by including stable compounds and controlling batch-to-batch variations. Out of 125 repeat control cultures, performed over a period of 1 year by two biotechnicians, relevant end-point parameters for folliculogenesis and oogenesis were set and evaluated for pro®ciency (Table III). The collected data demonstrated that the reproducibility of the selected parameters performed well for a bioassay. As can be appreciated from Table III the variability of the bioassay for all parameters was within the limits of acceptability for a biological test. Proof of concept
Proof of concept is still under analysis in the authors' laboratory by incorporation of different chemical compounds (e.g. taxol, atrazine, genistein, diazepam) over a range of doses in the follicle bioassay. Results revealed relevant dose-dependent outcomes for different parameters. Results are not shown in this review as they are an integral part of future publications. To validate the assay, results on de®ned end-points from invitro exposures were compared with ®ndings from in-vivo exposures which have been well documented in the current literature. Although comparisons are not always straightforward, ongoing analysis with selected chemicals testify that mouse follicle in-vitro culture is sensitive to reveal dose-dependent changes induced by physiologically relevant concentrations of compounds at the end organ.
Conclusion Today, society expresses an increased awareness for the wellbeing of living creatures and protection of the environment. This concern demands a more profound safety testing of NCE before they can be launched into the ecosystem. However, safety testing is carried out at the expense of sacri®cing millions of animals each year (Purchase, 1999). In the ®eld of toxicology, animal welfare is promoted with a pressing demand to reduce, replace and re®ne (`the 3Rs') the use
Follicle culture in reproductive toxicology
Figure 5. Comparison (`box and whisker' plots) of two systems for steroid production: follicle cultures in dishes with oil versus 96-well plates without oil. Panel (A): 17b-estradiol (E2) concentrations on days 8 and 12. Panel (B): production of E2 in Petri dishes (96 wells/pooled row) under oil coverage, showing amounts produced in individual wells of microtitre plates. Note the marked difference in E2 concentration between both systems. Panel C: progesterone (P) concentrations on days 8, 12 and 13 (post-ovulation) as measured in Petri dishes under oil. Panel D: progesterone concentrations in individual wells without oil. Note the differences in reference ranges between both systems.
Table III. Inter-assay variation of the bioassay (96-well without oil overlay) over a 1-year period (n = 125) Parameter
Mean 6 SD
Median
Minimum
Maximum
Lower limit/ % CVb no. rejecteda
Day 1: Follicle diameter (mm) Day 12: Differentiated follicle (%) Day 13 % survival Day 13 % muci®cation Oocyte diameter (mm) % Polar body extrusion
117.7 6 4.17 83 6 13
117.8 82
105.8 50
127.3 100
NA 57/5
3 14
92 100 71.7 86
64 55 67.9 33
100 100 74.6 100
72/5 73/2 NR 59/5
9 9 0 13
92 93 71.5 84
6 6 6 6
10 9 2.37 13
a
Number rejected = no. of cultures rejected on the basis of not reaching the lower limit (= mean ± 2SD). % CV = % coef®cient of variation (calculated after rejection of outliers). NA = not applicable; NR = not relevant. b
of laboratory animals in test protocols (Jackson, 1998), and of®cial bodies have been installed to validate in-vitro protocols to accommodate this goal [ECVAM (http://www.jrc.cec.eu.int); IICVAM (http://iicvam.niehs.nih.gov)]. In the case of reproductive function testing, in-vitro screening will never be able to cover all aspects of fertility because reproduction requires a complex of integrated functions. Although full-extent studies can only be performed in living animals, parts of the process can be mimicked
in in-vitro systems, and it is possible that a panel of well-designed and validated in-vitro tests could replace a substantial proportion of in-vivo testing procedures. In the case of female fertility testing, it is recognized that even the in-vivo tests cannot cover all aspects of female reproductive function (Ecobichon, 2001). Often it is dif®cult to pinpoint the site or predict the mechanism of action of the compound under investigation, and the ovary and the gametes it produces suffer
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R.G.Cortvrindt and J.E.J.Smitz especially from this problem. It is the inherent structure of the ovary that makes evaluation dif®cult, with the oocytes being inaccessible for analysis. The ovary is composed of a pool of follicles, all of which are at different stages of development and all vulnerable to a certain extent, depending on the stage of differentiation; this makes interpreting the effects of exposures very complicated. The value in studying folliculogenesis with in-vitro approaches has already been proven by several groups who have investigated the effects of physical factors (oxygen tension; Boland et al., 1993; Eppig and Wigglesworth, 1995; Smitz et al., 1996), antimetabolites including steroid synthesis pathway blockers (Murray et al., 1998) and toxic substances such as chlorvinphos (Nayudu et al., 1994) using in-vitro follicle culture methods. The follicle bioassay proposed in this review has been tested to ful®l the task of in-vitro ovarian function testing. A key feature of the test is that follicle growth and development of all follicle units enclosed in a single assay is perfectly synchronized by selection of a narrow class of follicles to start-off culture. The developing units can be observed daily, both morphologically and biochemically, throughout the entire culture period, at the end of which the oocytes can simply be collected for profound analysis. During culture, the follicles can be exposed to the compound under investigation either continuously or only at de®ned phases of development. Exact pinpointing and elucidation of the mode of action of the hazard (cell type, stage of follicular development, effect on functionality such as steroid or speci®c protein production, effect on meiosis, mechanism of cell death) is possible. In-depth mechanistic studies can be performed since the affected follicular structure is readily available and can be used as substrate in all kinds of technology for more profound analysis. Within a few weeks, very complex interactions can be evaluated and the size of the experiments permits a decently powered statistical interpretation on quantitative measurements. The choice of species for the in-vitro bioassay is sound, with rodents (mice, rats) having provided the bulk of reproductive toxicity data in in-vivo studies. Indeed, a massive database is available on chemical-induced reproductive toxicity for these species, and for risk assessment calculations can be taken into account for species differences (Ecobichon, 2001). Using mice for reprotoxicity studies offers the advantage that in-vitro protocols (in-vitro maturation, IVF, embryo culture, embryo transfer) are well de®ned and already applied world-wide. Moreover, there are numerous knockouts available for special in-depth studies on mechanisms. The issue of the metabolism of compounds under test is undoubtedly complex and inherent to all in-vitro testing protocols. Although compounds can be detoxi®ed, activated or eliminated in vivo, these complex mechanisms are clearly lacking in in-vitro systems. However, in the follicle bioassay, partial metabolism of the test compound by the follicle cells can be expected. The fact that the cultured follicle is an open remodelled structure allows free access of the substances under study to all cells. Pretreatment of the compound or co-culture with microsome fractions, as performed in mutagenicity tests, could also be implemented in the follicle bioassay system. The present authors suggest that the proposed bioassay could be a valuable in-vitro test for ovarian function and female gamete testing. In particular, it can be used in a straightforward manner to
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provide information on the mechanisms that would be both dif®cult and inef®cient to acquire using an in-vivo approach. Although still labour-intensive, in-vitro testing has the power to provideÐwithin a relatively short time period of analysisÐ precise answers to complicated questions by using quantitative data obtained with the use of routine techniques.
Acknowledgements The authors acknowledge the skilful follicle culture work of Lotte Arentoft and Isabelle Hellinckx, and Daniel de Matos, Daniela Nogueira and Jean Claire Sadu for performing IVF/ET of cultured oocytes. Kathy Billooye, Anne Gerard and Johan Schiettecatte of the Radioimmunoassay Laboratory provided excellent assistance in hormone analysis. This research was funded by the IWT (International Scienti®c and Technological Cooperation; Grant no. 980343). The authors thank Serono International, Geneva, Switzerland (Project GF9405) for the gift of recombinant gonadotrophin.
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Submitted on January 9, 2001; resubmitted on March 12, 2002; accepted on March 15, 2002