Cadmium: Toxic effects on the reproductive system and the embryo

Reproductive Toxicology 25 (2008) 304–315

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Reproductive Toxicology journal homepage: www.elsevier.com/locate/reprotox

Review

Cadmium: Toxic effects on the reproductive system and the embryo Jennifer Thompson ∗ , John Bannigan School of Medicine and Medical Science, Conway Institute of Biomolecular and Biomedical Research, University College Dublin, Ireland

a r t i c l e

i n f o

Article history: Received 5 October 2007 Received in revised form 6 February 2008 Accepted 12 February 2008 Available online 19 February 2008 Keywords: Cadmium Chick embryo Testis Ovary Blastocyst Cell adhesion Oxidative stress Ventral body wall defect

a b s t r a c t The heavy metal cadmium (Cd) is a pollutant associated with several modern industrial processes. Cd is absorbed in significant quantities from cigarette smoke, and is known to have numerous undesirable effects on health in both experimental animals and humans, targeting the kidneys, liver and vascular systems in particular. However, a wide spectrum of deleterious effects on the reproductive tissues and the developing embryo has also been described. In the testis, changes due to disruption of the blood–testis barrier and oxidative stress have been noted, with onset of widespread necrosis at higher dosage exposures. Incorporation of Cd into the chromatin of the developing gamete has also been demonstrated. Ovarian Cd concentration increases with age, and has been associated with failure of progression of oocyte development from primary to secondary stage, and failure to ovulate. A further mechanism by which ovulation could be rendered ineffective is by failure of pick-up of the oocyte by the tubal cilia due to suboptimal expansion of the oocyte–cumulus complex and mis-expression of cell adhesion molecules. Retardation of trophoblastic outgrowth and development, placental necrosis and suppression of steroid biosynthesis, and altered handling of nutrient metals by the placenta all contribute to implantation delay and possible early pregnancy loss. Cd has been shown to accumulate in embryos from the four-cell stage onwards, and higher dosage exposure inhibits progression to the blastocyst stage, and can cause degeneration and decompaction in blastocysts following formation, with apoptosis and breakdown in cell adhesion. Following implantation, exposure of experimental animals to oral or parenteral Cd causes a wide range of abnormalities in the embryo, depending on the stage of exposure and dose given. Craniofacial, neurological, cardiovascular, gastrointestinal, genitourinary, and limb anomalies have all been described in placentates, with axial abnormalities and defects in somite structure noted in fish and ventral body wall defect and vertebral malformation occurring in the chick. In this paper, we examine the mechanisms by which Cd can affect reproductive health, and consider the use of micronutrients in prevention of these problems. © 2008 Elsevier Inc. All rights reserved.

Contents 1. 2.

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Effects of cadmium on reproduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. The testis and spermatogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. The ovary and oogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. The pre-implantation embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Implantation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. The post-implantation embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanisms of reproductive toxicity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Altered cell adhesion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Oxidative stress and apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Ionic and molecular mimicry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. DNA damage and effect on the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author at: Room C212, Health Sciences Building, School of Medicine and Medical Science, University College Dublin, Belfield, Dublin 4, Ireland. Tel.: +353 1 7166634; fax: +353 1 7166649. E-mail address: [email protected] (J. Thompson). 0890-6238/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.reprotox.2008.02.001

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1. Introduction Cadmium (Cd; atomic number 48; relative atomic mass 112.40) is a toxic metal that belongs to group IIb in the periodic table. It occurs in nature at low concentrations, mainly in association with the sulfide ores of zinc (Zn), lead (Pb), and copper (Cu). However, due to the widespread nature of its occurrence, it is present in measurable amounts in almost everything that we eat, drink, and breathe [1]. Human activities adding to the Cd load in the environment include the combustion of fossil fuels, leechate from landfill sites, run-off from agricultural land, and mining residues, especially from Zn and Pb mines [2]. Electroplating to protect steel from corrosion, and the manufacture of Nickel–Cd batteries, pigments, stabilizers and alloys [3–5] also produce Cd as a by-product. A pollutant of worldwide concern, Cd has been reviewed by the International Register of Potentially Toxic Chemicals of the United Nations Environment Program, and included on the list of chemical substances considered to be potentially dangerous at the global level [6]. It is now recognized that human exposure to Cd must be minimized. The WHO “safe” level for human ingestion of Cd has been estimated at 500 ␮g/week [5]. Absorption of oral Cd tends to be erratic, with continuing presence of unabsorbed radio-Cd in the gut lumen for 3–5 weeks after a test meal in human subjects [7]. Dietary Cd intake estimates in European countries were 10–30 ␮g per day [8], with increased risk of consumption with certain foods such as shellfish, offal, and rice [9–12]. Individuals consuming dredge oysters in New Zealand were found to have a daily fecal excretion of Cd up to 580 ␮g, equivalent to more than 10 times the provisional tolerated weekly intake (PTWI). The dietary Cd absorption rate is about 5%, rising to 20–30% in some individuals [13]. In non-Cd-polluted areas, the most significant human route of Cd intake is cigarette smoking, with various estimates of 0.2–1.0 ␮g Cd assimilated with each cigarette smoked [13–17], accounting for approximately half the total human Cd intake. However, bioavailability of inhaled Cd oxide is relatively high, with 10% deposited in lung tissues and another 30–40% absorbed into the systemic blood circulation of smokers [13]. The whole-body biological half-life for Cd is very long, having a fast component of 75–128 days and a slow component of

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7.4–26.0 years [5,18,19]. This raises the possibility of cumulative effects. After absorption, Cd is transported to the liver, bound to albumin [20], where it induces the synthesis of metallothionein (MT), a class of small cysteine-rich heavy metal binding proteins. Changes within the liver itself following parenteral administration of CdCl2 are dose- and time-dependent, ranging from moderate diffuse hepatocellular degeneration through to multifocal necrosis [21,22]. Following release from the liver, MT-bound Cd enters the plasma. Cd is eliminated in the urine. MT-bound Cd appears in the glomerular filtrate, from where it is re-absorbed intracellularly by renal tubule cells. In the latter, the Cd is cleaved from the MT by lysosomal action, and Cd++ ions are re-excreted into the tubular fluid. The ability of Cd to induce hepatic and renal lesions exacerbates its toxic effects, and compounds its propensity to accumulate over years. A large body of work exists outlining the effects of Cd on gametogenesis in both males and females, and implicating its compounds in early embryolethality and implantation failure. In addition, animal studies have shown a wide range of anomalies following exposure at specific stages of embryogenesis, and emerging evidence has indicated that Cd may also be linked to pathological processes in late pregnancy and in the early postnatal period, causing third trimester complications and minor but significant problems in the offspring of exposed individuals. The focus of this review, therefore, is to examine the consequences of Cd exposure on reproductive processes and embryonic development, and to look at mechanisms by which these effects might occur. 2. Effects of cadmium on reproduction An overview of reproductive processes affected by Cd, along with suggested mechanisms of action, is given in Table 1. 2.1. The testis and spermatogenesis Testicular changes due to Cd toxicity have been seen in a variety of animal models at different stages of growth and maturity. Gonadal development in mouse embryos exposed to Cd in early organogenesis was studied by Tam and Liu [23]. Genital ridge size

Table 1 Summary of the early reproductive effects of Cd and suggested mechanisms by which these effects occur Reproductive effect

Histological finding

Probable underlying mechanism

References

1. Failure of maturation from leptotene to pachytene spermatocytes

Disruption of blood–testis barrier, testicular necrosis

[25–28,122–124]

2. Failure of maturation of spermatozoa; reduced viability 3. Failure to ovulate, defective steroidogenesis 4. Suppressed oocyte maturation

Disruption of blood–epididymal barrier, abnormal sperm morphology Damage to the endothelium in follicular arteries and to the ovarian interstitial stroma, ovarian necrosis Suppression of cumulus expansion, reduced numbers of oocytes reaching metaphase II, reduced fertilization of oocytes, increased oocyte degeneration

Substitution of Cd for Ca in adherens and desmosomal junction cadherins; disruption of tight junctions via the TGF␤3/p38 MAP kinase pathway, oxidative stress Junctional disruption Junctional disruption, ? oxidative stress

[48–56,59] [41,57–60]

5. Failure of developmental progression in pre-implantation embryo

Failure of progression from two-, four- and eight-cell stages to morula, failure of compaction/decompaction

6. Failure of implantation

Failure of oocyte pick-up by infundibulum, abnormal trophoblast outgrowth and transformation

Junctional disruption—due to suppression of synthesis of hyaluronic acid and/or inhibition of gap junctional intercellular communication and connexin phosphorylations, ? oxidative stress, ? genotoxicity ? Junctional disruption—due to inhibition of gap junctional intercellular communication and connexin phosphorylation, and disruption of tight junctions, oxidative stress Suppression of synthesis of hyaluronic acid, ? suppression of cell adhesion molecule expression, displacement of Ca by Cd on calmodulin

(?): Possible mechanisms that have not yet been proven definitively.

[41,125]

[62–71]

[58,64,73–81]

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was reduced in exposed animals, with retarded germ cell migration into the ridges, resulting in depleted populations of germ cells, defective maturation of gametes and subfertility in male offspring. Young rabbits dosed with 1.0–2.25 mg/kg body weight Cd had a significant decrease in germinal epithelial volume after 48 h, with signs of epithelial and basement membrane damage [24], along with reduced epithelial volume in the epididymis after 5 months of treatment. Similarly, adult male rats have been shown to develop gonadal damage following administration of Cd, either orally or subcutaneously (SC). Focal testicular necrosis and reduced spermatogenesis were seen in rats that received a single dose of 100–150 mg Cd/kg orally within 2 weeks of administration [25]. Rats injected with a large dose of CdCl2 (7 mg/kg SC) showed pronounced testicular hemorrhage and edema 24 h after treatment [21]. Fende and Niewenhuis [26] described endothelial cell damage in testicular blood vessels commencing 3 h after SC administration of Cd. Light and electron microscopy initially revealed edema, with widening of interstitial spaces and deposition of proteinaceous material 3 h after treatment, progressing to separation of endothelial cells 6 h after injection and extensive hemorrhage and histological disruption at 24 h. Testicular morphology 3 months after initial Cd exposure was greatly altered, with degenerated seminiferous tubules, abnormal Leydig cells, fibrosis, and reduced testicular size [27]. Similar effects were noted in a histological and ultrastructural study of gerbil testes [28]. Other cell populations within the testis have also been implicated as targets of Cd toxicity [29–31]. Sprague–Dawley rats injected SC with 0.6 mg/kg Cd daily over a 6-week period were found to accumulate Cd in the testes, mainly in spermatogonia and spermatocytes, with consequent reduction in both of these cell types [32]. Other studies have failed to demonstrate incorporation of Cd into sperm chromatin, suggesting instead that subfertility following Cd administration might result from damage to supporting testicular tissue [33]. Cd dose in the latter experiments was, however, considerably lower (0.1 mg/kg CdCl IP) than in those conducted by Aoyagi et al. [32] or by Toman et al. [24], and Dixon et al. [34] produced compelling data indicating that Cd is indeed incorporated into early and late spermatids and spermatogonia. Disruption of the blood–testis barrier (BTB) by Cd, with consequent reduction in germ cell numbers and infertility, has been linked to reduced occludin expression, thought to be mediated by the TGF␤3/p38 MAP kinase signaling pathway [35,36], indicating the involvement of occludens junction disturbance as well as adherens junction breakdown in BTB disruption [37]. Heavy smoking, the commonest source human intake of Cd, has been associated with low sperm count and motility. A strongly positive relationship has been identified between azoospermia and serum and seminal fluid Cd levels in infertile Nigerian males [38]. However, other studies have shown no differences in semen quality or fertility between smokers and non-smokers, even though significantly increased amounts of Cd were found in seminal fluid in those smoking 20 or more cigarettes per day [39]. In the rat model, increased rates of abnormal sperm morphology have been noted with increased seminal Cd levels [40], with positive correlation with TNF-␣ and IFN-␥, and a negative relationship with IL-4. All seminal parameters and cytokine anomalies in this model were reversible by administration of Zn with Cd. It has been suggested that absence of certain micronutrients in the diet contribute to male infertility associated with raised Cd levels [38]. In in vitro studies, Cd at 20 ␮M concentration has been shown to significantly decrease the viability of spermatozoa to 35.6% when compared with control viability of 54.4% [41]. Lower concentrations of Cd (2 ␮M), while having a sperm viability rate comparable with controls, increased acrosome-reacted spermatozoa (45.2%) com-

pared with both control and higher concentrations of Cd (31.9 and 32.5%, respectively). Polyspermy was significantly increased in this group (23.5%) when compared with controls and 20 ␮M Cd (6.7 and 0%, respectively), implying that Cd at higher concentrations affects cell viability, and, at lower concentrations, affects physiological function of spermatozoa. In addition to effects on fertility, Cd administration has been linked with tumors of the prostate and testis in experimental animals [3], preventable by Zn administration [20]. Waalkes et al. [42] demonstrated that SC or oral Cd induced prostatic neoplasms in rats, provided testicular function was maintained. A significant positive relationship was found between ingested Cd and prostate cancer in humans in a case control study conducted by West et al. [43]. However, cohort studies by Kazantzis [44] and Sorahan [45] failed to show any association. Although a positive relationship has been demonstrated between proliferative testicular lesions and Cd in rats [46,47], no definitive link exists in humans. 2.2. The ovary and oogenesis Oocyte development and associated events have been disrupted by Cd administration in numerous different species, including Xenopus laevis [48–50], hamsters [51,52], rats [53–56], mice [52], pigs [57,58], and sheep [41]. In Xenopus, exposure to various levels of Cd via the culture water over 30 days resulted in a decrease in number of developing oocytes in those exposed to ≥1.0 mg/l Cd. Reduction in the total number of oocytes and ovarian necrosis were seen at concentrations ≥2.5 mg/l, and reduction in ovarian weight, at concentrations ≥5 mg/l [48]. A reduction in breeding and fertilization rate was also noted ≥2.5 mg/l. Oocyte germinal vesicle breakdown was inhibited by Cd at higher concentrations [49]. Lienesch et al. [50] similarly found a reduction in the percentage of oocytes undergoing normal progression after Cd injection into the dorsal lymph sac every other day for 21 days, and a dramatic increase in the population of atretic oocytes. The effect of Cd on ovulation in the golden hamster was described by Saksena and Salmonsen [51]. Ovulation was inhibited at a dose of 5 mg/kg CdCl2 SC or higher, particularly if given close to the LH surge. Failure to ovulate was associated with ovarian necrosis and hemorrhage and a decreased progesterone level in the serum. Although ovarian lesions resolved within 4 days, prolonged periods of sterility were recorded in hamsters given 5–10 mg/kg CdCl2 , following which normal pregnancy with normal litter numbers was achieved. Rehm and Waalkes [52] investigated the effect of a single SC injection of CdCl2 at a dose of 20–47.5 ␮mol/kg on non-gravid Syrian Hamsters and rats and mice of different strains. Ovarian hemorrhagic necrosis occurred in hamsters treated with this dosage range of Cd, with administration to sexually immature animals and injection just prior to ovulation causing the most severe lesions at all doses. Assessment by light microscopy revealed that small arteries in developing follicles and the interstitial stroma were particularly susceptible to damage, while other ovarian tissue, such as primordial oocytes and corpora lutea, appeared resistant. Full morphological recovery occurred in hamster ovaries within 2 months, despite a loss of nearly 50% ovarian weight following the initial insult. Furthermore, pre-treatment with Zn acetate markedly reduced the extent of ovarian damage. Uterine and cervical hemorrhages occurred in immature hamsters at higher doses of CdCl2 . Rats similarly showed dose- and age-dependent toxicity in the ovaries, uterus and cervix. The genital tract in mice was more resistant to Cd-induced lesions, and both mice and rats demonstrated strain-dependent sensitivity to Cd. Ovarian steroidogenesis in the non-gravid rat [53], and in pregnant rats [54–56] has been profoundly influenced by Cd administration at various times of the estrus cycle. CFY rats given

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10–15 mg/kg CdCl2 on the day of diestrus II were found to have increased serum prolactin and decreased FSH, LH and basal progesterone levels the day after treatment, indicating an effect on the hypothalamic–pituitary–ovarian axis [53]. Concomitant treatment with four SC injections of 10–20 mg/kg ZnCl2 protected against sterility induced by CdCl2 at 10mg/kg body weight in in vivo experiments [54]. However, Zn failed to protect against Cd-induced drop in progesterone production in in vitro culture of rat ovarian granulosa cells. Sprague–Dawley rats given 2.5–5 mg/kg Cd SC on the day before ovulation similarly demonstrated suppression of ovarian steroidogenesis, with an initial reduction in progesterone and testosterone [59], and serum estradiol most severely affected 24 h after treatment [56]. Cd was also found to depress and delay progesterone secretion in early pregnancy [55]. Concentration of Cd in the oviducts following SC administration of 2.5–10 mg/kg CdCl2 on day 1 of pregnancy was found to rise in a time- and dose-dependent manner. However, animals receiving Cd later in pregnancy (day 16 or gestation) were relatively unaffected [56]. The role of Cd in suppressing FSH-induced cumulus expansion in oocyte–cumulus complexes (OCC) isolated from large antral porcine follicles has been described by Mlynarcikova et al. [57]. Incubation with micromolar concentrations of Cd decreased both the degree of cumulus expansion and progesterone production by the cumulus cells during 42 h of incubation of the OCC with FSH. High concentrations of Cd completely suppressed oocyte maturation, and significantly suppressed synthesis of hyaluronic acid, an integral component of expanded cumulus cells [58]. In experiments with the ovine oocyte, Leoni et al. [41] found that maturation rate was significantly affected at 2 and 20 ␮M CdCl2 , with a metaphase II (MII) rate of 63.8 and 32.0%, respectively when compared with controls (MII rate 96.8%). Both Cd concentrations also decreased numbers of oocytes resting in MII (29.0 and 19.8%, respectively; control rate 93.8%), reduced numbers of fertilized oocytes in culture (25.9 and 4.7%, respectively; control rate, 76.1%), and increased the rate of oocyte degeneration in both groups. Exposure to Cd of cultured human ovarian granulosa cells was found to cause reduction in progesterone production when compared with controls, and also caused morphological alterations in a time- and dose-dependent manner, with rounding up of cells after 4–8 h, retraction of cellular extensions, and detachment from neighboring cells [60]. 2.3. The pre-implantation embryo In addition to its effects on gametogenesis, Cd may also reduce the possibility of a successful pregnancy by interfering with the development of the pre-implantation embryo. Gamete fusion in the ovine model was found to be significantly reduced upon exposure to 2–20 ␮M Cd [41]. Fertilization of mouse oocytes was unaffected when cultured at 1.6 ␮M Cd, and treated ova went on to cleave into two-cell stage embryos at a similar rate to controls [61], although a reduced number of these embryos reached blastocyst stage. Gametes treated with 0.4–0.8 ␮M Cd were not inhibited from progressing to the blastocyst stage, but there was increased embryo loss at the implantation stage in the treated group. In vitro studies in which murine embryos were cultured at higher Cd concentrations (10–50 ␮M) at the two-cell stage, demonstrated a dose-dependent failure of developmental progression [62]. Although embryos cultured at this stage with 109 Cd showed little accumulation of radioisotope, there was marked incorporation into blastocysts [63]. In vivo exposure of the murine embryo to Cd at the two-cell stage by SC injection of the dam with 25–38 ␮M Cd/kg body weight on the morning of day 2 (D2) similarly failed to prevent initiation and maintenance of pregnancy when examined on D8 [63], but did delay implantation. Embryos that implanted went

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on to become normal fetuses [61], with resorption rates and body weights comparable to controls. Yu et al. [64] found that exposure of four-cell and morula stage mouse embryos to 5–10 ␮g/ml CdCl2 for 24 h caused degeneration and decompaction of treated embryos. In the rat, incubation for 24 h in 1 ␮g/ml CdCl2 prevented eight-cell embryos and morulae from developing to the blastocyst stage [65], with clear evidence of apoptosis following exposure. Previous papers have shown that Cd inhibits gap junction intercellular communication and connexin phosphorylation [66,67], both of which are essential processes in the progression from the eight-cell stage through to compaction [68,69]. Protection of the early embryo from Cd toxicity has been achieved by preventing Cd uptake by cells by pre- or co-incubation with Zn at 100× molar dose of Cd [63,70], and by co-culture with nifedipine at 0.01× molar excess [63]. Antioxidants are also known to be protective [71]. 2.4. Implantation An early step in the reproductive process is the movement of the secondary oocyte, arrested in the second meiotic metaphase (MII), into the Fallopian tube, where it will encounter spermatozoa and undergo the initial steps of fertilization. Thereafter, the oocyte completes meiosis II to become a definitive oocyte shortly before the fusion of male and female pronuclei [72]. This process requires intact meiotic mechanisms, including normal spindle formation, and correct sequential expression of cell adhesion molecules on both spermatozoa and oocyte [73]. The oocyte–cumulus complex (OCC) is picked up by cilia on the exterior surface of the infundibulum, which bind reversibly to the matrix of the OCC. The proteins and hyaluronan in this matrix link together and stabilize the OCC, thus facilitating pick-up of the entire complex by the cilia [74]. Cd-suppressed synthesis of hyaluronic acid in the OCC matrix [58] could be linked to the inability of the infundibular cilia to successfully introduce the oocyte into the Fallopian tube. Cell adhesion is an integral and important part of OCC pick-up [75], and although the precise sequence and pattern of expression of cell adhesion molecules during this process has yet to be elaborated, the involvement of gap junctions in cumulus expansion has already been identified [76], and the involvement of other junction types is possible. Cd-induced mis-expression of cell adhesion molecules, including various connexins [66,67], and also delocalization of molecules such as zonula occludens (ZO)-1 and N-cadherin at a cellular level has been associated with abnormalities in cell adhesion in gap, occludens and adherens junctions [77]. Accordingly, implantation of Cd-treated embryos is reduced when compared to controls, even at doses that fail to disturb in vitro development of the early embryo [64]. Trophoblast formation and invasion have also been identified as targets for Cd toxicity. Low concentrations (0.5 ␮g/ml) of CdCl2 presented to blastocysts in vitro significantly retarded the trophoblastic outgrowth areas and transformation of trophectoderm into giant cells [78]. Higher concentrations (1–5 ␮g/ml) caused trophoblastic detachment. Placental necrosis in rats injected with 40 ␮M/kg Cd SC on day 18 of pregnancy was preceded by an early toxic effect on trophoblast cells [79]. JAr choriocarcinoma cells, a neoplastic trophoblast cell line which is similar to early human trophoblast cells, was used by Powlin et al. [80] to determine possible effects of Cd on the trophoblast. Exposure of this cell line to 20–40 ␮M Cd inhibited cell proliferation. Prolonged exposure to lower concentrations of Cd (≥0.16 ␮M) caused changes in cellular morphology and detachment from neighbouring cells in association with reduced Ca influx [81]. Modification of the effect of Cd on the cell cycle by zaldaride, a calmodulin inhibitor, indicated that calmodulin, an intracellular calcium-binding protein, has a role in mediating Cdinduced toxicity in the trophoblast. Increased metallothionein (MT)

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Table 2 Cd exposure of the post-implantation embryo Stage of exposure

Species tested

Teratogenic effects observed

Gastrulation

Zebrafish, Xenopus, Rodents (hamster, rat, mouse)

Early neurulation

Chick, xenopus, rodents

Late neurulation

Chick, xenopus, rodents

Post-neurulation

Rodents

Growth retardation, craniofacial defects, ocular malformations, notochord/somite abnormality, gut malformations, cardiovascular anomalies, hypopigmentation, skeletal anomalies Abnormal body axis, ventral body wall (VBW) defects, upper limb defects, primary neural tube defects, facial clefting, diaphragmatic hernia, cardiovascular anomalies Abnormal body axis, VBW defects (abdominal), lower limb defects, retarded somite growth Secondary neural tube defects, limb reduction defects, urogenital abnormalities, umbilical hernia

The stage of exposure is related to the teratogenic effects observed. In all species treated, exposure at the time of gastrulation resulted in widespread damage to the embryo, with multiple structural malformations.

protein expression has been found in trophoblast cells exposed to Cd [80,81] and other placental sites [82,83]. The presence of MT in the placenta is generally accepted to be a protective mechanism against Cd toxicity within the cells in which it is produced, and is also thought to restrict Cd movement across the placenta, as the level of Cd is lower in fetal than in maternal blood [82]. However, it is thought by some [82,84] to have detrimental effects, by binding Zn or by interfering with the ability of trophoblast cells to handle Ca and Zn at the cytosolic level. Weir et al. [85], using the perfused human placenta model, demonstrated a reduction in transplacental transfer of Zn to the fetus, and also a drop in human chorionic gonadotrophin (hCG) production by the placenta, commencing 4 h from exposure. These findings were conformed by Breen et al. [86]. Piasek et al. [59] found that the placentas of smoking mothers contained double the Cd and approximately half the progesterone of those of non-smokers, and a reduction in placental iron in smokers in whom placental Cd was elevated [87]. Cd interference with progesterone biosynthesis in cultured human trophoblast cells has been linked with reduced activity of P450 cholesterol side-chain cleavage enzyme and decreased 3␤hydroxysteroid dehydrogenase enzyme mRNA [88]. 2.5. The post-implantation embryo In animal studies, Cd has confirmed teratogenic and embryotoxic effects in a wide variety of species [89–93]. The character of the changes induced has been found to be dependent upon the strain and species investigated, the dose of Cd given, the route of administration, and the stage of embryogenesis [92,94–96] (Table 2). Anomalies described in the various species investigated are also dependent on the timing of Cd administration. Alterations in gastrulation occur in a concentration-dependent manner, and are similar across the spectrum of animal models investigated. Effects include retarded growth, microcephaly, body axial deformity and ocular anomalies [97,98]. In the FETAX (Frog Embryo Teratogenesis Assay: Xenopus) experiment, Sunderman et al. [93] exposed Xenopus laevis embryos to various concentrations of Cd from 5 h post-fertilization, with exposure continuing through the gastrulation period up to 101 h post-fertilization. Anomalies described included gut malformations, heart lesions, facial anomalies and fin dysplasias, in addition to the aforementioned defects. In zebrafish, cardiac edema, tail malformations and hypopigmentation were also reported [99]. Hen Chow and Cheng [100] noted that zebrafish embryos exposed to Cd during the gastrulation period had defects in somite structure, which probably contributed to altered axial curvature, and also abnormal notochordal morphology, with the notochord failing to extend into the tail region. Alteration in body axial curvature was also a prominent finding in studies on the chick embryo. Menoud and Schowing [89] detected distortion of the longitudinal body axis and symmelia, a

condition in which the limbs are orientated backward and fused, in the chick following administration of Cd at 42 h, a stage at which neurulation is complete apart from the closure of the posterior neuropore. The upper limbs were predominantly affected when treated at this time point. In our more recent studies [101,102], chick embryos were treated with the same dose of Cd after 60 h incubation (corresponding to Hamburger and Hamilton stage 16–17), and developed a similar effect in the region of the lower limbs, associated with a large defect in the ventral abdominal wall. Review of Menoud and Schowing’s work revealed that strophosomy, a form of deficiency in the ventral body wall, is also described in their findings. Timed histological studies in our laboratory demonstrated that the ventral body wall defect, along with abnormal positioning of the limbs, was due to an abnormality in the direction of growth of the lateral plate mesoderm (LPM), with failure of embryonic folding in this region. Similar to the findings in the zebrafish, we found that Cd administration had profound effects on somite structure, with increased apoptosis in all three derivative embryonic tissues (sclerotome, myotome and dermatome), retarded growth and differentiation and reduced somite numbers 24 h after treatment [101–103]. The main thrust of our own work has been the investigation of Cd-induced ventral body wall defect, which is a highly reproducible and specific abnormality, similar in many respects to the human condition omphalocele, caused by direct exposure of the H–H stage 16–17 chick embryo blastodisc to 50 ␮l of 50–90 ␮M Cd [101]. The primary histological abnormality in the chick arises 3–4 h after treatment, with cell junction breakdown in the periderm as an initial event. Apoptotic changes in the ectoderm, neural tube, lateral plate mesoderm and particularly the somites occur within 4–8 h after Cd. As somite development is dependent on normal Wnt signaling from the ectoderm, we have been working on the hypothesis that disruption of peri-ectodermal tissue can interrupt this signaling pathway. The molecule ␤-catenin is pivotal in this respect, having roles to play in both the canonical Wnt pathway and being an intracellular associate of cadherin cellular adhesion molecules. We propose that breakdown of adherens and desmosomal junctions releases ␤-catenin into the cytosol with subsequent nuclear translocation, a theory borne out by recent immunohistochemical studies in our laboratory [104]. We have seen hyperproliferation in the peri-ectodermal tissue within this timeframe [102], which could be a local effect of ␤-catenin-mediated transcription of cyclin D1 or other cell cycle promoters. Therefore, it is feasible that somitic cell death and delay in differentiation could result from the Wntlike signaling effect of released ␤-catenin within the periderm and ectoderm, causing an imbalance in gene expression required for normal development of the somites. Further studies are in progress to examine these possibilities. In rodent models, axial anomalies appear to be less remarkable, but there are clear effects on neural tube closure, limb development and soft tissue formation, depending on the timing of administra-

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tion of Cd. Webster and Messerle [90] found that IP injection of 4 mg/kg CdCl2 in the mouse produced neural tube defects when given on day 7 or 8, that is to say, during neurulation. When Cd was given on day 9 or 10, exencephaly did not occur, but the neural tube re-opened secondary to marked cell death. This phenomenon was observed also by Schmid et al. [105] in mouse embryos cultured in vitro in the presence of Cd. Re-opening of the neural tube may occur when a critical level of cell necrosis in the wall of the neural tube is exceeded [106]. Mice treated with 4 mg/kg CdCl2 shortly after neurulation developed limb reduction abnormalities [91]. Cd administered IP to pregnant CD-1 mice on days 9–12 gestation induced postaxial forelimb defects in the offspring, indistinguishable from those produced by acetazolamide. This suggests a role for the enzyme carbonic anhydrase in the mechanism of teratogenesis [94]. In the hamster, Gale and Layton [107] injected a single dose 2 mg/kg CdSO4 intravenously on day 8 of gestation, and found a multiplicity of craniofacial, skeletal and soft tissue abnormalities upon examination of the fetuses on day 15. More recent studies using whole-embryo culture assay for hamster embryos reported 70% abnormality rate in embryos cultured at 0.25 ␮M Cd on day 8 of gestation, with animals exhibiting growth retardation, reduction in somite gain, axial anomaly and neural tube defect [108]. In a classic study, Ferm [109] administered Cd at multiple stages of embryonic development in the golden hamster, treating animals either early on day 8 of gestation, corresponding to the primitive streak stage, late on day 8, early on day 9 (when the neural tube has closed) or on the afternoon of day 9. The earliest treatment group had the highest rate of embryolethality, and exencephaly and eye defects were more frequent in this group. Rib and upper limb defects were most frequent in those treated late on day 8, whereas later groups had a preponderance of lower limb defects, reflecting the time-lag between upper and lower limb outgrowth and development. Sequential administration of Cd to pregnant rats yielded comparable time-related defects. Samarawickrama and Webb [110] found that hydrocephalus and eye defects occurred most frequently in pups of Wistar–Porton rats that received 1.25 mg Cd/kg body weight on days 9 and 10 of gestation. In the post-neurulation period, Holt and Webb [96] found that administration of Cd on gestation days 10, 12 or 14 increased the incidence of skeletal retardation, hydrocephalus, urogenital abnormalities, cleft palate, diaphragmatic hernia, and cardiovascular anomalies. Umbilical hernia and hydronephrosis did not occur before day 11 [110] and renal agenesis occurred exclusively within a narrow time window between days 9 and 11 of gestation. Although Cd has yet to be shown to be teratogenic in humans, an inverse relationship has been found between birth weight and cord blood Cd levels [111], birth weight and placental Cd concentration [112], and between birth length and maternal blood Cd [113]. There is, however, an association between maternal Cd exposure and early delivery, which may explain the association with smaller babies [114]. An inverse relationship has also been demonstrated between birthweight and Cd content in newborn hair [115,116]. Cd blood levels in newborn infants have been found to be 70% of maternal levels [117]. The difference in Cd level can probably be ascribed to placental function, as perfusion of human placentas with Cd [118] showed that the transfer rate from maternal to fetal side was very slow. The metal did not appear on the fetal side of the placenta until 40 min after perfusion started, and reached a steady state approximately 1 h later, at which time the concentration in the fetal perfusate was 1/20th that of the maternal. Microstructural changes, including edema and vacuolation followed by necrosis, were detected in placentas between 5 and 8 h after perfusion with Cd [85].

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Postnatal effects described in association with exposure to abnormally high levels of Cd in utero include significantly increased DNA synthesis in the bowel and bone marrow in 6-week old rats following administration of Cd orally to pregnant dams at a dose of 50 ppm in drinking water [119], and depressed immune function and sensorimotor abnormalities in animal studies and in children following prenatal exposure to Cd [120]. However, human studies to date are not conclusive due to confounding factors and problems in study design, and additional work needs to be done to elucidate the extent and type of damage to children following exposure. This is of particular relevance because of altered pharmacokinetics of metals in very young children, in whom absorption is more avid, excretion less efficient, and the blood–brain barrier less effective [120]. 3. Mechanisms of reproductive toxicity A number of mechanisms of Cd toxicity have been suggested, including ionic and molecular mimicry, interference with cell adhesion and signaling, oxidative stress, apoptosis, genotoxicity and cell cycle disturbance. Although the overall effect of Cd on any cell or tissue is likely to be due to a synergism of several mechanisms, it is possible that one mechanism will predominate in a specific cell type. 3.1. Altered cell adhesion Normal cell adhesion plays a role in driving meiosis and differentiation in spermatogenesis [37], oocyte maturation, formation of the OCC, pick-up of the oocyte by the cilia on the infundibulum of the Fallopian tube [74,75], compaction of the early embryo and development to the blastocyst stage [68,69]. Cell division, morphogenetic movements and apoptosis in the post-implantation embryo are also dependent on cell–cell contact and intercellular communication [121]. The interference by Cd with cell adhesion is complex, and is probably modified by the many other cellular effects of this metal. Desmosomal disruption in the testis secondary to Cd administration has been documented by Berliner and Jones-Witters [28] and Fende and Niewenhuis [26]. Functional adherens junctions have also been identified in the testis [122], although the inter-relationship between adherens junctions, tight junctions and desmosomes is very different to that described in other epithelia [123]. The integrity of the blood–testis barrier (BTB) is dependent on tight junctions between Sertoli cells located deep within the wall of the seminiferous tubule near the basal lamina, creating separate microenvironments for the development of pre-leptotene/leptotene spermatocytes on the basal aspect, and pachytene spermatocytes/spermatids on the luminal side. Wong et al. [36] have demonstrated the ability of Cd to disrupt the BTB via the TGF␤3/p38 pathway, with secondary loss of function at adherens junctions. Assembly and disassembly of inter-sertoli junctions is essential for normal migration of germ cells across the seminiferous epithelium and their concomitant differentiation [124]. Similarly, maintenance of a specific microenvironment in the epididymis is crucial to the further maturation of spermatozoa [125]. Intact mechanisms for cell adhesion are also essential for normal oogenesis, OCC expansion and oocyte pick-up by the Fallopian tube, as outlined in Sections 2.2 and 2.4. In the pre-implantation human embryo, gap junctions, expressing predominantly connexin-43, have been detected from the four-cell stage, becoming increasingly organized as development proceeds [68,126]. Construction of gap junctions in vertebrates depends on a family of transmembrane proteins called connexins [127]. Gap junction intercellular communication (GJIC), demon-

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strated by Jeong et al. [66] to be inhibited by Cd, is of utmost importance in transmission of small molecules and electrical impulses from cell to adjacent cell. Gap junction formation is heralded by the expression of various connexins, many of which have short lives of a few hours or less. Cd treatment impedes gap junction assembly by interfering with phosphorylation of certain connexins, thus preventing clonal expansion within the early embryo [67]. It also produces a decrease in expression of connexins 26 and 32 [66]. Interestingly, gap junctions co-localize with cadherins, and connexin-43 is regulated by ␤-catenin, already implicated as a possible player in Cd teratogenesis in the post-implantation embryo [104]. This again raises the possibility that ␤-catenin released upon disassociation of cadherin complexes could translocate into the nucleus and alter gene expression, in this case, connexin-43 expression. In the murine model, ␤-catenin accumulates in the cytoplasm between the two-cell and morula stages of development, but no evidence of concomitant nuclear staining has been observed [128]. Reactivation occurs between late morula and early blastocyst stages, a time when gap junction assembly is essential for compaction to occur, and deactivation occurs again when the fully expanded blastocysts hatch from the zona pellucida. Cd, through its action on cadherins and other adhesion molecules, can theoretically interfere with this tightly regulated activation of ␤catenin. Furthermore, tight junctions, another target for Cd [77], are known to be critical for blastocyst formation in mammalian embryos [129,130], in terms of establishing and maintaining watertight barriers within the formed blastocyst. Desmosomal junctions have also been detected in the pre-implantation human embryo at the blastocyst stage [68,131], and are probably vulnerable to Cd interference, although no experimental evidence of this has been presented to date. An interesting possible role for cell adhesion molecules is their role in signaling, remodeling and migration in the postimplantation embryo. In our previous work with Cd, we have noted a strong relationship between breakdown in cell adhesion in the periderm and failure of the lateral body folds to migrate ventrally in the post-gastrulation chick embryo [101,102], resulting in a large ventral body wall defect. Preliminary work in our laboratory has indicated that there is no change in distribution of cadherins in chicks with extensive peridermal desquamation following Cd, but associated molecules such as ␤-catenin and actin are profoundly affected, with marked clumping of actin and movement of ␤-catenin from its predominantly peripheral location adjacent to cellular junctions to a more generalized cytosolic distribution and, more importantly, into the nuclei of the periderm cells, indicating a distribution of ␤-catenin equivalent to that seen in activation of the canonical Wnt pathway [104]. Tight, adherens, and gap junctions, and desmosomes have been demonstrated in the periderm in developing humans and rats [132–135], and any or all of these molecules could be a target for interference by Cd, but a link between Cd-related junction disruption and aberrant signaling in the later embryo has yet to be confirmed. 3.2. Oxidative stress and apoptosis The role of Cd in inducing oxidative stress in adult tissue has been well documented. Increased levels of malondialdehyde, protein oxidation and reduced activity of superoxide dismutase (SOD) in rat testes have been reported following a single SC injection of 2 mg/kg CdCl2 [136], indicating an increase in lipid peroxidation and a decrease in the removal of superoxide radicals. Adequate Zn supplementation in the diet partially reversed these changes. Amara et al. [137] attributed decreased testicular growth rate, plasma testosterone, and reduced sperm count and motility to Cd-induced oxidative stress, as concurrent reduction

in glutathione peroxidase, catalase, mitochondrial Mn–SOD, and cytosolic CuZn–SOD, along with increased MT and malondialdehyde, were observed. Vitamins C and E can protect the rat testis from Cd-induced oxidative damage [138,139], and selenium [140–142], n-acetylcysteine [139], glycine [143], and glutathione-like compounds [144] have also been shown to rescue various cells and tissues from this mechanism. The pro-apoptotic transcription factor p53 is activated by oxidative stress, which aids in the generation of reduced glutathione and expression of sestrins, thus decreasing reactive oxygen species (ROS) and protecting the organism from resultant DNA damage [145]. However, although Cd induces abnormal levels of apoptosis in the testis, it does so via a p53-independent pathway [146] and in fact has been associated with a suppression of p53 expression. Conversely, similar doses produce apoptosis in the ventral prostate which is clearly p53-dependent [147]. In general, mechanisms by which Cd provokes apoptosis depend upon the dose given [148] and the cell type exposed [149]. Activation of p38 and disruption of the mitochondrial membrane appear to be the primary targets. However, Cd can also initiate apoptosis by intrinsic pathway activation, and at higher Cd exposure, necrosis may supervene. In view of the above, it is surprising that the literature lacks reports of experiments to detect oxidative stress in the whole embryo. So far, in our laboratory, efforts to detect oxidative stress in chick embryos have yielded ambiguous results. Similarly, we have failed to show a blockade to the teratogenic effects of Cd by antioxidants. Our preliminary results [150] have shown that selenium (Se) in doses of twice the molarity of Cd confers 100% protection against the teratogenic effects of Cd in the chick. Although Se is a known antioxidant [141,142], its protection against Cd may lie more simply in its ability, as a divalent cation, to block Cd uptake by embryonic cells. 3.3. Ionic and molecular mimicry Divalent cations, many of which are nutritive metals, have demonstrably protected embryonic and adult tissue from Cdinduced damage, indicating a mechanistic link [101,151,152]. The ability of Cd to imitate these nutrients in many biological pathways and processes is a very probable explanation for its toxicity. Experimental evidence indicates that Cd may interact with membrane transporters involved in the uptake of Ca, iron (Fe) and Zn through a process of “ionic mimicry” [153,154]. Transporters for other metals such as manganese (Mn) have also been implicated in Cd handling [153,155,156]. Ca channels are thought to be involved in Cd uptake in certain tissues, due to the similarity of ionic radii between Cd and Ca [157], and also because experiments with certain cell types demonstrated that the Ca channel antagonists nifedipine and verapamil significantly blocked the uptake of Cd [63,158]. It has been estimated that between 30 and 50% of Cd enters cells through Ca channels [158,159]. As well as interfering with cellular transport processes, Cd has also been found to form covalent complexes with sulfhydryl-containing biomolecules such as cysteine and glutathione (GSH), thus altering their behavior. It has also been proposed that Cd may substitute for nutritive metals in various enzymes and cell adhesion molecules [95,160–166]. Several studies indicate that Cd gains access to testicular cells by mimicking Zn at the site of Zn-transporters [167–169]. Passive and active mechanisms are involved in testicular transport of Cd. Addition of Zn can significantly inhibit the active component of Cd uptake [170], thus providing a mechanism by which Zn alleviates Cd toxicity in the testis. About 20% of Zn uptake by placental tissue is susceptible to competitive inhibition by Cd [171]. In addition to interference with Zn uptake, the importance of the presence of Cd within the cell becomes clear when considering the potential

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functions of Zn. Substitution of Cd for Zn in carbonic anhydrase may result in the inability to catalyze the hydration of CO2 to bicarbonate, thus creating an acidotic embryonic environment with adverse reproductive consequences [95]. Other enzymes from which Cd could potentially displace Zn include alcohol dehydrogenase, lactic dehydrogenase, malic dehydrogenase, alkaline phosphatase and carboxypeptidase [160,161]. Sunderman [172] proposed that the teratogenic effect is mediated by Cd substitution for Zn in finger-loop domains of certain DNA-binding proteins that regulate gene expression during embryogenesis. Experiments by Predki and Sarkar [173] confirmed that certain Zn-finger residues have a higher affinity for Cd than for Zn. The effects of Cd on Ca-regulated pathways are so diverse that a comprehensive account is outside the scope of this article. Important potential effects include replacement of Ca by Cd in the extracellular ligands of cadherin molecules [162], in desmosome and gap junction assembly [163–165], and in integrin function [166]. Mobilization of Ca from internal stores can be induced by extracellular Cd via the phosphatidylinositol pathway [174,175], upregulating the activity of Ca-dependent intracellular pathways and causing abnormal cell proliferation or cell death [175]. Intracellular Ca2+ is maintained at a low level (≤0.1 ␮M) due to the presence of Ca2+ pumps that export Ca2+ from the unstimulated cell. Activation of the phosphatidylinositol pathway mobilizes intracellular Ca2+ from the endoplasmic reticulum by binding to receptors that are ligand-gated Ca channels, thus increasing intracellular Ca2+ to about 1 ␮M. This, in turn, alters the activities of target proteins, many of which are enzymes [176]. Many of the effects of intracellular Ca2+ are mediated by the Ca-binding protein calmodulin, which is generally activated when cytosolic Ca2+ increases to above 0.5 ␮M, and is also known to be activated by Cd [121,177]. Therefore, several aspects of Ca movement and action, of vital importance in fertilization and subsequent embryo development, may be influenced by Cd. Metal homeostasis has to be carefully regulated by the cell to prevent production of toxic free radicals [178], thus triggering oxidative stress. Induction of metallothionein (MT) [179] by Zn and possibly other metals is an alternative mechanism by which protection against Cd toxicity could be conferred. This low-molecular weight protein binds to Cd, limits its availability to cells and tissues [82,180], and plays a role in transport, detoxification, and storage [181]. A possible role for MT in the oxidative stress reaction has also been documented [182]. It has been shown to be an effective free radical scavenger, important because the release of various species of oxygen metabolites is thought to be indirectly responsible for the initiation of apoptosis. However, MT has itself been implicated as a causal factor in apoptosis [180], and has intrinsically toxic effects in itself when bound to Cd [183–186]. 3.4. DNA damage and effect on the cell cycle Cd has been associated with increased chromosomal aberrations and decreased mitotic index in Chinese Hamster Ovary (CHO) cells [187], and non-disjunction in germ cells [188]. Inhibition of DNA repair [189] is thought to be an important mechanism by which Cd exerts its mutagenic effect in this cell type. Resistance to the toxic effects of Cd in CHO cells has been linked to the ability to synthesize metallothionein (MT)-1 protein and MT-1 gene amplification [190]. Administration of the ROS scavenger D-Mannitol was found to protect CHO cells against the mutagenicity of Cd, whereas the catalase inhibitor 3-amino-1,2,4-triazole potentiated this effect [191]. Increased mutation frequency after Cd was found to be associated with reduced activity of anti-oxidant enzymes, among them GSH peroxidase and catalase. This latter evidence indicates that the mutagenic effect of Cd is consequent upon its ability to gen-

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erate oxidative stress within the tissues. A dose-dependent effect was seen on the cell cycle in this culture system, with retardation of cell cycle progression of cells cultured with 1 ␮M Cd or more for 24 h [192], G2/M arrest in cells treated with lower doses (1 ␮M) and S-phase arrest at higher doses (4 ␮M).

4. Conclusions Cd has the potential to affect reproduction and development in many different ways, and at every stage of the reproductive process. Effects on the testis include disruption of the blood–testis barrier due to adverse effects on cell adhesion, oxidative stress, and necrosis at higher experimental doses. Incorporation into chromatin of the developing spermatozoa has also been described. In the ovary, oocyte development is inhibited, steroidogenesis reduced, and ovarian hemorrhage and necrosis supervene at higher Cd doses. Cumulus expansion and possibly oocyte pick-up by the tubal epithelium are also hampered, probably by interference with normal junction formation. Although the early pre-implantation embryo seems relatively resistant to Cd, increasing incorporation from the four-cell stage onwards predisposes to apoptosis in the blastocyst and implantation failure. Injurious effects on the placenta, including inhibition of trophoblastic invasion, decreased steroidogenesis, and adjusted handling of nutritive metals, have also been identified. The clinical implications of the effects of Cd on oocyte maturation, oocyte pick-up and development of the pre-implantation embryo through to implantation are obvious. An important source of this toxin in humans is inhalation through cigarette smoking. Cd thus absorbed has a relatively high bioavailability, and is readily assimilated into human tissues and transported via the bloodstream to more remote parts of the body, notably the reproductive tract. Cigarette smoking has been associated with subfertility in a number of studies [193–196]. Increased blood levels of Cd have been found in smokers [197], with levels correlating significantly with numbers of cigarettes consumed. The concentration of Cd in follicular fluid of female smokers undergoing in vitro fertilization [IVF] has been reported at 7.93 ± 0.16 ng/ml [198] compared with 6.73 ± 0.31 ng/ml in non-smokers. The concentration of Cd in human ovaries increases over time [199] with a linear increase with age in females between 30 and 65 years. Smokers were found to have increased ovarian Cd levels when compared with nonsmokers. Human ovarian granulosa cells exposed to Cd exhibited morphological alterations, breakdown of intercellular junctions and defective steroidogenesis. However, the precise mechanism by which Cd impedes oocyte development and granulosa cell function has yet to be identified, in particular at concentrations comparable to those found in the follicular fluid of smokers, and manipulations by which this damage might be prevented or reversed should be identified in vitro and in vivo. The role of oxidative stress and the possibility of preventing or reversing this effect in particular needs to be explored. Cigarette smoking has also been identified as a risk factor in ectopic pregnancy in several studies [200,201], possibly reflecting faulty movement of the oocyte through the uterine tubes, a process which is highly dependent on sequential cell junction formation. To date, the precise role of Cd in this sequence of events has not been defined. However, recurrent miscarriage and uterine fibroids have been directly linked with increased urinary Cd excretion in human females [202]. The effects of Cd on the post-implantation embryo are stageand dose-dependent. Effects caused by administration in the peri-gastrulation phase are myriad, with growth retardation, craniofacial defects, cardiovascular malformations, gut anomalies and body axial deformities reported in many of the species investigated.

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Neural tube defects could be produced by giving Cd before neurulation was complete, causing failure of fusion of the neural folds, or as a secondary phenomenon due to cell death in the neural tube. Limb and body wall defects tended to occur with Cd administration in the post-neurulation period. Mechanistic studies to date have indicated that teratogenesis can be linked to substitution of Cd for predominantly Zn or Ca in many biological processes, thus inactivating cellular adhesion and modifying in some way a wide selection of intracellular reactions, giving rise to oxidative stress and abnormal cell proliferation and apoptosis. This in turn opens the possibility of the involvement of signaling pathways, a line of inquiry currently being pursued in our laboratory, and the opportunity to look at means by which we might protect the developing embryo by signal modification, prevention of Cd uptake, competitive inhibition of intracellular Cd, or use of anti-oxidants. While the effects on gametogenesis, implantation and early embryo development seem to depend largely on impaired cell adhesion, other, less direct, effects of Cd can produce sub-optimal conditions for a safe pregnancy outcome, including iron and zinc deficiency in the fetus [85,203] and a possible association with the pathogenesis of pre-eclampsia [204]. In addition, pre-term delivery and cesarian section delivery were found to occur approximately four times more frequently in women with higher Cd burdens (urinary Cd ≥2 ␮g/g creatinine) than in their counterparts with lower Cd burdens [13]. Further experimental work in animal models and carefully designed case-controlled studies in humans are required to elucidate the consequences of Cd exposure on late pregnancy complications, neonatal health and long-term postnatal outcome. Whatever the effect, it is certainly prudent to recommend cessation of smoking in pregnancy and in the pre-conception period, and supplementation with micronutrients such as Zn, Fe, Se and vitamin C should be considered in ex-smokers having difficulty conceiving. References [1] WHO. Cadmium (Environmental Health Criteria No. 134). Geneva: WHO; 1992. [2] Muntau H, Baudo R. Sources of cadmium, its distribution and turnover in the freshwater environment. IARC 1992;118:133–48. [3] IARC. IARC monographs on the evaluation of carcinogenic risks to humans, vol. 58. Beryllium, Cd, mercury and exposures in the glass manufacturing industry. Lyons, France: IARC; February 9–16, 1993. [4] Martelli A, Rousselet E, Dycke C, Bouron A, Moulis J-M. Cadmium toxicity in animal cells by interference with essential metals. Biochimie 2006;88:1807–14. [5] WHO. Cadmium—environmental aspects (Environmental Health Criteria 135). Geneva: WHO; 1992. [6] IRPTC. IRPTC legal file 1986, vol. 1. Geneva: International Register of Potentially Toxic Chemicals, United Nations Environment Programme; 1987. [7] McLellan JS, Flanagan PR, Chamberlain MJ, Valberg LS. Measurement of dietary cadmium absorption in humans. J Toxicol Environ Health 1978;4:131–8. [8] Nasreddine L, Parent-Massin D. Food contamination by metals and pesticides in the European Union. Should we worry? Toxicol Lett 2002;127:29–41. [9] Watanabe T, Koizumi A, Fujita H, Kumai M, Ikeda M. Role of rice in dietary cadmium intake of farming population with no known man-made pollution in Japan. Tohoku J Exp Med 1984;144:83–90. [10] Vahter M, Berglund M, Slorach S, Jorhem L, Lind B. Integrated personal monitoring of cadmium exposure in Sweden. IARC 1992;118:113–20. [11] Jarup L, Berglund M, Elinder CG, Nordberg G, Vahter M. Health effects of cadmium exposure-a review of the literature and a risk estimate. Scand J Work Environ Health 1998;24(Suppl 1):1–51. [12] McKenzie-Parnell JM, Kjellstrom TE, Sharma RP, Robinson MF. Unusually high intake and fecal output of cadmium, and fecal output of other trace elements in New Zealand adults consuming dredge oysters. Environ Res 1988;46:1–14. [13] Satarug S, Moore MR. Adverse health effects of chronic exposure to lowlevel cadmium in foodstuffs and cigarette smoke. Environ Health Perspect 2004;112:1099–103. [14] Nandi M, Slone D, Jick H, Shapiro S, Lewis GP. Cadmium content of cigarettes. Lancet 1969;2:1329–30. [15] Lewis GP, Coughlin LL, Jusko WJ, Hartz S. Contribution of cigarette smoking to cadmium accumulation in man. Lancet 1972;1(7745):291–2. [16] Friberg L, Piscator M, Nordberg GF, Kjellstrom T. Cadmium in the environment. 2nd ed. Cleveland, OH: Chemical Rubber Co.; 1974.

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