Embryotoxicity of the nitric oxide donor sodium nitroprusside in preimplantation bovine embryos in vitro

Animal Reproduction Science 91 (2006) 225–236

Embryotoxicity of the nitric oxide donor sodium nitroprusside in preimplantation bovine embryos in vitro Nicolas M. Orsi ∗ Perinatal Research Group, Academic Unit of Paediatrics, Obstetrics and Gynaecology, D Floor, Clarendon Wing, Leeds General Infirmary, Belmont Grove, Leeds, LS2 9NS, UK Received 14 February 2005; received in revised form 21 April 2005; accepted 29 April 2005 Available online 16 June 2005

Abstract Many early pregnancy complications are associated with an imbalance in pro- and antiinflammatory cytokines, resulting in alterations in nitric oxide (NO) profile. Since very little is known about the modus operandi of this free radical in early embryos, this study characterised NO embryotoxicity in terms of bovine embryo development and metabolism. Embryos were generated by in vitro maturation and fertilisation of oocytes aspirated from abattoir-derived ovaries. Zygote to blastocyst rates were measured in SOFaaBSA in the presence and absence of the NO donor sodium nitroprusside (SNP) over the 0–50 ␮M range (n = 10 per group). Since concentrations <10 ␮M SNP depressed blastocyst rate, blastocyst cell numbers (determined by bisbenzimide staining; n = 22 and 20), glucose, pyruvate, lactate (measured ultramicrofluorometrically) and amino acid profiles (quantified by HPLC; n = 28 and 23) were assessed at 0 and 10 ␮M SNP. SNP depressed cell numbers, reduced pyruvate and glucose uptake, perturbed quantitative tyrosine, threonine, phenylalanine, lysine, glycine, tryptophan, methionine and valine profiles, and decreased retention into the negative range (P < 0.05). Qualitative asparagine and lysine profiles were affected by SNP, while proportional amino acid production and consumption were increased and decreased, respectively (P < 0.05). These findings indicate that SNP (presumably through increases in NO profile): (i) fails to improve bovine embryo development in vitro, (ii) exerts toxic effects, likely through ATP starvation induced by cytochrome c oxidase (oxidative phosphorylation) and glyceraldehyde-3-phosphate dehydrogenase (glycolysis) inhibition,

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and (iii) may affect albumin endocytosis/hydrolysis or protein biosynthesis, rather than causing a loss of intracellular amino acids or simply depressing their metabolism. © 2005 Elsevier B.V. All rights reserved. Keywords: Nitric oxide; Embryo metabolism; ATP starvation

1. Introduction A dysfunctional or untimely maternal inflammatory/immunological response underpins many early pregnancy complications, including poor embryo development, aberrant trophoblast invasion, failed implantation and recurrent miscarriage. These processes are typically characterised by an imbalance in T-helper type 1 (Th1; pro-inflammatory) and Th2 (anti-inflammatory) cytokines (Laird et al., 2003; Dey et al., 2004; Strandell et al., 2004; Tjoa et al., 2004). Many adverse effects of Th1 cytokines are mediated through alterations in nitric oxide (NO) profile (Athanassakis et al., 1999). Although embryos themselves do not produce toxic amounts of NO – unless exposed to embryotoxic factors – they are vulnerable to exogenous NO produced in response to maternal tract or systemic inflammatory/infectious processes (Lim and Hansel, 1998; Athanassakis et al., 2000; Hansen et al., 2004). NO is a free radical with a plethora of biological activities. Due to its brief half-life (t1/2 = 4 s), NO acts in an auto- and paracrine manner to transduce cellular signals by interacting with the haem prosthetic group of guanylate cyclase, increasing cyclic guanosine monophosphate, which then mediates its effects (Clementi et al., 1999; Inoue et al., 1999). NO is produced by NO synthase (NOS) through l-arginine oxidation to NO and citrulline in the presence of oxygen and NADPH (Wu and Morris, 1998). There are three NOS isoforms: neuronal, endothelial and inducible (nNOS, eNOS and iNOS), which have distinct functions, are independently regulated and differ in tissue localisation (Clementi et al., 1999). An inner mitochondrial membrane isoform (mtNOS) regulates respiration (Lopez-Figueroa et al., 2000). Both iNOS and eNOS have been identified in early mouse embryos and their implantation sites (Gouge et al., 1998; Purcell et al., 1999). NO production is tightly controlled and decay occurs principally through peroxynitrite formation and ubiquinol oxidation (Poderoso et al., 1999). Cells can both inhibit (e.g. by glucosamineinduced reduction of NADPH availability) and elicit NO production (e.g. in response to cytokines, bacterial endotoxin and ammonia) (Clementi et al., 1999; Kosenko et al., 1998; Wu et al., 2001). Nanomolar NO levels can inhibit respiration (and thereby ATP production) by reversible binding to the oxygen-binding site Fe2+ of cytochrome c oxidase (Blackmore et al., 1991; Brown, 1999). NO increases the apparent Km of respiration for oxygen, although inhibition is affected by oxygen, pyruvate, malate and succinate levels (Brown and Cooper, 1994; Takehara et al., 1995). Low NO profiles are antiapoptotic by reducing caspase activation, inducing bcl-2 (a mitochondrial permeability inhibitor) expression, and preventing cytochrome c release and cleavage of poly (ADP-ribose) polymerase (Genaro et al., 1995; Dimmeler et al., 1997; Leist et al., 1999; Li et al., 1999; Rossig et al., 1999). At micromolar concentrations, NO reacts with superoxide (O2 •− ) to generate peroxynitrite, which leads to iron–sulphur cluster destruction, lipid peroxidation, thiol nitrosylation

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and amino acid residue oxidation/nitration (Darley-Usmar and Halliwell, 1996; McBride et al., 1999). While nitration alters protein function and signal transduction pathways, lipid peroxidation results in membrane disruption (Darley-Usmar et al., 1995; Dixit and Parvizi, 2001; Hurst and Dobbie, 2001). Peroxynitrite also decays to cytotoxic products and reacts to generate yet more NO donors (e.g. nitrosothiols). NO and peroxynitrite induce iron and copper release, respectively, both of which promote oxidative damage (Darley-Usmar and Halliwell, 1996). Thus, NO:O2 •− balance is central to NO cytotoxicity (Keller et al., 1998; Darley-Usmar et al., 1995). Elevated NO levels can inhibit glycolysis, mitochondrial respiration and DNA replication through glyceraldehyde-3-phosphate dehydrogenase (GAPDH), cytochrome c oxidase and ribonucleotide reductase inhibition (Bolanos et al., 1994; Lepoivre et al., 1994; Brune and Mohr, 2001). This is exacerbated by NO and peroxynitrite deamination of nucleotide bases resulting in DNA fragmentation (Nguyen et al., 1992; Szab´o et al., 1996), which activates the nuclear repair enzyme poly (ADP ribose) synthetase. This enzyme uses NAD+ and involves further ATP consumption in order to regenerate this substrate (Hurst and Dobbie, 2001). Thus, variations in NO, O2 •− , oxygen and glutathione participate in the metabolic regulation (Foresti et al., 1997; Inoue et al., 1999). High NO levels are proapoptotic and increase expression of the tumour suppressor product p53 (Calmels et al., 1997; Haendeler et al., 1999) and the pore forming protein bax (Messmer et al., 1996) as well as disrupting transmembrane potential (Hortelano et al., 1999), causing cytochrome c release, a fall in glutathione level (Bustamante et al., 2000) and the onset of apoptotic pathways. Unlike NO, peroxynitrite irreversibly inhibits respiration through protein iron–sulphur centre displacement and oxidative damage to mitochondrial membranes/complexes I, II, IV and V, aconitase and superoxide dismutase, thereby resulting in loss of mitochondrial function and a fall in ATP:ADP ratio (Henry et al., 1993; Brown, 1999, 2000; Welter et al., 1996; Virag et al., 2003). Although NO promotes murine oocyte maturation, sperm capacitation/hyperactivation and trophoblast outgrowth (Hellstrom et al., 1994; Sengoku et al., 2001), its role in preimplantation development remains unclear. Although required for early embryo development (Fukuda et al., 1996; Gouge et al., 1998; Manser et al., 2004), NO can be cytotoxic to mouse and cattle embryos through ill-defined mechanisms. In view of the above, this study therefore examined the embryotoxicity of the NO donor sodium nitroprusside (SNP) in terms of bovine embryo development and metabolism. Such early embryos present good models due to their relative autonomy.

2. Materials and methods 2.1. Embryo production and culture All chemicals were purchased from Sigma (Poole, Dorset, UK), unless otherwise specified. Bovine embryos were generated by in vitro maturation and fertilisation of oocytes aspirated from abattoir-derived ovaries, as previously described (Orsi and Leese, 2004a, 2004b). Oocyte-cumulus complexes were matured for 24 h in tissue culture medium 199 supplemented with 10% fetal calf serum, 1 ␮g/ml oestradiol, 10.9 ng/ml fibroblast growth factor, 0.47 ␮g/ml epidermal growth factor, 0.025 IU luteinising hormone, 0.025 IU

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follicle-stimulating hormone (gonadotrophins from Ferring Pharmaceuticals, Berkshire, UK), 24 ␮g/ml apo-transferrin and 1 mM pyruvate. Fertilisation was carried out in Fertilisation Tyrode’s Albumin Lactate Pyruvate (Fert TALP) medium with 1 × 106 sperm/ml of freeze–thawed semen from a sire of proven fertility. Presumptive zygotes were cultured in groups of 14 in 20 ␮l microdrops of SOFaaBSA (Tervit et al., 1972) under a mineral oil overlay at 39 ◦ C and a humidified 5% O2 /5% CO2 /90% N2 atmosphere. The developmental effects of the NO donor SNP were investigated by determining blastocyst rate on day 8 of culture in the following concentration groups: control, 10 nM, 100 nM, 1 ␮M, 10 ␮M, 20 ␮M and 50 ␮M (n = 10 replicates per group). NO is generated from SNP through the reduction of nitroprusside by microsomes/NADPH, submitochondrial particles/NADH, thiols or cell lines in vitro (Rao and Cederbaum, 1995). SNP was specifically chosen to study the potentially embryotoxic effects of NO since this approach has already been validated in somatic cells in vitro (Tjuvajev et al., 1998). 2.2. Evaluation of blastocyst cell numbers In view of the findings from the concentration–response curve, cell counts were performed by bisbenzimide staining on day 8 expanded bovine blastocysts cultured in SOFaaBSA with or without 10 ␮M SNP (n = 22 and 20, respectively), as previously described (Orsi and Leese, 2001). 2.3. Metabolic profiling experiments Day 8 expanded bovine blastocysts of similar morphology were washed three times and held in incubation medium for 30 min (SOFaaBSA with or without 10 ␮M SNP) so as to allow equilibration of intracellular amino acid pools. Blastocysts were then transferred singly to 1 ␮l incubation drops of the above media for 12 h (n = 28 and 23 for the control and 10 ␮M SNP groups, respectively). Embryo-free drops were incubated alongside to account for any non-specific alterations in carbohydrate and amino acid profiles. Glucose, pyruvate and lactate profiles were determined ultramicrofluorometrically, as previously described (Thompson et al., 1996). By contrast, drops for amino acid profiling were diluted 1:40 with high-performance liquid chromatography (HPLC)-grade water (Fisher Scientific, Loughborough, Leicestershire, UK) and analysed by reverse-phase HPLC using o-phthaldialdehyde as a precolumn derivatising reagent, as previously described (Orsi and Leese, 2004a, 2004b). 0.5 mM d-␣-amino butyric acid (Sigma) was added as an internal, non-metabolisable standard to all SOFaaBSA formulations. This method did not allow the detection of proline or cysteine.

3. Data presentation and statistical analysis Blastocyst formation rates were expressed as a %. Glucose, pyruvate, lactate and amino acid profiles (appearance and disappearance from the medium) were expressed as pmol/embryo/h. The proportion of glucose conversion to lactate was expressed as glycolytic

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index (%). Amino acid data were permuted to yield overall appearance (sum of all amino acids appearing in the medium for each embryo), overall disappearance, turnover (sum of overall appearance and disappearance—this provides an indication of amino acid metabolic activity) and retention (difference between appearance and disappearance). Amino acid data were also expressed qualitatively, as a proportion (%) of turnover, to account for variations in metabolic activity between individual embryos. All data were presented ±S.E.M. Developmental data were compared by one-way analysis of variance, followed by Fisher’s LSD post hoc test. Differences in metabolism between groups were tested by Student’s t-test or, in the case of non-parametric distributions (assessed by Anderson–Darling tests), by Mann–Whitney U-tests. Percentages were arcsine log transformed for statistical analysis.

4. Results Concentrations of 10 ␮M SNP and above had deleterious effects on bovine embryo development (Fig. 1). Since 10 ␮M SNP was the lowest concentration to reduce development significantly, it was chosen – as the least pharmacological – for investigating the adverse effects of NO on blastocyst cell number and metabolism. Blastocyst cell number was significantly depressed by exposure to 10 ␮M SNP throughout preimplantation development (control: 118.47 ± 6.57; 10 ␮M SNP: 88.72 ± 8.17) (P < 0.05). The uptakes of pyruvate and glucose were significantly decreased by incubation in the presence of 10 ␮M SNP, whilst no significant differences were recorded in lactate production (Fig. 2). However, despite decreasing glucose consumption significantly, 10 ␮M SNP did not have a significant impact on glycolytic index (82.39 ± 7.09% and 94.45 ± 11.62% for the control and NO donor group, respectively). SNP significantly affected the quantitative profile of many amino acids, including: tyrosine, threonine, phenylalanine, lysine, glycine, tryptophan, methionine and valine

Fig. 1. Day 8 bovine blastocyst rate over a range of SNP concentrations (* P < 0.05; *** P < 0.001).

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Fig. 2. Pyruvate and glucose uptake and lactate production of lactate of single day 8 bovine blastocysts in the presence and absence of 10 ␮M SNP (* P < 0.05).

(Fig. 3). However, 10 ␮M SNP had no significant effects either on overall amino acid production or consumption. Overall profiles in the control and 10 ␮M SNP groups were: production, 9.46 ± 1.05 and 8.22 ± 0.51 pmol/embryo/h; consumption, −10.32 ± 0.92 and −12.09 ± 1.03 pmol/embryo/h, respectively. Similarly, SNP had no significant effect on amino acid turnover (control: 21.01 ± 1.16; SNP: 19.16 ± 1.06), but resulted in a significant decrease in retention into the negative range (control: 0.34 ± 1.27; SNP: −3.87 ± 1.22; P < 0.01), indicating that embryos cultured with NO donor had a net loss of measured

Fig. 3. Amino acid consumption/production in day 8 bovine blastocysts in the presence or absence of 10 ␮M SNP (*** P < 0.001; ** P < 0.01; * P < 0.05).

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Fig. 4. Qualitative amino acid profiles of day 8 bovine blastocysts in the presence or absence of 10 ␮M SNP (* P < 0.05).

endogenous amino acids into the culture medium. In contrast to quantitative profiles, only asparagine (whose consumption increased) and lysine (whose production decreased) qualitative profiles were significantly affected by SNP (P < 0.05) (Fig. 4). As a proportion of turnover, blastocyst amino acid production was significantly decreased and consumption increased by incubation with 10 ␮M SNP (Fig. 5). Qualitative amino acid retention, by contrast, remained unaffected.

5. Discussion Although mammalian embryos would not normally be expected to produce cytotoxic levels of NO, they may remain vulnerable to that derived directly/indirectly from the maternal tract (Lim and Hansel, 1998; Athanassakis et al., 2000). Moreover, embryonic NOS regulatory mechanisms likely differ from those of somatic cells. Thus, in early embryos, arginine is unlikely to participate in NG -hydroxyarginine synthesis to inhibit arginase (thus favouring iNOS activity) (Wu and Morris, 1998), since cattle embryos do not produce urea (Orsi and Leese, 2004a). A 10–20 ␮M SNP concentration decreased blastocyst rate and cell number, in agreement with Lim and Hansel (1998), suggesting that bovine embryos may be more sensitive than human embryos and gametes, which tolerate concentrations up to 100 ␮M (Joo et al., 1999). In the mouse, NO embryotoxicity varies with developmental stage and concentration: while embryo development is inhibited by 10 ␮M–1 mM SNP over the morula to hatching blastocyst transition, 100 nM SNP promotes trophoblast outgrowth. This phenomenon may be due to stage-specific alterations in NO/NOS system regulation (Sengoku et al., 2001).

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Fig. 5. Qualitative amino acid production and consumption (as a % turnover) of day 8 bovine blastocysts in the presence or absence of 10 ␮M SNP (* P < 0.05).

Thus, the metabolic assessment of blastocysts in the present study likely exposed embryos during their most sensitive period. The present data indicate that supplementation of the culture medium with SNP is not beneficial; this suggests that increasing NO profile does not improve bovine embryo development in vitro. The deleterious effects of SNP were particularly evident with respect to carbohydrate metabolism. The decrease in pyruvate and glucose consumption observed at 10 ␮M SNP may in part have resulted from a direct inhibition of cytochrome c oxidase (and consequently, oxidative phosphorylation) and of GAPDH (thus, glycolysis). Additional peroxynitrite formation may have compounded these effects by causing irreversible oxidative damage to the mitochondrial respiratory chain and other proteins. Mitochondrial injury during preimplantation development is likely to have a major impact on viability since there are no corrective/compensatory mechanisms until the late blastocyst/egg cylinder stage, when replication of existing mitochondria takes place (Cummins, 1998). This may explain the failure of these embryos to compensate for an inhibition of respiration by increasing their glycolytic flux. Thus, high levels of NO putatively derived from SNP likely compromised embryo viability through ATP starvation. This is supported by developmental data at 20–50 ␮M SNP: although many zygotes developed successfully to the morula stage, many (all at 50 ␮M SNP) subsequently arrested (data not shown). Such embryos may have generated sufficient ATP to support pre-cavitation development only; formation of the blastocoel cavity makes major demands on ATP provision in order to support Na+ /K+ ATPase activity (Donnay and Leese, 1999). Post-compaction bovine embryos principally convert exogenous glucose to lactate (Thompson et al., 1996). There was no difference in glycolytic index associated with exposure to 10 ␮M SNP, suggesting that the decrease in glucose consumption was relatively

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well matched by a fall in lactate production—this is consistent with GAPDH inhibition. Similarly, the proportion of glucose oxidised in the tricarboxylic acid (TCA) cycle will also have decreased for the reasons outlined earlier. Although the amount of glucose entering the pentose phosphate pathway may have remained unchanged, its impact on overall consumption will likely have been minimal (O’Fallon and Wright, 1986; Urner and Sakkas, 1999). The reasons behind the alterations in quantitative amino acid profiles are unclear, particularly for glycine, tyrosine, threonine, tryptophan and phenylalanine. Other profiles may be interpreted in terms of energy provision: the increase in methionine and valine in the presence of SNP might be explained by their entry into the TCA cycle at the level of succinyl CoA; the subsequent thioester bond cleavage catalysed by succinyl CoA synthetase is coupled to guanosine diphosphate phosphorylation to guanosine triphosphate. Its ␥phosphate group can be transferred to ADP to form ATP (via nucleoside diphosphokinase) (Berg et al., 2002), allowing ATP production independent of the electron transport chain. SNP had no effect on quantitative amino acid production and consumption, suggesting that these embryos struggled to compensate for a reduction in glucose and pyruvate consumption by increasing amino acid oxidation. In this respect, reliance on alternative substrates (e.g. amino acids, triacylglycerols, ketone bodies) still requires oxidative phosphorylation. Qualitatively, only asparagine and lysine profiles were affected by SNP. The notion that this reflects a depression in amino acid metabolism or a loss of intracellular amino acid pools is not supported by the similar turnover and overall production profiles in both groups. The negative amino acid retention of embryos cultured with NO donor could instead point to alterations in albumin endocytosis/hydrolysis and protein biosynthesis (Orsi and Leese, 2004b). This is supported by the effects of NO on qualitative lysine profile, since embryos cannot produce this amino acid from its ␣-ketoacid (Salway, 1994; Orsi and Leese, 2004b). By contrast, the qualitative differences in asparagine profile may have been an ATP-sparing strategy to minimise de novo asparagine synthesis from aspartate via asparagine synthetase (Salway, 1994). Finally, proportional production and consumption profiles were significantly decreased and increased by 10 ␮M SNP, respectively, possibly as a result of lower amino acid catabolism and/or alterations in protein synthesis/hydrolysis.

6. Conclusion In conclusion, these findings indicate that SNP can be embryotoxic, likely by inducing high NO profiles that depress energy generating pathways and perturb amino acid/protein metabolism.

Acknowledgements The author wishes to thank the European Commission for funding, the staff at ABP-York for the supply of bovine ovaries, Mr Peter Humpherson for HPLC advice and Professor Henry Leese for critical review of the manuscript.

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References Athanassakis, I., Aifantis, I., Baritakis, S., Farmakiotis, V., Koumantakis, E., Vassiliadis, S., 2000. Nitric oxide production by pre-implantation embryos in response to embryotoxic factors. Cell Phys. Biochem. 10, 169–176. Athanassakis, I., Aifantis, I., Ranella, A., Giouremou, K., Vassiliadis, S., 1999. Inhibition of nitric oxide production rescues LPS-induced fetal abortion in mice. Nitric Oxide 3, 216–224. Berg, J.M., Tymoczko, J.L., Stryer, L., 2002. Biochemistry, 5th ed. W.H. Freeman and Company, New York, USA. Blackmore, R.S., Greenwood, C., Gibson, Q.H., 1991. Studies of the primary oxygen intermediate in the reactions of fullt reduced cytochrome c oxidase. J. Biol. Chem. 266, 19245–19249. Bolanos, J.P., Peuchen, S., Heales, S.J., Land, J.M., Clark, J.B., 1994. Nitric oxide-mediated inhibition of the mitochondrial respiratory chain in cultured astrocytes. J. Neurochem. 63, 910–916. Brown, G.C., 1999. Nitric oxide and mitochondrial respiration. Biochim. Biophys. Acta 1411, 351–369. Brown, G.C., 2000. Nitric oxide as a competitive inhibitor of oxygen consumption in the mitochondrial respiratory chain. Acta Physiol. Scand. 168, 667–674. Brown, G.C., Cooper, C.E., 1994. Nanomolar concentrations of NO can reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. Fed. Eur. Biochem. Soc. Lett. 356, 295–298. Brune, B., Mohr, S., 2001. Protein thiol modification of glyceraldehyde-3-phosphate dehydrogenase and caspase-3 by nitric oxide. Curr. Protein Pept. Sci. 2, 61–72. Bustamante, J., Bersier, G., Romero, M., Badin, R.A., Boveris, A., 2000. Nitric oxide production and mitochondrial dysfunction during rat thymocyte apoptosis. Arch. Biochem. Biophys. 2, 239–247. Calmels, S., Hainaut, P., Ohshima, H., 1997. Nitric oxide induces conformational and functional modifications of wild-type p53 tumor suppressor protein. Cancer Res. 57, 3365–3369. Clementi, E., Brown, G.C., Foxwell, N., Moncada, S., 1999. On the mechanism by which vascular endothelial cells regulate their oxygen consumption. Proc. Natl. Acad. Sci. U.S.A. 96, 1559–1562. Cummins, J., 1998. Mitochondrial DNA in mammalian reproduction. Rev. Reprod. 3, 172–182. Darley-Usmar, V., Halliwell, B., 1996. Blood radicals—reactive nitrogen species, reactive oxygen species, transition metal ions, and the vascular system. Pharm. Res. 13, 649–662. Darley-Usmar, V., Wiseman, H., Halliwell, B., 1995. Nitric oxide and oxygen radicals: a question of balance. Fed. Eur. Biochem. Soc. Lett. 369, 131–135. Dey, S.K., Lim, H., Das, S.K., Reese, J., Paria, B.C., Daikoku, T., Wang, H., 2004. Molecular cues to implantation. Endocr. Rev. 25, 341–373. Dimmeler, S., Haendler, J., Nehls, M., Ziher, A.M., 1997. Suppression of apoptosis by nitric oxide via inhibition of interleukin-1beta-converting enzyme (ICE)-like and cysteine protease protein (CPP)-32-like proteases. J. Exp. Med. 185, 601–607. Dixit, V.D., Parvizi, N., 2001. Nitric oxide and the control of reproduction. Anim. Reprod. Sci. 65, 1–16. Donnay, I., Leese, H.J., 1999. Embryo metabolism during the expansion of the bovine blastocyst. Mol. Reprod. Dev. 53, 171–178. Foresti, R., Clark, J.E., Green, C.J., Motterlini, R., 1997. Thiol compounds interact with nitric oxide in regulating heme oxygenase-1 induction in endothelial cells. J. Biol. Chem. 272, 18411–18417. Fukuda, A., Hubbard, T.E., Breuel, K.F., 1996. Production of nitric oxide from mouse embryos and effect of nitrite on mouse embryonic development in vitro. Biol. Reprod. 54, 173 (Abstract). Genaro, A.M., Hortelano, S., Alvarez, A., Martinez, C., Bosca, L., 1995. Splenic B lymphocyte programmed cell death is prevented by nitric oxide release through mechanisms involving sustained Bcl-2 levels. J. Clin. Invest. 95, 1884–1890. Gouge, R.C., Marshburn, P., Gordon, B.E., Nunley, W., Huet-Hudson, Y.M., 1998. Nitric oxide as a regulator of embryonic development. Biol. Reprod. 58, 875–879. Haendeler, J., Zeiher, A.M., Dimmeler, S., 1999. Nitric oxide and apoptosis. Vitam. Horm. 57, 49–77. Hansen, P.J., Soto, P., Natzke, R.P., 2004. Mastitis and fertility in cattle—possible involvement of inflammation or immune activation in embryonic mortality. Am. J. Reprod. Immunol. 51, 294–301. Hellstrom, W.J., Bell, M., Wang, R., Sikka, S.C., 1994. Effect of sodium nitroprusside on sperm motility, viability and lipid peroxidation. Fertil. Steril. 61, 1117–1122. Henry, Y., Lepoivre, M., Drapier, J.C., Ducrocq, C., Boucher, J.L., Guissani, A., 1993. EPR characterisation of molecular targets for NO in mammalian cells and organelles. FASEB J. 7, 1124–1134.

N.M. Orsi / Animal Reproduction Science 91 (2006) 225–236

235

Hortelano, S., Alvarez, A.M., Bosca, L., 1999. Nitric oxide induces tyrosine nitration and release of cytochrome c preceding an increase in mitochondrial transmembrane potential in macrophages. FASEB J. 13, 2311– 2317. Hurst, R.D., Dobbie, M.S., 2001. NO and blood–brain barrier permeability. Biochemist 23, 15–19. Inoue, M., Nishikawa, M., Kasahara, E., Sato, E., 1999. Role of superoxide, NO and oxygen in the regulation of energy metabolism and suppression of senile diseases. Mech. Ageing Dev. 111, 89–95. Joo, B.S., Park, S.H., Park, S.J., Kang, H.S., Moon, H.S., Kim, H.D., 1999. The effect of nitric oxide on sperm cell function and embryo development. Am. J. Reprod. Immunol. 42, 327–334. Keller, J.N., Kindy, M.S., Holtsberg, F.W., St Clair, D.K., Yen, H.-C., Germeyer, A., Steiner, S.M., Bruce-Keller, A.J., Hutchins, J.B., Mattson, M.P., 1998. Mitochondrial manganese superoxide dismutase prevents neural apoptosis and reduces ischemic brain injury: suppression of peroxynitrite production, lipid peroxidation, and mitochondrial dysfunction. J. Neurosci. 18, 687–697. Kosenko, E., Kaminsky, Y., Lopata, O., Muravyov, N., Kaminsky, A., Hermenegildo, A., Felipo, V., 1998. Nitroarginine, an inhibitor of nitric oxide synthase, prevents changes in superoxide radical and antioxidant enzymes induced by ammonia intoxication. Metab. Brain Dis. 13, 29–41. Laird, S.M., Tuckerman, E.M., Cork, B.A., Linjawi, S., Blakemore, A.I., Li, T.C., 2003. A review of immune cells and molecules in women with recurrent miscarriage. Hum. Reprod. Update 9, 163–174. Leist, M., Single, B., Naumann, H., Fava, E., Simon, B., Kuhnle, S., Nicotera, P., 1999. Nitric oxide inhibits execution of apoptosis at two distinct ATP-dependent steps upstream and downstream of mitochondrial cytochrome c release. Biochem. Biophys. Res. Comm. 29, 215–221. Lepoivre, M., Flaman, J.M., Bobe, P., Lemaire, G., Henry, Y., 1994. Quenching of the tyrosyl free radical of ribonucleotide reductase by nitric oxide. Relationship to cytostasis induced in tumor cells by cytotoxic macrophages. J. Biol. Chem. 269, 21891–21897. Li, J., Bombeck, C.A., Yang, S., Kim, Y.M., Billiar, T.R., 1999. Nitric oxide suppresses apoptosis via interrupting caspase activation and mitochondrial dysfunction in cultured hepatocytes. J. Biol. Chem. 274, 17325–17333. Lim, J.M., Hansel, W., 1998. Improved development of in vitro-derived bovine embryos by use of a nitric oxide scavenger in a cumulus-granulosa cell co-culture system. Mol. Reprod. Dev. 50, 45–53. Lopez-Figueroa, M.O., Caamano, C., Morano, M.I., Ronn, L.C., Akil, H., Watson, S.J., 2000. Direct evidence of nitric oxide presence within mitochondria. Biochem. Biophys. Res. Comm. 272, 129–133. Manser, R.C., Leese, H.J., Houghton, F.D., 2004. Effect of inhibiting nitric oxide production on mouse preimplantation embryo development and metabolism. Biol. Reprod. 71, 528–533. McBride, A.G., Borutaite, V., Brown, G.C., 1999. Superoxide dismutase and hydrogen peroxide cause rapid nitric oxide breakdown, peroxynitrite production and subsequent cell death. Biochim. Biophys. Acta 30, 275–288. Messmer, U.K., Reed, U.K., Brune, B., 1996. Bcl-2 protects macrophages from nitric oxide-induced apoptosis. J. Biol. Chem. 271, 20192–20197. Nguyen, T., Brunson, D., Crespi, C.L., Penman, B.W., Wishnok, J.S., Tannenbaum, S.R., 1992. DNA damage and mutation in human cells exposed to nitric oxide in vitro. Proc. Natl. Acad. Sci. U.S.A. 89, 3030–3034. O’Fallon, J.V., Wright Jr., R.W., 1986. Quantitative determination of the pentose phosphate pathway in preimplantation mouse embryos. Biol. Reprod. 343, 58–64. Orsi, N.M., Leese, H.J., 2001. Protection against reactive oxygen species during mouse preimplantation development: role of EDTA, oxygen tension, catalase, superoxide dismutase and pyruvate. Mol. Reprod. Dev. 59, 44–53. Orsi, N.M., Leese, H.J., 2004a. Ammonium exposure and pyruvate affect the amino acid metabolism of bovine blastocysts in vitro. Reproduction 127, 131–140. Orsi, N.M., Leese, H.J., 2004b. Amino acid metabolism of preimplantation bovine embryos cultured with bovine serum albumin or polyvinyl alcohol. Theriogenology 61, 561–572. Poderoso, J.J., Lisdero, C., Schopfer, F., Riobo, N., Carreras, M.C., Cadenas, E., Boveris, A., 1999. The regulation of mitochondrial uptake by redox reactions involving nitric oxide and ubiquinol. J. Biol. Chem. 274, 37709–37716. Purcell, T.L., Given, R., Chwalisz, K., Garfield, R.E., 1999. Nitric oxide synthase distribution during implantation in the mouse. Mol. Hum. Reprod. 5, 467–475. Rao, D.N.R., Cederbaum, A.I., 1995. Production of nitric oxide and other iron-containing metabolites during the reductive metabolism of nitroprusside by microsomes and by thiols. Arch. Biochem. Biophys. 2, 363–371.

236

N.M. Orsi / Animal Reproduction Science 91 (2006) 225–236

Rossig, L., Fichtlscherer, B., Breitschopf, K., Haendeler, J., Zeiher, A.M., Mulsch, A., Dimmeler, S., 1999. Nitric oxide inhibits caspase-3 by S-nitrosation in vivo. J. Biol. Chem. 274, 6823–6826. Salway, J.G., 1994. Metabolism at a Glance. Blackwell Science Ltd., Oxford, UK. Sengoku, K., Takuma, N., Horikawa, M., Tsuchiya, K., Komori, H., Sharifa, D., Tamate, K., Ishikawa, M., 2001. Requirement of nitric oxide for murine oocyte maturation, embryo development, and trophoblast outgrowth in vitro. Mol. Reprod. Dev. 58, 262–268. Strandell, A., Thorburn, J., Wallin, A., 2004. The presence of cytokines and growth factors in hydrosalpingeal fluid. J. Assist. Reprod. Genet. 21, 241–247. Szab´o, C., Bryk, R., Zingarelli, B., Southan, G.J., Gahman, T.C., Bhat, V., Salzman, A.L., Wolff, D.J., 1996. Pharmacological characterization of guanidinoethyldisulphide (GED), a novel inhibitor of nitric oxide synthase with selectivity towards the inducible isoform. Br. J. Pharmacol. 118, 1659–1668. Takehara, Y., Kanno, T., Yoshioka, T., Inoue, M., Utsumi, K., 1995. Oxygen-dependent regulation of mitochondrial energy metabolism by nitric oxide. Arch. Biochem. Biophys. 323, 27–32. Tervit, H.R., Whittingham, D.G., Rowson, L.E.A., 1972. Successful culture in vitro of sheep and cattle ova. J. Reprod. Fertil. 30, 493–497. Thompson, J.G., Partridge, R.J., Houghton, F.D., Cox, C.I., Leese, H.J., 1996. Oxygen uptake and carbohydrate metabolism by in vitro derived bovine embryos. J. Reprod. Fertil. 106, 299–306. Tjoa, M.L., Oudejans, C.B., van Vugt, J.M., Blankenstein, M.A., van Wijk, I.J., 2004. Markers for presymptomatic prediction of preeclampsia and intrauterine growth restriction. Hypertens Pregnancy 23, 171–189. Tjuvajev, J., Kolesnikov, Y., Joshi, R., Sherinski, J., Koutcher, L., Zhou, Y., Matei, C., Koutcher, J., Kreek, M.J., Blasberg, R., 1998. Anti-neoplastic properties of human corticotropin releasing factor: involvement of the nitric oxide pathway. In Vivo 12, 1–10. Urner, F., Sakkas, D., 1999. Characterization of glycolysis and pentose phosphate pathway activity during sperm entry into the mouse oocyte. Biol. Reprod. 60, 973–978. Virag, L., Szabo, E., Gergely, P., Szabo, C., 2003. Peroxynitrite-induced cytotoxicity: mechanism and opportunities for intervention. Toxicol. Lett. 140–141, 113–124. Welter, R., Yu, L., Yu, C.-A., 1996. The effects of NO on electron transport complexes. Arch. Biophys. Biochem. 331, 9–14. Wu, G., Haynes, T.E., Li, H., Yan, W., Meininger, C.J., 2001. Glutamine metabolism to glucosamine is necessary for glutamine inhibition of endothelial nitric oxide synthesis. Biochem. J. 353, 245–252. Wu, G., Morris, S.M., 1998. Arginine metabolism: nitric oxide and beyond. Biochem. J. 336, 1–17.