Elder K. Human preimplantation embryo selection

Подождите немного. Документ загружается.

HUMAN PREIMPLANTATION EMBRYO SELECTION

double knockout animals were lost, and in the

absence of these data we cannot conclusively state

that inhibiting NO production results in arrested

embryo development at preimplantation stages. All

of the evidence suggests that NO clearly has a role in

embryonic development. Since NO signaling may

occur through multiple mechanisms, a great deal

remains to be investigated in order to understand

the pathways that may be involved in NO regulation

of the embryonic cell cycle, and whether other mole-

cules can subsume the regulation by NO. Hopefully,

a more complete picture should emerge with the

elucidation of pathways used by NO and with addi-

tional work on knockout animals to determine the

timing of embryos loss. In addition, information

regarding downstream pathways could potentially

be used to identify potential targets for novel IVF

selection technologies. Such information could also

allow for improvements in our ability to produce

culture media that more adequately supports opti-

mal levels of NO production and thus enhances

the activation of downstream pathways to improve

cleavage rates and quality of human IVF embryos.

ACKNOWLEDGMENTS

Much of the unpublished data listed in this chapter

is derived from work done by Dr Nury Steuerwald,

Simone Hendrickson, and Jennifer Seegers, and I

should like to express my gratitude to all members

of the laboratory past and present for their contri-

butions to it. Special thanks to Dr Laura W Schrum

for her review of the manuscript. The work described

has been supported by grants from the Charlotte

Mecklenburg Hospital Authority, The University of

North Carolina at Charlotte, and NIH.

REFERENCES

1. Thaler CD, Epel D. Nitric oxide in oocyte maturation, ovulation,

fertilization, cleavage and implantation: a little dab’ll do ya. Curr

Pharm Des 2003; 9(5): 399–409.

2 Gouge RC, Marshburn P, Gordon BE, Nunley W, Huet-Hudson YM.

Nitric oxide as a regulator of embryonic development. Biol Reprod

1998; 58(4): 875–9.

3 Manser RC, Houghton FD. Ca2-linked upregulation and mito-

chondrial production of nitric oxide in the mouse preimplantation

embryo. J Cell Sci 2006; 119(Pt 10): 2048–55.

4. Moncada S, Higgs EA. Endogenous nitric oxide: physiology, pathology

and clinical relevance. Eur J Clin Invest 1991; 21(4): 361–74.

5. Garthwaite J, Boulton CL. Nitric oxide signaling in the central nervous

system. Annu Rev Physiol 1995; 57: 683–706.

6. Griffith OW, Stuehr DJ. Nitric oxide synthases: properties and

catalytic mechanism. Annu Rev Physiol 1995; 57: 707–36.

7. Rosselli M, Keller PJ, Dubey RK. Role of nitric oxide in the biology,

physiology and pathophysiology of reproduction. Hum Reprod

Update 1998; 4(1): 3–24.

8. Forstermann U, Trogisch G, Busse R. Species-dependent differences in

the nature of endothelium-derived vascular relaxing factor. Eur J

Pharmacol 1984; 106(3): 639–43.

9. Denninger JW, Marletta MA. Guanylate cyclase and the NO/cGMP

signaling pathway. Biochim Biophys Acta 1999; 1411(2-3): 334–50.

10. Nathan C, Xie QW. Regulation of biosynthesis of nitric oxide. J Biol

Chem 1994; 269(19): 13725–8.

11. Wetzka B, Schafer WR, Stehmans A, Zahradnik HP. Effects of nitric

oxide donors on the contractility and prostaglandin synthesis of

myometrial strips from pregnant and non-pregnant women. Gynecol

Endocrinol 2001; 15(1): 34–42.

12. Alderton WK, Cooper CE, Knowles RG. Nitric oxide synthases:

structure, function and inhibition. Biochem J 2001; 357(Pt 3):

593–615.

13. Corson MA, James NL, Latta SE et al. Phosphorylation of endothelial

nitric oxide synthase in response to fluid shear stress. Circ Res 1996;

79(5): 984–91.

14. Fisslthaler B, Dimmeler S, Hermann C, Busse R, Fleming I.

Phosphorylation and activation of the endothelial nitric oxide syn-

thase by fluid shear stress. Acta Physiol Scand 2000; 168(1): 81–8.

15. Hayashi Y, Nishio M, Naito Y et al. Regulation of neuronal nitric-oxide

synthase by calmodulin kinases. J Biol Chem 1999; 274(29): 20597–602.

16. Komeima K, Hayashi Y, Naito Y, Watanabe Y. Inhibition of neuronal

nitric-oxide synthase by calcium/calmodulin-dependent protein

kinase IIalpha through Ser847 phosphorylation in NG108-15 neu-

ronal cells. J Biol Chem 2000; 275(36): 28139–43.

17. Nakane M, Mitchell J, Forstermann U, Murad F. Phosphorylation by

calcium calmodulin-dependent protein kinase II and protein kinase C

modulates the activity of nitric oxide synthase. Biochem Biophys Res

Commun 1991; 180(3): 1396–402.

18. Zembowicz A, Tang JL, Wu KK. Transcriptional induction of endothe-

lial nitric oxide synthase type III by lysophosphatidylcholine. J Biol

Chem 1995; 270(28): 17006–10.

19. Hirata K, Miki N, Kuroda Y et al. Low concentration of oxidized low-

density lipoprotein and lysophosphatidylcholine upregulate constitu-

tive nitric oxide synthase mRNA expression in bovine aortic

endothelial cells. Circ Res 1995; 76(6): 958–62.

20. Freeman JE, Kuo WY, Drenger B et al. Analysis of lysophophatidyl-

choline-induced endothelial dysfunction. J Cardiovasc Pharmacol

1996; 28(3): 345–52.

21. Zhang J, Patel JM, Li YD, Block ER. Proinflammatory cytokines down-

regulate gene expression and activity of constitutive nitric oxide syn-

thase in porcine pulmonary artery endothelial cells. Res Commun Mol

Pathol Pharmacol 1997; 96(1): 71–87.

22. Arnal JF, Yamin J, Dockery S, Harrison DG. Regulation of endothelial

nitric oxide synthase mRNA, protein, and activity during cell growth.

Am J Physiol 1994; 267(5 Pt 1): C1381–8.

23. Braude P, Bolton V, Moore S. Human gene expression first occurs

between the four- and eight-cell stages of preimplantation develop-

ment. Nature 1988; 332(6163): 459–61.

24. Delouis C, Bonnerot C, Vernet M, Nicolas JF. Expression of microin-

jected DNA and RNA in early rabbit embryos: changes in permissiveness



HPE_Chapter14.qxp 7/13/2007 4:52 PM Page 176

NITRIC OXIDE REGULATION OF THE PREIMPLANTATION EMBRYO

for expression and transcriptional selectivity. Exp Cell Res 1992;

201(2): 284–91.

25. Flach G, Johnson MH, Braude PR, Taylor RA, Bolton VN. The transi-

tion from maternal to embryonic control in the 2-cell mouse embryo.

EMBO J 1982; 1(6): 681–6.

26. Jarrell VL, Day BN, Prather RS. The transition from maternal to

zygotic control of development occurs during the 4-cell stage in the

domestic pig, Sus scrofa: quantitative and qualitative aspects of

protein synthesis. Biol Reprod 1991; 44(1): 62–8.

27. Memili E,Dominko T, First NL. Onset of transcription in bovine oocytes

and preimplantation embryos. Mol Reprod Dev 1998; 51(1): 36–41.

28. Zernicka-Goetz M. Activation of embryonic genes during preimplan-

tation rat development. Mol Reprod Dev 1994; 38(1): 30–5.

29. Pines J. Cyclins and cyclin-dependent kinases: a biochemical view.

Biochem J 1995; 308 (Pt 3): 697–711.

30. Morgan DO. Principles of CDK regulation. Nature 1995; 374(6518):

131–4.

31. Smits VA, Medema RH. Checking out the G(2)/M transition. Biochim

Biophys Acta 2001; 1519(1-2): 1–12.

32. Ma T, Van Tine BA, Wei Y et al. Cell cycle-regulated phosphorylation

of p220(NPAT) by cyclin E/Cdk2 in Cajal bodies promotes histone

gene transcription. Genes Dev 2000; 14(18): 2298–313.

33. Guo K, Andres V, Walsh K. Nitric oxide-induced downregulation of

Cdk2 activity and cyclin A gene transcription in vascular smooth

muscle cells. Circulation 1998; 97(20): 2066–72.

34. Nigg EA. Cyclin-dependent protein kinases: key regulators of the

eukaryotic cell cycle. Bioessays 1995; 17(6): 471–80.

35. Hartwell LH, Weinert TA. Checkpoints: controls that ensure the order

of cell cycle events. Science 1989; 246(4930): 629–34.

36. Hogan B, Constantini, F, Lacy E. Manipulating the Mouse Embryo: A

Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Laboratory,

1986.

37. Brinster RL. Nutrition and metabolism of the ovum, zygote, and

blastocyst. Washington, DC: American Physiological Society, 1973.

38. Whitten WK. Nutrition Requirements for the Culture of Preimp-

lantation Embryos in vitro. Adv Biosci 1971; 6: 129-139.

39. Novaro V, Rettori V, Gonzalez ET et al. Interaction between uterine

PGE and PGF2 alpha production and the nitridergic system during

embryonic implantation in the rat. Prostaglandins 1996; 51(6): 363–76.

40. Sengoku K, Takuma N, Horikawa M et al. Requirement of nitric oxide

for murine oocyte maturation, embryo development, and trophoblast

outgrowth in vitro. Mol Reprod Dev 2001; 58(3): 262–8.

41. Chen HW, Jiang WS, Tzeng CR. Nitric oxide as a regulator in pre-

implantation embryo development and apoptosis. Fertil Steril 2001;

75(6): 1163–71.

42. Nishikimi A, Matsukawa T, Hoshino K et al. Localization of nitric

oxide synthase activity in unfertilized oocytes and fertilized embryos

during preimplantation development in mice. Reproduction 2001;

122(6): 957–63.

43. Orsi NM. Embryotoxicity of the nitric oxide donor sodium nitroprus-

side in preimplantation bovine embryos in vitro. Anim Reprod Sci

2006; 91(3–4): 225–36.

44. Tranguch S, Steuerwald N, Huet-Hudson YM. Nitric oxide synthase

production and nitric oxide regulation of preimplantation embryo

development. Biol Reprod 2003; 68(5): 1538–44.

45. Chung HT, Pae HO, Choi BM, Billiar TR, Kim YM. Nitric oxide as a

bioregulator of apoptosis. Biochem Biophys Res Commun 2001;

282(5): 1075–9.

46. Nguyen T, Brunson D, Crespi CL et al. DNA damage and mutation in

human cells exposed to nitric oxide in vitro. Proc Natl Acad Sci USA

1992; 89(7): 3030–4.

47. Dinerman JL, Lowenstein CJ, Snyder SH. Molecular mechanisms of

nitric oxide regulation. Potential relevance to cardiovascular disease.

Circ Res 1993; 73(2): 217–22.

48. Murad F. Regulation of cytosolic guanylyl cyclase by nitric oxide: the

NO-cyclic GMP signal transduction system. Adv Pharmacol 1994; 26:

19–33.

49. Nunokawa Y, Tanaka S. Interferon-gamma inhibits proliferation of rat

vascular smooth muscle cells by nitric oxide generation. Biochem

Biophys Res Commun 1992; 188(1): 409–15.

50. Kawahara K, Gotoh T, Oyadomari S et al. Nitric oxide inhibits the pro-

liferation of murine microglial MG5 cells by a mechanism involving

p21 but independent of p53 and cyclic guanosine monophosphate.

Neurosci Lett 2001; 310(2-3): 89–92.

51. Denham S, Rowland IJ. Inhibition of the reactive proliferation

of lymphocytes by activated macrophages: the role of nitric oxide.

Clin Exp Immunol 1992; 87(1): 157–62.

52. Nisoli E, Clementi E, Tonello C et al. Effects of nitric oxide on prolif-

eration and differentiation of rat brown adipocytes in primary

cultures. Br J Pharmacol 1998; 125(4): 888–94.

53. Buga GM, Wei LH, Bauer PM, Fukuto JM, Ignarro LJ. NG-hydroxy-L-

arginine and nitric oxide inhibit Caco-2 tumor cell proliferation by

distinct mechanisms. Am J Physiol 1998; 275(4 Pt 2): R1256–64.

54. Pervin S, Singh R, Chaudhuri G. Nitric oxide-induced cytostasis and

cell cycle arrest of a human breast cancer cell line (MDA-MB-231):

potential role of cyclin D1. Proc Natl Acad Sci USA 2001; 98(6):

3583–8.

55. Hajri A, Metzger E, Vallat F et al. Role of nitric oxide in pancreatic

tumour growth: in vivo and in vitro studies. Br J Cancer 1998; 78(7):

841–9.

56. Ulibarri JA, Mozdziak PE, Schultz E, Cook C, Best TM. Nitric oxide

donors, sodium nitroprusside and S-nitroso-N-acetylpencillamine,

stimulate myoblast proliferation in vitro. In Vitro Cell Dev Biol Anim

1999; 35(4): 215–8.

57. Du M, Islam MM, Lin L et al. Promotion of proliferation of murine

BALB/C3T3 fibroblasts mediated by nitric oxide at lower concentra-

tions. Biochem Mol Biol Int 1997; 41(3): 625–31.

58. Luczak K, Balcerczyk A, Soszynski M, Bartosz G. Low concentration of

oxidant and nitric oxide donors stimulate proliferation of human

endothelial cells in vitro. Cell Biol Int 2004; 28(6): 483–6.

59. Sauzeau V, Rolli-Derkinderen M, Marionneau C, Loirand G, Pacaud P.

RhoA expression is controlled by nitric oxide through cGMP-dependent

protein kinase activation. J Biol Chem 2003; 278(11): 9472–80.

60. Katsuyama K, Shichiri M, Marumo F, Hirata Y. NO inhibits cytokine-

induced iNOS expression and NF-kappaB activation by interfering

with phosphorylation and degradation of IkappaB-alpha. Arterioscler

Thromb Vasc Biol 1998; 18(11): 1796–802.

61. Fiedler B, Lohmann SM, Smolenski A et al. Inhibition of calcineurin-

NFAT hypertrophy signaling by cGMP-dependent protein kinase

type I in cardiac myocytes. Proc Natl Acad Sci USA 2002; 99(17):

11363–8.

62. Pilz RB, Suhasini M, Idriss S, Meinkoth JL, Boss GR. Nitric oxide and

cGMP analogs activate transcription from AP-1-responsive promoters

in mammalian cells. FASEB J 1995; 9(7): 552–8.

63. Thiriet N, Esteve L, Aunis D, Zwiller J. Immediate early gene induction

by natriuretic peptides in PC12 phaeochromocytoma and C6 glioma

cells. Neuroreport 1997; 8(2): 399–402.

64. Sharma RV, Tan E, Fang S, Gurjar MV, Bhalla RC. NOS gene transfer

inhibits expression of cell cycle regulatory molecules in vascular

smooth muscle cells. Am J Physiol 1999; 276(5 Pt 2): H1450–9.

65. Fukumoto S, Koyama H, Hosoi M et al. Distinct role of cAMP and

cGMP in the cell cycle control of vascular smooth muscle cells: cGMP



HPE_Chapter14.qxp 7/13/2007 4:52 PM Page 177

HUMAN PREIMPLANTATION EMBRYO SELECTION

delays cell cycle transition through suppression of cyclin D1 and

cyclin-dependent kinase 4 activation. Circ Res 1999; 85(11): 985–91.

66. Hanada S, Terada Y, Inoshita S et al. Overexpression of protein kinase

G using adenovirus inhibits cyclin E transcription and mesangial cell

cycle. Am J Physiol Renal Physiol 2001; 280(5): F851–9.

67. Kronemann N, Nockher WA, Busse R, Schini-Kerth VB. Growth-

inhibitory effect of cyclic GMP- and cyclic AMP-dependent vasodila-

tors on rat vascular smooth muscle cells: effect on cell cycle and cyclin

expression. Br J Pharmacol 1999; 126(1): 349–57.

68. Chan GK, Yen TJ. The mitotic checkpoint: a signaling pathway that

allows a single unattached kinetochore to inhibit mitotic exit. Prog

Cell Cycle Res 2003; 5: 431–9.

69. McGowan CH, Russell P. The DNA damage response: sensing and

signaling. Curr Opin Cell Biol 2004; 16(6): 629–33.

70. Sibon OC, Laurencon A, Hawley R, Theurkauf WE. The Drosophila

ATM homologue Mei-41 has an essential checkpoint function at the

midblastula transition. Curr Biol 1999; 9(6): 302–12.

71. Gagioti S, Scavone C, Bevilacqua E. Participation of the mouse

implanting trophoblast in nitric oxide production during pregnancy.

Biol Reprod 2000; 62(2): 260–8.

72. Ariel I, Hochberg A, Shochina M. Endothelial nitric oxide synthase

immunoreactivity in early gestation and in trophoblastic disease.

J Clin Pathol 1998; 51(6): 427–31.

73. Abe KM, Inoue N, Taga M, Kato T. Messenger RNA of neuronal nitric

oxide synthase is expressed and possibly functions in mouse oocytes

and embryos during preimplantation development. Biomed Res

(Tokyo) 1999; 20: 61–65.

74. Son H, Hawkins RD, Martin K et al. Long-term potentiation is

reduced in mice that are doubly mutant in endothelial and neuronal

nitric oxide synthase. Cell 1996; 87(6): 1015–23.

75. Phillips TJ, Hen R, Crabbe JC. Complications associated with genetic

background effects in research using knockout mice. Psychophar-

macology (Berl) 1999; 147(1): 5–7.

76. Huber A, Saur D, Kurjak M, Schusdziarra V, Allescher HD. Charac-

terization and splice variants of neuronal nitric oxide synthase in rat

small intestine. Am J Physiol 1998; 275(5 Pt 1): G1146–56.

77. Rothe F, Huang PL, Wolf G. Ultrastructural localization of neuronal

nitric oxide synthase in the laterodorsal tegmental nucleus of wild-

type and knockout mice. Neuroscience 1999; 94(1): 193–201.

78. Meng W, Ma J, Ayata C et al. ACh dilates pial arterioles in endothelial

and neuronal NOS knockout mice by NO-dependent mechanisms.

Am J Physiol 1996; 271(3 Pt 2): H1145–50.



HPE_Chapter14.qxp 7/13/2007 4:52 PM Page 178

INTRODUCTION

Selection of human embryos for transfer following

in vitro fertilization (IVF) programs is routinely

based on pronuclear or embryo morphology and

cleavage rate.

Despite numerous attempts to develop

embryo scoring techniques that can predict embryo

viability, implantation rates remain low.

Not all

embryos with good morphology will implant, and

fragmented embryos can develop to the blastocyst

stage in vitro.

In fact, embryo selection based on

morphology fundamentally eliminates only severely

impaired embryos. To achieve acceptable pregnancy

rates more than one embryo is routinely trans-

ferred, with an unavoidable risk of multiple gesta-

tions. Micromethods for assessing the metabolism

of embryos were developed in order to improve the

selection of embryos with the best developmental

potential.

Measurements of metabolite uptake and produc-

tion assist in understanding how preimplantation

embryos interact with their in vitro or in vivo condi-

tions, and also give important information about

components that can have beneficial or deleterious

effects on embryo viability. In vitro culture condi-

tions have been shown to affect the embryo during

early and late development. Alterations of in vitro

culture conditions can modify gene expression,

3,4

cellular metabolism,

or imprinting status

during

early development, and may also have profound

effects on prenatal or postnatal development, such as

the large offspring syndrome observed when bovine

embryos are cultured in the presence of serum.

Preimplantation embryo development is a com-

plex process that produces a multicellular organism

from a single cell. Cellular division and differentia-

tion are associated with different metabolic pathways

that mainly focus on energy production and syn-

thesis of metabolic precursors for macromolecules.

Understanding early embryo metabolism is also

essential in order to set up culture media and condi-

tions better adapted to preimplantation development

in vitro, reducing possible adverse effects during

in vitro culture.

Metabolic measurements may also

help in the selection of embryos for transfer in IVF

programs.

This chapter focuses on embryo metabolism that

can be measured by non-invasive techniques. It

describes the main pathways that produce ATP from

glucose or amino acids once they enter the cell, and

presents the different techniques used to evaluate

the uptake or release of metabolites by embryos

in vitro and how this uptake or release can be corre-

lated to embryo viability.

EARLY PREIMPLANTATION EMBRYO

DEVELOPMENT

The preimplantation period begins with oocyte

fertilization and finishes with the formation of a

hatched blastocyst ready to implant. In humans, this

period lasts for approximately 5–7 days. The mature

oocyte contains sufficient maternal transcripts and

proteins to support the process of fertilization and

the first two cell divisions. The activation of the

embryonic genome, which represents the transition

from maternal to embryonic genome control, is

thought to occur during the third cell division. At

compaction, the first epithelium is formed and two

distinct cell lines can be distinguished, one that will

form the trophectoderm (TE) and the other that

will form the fetus. These major events are associ-

ated with a drastic increase in embryo metabolism

15. Uptake and release of metabolites in human

preimplantation embryos

Fabienne Devreker



HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 179

HUMAN PREIMPLANTATION EMBRYO SELECTION

that involves the production of ATP, mRNA, and

proteins.

Several experiments performed on preimplanta-

tion embryos from different species, including

mouse, cow, rat, pig, sheep, and human, showed that

embryo metabolism is different in the pre- and

postcompaction stages, corresponding with before

and after the fourth cell division. Before com-

paction, blastomeres are loosely joined together

and therefore equally exposed to their environment.

Precompaction stage embryos rely on maternal

storage and have a relatively low level of biosynthe-

sis,

low respiratory rates,

8,9

and a limited ability to

metabolize glucose as a source of energy.

On the

other hand, postcompaction stage embryos have

developed important gap-junctional complexes in

the outer embryonic cells

and cytoplasmic and

membrane polarization have been observed. Two

distinct apical and basal membranes, that are kept

separated by zonular tight junctions, are formed.

The formation of tight cellular junctions between

the outer cells creates two different microenviron-

ments for the inner and outer cells. The inner cells

exposed to the inside microenvironment will form

the inner cell mass (ICM), while outer cells will

form the trophectoderm.

Embryos then have a

high rate of biosynthesis and an exponential increase

in their demand for energy. Indeed, blastocyst for-

mation requires new proteins or enzymes for the

formation of cellular junctional complexes, cell

differentiation, and the activation of a Na

⫹

ATPase pump for the formation of the blastocoel

cavity.

In mouse and rabbit embryos, the formation of

the blastocoel is associated with a significant increase

in the synthesis and activity of Na

⫹

ATPase.

Postcompaction stage embryos become able to

metabolize glucose and to actively control their

cytoplasmic pH and composition (Table 15.1).

10,14,15

TECHNIQUES FOR MEASURING EMBRYO

METABOLISM IN VITRO

The metabolism of mammalian embryos from several

species has been extensively studied, including

mouse,

10,16–21

hamster,

cow,

23,24

and rabbit.

25,26

Much less is known about other species such as

sheep,

9,27

rat,

pig,

and human.

2,10,14,18

Carbo-

hydrate metabolism of embryos has been evaluated

by measurements of substrate uptake or release

using either radiolabeled substrates

16–18,29,31

or non-

invasive microfluorescence assays,

1,2,10,14

and by

comparing embryo development in vitro in the

presence or absence of energy substrates in the

culture medium (the ‘nutritional’ approach).

Radioisotopic studies use different markers as

tracers, including [

C]-glucose, [

C]-pyruvate, or

[

C]-lactate and their products, such as

C-lactate, that can be collected or isolated

chromatographically before measuring their

radioactivity.

This technique made it possible to

both quantify substrate utilization and to identify

the metabolic pathways used by the embryos. The

disadvantage of this technique is that measurements

are made for a group of embryos, and it is therefore

not possible to correlate embryo metabolism with

embryo development or viability post-transfer.

Furthermore, radiolabeled substrates can have toxic

effects on the embryo. Radiolabeled experiments

have, for example, demonstrated that CO

produc-

tion is both stage and substrate specific. Incubating

mouse embryos with [

C]-pyruvate or [

C]-lactate

as the sole substrate showed that

production

by unfertilized oocytes, fertilized, and 8-cell embryos

was low, and increased oocytes from the morula to

blastocyst stages with both substrates. However, the



Table 15.1 Metabolic differences between cleavage

stage and postcompaction stage embryos

Cleavage stage Postcompaction stage

Inability to metabolize glucose Switch to glucose metabolism

Pyruvate is essential Pyruvate is less important

Activation of anaerobic glycolysis

Low level of ATP production High level of ATP production

Low level of mRNA production High level of mRNA

Low respiratory rate High respiratory rate

Low protein synthesis Increase in protein synthesis

Beneficial effect of Requirements for the 20 Eagle’s

non-essential amino acids amino acids

High sensitivity to pH variations Less sensitivity to pH variations

HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 180

UPTAKE AND RELEASE OF METABOLITES IN HUMAN PREIMPLANTATION EMBRYOS

production of CO

from lactate by post compaction

embryos was 50% lower compared with the CO

from pyruvate.

The nutritional approach analyzes embryo

development in vitro in the presence or absence of

determined components in the culture medium,

including carbohydrates, amino acids, oxygen, etc.

In order to determine the toxicity or the beneficial

effects of the tested component, several parameters

are compared, such as the percentage of embryos

reaching the blastocyst stage, implantation post-

transfer, or more invasively, the cell numbers in the

TE and ICM

in resultant blastocysts. Once the new

culture conditions have been proven to adequately

support embryo development in vitro, these con-

ditions can be used in clinical programs and the

outcome of the resultant offspring can be assessed.

Non-invasive microfluorescence assays make it

possible to measure substrate depletion or release by

individual embryos in culture media and to corre-

late metabolism with embryo viability.

2,10,14,33

This

method relies on the generation or consumption

of the reduced pyridine nucleotides NADH and

NADPH.

These nucleotides have the property of

absorbing light at 340 nm, whereas the oxidized

forms, NAD

⫹

and NADP

⫹

, emit fluorescence when

activated with light at 340 nm. By choosing appro-

priate enzymatic reactions, the increase or decrease

in the absorbance of fluorescence due to NADH or

NADPH formation or oxidation can be related to

the concentration of a metabolite or to the rate of

an enzymatic reaction.

14,20

The measurements have

been facilitated by adapting these fluorometric assays

to a Cobas-Bio autoanalyzer (Roche Instruments,

UK).

In the human, spare embryos are incubated

singly in 4 or 5 ␮l droplets of culture medium for 24

hours, and 1 or 2 ␮l samples of the medium are then

analyzed for nutritional content.

32,35,36

By choosing

appropriate enzymatic reactions, this method can

be used to measure enzyme activities,

glucose,

pyruvate, lactate, or ammonium.

For amino acids, the spent medium is analyzed

by reverse-phase high performance liquid chro-

matography (HPLC).

Ammonium excretion is

measured in the spent medium by a quantitative

enzymatic reaction using a Cobas-Bio autoanalyzer.

The method is based on the reductive amination of

2-oxoglutarate, using glutamate dehydrogenase and

NADPH.

To measure oxygen uptake, embryos are incu-

bated in groups in 5 ␮l polymerase chain reaction

micropipettes. The reaction is based on the fluores-

cence emitted by pyrene. Pyrene is a highly fluores-

cent non-toxic compound that is excited at 340 nm

and emits light at 450 nm. The oxygen consumption

is assessed by the change in pyrene’s fluorescence,

measured by a fluorescence microscope with photo-

multiplier and photometer attachment.

After the

period of incubation, embryos are replaced in drops

of fresh medium and cultured to later stages. Other

methods are available that are less easy to perform,

based on the conversion of oxyhemoglobin to hemo-

globin or on the use of a microelectrode.

Evalua-

tion of ATP consumption is either measured through

HPLC,

or calculated on the basis that one mole of

oxygen produces 6 moles of ATP, or that one mole

of glucose produces 2 moles of ATP by anaerobic

glycolysis.

CARBOHYDRATE METABOLISM

Embryos use two main pathways to generate the

ATP that is necessary for cellular metabolism: aero-

bic glycolysis or the tricarboxylic acid cycle (TCA or

Krebs cycle) and anaerobic glycolysis by the Embden-

Meyerhof pathway (EMP).

Glucose, pyruvate, and lactate occupy a central

position as substrates for both energy production

and for synthesis of complex molecules. Glucose

enters the cell by either diffusion or facilitated trans-

port (GLUT1).

Once it enters the cell, glucose is

directly phosphorylated into glucose 6-phosphate

(G-6-P). At this stage glucose can enter into four

major pathways (Figure 15.1).

The first is the

EMP, anaerobic glycolysis that converts one mole-

cule of glucose into two molecules of pyruvate (a

3-carbon molecule) with the production of two

molecules of ATP. In the presence of oxygen and the

relative absence of lipids or other substrates, pyru-

vate enters the Krebs cycle for oxidative metabolism

in mitochondria and is converted into acetyl-CoA,



HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 181

HUMAN PREIMPLANTATION EMBRYO SELECTION

yielding 38 molecules of ATP for each molecule of

glucose consumed. In the absence of oxygen, pyru-

vate is converted into lactate (Figure 15.1).

The second pathway is the pentose phosphate

shunt which produces NADPH and ribose 5-phos-

phate (Figure 15.1). The reduced equivalents of

NADP

⫹

are transferred to glutathione for the pro-

tection of the cell against peroxidation, and used in

the synthesis of lipids or other complex molecules.

Ribose 5-phosphate is an obligatory precursor for

the synthesis of all nucleotides.

The third pathway involves the transfer of one

amine group (transamination) from glutamine to

fructose, to form glucosamine 6-phosphate and

glutamate. Glucose is therefore a precursor for

glycoproteins, mucoproteins, and mucopolysaccha-

rides (Figure 15.1).

The fourth pathway is used when the glucose

supply is in excess of requirements, and thus glucose

is stored within the cell in the form of glycogen, a

branched complex polymer of glucose molecules.

In the mouse, pyruvate enters embryos by a

combination of diffusion and facilitated trans-

port.

20,31

Pyruvate then enters into the TCA cycle to

produce ATP. Lactate appears to enter into the

embryo by rapid diffusion.

Lactate has been shown



Hexosamines

Hexose

phosphate

Glyceraldehyde 3-phosphate

Ribose 5-phosphate

NADPH

Pentose

phosphate

Pyruvate Lactate

Triose

phosphate

Glycoproteins

Glycolipids

Glycosaminoglycans

TCA Cycle

NAD

+ H

NADH

FAD

2Fe

NADH

FADH

Flavoprotein Cytochrome

Electron transport chain

Glucose

32 ATP

4 ATP

4 ADP + Pi

2 ADP + Pi

2 ATP

Cytosol

Mitochondria

Acetyl-CoA

Figure 15.1 Diagram illustrating three catabolic pathways of glucose. (1) Through glycolysis, glucose is degraded into pyruvate with

the production of 2 molecules of ATP. (2) Through transamination with amino acids, glucose participates in the synthesis of many

glycoproteins and glycolipids. (3) The pentose-phosphate shunt uses glucose for the synthesis of DNA and RNA molecules. The

storage of glucose under the form of glycogen is not represented in this diagram. Adapted from Rieger et al.

HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 182

to be the preferred substrate for 1-cell hamster

embryos.

In the mouse, lactate can be utilized as

an energy source from the 2-cell stage onwards,

and acts synergistically with pyruvate.

Lactate is,

however, mainly a product of glucose oxidation

(Figure 15.1).

PYRUVATE

Pyruvate is usually preferred to glucose during the

early stages of development, and, using a nutritional

approach, has been shown to be essential for the

early preimplantation embryo development of

mouse

and human embryos.

Pyruvate uptake by

mouse embryos is already present at the 1-cell stage,

increases throughout development and declines

slightly at the blastocyst stage.

14,31,35,36

Pyruvate is a

major energy source during human oocyte matura-

tion (Figure 15.2).

47,48

After the activation of the embryonic genome,

pyruvate uptake by mouse or human embryos

exceeds glucose uptake from cleavage stage up to the

blastocyst stage, to reach a maximum of 40–45

pmol/l per embryo/hour at the morula stage.

2,35,49

glucose is available in the culture medium, pyruvate

uptake declines slightly at the morula stage.

2,10,35

Pyruvate uptake has been correlated to embryo

viability: embryos that developed to the blasto-

cyst stage took up more pyruvate than those that

arrested.

2,35

Consumption of pyruvate was related

to embryo viability in patients with tubal or poly-

cystic ovary (PCO) infertility. Suboptimal ovulation

regimens resulted in embryos with lower pyruvate

uptake and lower blastocyst cell numbers.

Pyruvate

uptake is lower when embryos are cultured in sub-

optimal conditions that compromise their viability

(Figure 15.3).

The uptake of pyruvate during the first 24 h fol-

lowing fertilization in single human embryos that

were derived from natural cycles was correlated to

embryo morphology. Since a single embryo was

transferred, the uptake of pyruvate could be directly

correlated with the embryo. The results showed that

embryos had a wide range of pyruvate uptake values

(2–53 pmol/l per embryo/h), but that this variation

was reduced significantly to an intermediate range

of values in those embryos that were able to implant

(10–30 pmol/l per embryo/h).

Similarly, pyruvate

uptake by human embryos during the second and

third day of development displays a wide range of

values. Embryos that implanted, however, took up

less pyruvate than those that did not (22.9 ⫾ 1.0 and

UPTAKE AND RELEASE OF METABOLITES IN HUMAN PREIMPLANTATION EMBRYOS



Unfertilized

Activated

Fertilized

Oocyte fertilization First cleavage division

Pyruvate uptake

(pmol/l per embryo/h)

Figure 15.2 Pyruvate consumption during oocyte fertilisation

and first cleavage division. First period represents measurements

during 24H from the time of insemination until the assessment

of fertilization status. Second period represents measurements

during 24H from Day 1 until Day 2 post insemination, at the time

of transfer. Pyruvate uptake is measured in pmoles/ embryo/

hour.

45 *

: p ⬍ 0.01. Adapted from Devreker et al.

4–7 cell 8–16 cell M B

Stage

Pyruvate uptake

(pmol/l per embryo/h)

With glutamine

Without glutamine

Figure 15.3 Pyruvate uptake by individual human embryos in

the presence or absence of glutamine. M, morula; B, blastocyst.

Adapted from Devreker et al.

51 *

: p ⬍ 0.05.

HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 183

HUMAN PREIMPLANTATION EMBRYO SELECTION

27.1 ⫾ 0.6 pmol/l per embryo/h, respectively, on

day 2; and 22.4 ⫾ 1.5 and 26.9 ⫾ 0.8 pmol/l per

embryo/h, respectively, on day 3).

However, the

wide range of values precluded the use of pyruvate

uptake to predict embryos that will implant. More

recently, pyruvate uptake by human oocytes can be

detected earlier, during fertilization and the first

divisions.

Fertilization and activation increased

pyruvate uptake by the human embryo by approxi-

mately 30%. However, it was not related to subse-

quent morphology, developmental stage at the time

of transfer, or to implantation (Table 15.2). The low

number of embryos that effectively implanted may

explain the fact that measurements of pyruvate

uptake by human embryos during the first 2 days of

development did not help to predict embryo viability

post-transfer.

Culture of isolated human blastomeres in vitro

showed that those that underwent cell division had

a significantly higher pyruvate uptake than those

that did not cleave (Hardy, personal communica-

tion). Thus, pyruvate uptake appears to be related to

cell division, and in the zygote it is possible that the

cytoskeletal changes associated with expulsion of

the second polar body and movement of pronu-

clei

48,53

require energy, which is provided by pyru-

vate. In another study, although pyruvate uptake by

human embryos cultured in sequential media was

higher for embryos that developed up to the blasto-

cyst stage, this could not be correlated to embryo

morphology.

GLUCOSE

In contrast to other cell lines, cleavage stage

embryos have little capacity to metabolize glucose

prior to the activation of the embryonic genome.

Using radiolabeled [

C]-glucose

or microfluores-

cence assays,

2,35,36,47

several authors have demon-

strated that glucose uptake by early human embryos

is very low before the 4-cell stage, less than 10

pmol/l per embryo/h,

and reached 20–30 pmol/

l per embryo/h at the blastocyst stage.

10,14,30,35,49

similar pattern for glucose uptake has been observed

for other mammalian species, including mouse,

14,31,33

pig,

rabbit,

and bovine.

20,23

For example, the

total glucose metabolism almost doubles between

the morula and expanded blastocyst stages in the

cow

and increases linearly with the volume of

horse blastocysts.

Glucose uptake by mouse

embryos is low before the 8-cell stage and increases

sharply at the transition from morula to blasto-

cyst stage.

14,33

The consumption of glucose was

later shown to help in predicting the resistance

of bovine embryos to the freezing–thawing pro-

cedure.

Glucose uptake by human embryos prior to acti-

vation of the embryonic genome was undetectable

using fluorescence assays.

Gardner et al

reported

that day 4 human embryos that developed up to the

blastocyst stage in sequential media culture took up

significantly more glucose during blastocyst forma-

tion. Glucose uptake could also be correlated to

blastocyst morphology, being higher in those

blastocysts of highest grade. The absence of detec-

tion of glucose uptake by early human embryos

does not necessarily mean that these embryos do

not metabolize small quantity of glucose. Low levels

of glucose in culture media are probably necessary

for embryo development. However, as glucose can

easily enter the cell by diffusion, a high level of glu-

cose in culture media for precompaction embryos

can have deleterious effect on embryo viability.

Indeed, part of the glucose can be stored under

the form of glycogene. Too much intracellular

glycogene can disturb the cytoplasmic architecture

and would be toxic for the cell.



Table 15.2 Mean pyruvate uptake (pmol/embryo/h)

by normally fertilized embryos according to the

stage reached at the time of transfer. Values are

mean ⫾ SEM. From Devreker et al

Stage reached by day 2 n Day 0–1 Day 1–2

One cell stage 13 10.0 ⫾ 2.2 11.8 ⫾ 2.0

2–3-cell stage 76 15.6 ⫾ 1.4 15.0 ⫾ 1.3

4-cell stage 106 13.4 ⫾ 0.9 16.0 ⫾ 1.0

⬎4-cell stage 38 19.7 ⫾ 1.9

a,b

12.5 ⫾ 1.5

Highly fragmented 23 15.1 ⫾ 2.2 13.3 ⫾ 2.6

embryos

Significantly higher compared with the 1-cell stage;

*p ⫽ 0.010 (Mann–

Whitney).

HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 184

UPTAKE AND RELEASE OF METABOLITES IN HUMAN PREIMPLANTATION EMBRYOS

LACTATE

Lactate production by preimplantation embryos is

low before compaction and increases thereafter,

coincident with the rise in glucose consumption.

Lactate production has been observed even in the

absence of exogenous substrate, suggesting that

embryos use some endogenous source of

energy.

2,12,14,33,35

Radiolabeled experiments showed

that embryos produce lactate from glucose or pyru-

vate.

18,48

A high lactate production, however, could

be a sign of impaired embryo viability. Some

authors

have reported that mouse embryos with

lower viability produced a higher amount of lactate

in vitro. Alteration of embryo viability is compro-

mised as early as 6 h of culture in vitro, and is asso-

ciated with a net increase in lactate production.

This increase in lactate production can be sup-

pressed if amino acids and vitamins are added to the

culture medium. Moreover, mouse embryos grown

in vitro produced twice as much lactate as those

grown in vivo.

Measurements of both glucose

consumption and lactate production by mouse

embryos allowed their glycolytic activity to be eval-

uated. It was shown that blastocysts with the lowest

level of anaerobic glycolysis had the highest poten-

tial to develop post-transfer.

Van den Bergh et al

reported a similar observation for human embryos

cultured in sequential media. However, to date,

lactate production has not been used to predict or

evaluate human embryo viability before transfer.

AMINO ACIDS

Amino acids are one of the key constituents of

cellular metabolism, and embryos are equipped

with an array of sodium-dependent and -independ-

ent transporter systems that regulate amino acid

flux and availability.

Amino acids not only form the

basic components of proteins, but also fulfil a num-

ber of other functions. They contribute to ATP

production via deamination reactions, producing

intermediary metabolites that can enter the TCA

cycle, and also stimulate activation of the embryonic

genome, as well as blastocyst formation and hatch-

ing. Amino acids have been shown to act as

osmolytes, and help to maintain intracellular

pH.

6,22,56

Glutamine, taurine, and glycine have been

reported to protect cleavage stage embryos against

osmotic shock.

Glycine maintains embryo develop-

ment in the presence of high osmolarities,

and

amino acids are thought to have a significant role in

signal transduction cascades.

By providing sub-

strates to the TCA cycle, in mouse and hamster

embryos amino acids can protect against the delete-

rious effects of glucose by inhibiting glycolytic

activity.

22,56

The oxidation of amino acids causes

allosteric inhibition of the glycolytic enzyme phospho-

fructokinase through an increase in ATP and/or

citrate, or by the direct inhibition of pyruvate kinase

by alanine.

Amino acids are present in large amounts in

female reproductive tract fluids, zygotes, and embryos,

and are therefore likely to be important for pre-

implantation embryo development.

22,57

For exam-

ple, glycine content is the highest in rabbit

oviductal fluids, along with taurine, and is cycle

dependent.

61,62

Alanine, threonine, glutamate, serine,

taurine, and glycine represent 85% of the amino

acid content of rabbit oviductal fluids.

61,62

In the

mouse, the most abundant amino acids in oviduct

and uterine fluids are glutamine, taurine, glycine,

and alanine, while alanine, aspartate, glutamate, tau-

rine, and glycine are among the most abundant

amino acids in oocytes and embryos.

Human uter-

ine fluids contain large amounts of taurine, gluta-

mate, glycine, and alanine.

These four amino acids

are also predominant in bovine uterine fluids.

Amino acids singly or in combination have been

shown to promote the early development of embryos

from various species, including mouse,

cow,

ham-

ster,

sheep,

rabbit,

rat,

and human.

51,56,68,69

However, they have differential effects on embryo

development; some promote, while others can inhibit

embryo development.

Several authors therefore

measured amino acid depletion or appearance in

the culture medium during embryo development

in vitro. The profile of amino acid uptake or secretion

differs between species, and also at different embryo



HPE_Chapter15.qxp 7/14/2007 5:54 PM Page 185