Elder K. Human preimplantation embryo selection

Подождите немного. Документ загружается.

HUMAN PREIMPLANTATION EMBRYO SELECTION

with respect to the number of fully grown antral fol-

licles.

Failure to retrieve oocytes has often been

attributed to patient error in the timing of ovulation

induction by hCG administration. In these instances,

the self-administration of hCG may be mistimed

and so late that the subsequent oocyte retrieval was

too premature with respect to the preovulatory

development of the follicle. Failures in these retrievals

may be due to oocytes that remain firmly adhered to

the follicular wall during aspiration. In addition to

timing errors, failure to administer hCG, or to inject

it correctly (and entirely), as well as potential differ-

ences in hCG bioactivity between lots, have been

suggested as iatrogenic rather than biological etiolo-

gies of this phenomenon. In these cycles, especially

when the follicular aspirates are devoid of the usual

masses of cumulus granulosa cells, it is prudent to

assay serum ␤hCG level. In practice, when serum

levels are ⬍10 mIU/ml, the procedure is suspended,

a second hCG injection is given, and a second aspi-

ration attempted some 36 hours later. The low cir-

culating levels of hCG are indicative of an iatrogenic

origin of the ‘empty’ follicles and a timely follow-up

ovulation induction can produce retrievable oocytes

that are viable, as evidence by successful outcomes.

However, many cases do not have an iatrogenic

origin, and in these instances, follicles that have

adequate exposure to hCG do not appear to contain

retrievable oocytes. Analysis of steroid levels in

follicular fluid aspirates from ‘empty’ follicles shows

high estradiol : progesterone ratios and high andro-

stenedione levels, which indicates that they are not

simply cysts, or prematurely luteinized, or atretic

follicles.

One biological explanation for their occur-

rence suggests an intrinsic defect in folliculogenesis

that leads to early oocyte atresia.

This notion is

supported by the finding that empty follicles recur

in subsequent IVF cycles for some affected women.

However, case reports of successful outcomes with

a second round of ovulation induction when cir-

culating hCG levels are well above the 10 mIU/ml

threshold noted above have been reported. In these

instances, empty follicles may not reflect oocyte dege-

neration but rather an inability of the oocyte–cumulus

complex to respond to hCG, or possibly, the need

for a longer period to separate the complex from the

follicular wall.

97,98

Although a relatively rare phe-

nomenon, additional research is needed to determine

the exact cause(s) of this syndrome and whether

factors that indicate a normal follicle such as peri-

follicular blood flow values, can predict the occur-

rence of ‘empty’ follicles.

THE INFLUENCE OF MATERNAL AGE AND

LIFESTYLE FACTORS ON FOLLICULAR

BIOLOGY AND OOCYTE COMPETENCE

Competence assessments at the follicular level may

be particularly relevant in the evaluation of women

of advanced reproductive (i.e. biological) age and

those whose life-style factors may negatively con-

tribute to fertility. Maternal age is the single most

important biological determinant of a woman’s fer-

tility potential, and has an enormous impact on the

quantity of follicles and the quality of the oocytes

she ovulates towards the end of her reproductive

life. The total number of oocytes peaks at approxi-

mately 5 million around month 5 of fetal age and

about 2 million remain at birth.

This number is

further reduced during a woman’s reproductive life;

perhaps a thousand or fewer primordial follicles

remain when menstrual cycles cease.

100

The physical

‘expulsion’ of oocytes from the ovarian cortex and

programmed cell death by apoptosis accounts for

oocyte loss during the fetal and neonatal stages (for

review see reference 4), while apoptosis and other

mechanisms of cell death (e.g. atresia) are involved

during reproductive life. In a woman’s normal repro-

ductive lifespan, approximately 400–500 primordial

follicles will progress to ovulation.

Although there are usually sufficient primordial

follicles remaining to allow menstrual cycles to con-

tinue until the sixth decade of life, fertility rates

begin to fall much earlier. As shown in Figure 24.2, a

gradual decline is first observed in the late 20s

102

but the rate accelerates considerably around age

37.

101,103,104

This progressive and natural reduction

in ovarian reserve is one factor contributing to the

age-related decline in fertility, but the quality of the

corresponding oocytes also declines. Between the time

of their formation in the fetus and the few hours



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 316

OOCYTE SELECTION IN CONTEMPORARY CLINICAL IVF

demonstrate that the incidence of numerical (hypo-

haploidy, hyperhaploidy), structural (premature

chromatid separation), and organizational abnor-

malities (scattering or malalignment on the spindle)

increases with age, as do malformations in meiotic

spindle organization.

2,3,53

How these defects arise

and their specific association with age remains

controversial.

In 1968, Henderson and Edwards

105

proposed

what has become known as the ‘first in, first out’

theory, whereby chromosomal defects common in

oocytes ovulated late in life occur in those gametes

which formed late during fetal oogenesis. According

to this notion, the longer an oocyte resides in the

ovary, the greater is the probability that it will have

been exposed to developmentally toxic or adverse

influences, such as oxidative free radicals or lytic

enzymes released from proximal atretic oocytes.

106,107

Others suggest that the prolonged interval of mei-

otic arrest compromises the capacity of the oocyte

to resume meiosis

108

or to form normally function-

ing metaphase spindles that can capture and retain

all chromosomes, or once formed properly segre-

gate chromosomes in a numerically equivalent man-

ner between the oocyte and the polar bodies.

13,53

The

normal cycle checkpoints that prevent cell division

when chromosomal capture by kinetochore micro-

tubules is incomplete or chromosomal alignment on

the metaphase spindle is aberrant do not occur in

meiotic oocytes, and those which are operational

may decline in efficiency with advanced maternal

age.

The occurrence of aneuploidies associated with

spindle and segregation disorders (premature cen-

tromeric separation, anaphase lag, non-disjunction)

not only lowers fertility potential and increases mis-

carriage rates, but also is clearly associated with a

higher incidence of chromosomally abnormal

offspring.

Patterns of follicular growth change as women

age; during the follicular phase, early growth is ini-

tially more rapid but then slows to produce termi-

nally developed follicles the average diameter of

which at ovulation is smaller than is typical for their

younger counterparts. Older women also tend to have

frequent cycles in which more than the usual single

large (dominant) follicle develops. Both phenomena



prior to ovulation, the cell cycle has been arrested at

prophase of the first meiotic division. The resump-

tion of arrested meiosis is initiated by the sponta-

neous LH surge (or hCG administration after COHS)

and as a result, over 4 decades can elapse between

the beginning and end of the first meiotic division

in women.

After the LH-induced completion of meiosis I,

meiosis II progresses to and arrests at metaphase II.

The second meiotic division is resumed after sperm

penetration and completed with the extrusion of the

second polar body. Chromatin condensation into

bivalent MI and monovalent MII chromosomes per-

mits cytogenetic analysis of ploidy at each metaphase

during the preovulatory maturation of the oocyte.

The availability of unfertilized oocytes from IVF

procedures has provided an abundance of gametes

for cytogenetic analysis, and the results consistently

25 30

Age (years)

Antral follicle count

35 40 45

12%

4.8%

Figure 24.2 The relation between antral follicle counts and age

shows a steeper drop after age 37 years. From Faddy et al

101

with

permission.

HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 317

HUMAN PREIMPLANTATION EMBRYO SELECTION

are probably the consequence of unusually high

FSH levels that produce a pattern of follicular stim-

ulation not unlike the situation observed in cycles of

COHS.

109

While the capacity of follicles to make

estradiol and inhibin A does not appear to be age

related, against a background of equivalent FSH lev-

els, ‘younger’ follicles tend to produce more inhibin

110

Follicles from older women are less likely to

transform into functional corpora lutea; despite ultra-

sonic and hormonal evidence of follicular devel-

opment and ovulation, these follicles often fail to

synthesize progesterone after ovulation.Because estra-

diol production is sufficient to prevent menses for

as many as 12 days following ovulation, these aluteal

cycles are clinically unrecognized.

109

Rates of ovar-

ian stromal blood flow are reduced in women over

40.

111

Gaulden

showed that older women experi-

ence inadequate expansion of the perifollicular vas-

culature during the follicular phase of the natural

menstrual cycle, and proposed that perifollicular

under-vascularization could be associated with intra-

follicular under-oxygenation. According to her novel

hypothesis on the relationship between intrafollic-

ular hypoxia and trisomy (e.g. Down syndrome),

increased frequencies of chromosomal aneuploidy

may result from a modest acidification of the ooplasm

such that even a slight reduction in intracellular pH

could adversely influence the microtubular organi-

zation of the spindle and kinetic capacity of spindle

microtubules to segregate chromosomes normally.

Preliminary Doppler ultrasound findings indicate

that in natural menstrual cycles, women ⬎40 years

of age rarely produce follicles the perifollicular blood

flow characteristics of which are consistent with

higher levels of intrafollicular oxygenation and com-

petent, euploid oocytes (Van Blerkom, unpublished).

This possible mechanism of aneuploidy generation

requires confirmation, but if validated, may pro-

vide one of the most important reasons to include

Doppler ultrasonography in fertility treatments that

require COHS in general, and in women of advanced

reproductive age, in particular.

Tobacco smoking is one of the most common and

significant ‘lifestyle’ factors that can reduce a woman’s

fertility potential. Polycyclic aromatic hydrocar-

bons, heavy metals such as cadmium, and nicotine

metabolites are detected in follicular fluid and are

currently thought to interfere with normal biosyn-

thetic and regulatory activities during the follicu-

lar and preovulatory phases that may also involve

the oocyte.

112–115

Smoking causes an increase in

intrafollicular levels of reactive oxygen species the

pleiotropic effects of which on the oocyte and gran-

ulosa cell complement can include DNA fragmenta-

tion, chemical modification of proteins and certain

nucleotides, and lipid peroxidation.

116

Biochemical

alterations of this type adversely influence cell

function, and in severe cases, cause irreversible cell

damage or death. Follicular aspirates obtained from

smokers during ovum retrieval for IVF show higher

levels of oxidation and lower levels of activity for nor-

mal follicular antioxidative mechanisms.

117

Smokers

also produce a high number of diploid oocytes,

which may explain the unusually high frequency of

triploidy reported for these women, despite a reduced

fertilization capacity of their oocytes.

118

Significantly,

fewer follicles exist in the ovaries of current or

previous smokers,

119

and women who smoke go

through an earlier menopause.

120

Although other

agents such as caffeine and alcohol have been shown

to decrease fertility,

121

it remains unclear whether

the follicle or oocyte, or both, are directly affected.

Weight is another lifestyle factor that can influ-

ence fertility, but the ovary is not directly involved;

women who are significantly underweight or over-

weight are more likely to be anovulatory because of

hormonal changes associated with altered hypothal-

amic and pituitary function. However, for ovulatory

women undergoing COHS and IVF, weight is nega-

tively associated with outcome.After ovulation induc-

tion, intrafollicular levels of hCG have been reported

to be inversely related to body mass index (BMI).

High BMI was not only associated with low intrafol-

licular hCG levels, but also with poor fertilizability,

embryo quality, and a significant reduction in the

likelihood of a positive outcome.

122

For most women, however, smoking currently

remains the single most important and ubiquitous

lifestyle factor that negatively influences fertility.

This factor becomes especially acute when it is con-

sidered that over the reproductive life of a woman,

only 400–500 follicles will progress to ovulation,

123



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 318

OOCYTE SELECTION IN CONTEMPORARY CLINICAL IVF

and with age, the ability of follicles to produce a

normal oocyte undergoes a natural and progressive

decline. Studies of perifollicular blood flow that may

correlate with levels of glucocorticoid and growth

factor expression during the follicular phase (e.g.

VEGF, leptin, AMH, inhibin B) may provide a molec-

ular basis for reduced fertility in these women.

DO REPEATED CYCLES OF COHS

NEGATIVELY INFLUENCE

FERTILITY POTENTIAL?

It is common for infertile women to undergo multi-

ple cycles of ovarian stimulation prior to initiating

IVF, and to then have multiple IVF procedures per-

formed before a viable pregnancy occurs or treat-

ment is abandoned. Animal studies suggest that

repeated ovarian stimulation can have adverse effects

on follicular development and oocyte competence,

especially with respect to the formation of a normal

meiotic spindle with appropriately aligned chromo-

somes (for review see reference 124). However, at

present there is no compelling evidence to suggest

that a similar decline in oocyte competence accom-

panies multiple cycles of COHS and ovulation

induction in the human. While women who undergo

repeated cycles of stimulation are more likely to

require progressively higher dosages of gonado-

tropins, when compared with levels used in the first

cycle, the number of follicles produced and peak

estradiol levels detected are similar.

125,126

Repeated

IVF cycles involve multiple follicular punctures and

aspirations, and therefore can theoretically damage

the ovary. There is no microscopic evidence to date

to support potential adverse downstream conse-

quences for human oocyte competence associated

with multiple COHS/IVF procedures, although this

issue does require further investigation.

SUMMARY

While the complex cellular and molecular biology

of the ovarian follicle has long been recognized as a

factor in the establishment of normal developmental

competence for the oocyte, understanding its rela-

tionship to the success of assisted reproduction in

the human has increased in importance where restric-

tions on the number of embryos that can be replaced,

or the stage of preimplantation embryogenesis

permissible for transfer, are now legally mandated.

These limits make the identification of follicle-

specific biochemical properties and physiological

characteristics that may be associated with oocyte

competence necessary in clinical practice, as well as

prerequisites for the aforementioned ‘holy grail’ of

clinical IVF: accurate gamete selection. This is all

the more important where restrictions extend to

the number of oocytes that can be inseminated

for IVF. Ultimately, the most practical means of

follicular evaluation for oocyte selection are those

that are non-invasive and can be readily introduced

in assisted reproduction programs. The types of

biochemical analyses and Doppler ultrasound char-

acterizations described here meet these criteria, and

remain promising leads in this regard. However,

much remains to be understood about intra-

follicular conditions that may influence the human

oocyte, including the origin of follicle-specific dif-

ferences and whether their developmental effects on

the oocyte are direct or indirect. While the tone of

this review is sanguine, the well-used caveat ‘more

research is required’ applies because unambiguous

associations with outcome will continually be req-

uired to validate this approach in the assessment of

oocyte competence. Ideally, as investigations progress

and clinical correlations with outcome are obtained,

a more complete understanding of the molecular

mechanisms that operate within the follicle, and

how they may influence oocyte competence, will

emerge.

REFERENCES

1. Edwards R. Causes of early embryonic loss in human pregnancy.

Hum Reprod 1986; 1: 185–98.

2. Van Blerkom J. Developmental failure in human reproduction asso-

ciated with chromosomal abnormalities and cytoplasmic pathologies

in meiotically mature oocyes. In. Van Blerkom J, ed. The Biological

Basis of Early Human Reproductive Failure. Oxford: Oxford University

Press, 1994: 283–326.

3. Plachot M. Genetic analysis of the oocyte – a review. Placenta 2003;

24 (Suppl 2): S66–72.



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 319

HUMAN PREIMPLANTATION EMBRYO SELECTION

4. Makabe S, Van Blerkom J. An Atlas of Human Female Reproductive

Function: Ovarian Development to Early Embryogenesis after in

vitro Fertilization. New York: Taylor and Francis, 2006.

5. Motta PM, Nottola SA, Familiari G et al. Morphodynamics of the

follicular-luteal complex during early ovarian development and

reproductive life. Int Rev Cytol 2003; 223: 177–288.

6. Sandalinas M, Sadowy S, Alikani M et al. Developmental ability. of

chromosomally abnormal human embryos to develop to the blasto-

cyst stage. Hum Reprod 2001; 16: 1954–8.

7. Verlinsky Y, Kuliev A.An Atlas of Preimplantation Genetic Diagnosis.

New York; Parthenon Publishing, 2000.

8. Van Blerkom J, Henry G. Oocyte dysmorphism and aneuploidy in

meiotically mature human oocytes after controlled ovarian stimula-

tion. Hum Reprod 1992; 7: 379–90.

9. Meriano J, Alexis J, Visram-Zaver S et al. Tracking of oocyte dysmor-

phisms for ICSI patients may prove relevant to the outcome in sub-

sequent patient cycles. Hum Reprod 2001; 16: 2118–23.

10. Mikkelsen A, Lindenberg S. Morphology of in-vitro matured oocytes:

impact on fertility potential and embryo quality. Hum Reprod 2001;

16: 1714–18.

11. Otsuki J, Okada K, Morimoto Y et al. The relationship between preg-

nancy outcome and smooth endoplasmic reticulum clusters in MII

human oocytes. Hum Reprod 2004; 19: 1591–7.

12. Ciotti P, Notarangelo A, Morselli-Labate V et al. First polar body

morphology before ICSI is not related to embryo quality or preg-

nancy rate. Hum Reprod 2004; 19: 2334–9.

13. Eichenlaub-Ritter U, Sun F. Maternal age and oocyte competence. In

Van Blerkom J, Gregory L, eds. Essential IVF: Basic Research and

Clinical Applications. Boston: Kluwer Academic Publishers, 2004:

201–30.

14. Akande V, Flemming C, Hunt L, Keay S, Jenkins J. Biological versus

chronological ageing of oocytes, distinguishable by raised FSH levels

in relation to the success of IVF treatment. Hum Reprod 2002; 17:

2002–8.

15. Simonetti S, Veeck L, Jones H, Jr. Correlation of follicular fluid vol-

ume with oocyte morphology from follicles stimulated by human

menopausal gonadotropin. Fertil Steril 1985; 44: 177–80.

16. Wittmaack F, Kreger D, Blasco L et al. Effect of follicular size on

oocyte retrieval, fertilization, cleavage, embryo quality in in vitro fer-

tilization cycles: a 6-year data collection. Fertil Steril 1994; 62: 1205–10.

17. Dubey A, Wang H, Duffy P, Penzias A. The correlation between follic-

ular measurements, oocyte morphology and fertilization rates in an

in vitro fertilization program. Fertil Steril 1995; 64: 787–90.

18. Arnot A,Vanderckhove P, DeBono M, Rutherford A. Follicular volume

and number during in vitro fertilization: association with oocyte devel-

opmental capacity and pregnancy rate. Hum Reprod 1995; 10: 256–61.

19. Miller K, Goldberg J, Falcone T. Follicle size and implantation of

embryos from in vitro fertilization. Obstet Gynecol 1996; 88: 583–6.

20. Ectors F, Vanderzwalmen P, Van Hoeck J et al. Relationship of human

follicular diameter with oocyte fertilization and development after

in-vitro fertilization or intracytoplasmic sperm injection. Hum

Reprod 1997; 12: 2002–5.

21. Bergh C, Broden H, Lundin K, Hamberger L. Comparison of fertil-

ization, cleavage and pregnancy rates of oocytes from large and small

follicles. Hum Reprod 1998; 13: 1912–5.

22. Salha O, Nugent D, Dada T et al. The relationship between follicular

fluid aspirate volume and oocyte maturity in in-vitro fertilization

cycles. Hum Reprod 1998; 13: 1901–6.

23. Triwitayakorn A, Suwajanakorn S, Pruksananonda K et al. Correlation

between human follicular diameter and oocyte outcome in an ICSI

program. J Assist Reprod Genet 2003; 20: 143–7.

24. Gerris J. Reducing the number of embryos to transfer after IVF/ICSI

In Van Blerkom J, Gregory L, eds. Essential IVF: Basic Research and

Clinical Applications. Boston: Kluwer Academic Publishers, 2004:

505–75.

25. Benagiano G, Gianaroli L. The new Italian IVF legislation. Reprod

Biomed Online 2004: 9: 117–25.

26. Papanikolaou E, Camus M, Kolibiankakis E et al. In vitro fertilization

with single blastocyst-stage versus single cleavage-stage embryos.

N Engl J Med 2006; 354: 1139–46.

27. Borini A. Predictive factors for embryo implantation potential. Reprod

Biomed Online 2005; 10: 653–68.

28. Manna C, Nardo N. Italian law on assisted conception: clinical and

research implications. Reprod Biomed Online 2005;11:532–4.

29. Strohmer H, Herczeg C, Plockinger B et al. Prognostic appraisal of

success and failure of in vitro fertilization program by transvaginal

Doppler ultrasound at the time of ovulation induction. Ultrasound

Obstet Gynecol 1991; 1: 272–4.

30. Steer C, Campbell T, Tan S et al. The use of color flow imaging after

in vitro fertilization to identify optimum uterine conditions before

embryo transfer. Fertil Steril 1992; 57: 372–6.

31. Oyesanya O, Parsons J, Collins W et al. Prediction of oocyte recovery

rate by transvaginal ultrasonography and color Doppler imaging

before human chorionic gonadotropin administration in in vitro

fertilization cycles. Fertil Steril 1996; 65: 806–9.

32. Nargund G, Doyle P, Bourne T et al. Association between ultrasound

indices of follicular blood flow, oocyte recovery, and preimplantation

embryo quality. Hum Reprod 1996; 11: 109–12.

33. Nargund G, Doyle P, Bourne T et al. Ultrasound derived indices of

follicular blood flow before HCG administration and the prediction

of oocyte recovery and preimplantation embryo quality. Hum Reprod

1996; 11: 2515–7.

34. Chui D, Jurovic D, Bourne T et al. Follicular vascularity – the predic-

tive value of transvaginal power Doppler ultrasonography in an in

vitro fertilization programme. A preliminary study. Hum Reprod 1997;

12: 191–6.

35. Van Blerkom J, Antczak M, Schrader R. The developmental potential

of the human oocyte is related to the dissolved oxygen content of

follicular fluid: association with vascular endothelial growth factor

levels and perifollicular blood flow characteristics. Hum Reprod

1997; 12: 1947–55.

36. Bahl P, Pugh N, Chui D et al. The use of transvaginal power Doppler

ultrasonography to evaluate the relationship between perifollicular

vascularity and outcome in in-vitro fertilization treatment cycles.

Hum Reprod 1999; 14: 939–45.

37. Coulam C, Goodman C, Rinehart J. Color Doppler indices of follic-

ular blood flow as predictors of pregnancy after in-vitro fertilization

and embryo transfer. Hum Reprod 1999; 14: 1979–82.

38. Huey S,Abuhamad A, Barrosa G et al. Perifollicular blood flow Doppler

but not follicular pO2, pCo2, or pH, predict oocyte developmental

competence in in vitro fertilization. Fertil Steril 1999; 72: 707–12.

39. Borini A, Maccolini A, Tallarini A et al. Perifollicular vascularity and

its relationship with oocyte maturity and IVF outcome. Ann NY

Acad Sci 2001; 943: 64–7.

40. Gregory L. Perifollicular vascularity: a marker of follicular hetero-

geneity and oocyte competence and a predictor of implantation in

assisted conception cycles. In Van Blerkom J, Gregory L, eds. Essential

IVF: Basic Research and Clinical Applications. Boston: Kluwer

Academic Publishers, 2004: 59–80.

41. Van Blerkom J. Follicular influences on oocyte and embryo compe-

tence In DeJonge C, Barratt C, Eds.Assisted Reproductive Technology,

Cambridge: Cambridge University Press, 2002: 81–105.



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 320

OOCYTE SELECTION IN CONTEMPORARY CLINICAL IVF

42. Bahl P, Pugh N, Gregory L et al. Perifollicular vascularity as a poten-

tial variable affecting outcome in stimulated intrauterine insemina-

tion treatment cycles: a study using transvaginal power Doppler.

Hum Reprod 2001: 16; 1682–9.

43. Shrestha S, Costello M, Sjoblom P et al. Power Doppler ultrasound

assessment of follicular vascularity in the early follicular phase and

its relationship with outcome in in vitro fertilization. J Assist Reprod

Genet 2006; 23: 161–9.

44. Palomba S, Russo T, Falbo A et al. Clinical use of the perifollicular

vascularity assessment in IVF cycles: a pilot study. Hum Reprod 2006;

21: 1055–61.

45. Merce L, Bau S, Barco M et al. Assessment of the ovarian volume,

number and volume of follicles and ovarian vascularity by three-

dimensional ultrasonography and power Doppler angiography on

the HCG day to predict the outcome in IVF/ICSI cycles. Hum

Reprod 2006; 21: 1218–26.

46. Kim K, Oh D, Jeong J et al. Follicular blood flow is a better predictor

of the outcome of in vitro fertilization-embryo transfer than follicu-

lar fluid vascular endothelial growth factor and nitric oxide concen-

trations. Fertil Steril 2004; 82: 586–92.

47. Pan H-A, Wu H-H, Cheng Y-C et al. Quantification of ovarian

stromal Doppler signals in poor responders undergoing in vitro fer-

tilization with three-dimensional power Doppler ultrasonography.

Am J Obstet Gynecol 2004; 190: 338–44.

48. Bergqvist A, Bergqvist D, Ferno M. Estrogen and progesterone recep-

tors in vessel walls: biochemical and immunochemical assays. Acta

Obstet Gynecol Scand 1993; 72: 10–6.

49. Gislard V, Miller V, Van Houte P. Effects of 17␤-oestradiol on

endothelium-dependent responses in the rabbit. J Pharmacol Exp

Ther 1998; 244: 19–22.

50. Kan A, Ng EH, Yeung WS, Ho PC. Perifollicular vascularity in poor

ovarian responders during IVF. Hum Reprod 2006; 21: 1539–44.

51. Van Blerkom J. Epigenetic influences on oocyte developmental com-

petence: Follicular oxygenation and perifollicular vascularity. J Assist

Reprod Genet 1998; 15: 226–34.

52. Gaulden M. The enigma of Down syndrome and other trisomic

conditions. Mutat Res 1992; 269: 68–88.

53. Battaglia D, Goodwin P, Klein N et al. Influence of maternal age on

meiotic spindle assembly in oocytes from naturally cycling women.

Mol Hum Reprod 1996; 11: 2217–22.

54. Clark A, Stokes Y, Lane M, Thompson J. Mathematical modelling

of oxygen concentration in bovine and murine cumulus-oocyte

complexes. Reproduction 2006; 131: 999–1006.

55. Dickey R, Matulich E. Ultrasound imaging at the beginning of the

second millennium. In DeJonge C, Barratt C, eds.Assisted Reproductive

Technology, Cambridge: Cambridge University Press, 2002: 282–301.

56. Antczak M. The synthetic and secretory behaviors (nonsteroidal) of

ovarian follicular granulosa cells: parallels to cells of the endothe-

lial cell lineage. In Van Blerkom J, Gregory L, Eds. Essential IVF:

Basic Research and Clinical Applications. Boston: Kluwer Academic

Publishers, 2004: 1–41.

57. Michael A. Do biochemical predictors of outcome exist? In Van

Blerkom J, Gregory L, Eds. Essential IVF: Basic Research and Clinical

Applications. Boston: Kluwer Academic Publishers, 2004: 81–110.

58. Malamitsi-Puchner A. Novel follicular fluid factors influencing

oocyte developmental potential in IVF: a review. Reprod Biomed

Online 2006; 12: 500–6.

59. Otani N, Minami S, Yamoto M et al. The vascular endothelial growth

factor/fms-like tyrosine kinase system in human ovary during the

menstrual cycle and early pregnancy. J Clin Endocrinol Metabol

1999; 84: 3845–51.

60. Yan Z, Neulen J, Raczek S et al. Vascular endothelial growth factor

(VEGF)/vascularity permeability factor (VPF) production by lutein-

ized human granulosa cells in vitro; a paracrine signal in corpus

luteum formation. Gynecol Endocrinol 1993; 12: 145–53.

61. Antczak M, Van Blerkom J, Clark A. A novel mechanism of vascular

endothelial growth factor, leptin and transforming growth factor

beta2 sequestration in a subpopulation of human ovarian follicle

cells. Hum Reprod 1997; 12: 2226–34.

62. Cioffi J,Van Blerkom J, Antczak M et al. The expression of leptin and

its receptors in pre-ovulatory human follicles. Mol Hum Reprod

1997: 3: 467–72.

63. Bouloumie A, Drexler H, Lafontan M, Busse R. Leptin, the product of

the ob gene, promotes angiogenesis. Circ Res 1998; 83: 1059–66.

64. Kamut B, Brown L, Mannseau E et al. Expression of vascular endothe-

lial growth factor by human granulosa and theca lutein cells. Role in

corpus luteum development. Am J Pathol 1995; 146: 157–65.

65. Shweiki D, Itin A, Soffer D, Keshet E. Vascular endothelial growth fac-

tor induced by hypoxia may mediate hypoxia-initiated angiogenesis.

Nature 1992; 359: 843–5.

66. Smeneza G. Regulation of mammalian O

homeostasis by hypoxia-

inducible factor 1. Ann Rev Cell Dev Biol 1999; 15: 551–78.

67. Bruick RK. Oxygen sensing in the hypoxic response pathway: regula-

tion of the hypoxia-inducible transcription factor. Genes Dev 2003;

17: 2614–23.

68. Bell E, Brooke M, Emerling B, Navdeep S. Mitochondrial regulation

of oxygen sensing. Mitochondrion 2005; 5: 322–32.

69. Bunn H, Poynton R. Oxygen sensing and molecular adaptations to

hypoxia. Physiol Rev 1996; 76: 839–85.

70. Antczak M. Possible ramifications of the identification of ovarian

follicular granulosa cells as specialized endothelial-like cells: a specu-

lative treatise. Reprod Biomed Online 2001; 2: 188–97.

71. Artini P, Monti M, Fasciani A et al. Vascular endothelial cell growth

factor, interleukin-6 and interleukin-2 in serum and follicular fluid

of patients with ovarian hyperstimulation syndrome. Eur J Obset

Gynecol Reprod Biol. 2002; 101: 169–74.

72. Friedman C, Seifer D, Kennard E et al. Elevated levels of follicular fluid

vascular endothelial growth factor is a marker of diminished pregnancy

73. Tetsuka M, Haines L, Milne M et al. Regulation of 11␤-hydroxy-

steroid dehydrogenase type 1 gene expression by LH and inter-

leukin-1␤ in cultured rat granulosa cells. J Endocrinol 1999; 163:

417–23.

74. Keay S, Harlow C, Wood P et al. Higher cortisol : cortisone ratios in

the preovulatory follicle of unstimulated IVF cycles indicate oocytes

with increased pregnancy potential. Hum Reprod 2002; 17: 2003–8.

75. Michael A, Gregory L, Thaventhiran L et al. Follicular variation in

ovarian 11␤-hydroxysteroid dehydrogenase (11␤HSD) activities:

evidence for the paracrine inhibition of 11␤HSD in human granu-

losa-lutein cells. J Endocrinol 1996;148: 419–25.

76. Durlinger A, Gruijeters M, Kramer P et al. Anti-Mullerian hormone

inhibits initiation of primordial follicle growth in the mouse ovary.

Endocrinology 2002; 143: 1976–84.

77. van Rooij I, Broekmans F, te Velde E et al. Serum anti-Mullerian hor-

mone levels: a novel measure of ovarian reserve. Hum Reprod 2002;

17: 3065–71.

78. Hazout A, Bouchard P, Seifer D et al. Serum antimullerian hor-

mone/mullerian-inhibiting substance appears to be a more discrim-

inatory marker of assisted reproductive technology outcome than

follicle-stimulating hormone, inhibin B, or estradiol. Fertil Steril

2004; 82: 1323–9.

79. Seifer D, MacLaughlin D, Christian B et al. Early follicular serum

mullerian-inhibiting substance levels are associated with ovarian



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 321

HUMAN PREIMPLANTATION EMBRYO SELECTION

response during assisted reproductive technology cycles. Fertil Steril

2002; 77: 468–71.

80. Hirobe S, He W, Gustafson M, MacLaughlin D, Donahoe P. Mullerian

inhibiting substance gene expression in the cycling rat ovary corre-

lates with recruited or graafian follicle selection. Biol Reprod 1994;

50: 1238–43.

81. Fallat M, Siow Y, Marra M et al. Mullerian-inhibiting substance in

follicular fluid and serum: a comparison of patients with tubal factor

infertility,polycystic ovary syndrome, and endometriosis. Fertil Steril

1997; 67: 962–5.

82. Franchin R, Mendez Lozano D, Louafi N et al. Dynamics of serum

anti-mullerian hormones levels during the luteal phase of controlled

ovarian hyperstimulation. Hum Reprod 2005; 20: 747–51.

83. Silberstein T, MacLaughlin D, Shai I et al, Mullerian inhibiting sub-

stance levels at the time of HCG administration in IVF cycles pre-

dicts both ovarian reserve and embryo morphology. Hum Reprod

2006: 21: 159–63.

84. Eldar-Geva T, Ben-Chetrit A, Spitz I et al. Dynamic assays of inhibin

B, anti-mullerian hormone and estradiol following FSH stimulation

and ovarian ultrasonography as predictors of IVF outcome. Hum

Reprod 2005; 20: 3178–83.

85. Groome N, Illingworth P, O’Brien M et al. Measurement of dimeric

inhibin B throughout the human menstrual cycle. J Clin Endocrinol

Metab 1996; 814: 1401–5.

86. Fowler P, Fahy U, Culler M et al. Gonadotrophin surge-attenuating

factor bioactivity is present in follicular fluid from naturally cycling

women. Hum Reprod 1995; 10: 68–74.

87. Enskog A, Nilsoon L, Brannstrom M. Peripheral blood concentra-

tions of inhibin B are elevated during gonadotrophin stimulation in

patients who later develop ovarian OHSS and inhibin A concentra-

tions are elevated after OHSS onset. Hum Reprod 2000; 15: 532–8.

88. Lau C, Ledger W, Groome NP et al. Dimeric inhibins and activin A in

human follicular fluid and oocyte-cumulus culture media. Hum

Reprod 1999; 14: 2525–30.

89. Chang C, Wang T, Horng S et al. The concentration of inhibin B in

follicular fluid: relation to oocyte maturation and embryo develop-

ment. Hum Reprod 2002; 17: 1724–8.

90. Urbancsek J, Hauzman E, Klinga K et al. Use of serum inhibin B

levels at the start of ovarian stimulation and at oocyte pickup in the

prediction of assisted reproductive treatment outcome. Fertil

Steril 2005; 83: 341–8.

91. Mendoza C, Ruiz-Requena E, Ortega E et al. Follicular fluid markers

of oocyte developmental potential. Hum Reprod 2002; 17: 1017–22.

92. Coulam C, Bustillo M, Schulman J. Empty follicle syndrome. Fertil

Steril 1986; 46: 1153–5.

93. Ndukew G, Thornton S, Fishel S et al. Curing empty follicle syn-

drome. Hum Reprod 1997; 12: 21–3.

94. Tsuiki A, Rose B, Hung T. Steroid profiles of follicular fluids from a

patient with empty follicle syndrome. Fertil Steril 1988; 49: 104–7.

95. Bustillo M. Unsuccessful oocyte retrieval: technical artifact or

genuine ‘empty follicle syndrome’. Reprod Biomed Online 2004;

8: 59–67.

96. Zreik TG, Garcia-Velasco J, Vergara T et al. Empty follicle syndrome:

evidence for recurrence. Hum Reprod 2000; 15: 999–1002.

97. Hassan H, Saleh H, Khalil O et al. Double oocyte aspiration may be a

solution for empty follicle syndrome. Case reports. Fertil Steril 1998;

69: 138–9.

98. Awoniyi A, Govindbhai J, Zierke S et al. Continuing the debate on

empty follicle syndrome: can it be associated with normal bioavail-

ability of ␤-human chorionic gonadotropin on the day of oocyte

recovery? Hum Reprod 1998: 13: 1281–4.

99. Yen S, Jaffe R, Barbieri R. Reproductive Endocrinology, 4th edn.

WB Saunders, 1999, Philadelphia.

100. Faddy M, Gosden R. A model conforming the decline in follicle

numbers to the age of menopause in women. Hum Reprod 1996; 11:

1484–6.

101. Scheffer G, Broekmans F, Dorland M et al. Antral follicle counts by

transvaginal ultrasonography are related to age in women with

proven natural fertility. Fertil Steril 1999; 72: 845–51.

102. Dunson D, Colombo B, Baird D. Changes with age in the level and

duration of fertility in the menstrual cycle. Hum Reprod 2002; 17:

1399–403.

103. Piette C, de Mouzon J, Bachelot A et al. In-vitro fertilization: influ-

ence of women’s age on pregnancy rates. Hum Reprod 1990; 5: 56–9.

104. Faddy M, Gosden R, Gougeon A et al. Accelerated disappearance of

ovarian follicles in mid-life-implications for forecasting menopause.

Hum Reprod 1992; 7: 1342–6.

105. Henderson D, Edwards R. Chiasma frequency and maternal age in

mammals. Nature 1968; 218: 22–8.

106. Volarcik K, Sheean L, Goldfarb J et al. The meiotic competence of in

vitro matured human oocytes is influenced by donor age: evidence

that folliculogenesis is compromised in the reproductively aged

ovary. Hum Reprod 1998; 13: 154–60.

107. Tarin J. Aetiology of age-associated aneuploidy: a mechanism based

of the ‘free radical theory of aging’. Mol Hum Reprod 1995; 1:

1563–5.

108. Crowley P, Gulah D, Hayden T et al. A chiasma-hormonal hypothesis

relating Down’s syndrome and maternal age. Nature 1979; 280:

417–8.

109. Santoro N, Isaac B, Neal-Perry G et al. Impaired folliculogenesis and

ovulation in older reproductive age women. J Clin Endocrinol Metab

2003: 88: 5502–9.

110. Hansen K, Thyer A, Sluss P et al. Reproductive ageing and ovarian

function: is the early follicular phase FSH rise necessary to maintain

adequate secretory function in older ovulatory women? Hum Reprod

2005; 20: 89–95.

111. Ng E, Chan C, Yeung W et al. Effect of age on ovarian stromal flow

measured by three-dimensional ultrasound with power Doppler in

Chinese women with proven fertility. Hum Reprod 2004; 19: 2123–37.

112. Zenzes M, Wang P, Casper R. Cigarette smoking may affect meiotic

maturation of human oocytes. Hum Reprod 1995; 10: 3213–7.

113. Zenzes M, Krishnan S, Krishnan B et al. Cadmium accumulation in

follicular fluid of women in in vitro fertilization-embryo transfers is

higher in smokers. Fertil Steril 1995; 64: 599–603.

114. Zenzes M, Reed T, Wang P et al. Cotinine, a major metabolite of

nicotine, is detectable in follicular fluids of passive smokers in

in vitro fertilization therapy. Fertil Steril 1996; 66: 614–19.

115. Zenzes M, Puy L, Bielecki R. Immunodetection of benzo[a]pyrene

adducts in ovarian cells of women exposed to cigarette smoke. Mol

Hum Reprod 1998; 4: 159–65.

116. Finkel T, Holbrook N. Oxidants, oxidative stress and the biology of

ageing. Nature. 2000; 408: 239–47.

117. Paszkowski T, Clarke RN, Hornstein MD. Smoking induces oxidative

stress inside the Graafian follicle. Hum Reprod 2002; 17: 921–5.

118. Rosevear S, Holt D, Lee T et al. Smoking and decreased fertilisation

rates in vitro. Lancet 1992; 340: 1195–6.

119. Westhoff C, Murphy P, Heller D. Predictors of ovarian follicle

number. Fertil Steril 2000; 74: 624–8.

120. Midgette A, Baron J. Cigarette smoking and the risk of natural

menopause. Epidemiology 1990; 6: 474–80.

121. Hassan M, Killick S. Negative lifestyle is associated with a significant

reduction in fecundity. Fertil Steril 2004; 81: 384–92.



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 322

OOCYTE SELECTION IN CONTEMPORARY CLINICAL IVF

122. Carrell D. Body mass index is inversely related to intra-follicular

HCG concentrations, embryo quality and IVF outcome. Reprod

BioMed Online 2001; 3: 109–11.

123. Peters H. Intrauterine gonadal development. Fertil Steril 1976; 27:

493–500.

124. Van Blerkom J, Davis P. Differential effects of repeated ovarian stim-

ulation on cytoplasmic and spindle organization in metaphase II

mouse oocytes matured in vivo and in vitro. Hum Reprod 2001; 16:

757–64.

125. Doldi N, Persico P, De Santis L et al. Consecutive cycles in in vitro

fertilization–embryo transfer. Gynecol Endocrinol 2005; 20: 132–6.

126. Hoveyda F, Engmann L, Steele J et al. Ovarian response in three con-

secutive in vitro fertilization cycles. Fertil Steril 2002; 77: 706–10.



HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 323

HPE_Chapter24.qxp 7/13/2007 4:54 PM Page 324

INTRODUCTION

The potential impact of an abnormal paternal

genome on reproductive outcome is unquestion-

ably less than that of its female counterpart. The

importance of the egg is best illustrated by the

plethora of studies relating female age to pregnancy

outcome.

1,2

Furthermore, the impact of egg quality

on reproductive success has been powerfully estab-

lished by the continued success of donor oocyte

programs. In the USA, the Assisted Reproductive

Technology Report for 2003 indicated that the

national data from donor oocytes gave rise to a 50%

live birth rate, with some clinics reporting live birth

rates of over 70%.

In contrast, the influence of the human sperm on

reproductive outcome has not yet been well charac-

terized. In animal studies, evidence of a paternal

effect on reproductive outcome is clear. Many

animal studies have centered around examining the

effect of radiation or toxic chemicals on spermato-

zoa and their subsequent effects on reproductive

outcome. An example of some of these intriguing

studies includes data from Robaire’s group, who

have shown clear evidence that cyclophosphamide

treatment affects rodent sperm in such a way that it

leads to dysregulation of gene expression in the

preimplantation embryo, increased postimplanta-

tion losses, fetal malformations, and transmission of

fetal abnormalities to the subsequent generation.

4–6

One of the most worrying traits observed in these

studies is the inheritance of a paternal component

over generations. This observation has recently been

dramatically highlighted in a study examining the

transient exposure of a gestating female rat to

endocrine disruptors during the period of gonadal

sex determination. The endocrine disruptors induced

an adult male phenotype of decreased spermatogenic

capacity (cell number and viability) and increased

incidence of male infertility in the F1 generation.

These effects were transferred through the male germ

line to nearly all males of all subsequent generations

examined (that is, F1 to F4).

In the human, pater-

nal effects on reproductive outcome are not as clear

cut. One study by Parker et al

suggested an associa-

tion between the risk of stillbirth and the father’s

exposure to external ionizing radiation before

conception. However this epidemiological study has

also been criticized.

A number of studies have also

linked advanced paternal age with the inheritance of

adverse traits, including birth defects, mental retar-

dation, Apert’s syndrome, autism, and schizophre-

nia.

10–14

All of these studies further suggest that

biological mechanisms exist in sperm, and their

disturbance will disrupt the delicate equilibrium

needed for successful fertilization and embryo

growth, with various adverse reproductive outcomes

as a result.

The factors present in the paternal genome that

may have an impact on poor reproductive outcome

are still hypothetical. However, in the past decade

the quality of the sperm nuclear DNA has been

more rigorously examined. This has also coincided

with an increased use of intracytoplasmic sperm

injection (ICSI), for which the quality of spermato-

zoa is generally accepted to be poorer. An abun-

dance of articles have been published indicating

that DNA strand breaks are clearly detectable in

ejaculated spermatozoa, and their presence is

increased in the ejaculates of men with poor semen

parameters.

15–21

This chapter therefore examines

increasing evidence that paternal factors exist which

may impact on reproductive outcome, with partic-

ular emphasis on abnormal sperm nuclear DNA.



25. Sperm DNA and embryo development

Denny Sakkas and Emre Seli

HPE_Chapter25.qxp 7/14/2007 5:52 PM Page 325