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From the * Department of Farm Animal Health,
Department of Biochemistry and Cell Biology,
Faculty of Veterinary Medicine, Utrecht University, Utrecht, The
Netherlands.
| Correspondence to: Dr B. M. Gadella, Department of Biochemistry and Cell Biology, Department of Farm Animal Health, Faculty of Veterinary Medicine, PO Box 80.176, Yalelaan 2, 3584 CM, Utrecht, The Netherlands (e-mail: B.Gadella{at}vet.uu.nl). |
| Received for publication September 29, 2004; accepted for publication July 14, 2005. |
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Abstract |
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Key words: Mitotic spindles, blastocyst, ICSI
Although initial data suggest that the normality of babies born after in vitro fertilization (IVF)/ICSI is not jeopardized, concerning reports of genetic/chromosomal defects in children born by ICSI have been published. The results of controlled studies indicate that the risk of sex chromosome anomalies (Bonduelle et al, 1998, 2002a) or congenital malformations (Ericson and Kallen, 2001; Ludwig and Katalinic, 2003) in babies born by ICSI, as compared with natural conception, is increased. Moreover, an increased risk of imprinting errors in offspring born following ICSI with testicular sperm has been reported (Arney et al, 2002; Cox et al, 2002). It is important to mention that recent human fertility studies strongly indicate that the risk of major birth defects between ICSI and conventional IVF are not different (Bonduelle et al, 2002b), and both are 2 times higher (Hansen et al, 2002) or at least increased (De Vroey and van Steirteghem, 2004) compared with naturally conceived infants.
In the context of assisted conception both in animal models and in clinical studies, the degree of DNA aberrations or damage in sperm cells has been linked to impairment of fertilization, embryo development (Egozcue et al, 2000; Hargreave, 2000a; Shi and Martin, 2000), and a reduced chance of producing live offspring (Sakkas et al, 2000; Shen and Ong, 2000; Hales and Robaire, 2001). However, the potential ability of sperm with damaged DNA to fertilize oocytes and consequences on embryo development have not been widely studied. Besides, although a considerable quantity of data on paternally mediated adverse effects in the offspring has been accumulated, the regarding mechanism is still not clarified. The results of different studies indicate that 2 principal mechanisms by which such an effect may be induced are the induction of germ line genomic instability or the suppression of germ cell apoptosis (Brinkworth, 2000).
In the present study, we have exclusively damaged the DNA of bovine sperm by irradiation. The DNA-damaged cells did not show signs of functionally affected integrity of membranes, organelles, and motility. The objective of this study was to investigate the fertilizing ability of DNA-damaged sperm in a bovine IVF model and to follow the consequences of introducing paternal damaged DNA into the fertilized oocyte for further embryo development. Furthermore, our study was aimed at elucidating the mechanism by which such effects may be induced. We hypothesized that posttesticular sperm DNA damage (as long as no severe DNA decondensation takes place affecting the sperm head volume) will only marginally affect sperm physiology due to the complete block of gene expression in those cells. However, we expect that this DNA damage will affect embryo development at the stage that paternal gene expression is initiated (<8-cell stage; Viuff et al, 1996). In this scenario, either the damaged sperm DNA can be repaired by the fertilized oocyte before onset of embryonic genome expression (Generoso et al, 1979; Brandriff and Pedersen, 1981) or, in case the DNA damage persists, it will then become sensitive for the apoptotic machinery of the developing embryo. Therefore, we measured the rate of embryo cleavage as well as blastocyst formation and the incidence of embryonic cell apoptosis in oocytes that were IVF treated with 0-10 Gy irradiated sperm. Possible implications for oocytes fertilized by the use of assisted reproductive techniques (ART) are discussed.
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Materials and Methods |
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Induction of DNA Strand Breaks in Spermatozoa
Two thawed straws were subjected to x-ray irradiation using a RT 250 at 200
KV, 20 MA (Philips, Eindhoven, The Netherlands) at a single dose of 0.6, 1.25,
2.5, 5, or 10 Gy, while 2 other straws were kept at room temperature as
control. In experiments in which higher doses of irradiation were required,
bull semen straws were subjected to Gamma rays at a single dose of 5, 10, 15,
20, 25, 30, 35, 40, 45, 50, 75, 100, 150, 225, and 300 Gy from a
60Co isotope source (Gamma cell 1000; Atomic Energy of Canada,
Limited, Mississanga, Canada). Irradiation dose rate at room temperature was
12 Gy/min. Dosage output of irradiation machines was regularly controlled.
Sperm-Motility Assessment
After irradiation, the semen straws were transferred to the laboratory and
diluted in 1:1 volume of prefiltered (Millex-GS, 0.22 µm) egg-yolk extender
and were kept in an oven at 37°C. At 0, 2, 4, and 6 hours after
irradiation, the sperm total motility was assessed by using a HTM-IVOS
Motility Analyzer version 8.1 (Hamilton Thorn Research, Beverly, Mass). To
obtain accurate cell concentration and motility measurements, some of the
computer program options were adjusted. Briefly, minimum contrast was set to
9, minimum size to 6, low size gate to 1.0, high size gate to 1.6, low
intensity gate to 0.6, and high intensity gate to 1.2. For each treatment, 3
replicates and at least 300 sperm cells in each replicate were evaluated.
Detection of Single-Stranded DNA by Acridine Orange Staining
Staining of single-stranded DNA in irradiated sperm samples was performed
as previously described (Evenson et al,
2002). Sperm samples were centrifuged for 3 minutes at 1000
x g and the pellet was washed once and resuspended in TNE
buffer (containing 0.15 M NaCl, 1 mM EDTA, and 10 mM Tris, pH 7.2) at 4°C
to a final concentration of 5 x 106 cells/mL. Two hundred
microliters of this sperm suspension was mixed with 400 µL of a
detergent/acid solution of 0.1% Triton X-100, 0.15 M NaCl, and 0.08 M HCl (pH
1.4). After 30 seconds, cells were stained by adding 1.2 mL of a solution
containing 6 µg/mL of acridine orange in staining buffer (0.15 M NaCl, 1 mM
EDTA, 10 mM Tris, 0.2 M Na2HPO4, 0.1 M citric acid [pH
6.0]).
Stained cells were analyzed by a FACSCalibur flow cytometer (Becton Dickinson, San Jose, Calif) equipped with an Argon ion laser, tuned at 488 nm. Events (10 000) were accumulated for each measurement. Under these experimental conditions, when excited with a blue-light source, acridine orange (AO) intercalates with double-stranded DNA and emits green fluorescence (530 ± 15 nm) (detected in a fluorescence detector FL1). AO associated with single-stranded DNA (or RNA) emits red fluorescence (>630 nm) detected in another fluorescence detector FL3. Because it has been demonstrated that residual RNA molecules in mature sperm do not interfere with the measurement (Evenson et al, 1991), the ratio of red to green fluorescence reflects the level of single- vs double-stranded DNA. Sperm DNA damage was quantified by flow-cytometry measurements (FCM) of the emission shift from green (native, double-stranded DNA) to red (denatured, single-stranded DNA) fluorescence and displayed as red vs green fluorescence intensity cytogram patterns. The flow cytometer can resolve 1024 X-channels vs Y-channels of spermatozoa that emit green (530 ± 15-nm band-pass filter) and red (>630-nm long-pass filter) fluorescence to precisely characterize each of the 10 000 cells measured. Three minutes after the start of the procedure, allowing time for hydrodynamic equilibration, signal acquisition to a computer list mode file was initiated. By using a double scatter plot for forward and sideways scatter and based on different scatter properties, nonsperm events were gated out. The double scatter plot analysis of raw data with each point representing the coordinate of red and green fluorescence intensity values for every individual sperm was performed with use of win MDI version 2.8 software (J. Trotter, free software). Nonirradiated samples were stained and used as negative control.
Deoxynucleotidyl Transferase-Mediated Nick End Labeling Detection of DNA Damage
DNA strand breaks (single-stranded as well as double-stranded DNA with
3'-OH overhang) are detected in permeabilized fixed sperm cells by
subjecting them to exogenous TdT in the presence of biotin-dUTP. The
3'-OH ends of DNA strand breaks serve as primers for the incorporation
of biotin-dUTP, which is detected through the use of fluoresceinated avidin
(Gorczyca et al, 1993). In our
study, an in situ DNA strand-break detection kit (Roche Diagnostics, Mannheim,
Germany) using deoxynucleotidyl transferase (TdT)-mediated dUTP nick end
labeling (TUNEL) was used. Briefly, sperm cells were vortexed in phosphate
buffered saline (PBS) containing 0.1% w/v polyvinylalcohol (PBS-PVA) and
washed twice in PBS-PVA (500 x g, 5 minutes) and resuspended in
PBS with 0.1% Triton X-100 plus 0.1% sodium citrate for 5 minutes on ice for
cell permeabilization. The resulting cell suspension was washed twice in
PBS-PVA and resuspended and incubated in 50 µL of TUNEL reaction mixture
for 1 hour at 37°C in a dark and humidified atmosphere. Finally, sperm
cells were rinsed in PBS-0.1% w/v bovine serum albumin (PBS-BSA) and subjected
to flow cytometry. Control cells for each sample were treated identically
except for the omission of the TdT enzyme from TUNEL reaction mixture
(supplier's protocol). Red fluorescence signal was used to gate by computer
the cells of interest and measurements were recorded 3 minutes after
initiation of staining. Based on different scatter properties, nonsperm events
gated out and the incorporation of fluorescein-conjugated dUTP into the sperm,
DNA was quantitated on the FL-1 detector (530 ± 15 nm band-pass
filter). Similarly, processed granulosa cells from atretic and growing-phase
follicles were used as positive control to validate the methodology (for
protocol details, see Hendriksen et al,
2003).
Detection of Sperm Cell-Membrane Integrity
Semen samples were pelleted (3 minutes at 1000 x g), washed
and diluted in PBS to a concentration of 10 x 106 cells/mL
and then stained for 18 minutes at 38°C with a combination of 2.5 nM
Mitotracker red (chloromethyldihydroX-rosamine), 25 nM Yo-Pro-1, and 10
µ/mL PNA-FITC (Molecular Probes Europe, Leiden, The Netherlands). Yo-Pro-1,
a membrane-impermeable DNA dye, was used to detect plasma membrane
deterioration, PNA-FITC (a lectin that specifically binds to the glycosylated
proteins of the outer acrosomal membrane) was used to detect acrosome leakage
(Flesch et al, 1998), and
Mitotracker red (a dye that specifically intercalates in the polarized inner
mitochondrial membrane) was used to detect functional ATP-producing
mitochondria (Flesch et al,
1998; Gadella and Harrison,
2002). After staining, the samples were subjected to FCM
(FACSVantage; Becton Dickinson, San Jose, Calif). Only sperm-specific scatter
events were collected and nonsperm events were gated out for analysis.
Mitotracker red fluorescence was detected on FL-3 (>630 long-pass filter)
while Yo-Pro and/or PNA-FITC fluorescence was detected on FL-1 (530 ±
15 nm band-pass filter). Mitotracker red positive sperm events indicated cells
with active inner membrane-polarized mitochondria, whereas cells with reduced
Mitotracker red fluorescence were those whose mitochondria had lost their
inner transmembrane potential. Viable (plasma membrane and/or
acrosome-intact)cells were Yo-Pro-1 and/or PNA-FITC negative, whereas
degenerated cells were Yo-Pro-1 and/or PNA-FITC positive; see also Gadella and
Harrison (2002).
Collection of Oocytes
Bovine ovaries were collected at a slaughterhouse and were stored in a
thermo-flask during transportation to the laboratory within 1 hour. Excised
ovaries were rinsed in physiological saline (0.9% NaCl) containing antibiotics
(100 IU penicillin and 100 mg streptomycin per mL) and dried with a paper
towel. Cumulus-oocyte complexes (COCs) were aspirated from small antral
follicles (2-8 mm diameter) using an 18-gauge needle attached to a tube in
line with a vacuum pump. COCs were selected on the presence of a multilayered
compact cumulus investment and homogeneous ooplasm and randomly assigned to
the various treatments.
In Vitro Maturation, Fertilization, and Embryo Culture
Selected COCs were rinsed in HEPES buffered M199 (Gibco BRL, Paisley,
United Kingdom) supplemented with 10% fetal calf serum (FCS) and, in groups of
35, were randomly allocated to each well of 500 µL of culture medium M199
(Gibco, Life Technologies, Breda, The Netherlands) supplemented with 10% FCS
and 0.05 units/mL recombinant human FSH (Organon, Oss, The Netherlands). After
24 hours of maturation, COCs were transferred to IVF medium (Fert-TALP) as
described by Parrish et al
(1988) and modified by Izadyar
et al (1996). For IVF,
frozen/thawed spermatozoa were first irradiated and, along with nonirradiated
control samples, were centrifuged over a Percoll gradient for 30 minutes at
700 x g at 25°C as described by Somfai et al
(2002). Sperm (final
concentration 0.5 x 106 spermatozoa/mL), heparin, and PHE
(penicillamin, hypotaurine, and epinephrine) were added to each well of IVF
medium, as we have described previously
(Fatehi et al, 2002). After 20
hours of incubation, the presumptive zygotes were freed from cumulus cells by
vortexing for 3 minutes and placed in a coculture system of 0.5 mL M199
supplemented with 10% FCS on a monolayer of buffalo rat liver (BRL) cells, as
described by Izadyar et al
(1996). The BRL cells were
separated from the BRL cell line from the American Type Culture Collection and
were cultured in a 1:1 mixture of Ham F12 medium and Dulbecco modified Eagle
medium (Gibco) supplemented with 7.5% FCS (Gibco) and antibiotics. On the
fourth and eighth days of culture, embryos were transferred to fresh coculture
wells.
Assessment of Embryo Development
Embryos were scored morphologically and evaluated for 1) the percentage of
cleaved embryos 4 days after fertilization; 2) the percentage of blastocysts
on days 7, 9, and 11; and 3) the percentage of hatched blastocysts on day 11,
expressed on the basis of the number of oocytes at the onset of culture.
Embryo Analysis
Groups of 5 fertilized oocytes were cultured until day 7, as described
above, and were subsequently permeabilized by incubation for 15 minutes in a
glycerol-based microtubule-stabilizing solution at 37°C
(Simerly and Schatten, 1993)
and then fixed at 4°C in PBS containing 3% (w/v) paraformaldehyde
overnight. The next day, embryos were washed in PBS containing 150 mM glycine
and 0.1% (w/v) BSA 3 times. Microtubules were labeled by incubating the
embryos with a monoclonal anti-tubulin antibody (Sigma, T-5168) diluted in
PTBA (PBS containing 0.5% [v/v] Triton X-100, 0.1% [w/v] BSA, and 0.02% sodium
azide) for 90 minutes at 37°C. The embryos were washed 3 times in PBTA,
blocked in PBTA containing 0.1 M glycine and 1% (v/v) goat serum, and were
incubated for 1 hour in PBTA with a TRITC-labeled goat anti-mouse antibody
(Sigma, T-5393) diluted 1:100. Subsequently, the embryos were washed 3 times
in PBTA and 2 times in PBS and then were incubated with 5 µM ToPro-3
(Molecular Probes Europe) in PBS and washed 2 times with PBS. ToPro-3 is used
to detect nuclear DNA fragmentation and formation of apoptotic bodies. Tubulin
staining is an indirect immunostaining to detect spindle formation and
aberrations thereof, which could mark the mitosis. Stained embryos were
mounted on glass slides using antifading mounting medium (Vectashield, Vector
Lab, Burlingame, Calif) and were examined under an inverted spectral confocal
laser scanning microscope (Leica TSC-SP; Leica, Mannheim, Germany) equipped
with a Kr laser (the 568-nm laser line was use for excitation of tubulin) and
an He/Ne laser (the 628-nm laser line was used for ToPro-3 excitation). The Ar
laser (488 nm) was used for bright-field images. Optical sections were made
with discrete steps of 1 µm in depth and pinhole set at 1 µm (oil
immersion objective of 40x). The images were averaged by accumulating 4
images per section either in a nonzoomed or in a 40x zoomed mode.
Extended focus images were made in TCS software (Leica).
Statistics
All the experiments consisted of at least 3 independent experimental runs.
The percentages of fertilization, cleavage rate, blastocyst formation, and
fragmented embryos were compared using the chi-square test. Differences
between means were judged by 1-way analysis of variance (ANOVA). Values are
indicated as mean ± SEM. Significant differences were defined as
P < .05. Linear (Pearson) regressions of data were performed and
the mean slope ± SE as well as the mean calculated point of
intersection with the y-axis ± SE are indicated. The P values
for significance of difference with a slope of 0 are also calculated from the
linear regression data.
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Results |
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Effects of Irradiation on Sperm DNA Integrity
In contrast with the absence of sperm-membrane deterioration, irradiation
caused significant and dose-dependent damage to sperm DNA. The lowest dose of
irradiation (0.6 Gy) already caused significant DNA damage in 25% of the sperm
cells, as detected by TUNEL (Figure
2A). Increasing doses of irradiation resulted in a further
increase in the number of sperm cells with DNA damage and, at 10 Gy, 75% of
the sperm cells were TUNEL positive (Figure
2A).
The result of sperm irradiation on DNA damage, detected by AO staining and FCM (the amount of single-stranded DNA) is illustrated in Figure 2A. Two-dimensional dot plots are all events including yolk particles that are present in frozen-thawed sperm specimens. The yolk particles vary in size and do not contain DNA; nevertheless, because of their affinity for AO, they provide a nice diagonal calibration pattern (for FCM, it is not required to remove unbound stain). The pattern of yolk particles is used as a reference signal to visualize the fluorescence ratio change in AO staining of the sperm cells. Each panel depicts 10 000 sperm events (plus a varying amount of yolk particles).
The ratio FL-1/FL-3 signifies the scale of single-stranded DNA, which, in nonirradiated sperm cells, equals 0.41 ± 0.03 (see Figure 2B). At the lowest dose of 0.6 Gy irradiation, this ratio was almost doubled (0.74 ± 0.04), which indicates significant DNA damage. Increasing the dose of irradiation resulted in a higher FL-1/FL-3 ratio up to 1.03 ± 0.06 at 10 Gy (Figure 2B). The DNA damage highly and significantly correlated with the logarithm of the irradiation dose used (P < .005 for TUNEL and P < .001 for AO ratios), demonstrating that x-ray irradiation induced sperm DNA damage in a dose-dependent fashion (Figure 2B and C).
Effects of Sperm Irradiation on Fertilization Rate and Embryo Development
Irradiated bull sperm (0-10 Gy) were used in IVF of in vitro-cultured
bovine oocytes. IVF with nonirradiated sperm as control resulted in a cleavage
rate of 68% ± 2.7% on day 4
(Table). Irradiation doses of
0.6-5 Gy to sperm samples did not significantly reduce the cleavage rates.
Only the highest dose of 10 Gy caused a significant decrease in cleavage rate
at day 4 (46% ± 6.6%, n = 3;
Table).
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The embryo development of fertilized oocytes was followed up to 11 days. Blastocyst formation at days 7, 9, and 11 was assessed and hatching of the embryos from the zona pellucida was recorded at day 11. IVF results with nonirradiated sperm were comparable with those described in the literature (Beker-van Woudenberg et al, 2004). In sharp contrast with the absence of an effect of sperm irradiation on the day-4 cleavage rate, further embryo development was significantly reduced and, at irradiation doses greater than 2.5 Gy, even blocked (Table). Significant effects were already noted at 1.25 Gy of irradiation (P < .01). Hatching rates were even more affected by irradiation and were almost half of control values at the lowest irradiation dose (0.6 Gy) (Table).
At day 7, embryos were stained by ToPro-3 and tubulin. In total, 24 embryos from the irradiated group and the same number from the nonirradiated group were imaged for morphology, as well as for tubulin and DNA organization. This was done to estimate the rate of apoptosis in the developing embryos with the notion that, during the execution phase of apoptosis, tubulin polymers will become disorganized and DNA as well as the cell nucleus will fragment (for reviews, see Nagata et al, 2003; Scovassi et al, 2003). Without exception, all embryos derived from IVF of irradiated sperm underwent apoptosis at the 4-8-cell stage, resulting in fragmentation and condensation of their DNA into apoptotic bodies (see Figure 3A through J). In half (11 of 24) of the embryos, apoptosis was complete (DNA fragmentation and apoptotic bodies with no sign of spindle formation, indicating the absence of mitosis; Figure 3A through D), while in the other half (13 of 24), signs of apoptosis, along with the spindle formation (mostly aberrant), in different parts of the embryos were detected (Figure 3E through J).
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Discussion |
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In our view, these results demonstrate that the damaged sperm DNA does only affect embryo development after initiation of embryonic gene expression. It is well known that the first cleavages of the embryo are not dependent on the embryonic genome but on a large mRNA pool, which is already present in the cytosol of the unfertilized oocyte. We show for the first time that, in a bovine IVF model, functional sperm with damaged DNA indeed normally fertilize the oocyte and have no significant effect on the first cleavages of the fertilized oocyte.
In bovine embryos, gene transcription does not initiate at the 1- or 2-cell stage but a major burst of gene expression occurs at the 8-16-cell stage (Viuff et al, 1996). Our study suggests that, at the time of onset of embryonic gene expression (4-8-cell stage), the paternal DNA damage becomes sensible for the apoptotic machinery of the early embryo and, by blocking the mitosis, it arrests further embryonic development. The study also shows that repair of paternal DNA damage by the fertilized oocyte is relatively limited and, when the sperm cells were irradiated with doses greater than 1.25 Gy, repair of damaged paternal DNA was almost impossible.
Effects of Irradiation on Sperm Cells
Our results show that irradiation with gamma or x-rays did not impair sperm
functioning but induced DNA damage. These results are supported by Ahmadi and
Ng (1999), who showed that
irradiation of mouse sperm rendered functional and motile sperm with signs of
DNA damage. In extension, we showed that irradiated bovine sperm did not show
any signs of membrane deterioration and that this IVF of bovine oocytes with
this treated sperm resulted in normal cleavage rates but caused complete
apoptosis at the 4-8-cell stage of the embryo and thus blocked further embryo
development. Exposure of somatic cells to ionizing radiation results in DNA
and chromosomal damage (Cleaver,
1983; Carrano,
1986). Irradiation of sperm cells, however, does not induce great
structural chromosomal aberrations in spermatozoa
(Tateno et al, 1996) and DNA
is the principle molecular target of irradiative damage, causing cell-killing
mutations and carcinogenesis (Painter,
1980; Makrigiorgos et al,
1990). DNA damage often results in cellular apoptosis in which the
cell nucleus fragments into apoptotic bodies that bleb from the plasma
membrane and are phagocytosed by neighboring cells
(Gamen et al, 2000;
Yamazaki et al, 2000; Nagata et al, 2003; Scovassi
et al, 2003). Sperm cells do not have any gene expression, as DNA
transcription is completely silenced, and therefore contain a minimal amount
of RNA (O'Brien et al, 1994).
Sperms cells also lack ribosomes and an endoplasmic reticulum, which is
required for eventual RNA translation
(O'Brien et al, 1994).
Therefore, de novo protein synthesis is absent in mature sperm cells. In fact,
mature sperm cells have been reported to lack caspases
(De Vries et al, 2003) and
fail to induce apoptosis (Gadella and
Harrison, 2002; De Vries et
al, 2003). Expression of active caspases is the general way to
execute apoptosis, which is manifest in cells showing nuclear fragmentation
(Spanos et al, 2002). The
absence of protein synthesis and of a proper apoptotic machinery explains why
the specific sperm DNA damage imposed by irradiation did not result in the
induction of depolarization of the mitochondrial inner membranes or
facilitated phospholipid scrambling (both signs of apoptosis;
Gulbins et al, 2003;
Kaina, 2003). In analogy, it
should also be mentioned that DNA damage cannot be repaired by the sperm cell
(Chandley and Kofman-Alfaro,
1971; Gledhill and
Darzynkiewicz, 1973).
In this study, DNA damage was artificially induced by the irradiation, but abnormalities in sperm DNA are also well documented in sub/infertile clinical cases: only 0.1% of the sperm cells from healthy fertile males contain damaged DNA (Baccetti et al, 1996). This proportion dramatically increases in patients with varicocele and round-headed sperm to 10% and in men with cryptorchidism reaches to 15%-20%. In unexplained infertility cases, up to 25% of the sperm cells have DNA damage. Testicular seminoma results in DNA damage in up to 50% of the sperm cells. Moreover, asthenozoospermia has been shown to coincide with high levels of chromatin abnormalities (Lopes et al, 1998).
Although the extent of sperm DNA damage is closely related to male infertility (Aitken, 1999; Sakkas et al, 1999), its origin is still largely controversial. DNA damage can be caused by improper packaging and ligation during spermatogenesis and epididymal sperm maturation (Sailer et al, 1995). Oxidative stress by elevated reactive oxygen species (ROS) may also cause DNA damage. Higher levels of ROS and thus of induction of sperm DNA damage can be expected by genital infections (leukocytes produce high amounts of ROS; Jarow, 1998; Moustafa et al, 2004) and in smoking patients (Shen et al, 1997; Jarow, 1998). Sperm cells have highly condensed DNA that is densely packed to protamines, replacing histones (Oliva and Dixon, 1991). Effects on reduced DNA protamination and DNA fragmentation have been observed resulting in poor semen quality (Aitken et al, 1993; Gomez and Aitken, 1996) and increased risks of birth defects.
Effects of Sperm DNA Damage on Fertilization and Early Embryonic Development
Besides the normal morphology, motility, and membrane integrity, the
irradiated sperm cells were also able to fertilize the oocyte and the
fertilized oocyte can develop normally to 2- and 4-cell embryo stages,
apparently after pronucleus formation and syngamy of the 2 haploid pronuclei.
However, in contrast with IVF with nonirradiated sperm cells, fertilization
with irradiated sperm resulted in apoptosis of the embryonic cells after the
second and third cleavage and thus blocked further embryo development. This
was evidenced by a failure of blastocyst formation, nuclear fragmentation into
apoptotic bodies, and by failures in spindle formation. At low irradiation
doses, apoptosis was not induced at a significant level despite the fact that
DNA damage was observed. This may indicate that the oocyte is able to repair
damaged paternal DNA up to a limited degree. Indeed, somatic cells
(Sancar and Sancar, 1988), in
contrast with sperm cells (Chandley and
Kofman-Alfaro, 1971; Gledhill
and Darzynkiewicz, 1973), have a nucleotide excision repair
mechanism that is used to remove and replace affected bases and correct the
effect of irradiation. Oocytes do share with somatic cells an operative
excision repair capacity (Masui and
Pedersen, 1975), which can also repair sperm DNA that was damaged
before fertilization (Generoso et al,
1979; Brandriff and Pedersen,
1981). The oocyte repair mechanism is limited and, in our study,
sperm DNA damage induced by irradiation doses of greater than 1.25 Gy could
most likely not be repaired. The irradiated sperm were still able to fertilize
and to induce early embryo development. After 2-3 embryo cleavages, abnormal
spindle formation as well as nuclear fragmentation took place and resulted in
a developmental block of the embryos prior to blastocyst formation. The
formation of a functional mitotic spindle requires coordination between
centrosomes and chromosomes (Kellogg et
al, 1994; Zhang and Nicklas,
1995). In most mammalian species, including humans, the sperm
contributes a centriole during fertilization of an oocyte and helps to
organize the assembly of the first mitotic spindle in the zygote
(Navara et al, 1995). In fact,
spindle formation during the first 2 cleavages of the fertilized oocytes was
normal, suggesting that the sperm centriole remained functionally intact. An
electron microscopy analysis can challenge this hypothesis. Furthermore, the
nongenomic sperm factors involved in activation of the fertilized oocytes
remained functionally competent.
The induction of apoptosis at the 4-8-cell stage of the embryo coincides exactly with the developmental phase where embryonic gene expression is initiated (De Sousa et al, 1998; Bordignon and Smith, 1999). During the first cleavages, the fertilized oocyte translates RNA that was stored in the oocyte prior to ovulation (Gandolfi and Gandolfi, 2001). Our results further demonstrate that the paternal genome is not involved in the first 2 cleavages, suggesting that a functional apoptotic pathway is suppressed in early embryos and that these embryos are more resistant to induced apoptosis than blastocysts (Weil et al, 1996). In this respect, we believe that enzymes involved in DNA and nuclear fragmentation are not operative in the oocyte and only become operative after the second cleavage of the embryo. DNA fragmentation normally is initiated by effector caspases 3, 6, 7, which become activated during the execution phase of cell apoptosis (for reviews, see Nagata et al, 2003; Scovassi et al, 2003). Briefly, the nucleus condenses and falls apart into apoptotic bodies due to caspase 6-mediated breakdown of the nuclear lamina (proteolysis of lamin) and to other effector caspases involved in degrading poly(ADP-ribose) polymerase activating nucleases and DNases.
The DNA repair machinery has also been shown to be defective in arrested germ cells and in spontaneously aborted embryos (Spandidos et al, 1998; Nudell et al, 2000). It is therefore well possible that more subtle DNA aberrations in sperm insufficient to induce gross responses in cell cycle arrest and apoptosis (like shown in this study) may be expressed in a later embryonic or even postnatal phase (Hales and Robaire, 1990; Sakkas et al, 2000).
The development of new ART and especially of IVF and ICSI has improved infertility treatments. However, IVF as well as ICSI overcome a number of factors normally regulating the fertilization of the oocytes by more directly introducing the male haploid genome of 1 sperm cell into the oocyte. Concern has been expressed that ART (and especially ICSI) bypasses natural sperm selection and could inadvertently introduce a defective paternal genome into the oocyte (Campbell and Irvine, 2000; Hargreave, 2000a,b,c; Mortimer, 2000; Sakkas et al, 2000). Recent extensive human case studies essentially showed that conventional IVF and ICSI are associated with similar frequencies of congenital malformations (Devroey and van Steirteghem 2004; van Landuyt et al, 2005), but these abnormalities were about twice as high when compared with naturally conceived infants (Hansen et al, 2002). Despite these figures, utilization of ART has rapidly increased over the past decades. The introduction of new procedures brings with it a responsibility to evaluate and monitor short- and long-term outcomes of the intervention (Kondro, 1993; Buitendijk, 1999; Doyle and McNeilly, 1999). Our study may provide new insights in how aberrant paternal DNA introduced by ART into the oocyte causes aberrations in embryo development.
In consequence of our study, it may be recommended to follow embryo development of oocytes fertilized by ART up to a blastocyst stage before embryo transfer into the uterus. Although this option has been proposed by some IVF clinics, it is not a common practice and, due to the lack of adequate media that could support in vitro embryo development in early stages, day 2 transfer has been a dominant procedure (Milki et al, 2000; Karaki et al, 2002). Prolonging the duration of culture may allow chromosomally competent embryos to develop to the blastocyst stage and permit the selection of embryos that have the potential of continued development under embryonic genome control rather than embryos that carry aberrant (paternal) DNA. Further studies are required to establish whether ART leads to an increase in developmental failure of embryos at the preblastocyst stage.
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