Developmental capacity of damaged spermatozoa

Ali Ahmadi and Soon-Chye Ng1

Department of Obstetrics and Gynaecology, National University of Singapore, Lower Kent Ridge Road, Singapore


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We report the first detailed and systematic study in a mammalian system to unravel the mystery of the beginnings of life. The fertilizing ability of damaged spermatozoa at various levels of disintegration (cellular and molecular) has been investigated in homologous (mouse) and heterologous (human spermatozoon, hamster oocyte) models. Live pups were produced after destruction of spermatozoa at various cellular and molecular levels followed by injection into oocytes. We demonstrate that with damaged spermatozoa, the key point in the fertilization process is the activation of the oocyte by injection of cytosolic sperm factor. A similar fertilization rate as that using live intact spermatozoa can be achieved following activation. However, the integrity of the genetic material influenced in-vitro development of the embryos and live fetuses. This study contributes to a better understanding of the fertilizing ability of damaged spermatozoa. These findings can be applied clinically to patients with necrozoospermia or very severe oligozoospermia and in wildlife research where damaged spermatozoa from rare species can be used to regenerate young, and hence propagate the species. Also implied is the possible contribution of sperm DNA strand breakage to early pregnancy loss.

Key words: apoptosis/cytosolic sperm factor/DNA damage/necrosis/oocyte activation


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
A problem commonly encountered in severe male factor patients is the low rate of viability of spermatozoa. Necrozoospermia, a poorly documented cause of male infertility, is defined as a condition in which spermatozoa in the ejaculated semen are apparently dead, occurring in 0.2–0.48% of infertile couples (Phadke et al., 1978Go). Fertilization using initially immotile and presumably `dead' spermatozoa is reported to be very low (Dozortsev et al., 1995Go; Hoshi et al., 1995Go; Nagy et al., 1995Go). However, there are only two reports (in cow and human) of live offspring after injection of immobilized, killed spermatozoa (Goto et al., 1990Go; Hoshi et al., 1994Go). A systematic study of this possibility is therefore of great importance both academically and clinically.

It has been generally accepted that apoptosis and necrosis are two distinct modes of cell death. Apoptosis, or programmed cell death, is an active and physiological mode of cell death, in which the cell itself designs and executes the programme of its own demise and subsequent disposal. Early changes during apoptosis are loss of intracellular water and increase in the concentration of ionized calcium in the cytoplasm (Fesus et al., 1991Go). Chromatin condensation, followed by nuclear disintegration and formation of apoptotic bodies represent other typical features of apoptosis. The integrity of the plasma membrane is preserved to the late stages of apoptosis.

Necrosis, in contrast, is a passive, catabolic and degenerative process. It generally represents a cell's response to gross injury and can be induced by an overdose of a cytotoxic agent. The early events of necrosis include swelling of mitochondria as well as swelling of the whole cell, combined with marginal chromatin condensation, followed by rupture of the plasma membrane and release of cytoplasmic constituents (Majno and Joris, 1995Go). DNA degradation is not so extensive during necrosis as in the case of apoptosis.

The aim of this investigation is to study the developmental capacity of spermatozoa at various levels of disintegration. Recently we have shown that fertilization and embryonic development can be achieved by spermatozoa at an early stage of disintegration in which only plasma membrane, mitochondria, or protamine have been lost (Ahmadi and Ng, 1997aGo,bGo; 1999aGo,bGo1999a,b). On the other hand, apoptotic spermatozoa can fertilize the oocyte, but embryonic development is strongly related to the degree of DNA damage (Ahmadi and Ng, 1999cGo). The key factor involved in bringing about fertilization by damaged spermatozoa is activation of the oocyte, as shown here, by the co-injection of cytosolic sperm factor (CSF) and spermatozoa into the oocyte (Dale et al., 1985Go; Stice and Robl, 1990Go; Swann, 1990Go). Although fertilization can be achieved using damaged spermatozoa, development of the resultant embryo and live fetuses is related to the integrity of the genetic material.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We used EBSS medium in the hamster-ICSI assay experiment (Ahmadi et al., 1996aGo) and Whittingham's T6 medium in the mouse experiment.

Preparation of spermatozoa
Frozen-thawed human semen samples from proven donors were used for this study. The spermatozoa were processed by a discontinuous Percoll gradient (40, 70, 90%) separation method (Ahmadi et al., 1996bGo). C57BLxCBA (F1 hybrid) male mice 10–12 weeks old were used as sperm donors. Recovery of spermatozoa from the epididymides of mice has been described in detail previously (Ahmadi et al., 1995Go).

Disintegration at the cellular level
Sperm samples were twice frozen (without cryoprotectant) to –80°C and thawed, and then stored at –80°C for at least 1 month. With this treatment all cellular components including mitochondria and membrane were destroyed. Disintegration of the cell was confirmed by staining with 0.1 µmol/l rhodamine 123 (for study of integrity of mitochondria) for 30 min followed by counterstaining with 1 µmol/l propidium iodide (for evaluation of plasma membrane integrity) for 15 min. Live spermatozoa fluoresced green in the midpiece region, while damaged spermatozoa fluoresced red in the head region.

Induced damage at the molecular level
Nuclear protein (protamine) level
The spermatozoa were treated with 5 mmol/l dithiothreitol (DTT) for 30 min before the freezing-thawing procedure. In addition to cellular death, there was destruction of the disulphide bonds of the protamine, confirmed by acridine orange staining. A 10 µl sample of sperm suspension was mixed with 20 µl of a low-pH detergent solution (0.15 mol/l NaCl, 0.08 N HCl, 0.01% Triton X-100, pH 1.4). After 30 s, 60 µl of acridine orange staining solution (6 µg/ml acridine orange in 0.2 mol/l Na2HPO4, 1 mmol/l disodium EDTA, 0.15 mol/l NaCl, 0.1 mol/l citric acid monohydrate, pH 6) was added, and 10 µl of sample was placed on the slide and covered with a coverslip. About 250 spermatozoa per sample were evaluated with a fluorescence microscope. The nuclei of spermatozoa with intact protamine fluoresced green, while those with damaged protamine fluoresced orange-red.

DNA
In another experiment, spermatozoa were exposed to gamma radiation (5 Gy and 10 Gy) prior to DTT treatment, freezing and thawing (`totally damaged'). Induction of DNA damage was evaluated by the terminal deoxynucleotidyl transferase (TdT)-mediated dUPD nick-end labelling (TUNEL) test using an Apoptag Plus kit (Oncor, Gaithersburg, MD, USA). The samples were stained according to the instructions and DNA strand breaks were evaluated by flow cytometry. The presence of DNA strand breaks was detected by labelling the 3' OH termini in DNA breaks with digoxigenin-conjugated nucleotide, in the enzymatic reaction catalysed by exogenous TdT. The 3'-OH ends of the DNA strand breaks serve as primers for the incorporation of biotin- or digoxigenin-16-dUTP, which is detected through the use of fluoresceinated avidin (Gorczyca et al., 1993Go). TdT enzyme adds multiple biotin- or digoxigenin-16-dUTP molecules per single DNA strand break, and the length of the synthesized polynucleotides is determined by the ratio of monomer (digoxigenin-16-dUTP) to primer (3'-OH end) (Chang and Bolum, 1986Go).

Cytosolic sperm factor (CSF) preparation
The mouse and human spermatozoa were washed with Dulbecco's phosphate-buffered saline containing 0.1 g/l MgSO4,6H2O, 0.2 g/l KCl, 0.2 g/l KH2PO4, 8.0 g/l NaCl, and 1.15 g/l Na2HPO4 (DPBS, Sigma Chemical Co, St Louis, MO, USA). The suspension was then lysed by two freeze-thaw cycles in liquid nitrogen and was kept for 1 h at room temperature. The sample was centrifuged at 1600 g for 10 min to remove sperm cell remnants. The supernatant was further centrifuged at 100 000 g for 1 h at 4°C. The clean supernatant was collected as cytosolic sperm factor and aliquoted and kept at –80°C until used. Just before injection the cytosolic sperm factor was added to sperm-polyvinylpyrrolidone to obtain a final concentration of about five and ten sperm equivalents in 10 pl for human and mouse respectively.

Preparation of oocytes
Frozen-thawed hamster oocytes were used for this study as described in detail elsewhere (Ahmadi et al., 1996bGo).

C57BLxCBA (F1 hybrid) female mice 4–5 weeks old were used as oocyte donors in this experiment. Ovulation induction in mice has been described elsewhere (Ahmadi et al., 1995Go). Oocytes were retrieved from the oviducts 18 h post-HCG and they were freed of cumulus by treating with 0.1% hyaluronidase.

Microinjection procedure
The holding and microinjection pipettes were made by drawing glass capillary tubes with the horizontal Sutter puller (P-87, Sutter Instruments, Novato, CA, USA), and were further processed on a Narishige microgrinder (EG-4, Narishige, Tokyo, Japan) and De Fonbrune microforge (MF-1, Technical Products International Inc., St Louis, MO, USA). Injection pipettes had an internal diameter of 5–6 µm and outer diameter of 7–8 µm in hamster intracytoplasmic sperm injection (ICSI) experiments. These ranged from 7 to 8 µm and 9 to 10 µm for mouse experiments respectively. The injection pipette had a bevel angle of 40 with a sharp spike on the tip. The holding pipettes had an outer diameter of 80 µm and an inner diameter of 40 µm.

Microinjection and oocyte holding systems were controlled pneumatically using a simple air-based system (Ahmadi et al., 1996bGo). We connected a 3 ml syringe to the pipette holder using Teflon tubing. ICSI was carried out on the 37°C heated stage of a Zeiss Axiovert 135 inverted microscope mounted with a fully motorized Eppendorf micromanipulator (Netheler-Hinz, Hamburg, Germany). The details of the procedure have been described elsewhere (Ahmadi et al., 1995Go, 1996aGo).

Evaluation of oocytes and culture of embryos
In the hamster-ICSI assay experiment, the oocytes were checked 18–20 h after injection for pronucleus formation and fixed in 1% glutaraldehyde, stained with aceto-orcein and examined for sperm head decondensation and male pronuclei formation under x100 magnification. The state of the sperm nucleus within the cytoplasm of oocyte was classified into five stages (Ahmadi and Ng, 1997cGo): (i) intact sperm nucleus; (ii) decondensing sperm nucleus; (iii) completely decondensed sperm nucleus; (iv) round sperm nucleus enclosed within a nuclear membrane; (v) an enlarged nucleus with nucleoli; Ab = abnormal decondensation.

The mouse oocytes were checked 6–8 h after ICSI for fertilization. Oocytes with two pronuclei and a second polar body were considered to be fertilized. Oocytes with one pronucleus and a second polar body were considered as activated oocytes. The fertilized oocytes were separated from the rest and allowed to develop to the blastocyst stage. The blastocysts were transferred to the uterine horns of recipients on day 3 of pseudopregnancy. The recipients were autopsied on day 16–18 of gestation and the number of implantation sites and fetuses were recorded. One female recipient in each group was allowed to proceed to term.

Statistical analysis
All data were analysed by analysis of variance. The calculations were performed by one-way analysis of variance (ANOVA) using the SPSS package computer software.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Suspensions of mouse and human spermatozoa were subjected to different treatments, inducing damage at the various (cellular and molecular) stages before injection into oocytes (Table IGo). For cellular damage, spermatozoa were frozen-thawed without using a cryoprotectant and kept at –80°C for at least 1 month. At the molecular level we focused on nuclear proteins (protamine) and DNA. In order to induce nuclear protein damage the sperm suspensions were treated with DTT prior to freezing and thawing. Acridine orange staining revealed that protamine was destroyed in all spermatozoa after DTT treatment (data not shown). For induction of DNA damage, the spermatozoa were exposed to gamma-radiation at the doses of 5 and 10 Gy prior to DTT and freeze-thaw treatments (`totally damaged'). DNA damage was evaluated by the TUNEL test (Figure 1Go).


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Table I. Overview of experiments involving damaged spermatozoa
 


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Figure 1. Detection of DNA strand breaks using TUNEL test in irradiated spermatozoa. The intensity of FITC labelling of cells with DNA strand breaks increases with radiation dose.

 
Human sperm head decondensation and male pronucleus formation by damaged spermatozoa in hamster oocytes
In order to study sperm head decondensation and male pronucleus (PN) formation, human spermatozoa were injected into the hamster oocytes. Results of experiments in which damaged spermatozoa were injected into hamster oocytes are shown in Table IIGo. The sperm head decondensation and male pronucleus formation rates were 74.5 and 46.6% respectively in live intact sperm group (Experiment 1), while these rates were significantly lower at 36 and 18% respectively (P < 0.01) when the spermatozoa were subjected to freeze-thawing and 1 month storage at –80°C (Experiment 2) prior to ICSI (Table IIIGo). These low rates could be due to the absence of cytosolic sperm factor to trigger the activation process of oocytes. Hence, in the next experiment (Experiment 3), spermatozoa which were subjected to freeze-thawing and 1 month storage were injected together with cytosolic sperm factor (CSF). Sperm head decondensation and PN formation were increased by CSF injection to 83.3 and 51.9% respectively. Hence, for Experiments 4 to 6, the treated spermatozoa were injected with CSF in order to initiate the fertilization process. The sperm head decondensation and male pronucleus formation were 85.3 and 50.9%, 87.8 and 56.6%, and 85.7 and 59.2% in Experiments 4–6 respectively, with no significant differences from controls. However, there was a significant difference in sperm head decondensation and male PN formation between Experiment 2 and those with CSF injection (P < 0.01). The spermatozoa could undergo decondensation and PN formation within the hamster oocytes, in spite of damage to the protamine and DNA, if the oocyte were activated by CSF.


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Table II. Results of intracytoplasmic injection of human damaged sperm into the hamster oocytes
 

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Table III. Sperm head decondensation and male pronucleus formation following injection of damaged human spermatozoa into hamster oocytes
 
Fertilization and in-vitro development of mouse embryos derived from damaged spermatozoa
Our results of fertilization using damaged spermatozoa in the heterologous model (human spermatozoa/hamster oocyte) revealed that the key point in fertilization by damaged spermatozoa is activation of the injected oocytes, i.e. injection of CSF. Furthermore, the integrity of nuclear protein and DNA did not influence sperm head decondensation and male pronucleus formation. Further development was tested using the same approach in a homologous model (mouse) and following embryonic development. The results of experiments in which damaged spermatozoa at various stages of disintegration were injected into mouse oocytes are summarized in Table IVGo. The fertilization rate was 55.1% with live intact spermatozoa (Experiment 1), while this rate decreased to 17.5% when the spermatozoa were frozen-thawed and stored at –80°C (Experiment 2) prior to ICSI. The fertilization rate increased to 62.8% in Experiment 3 when the spermatozoa were injected together with CSF (P < 0.01) when compared to Experiment 2. However, this difference was not statistically significant between Experiment 3 and control (P > 0.01). Fertilization rates were 65.4, 61.3 and 57.1% in Experiments 4–6 in which spermatozoa were treated with DTT (for destruction of protamine) prior to freeze-thawing and 1 month storage, and exposure to 5 Gy and 10 Gy in addition to DTT, freeze-thawing and 1 month storage treatment respectively (P > 0.01). There was a significant difference in fertilization rate between Experiment 2 and all the CSF-injected group (P < 0.01). All of the fertilized oocytes cleaved to 2-cell embryos. Further development to the blastocyst stage was achieved in 58.5% of the cultured 2-cell embryos in the live intact sperm group (Experiment 1). However, this rate decreased to 40% in Experiment 2 when spermatozoa were subjected to freeze-thawing and 1 month storage at –80°C. Injection of these spermatozoa together with CSF (Experiment 3) resulted in an increased blastocyst formation rate to 56.3% (P > 0.01), suggesting that CSF not only was responsible for fertilization, but also supported the in-vitro development of the resultant embryos up to at least the blastocyst stage. The blastocyst formation rate was 52.7, 24.6 and 8.3% in Experiments 4–6 respectively (P < 0.01). This suggested that the protamine integrity of the spermatozoa did not affect development to the blastocyst, but that this development was very much dependent on its DNA integrity.


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Table IV. Fertilization and in-vitro development of mouse oocytes injected with spermatozoa at various stages of disintegration
 
Postimplantation development of mouse embryos resulting from damaged spermatozoa
The implantation rates of blastocysts derived from intracytoplasmic injection of spermatozoa at various stages of disintegration after transfer to recipients are shown in Table VGo. There were no blastocysts transferred in Experiment 2 and Experiment 6 as only a few blastocysts were available. Of 33 transferred blastocysts in the live intact group (Experiment 1), 60.6% (20/33) implanted and 36.4% (12/33) developed to live fetuses. These rates were 54.3% (19/35) and 31.4% (11/35), 51.5% (17/33) and 26.7% (8/33), 54.5% (6/11) and 18.2% (2/11) in Experiments 3–5 respectively. There were no significant differences in implantation sites and rates between all experimental and control groups (P > 0.01). However, there was a significant difference in live fetus development between Experiments 4 and 5 and Experiments 1 and 3; suggesting that the integrity of protamine and DNA affects the live birth rate. Furthermore, five blastocysts were transferred to the uterine horn of pseudopregnant mice in Experiments 1, 3, 4 and 5, which resulted in three (one male and two females), two (one male and one female), two (two males) and one (male) normal pups respectively. Overall rates of development to live fetus and offspring of transferred blastocysts were 39.5% (15/38), 32.5% (13/40), 26.3% (10/38) and 18.7% (3/16) in Experiments 1, 3, 4 and 5 respectively.


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Table V. Implantation and development to live fetuses of mouse blastocysts derived from intracytoplasmic sperm injection (ICSI) of damaged spermatozoa at various stages of disintegration after transfer to pseudopregnant recipients
 

    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Sperm death could be due to primary necrosis that resulted from infection with subsequent inflammation either during passage and storage in the epididymis (epididymal necrozoospermia) or originating from the testis. Epididymal necrozoospermia appears to be caused by either a hostile luminal environment in the epididymis or an inherent structural instability in the spermatozoa (Wilton et al., 1988Go). This may result in rapid degeneration of spermatozoa during passage through the epididymis and production of dead and severely degenerated ejaculated spermatozoa. This problem can be overcome by frequent ejaculation, which reduces the time lag between sperm production and ejaculation. However, when the patient has oligozoospermia and necrozoospermia, frequent ejaculation may result in very low sperm counts in the ejaculate. The alternative could be extraction of spermatozoa directly from the testis by biopsy; but this is an invasive procedure that may not be acceptable to the patient. If the origin of sperm death is in the testis, a logical strategy is to consider fertilization from damaged spermatozoa.

In mammals, including man, germ cell death is conspicuous during spermatogenesis and plays a pivotal role in sperm output. Apoptosis may be the result of hormonal disorders. Presence of hormonally-induced apoptosis has been demonstrated in rodent testis (Nantel et al., 1996Go). Apoptosis is found essentially in the spermatogenesis stages VII–XIV in the rodent and also in the postmeiotic stages of the dogfish (Callard et al., 1995Go). Recently the presence of apoptosis has been reported in human spermatozoa (Gorczyca et al., 1993Go; Baccetti et al., 1996Go). The characteristics of apoptotic spermatozoa are consistent regardless of the pathology. However, the prevalence of apoptosis in spermatozoa varies in various pathologies: it is seen at a rate of 0.1% in fertile controls up to 10% in varicocele, infection (including AIDS) and globozoospermia patients; 20% in cryptorchid men; 25% in unexplained infertility, and 50% in testicular seminoma carriers (Baccetti et al., 1996Go). Although our data show that apoptotic spermatozoa can fertilize oocytes at the same rate as intact spermatozoa, the development of the resultant embryos to blastocyst and to term is very much related to the integrity of the DNA. However, the oocyte has the ability to repair the DNA damage as oocytes fertilized by DNA-damaged spermatozoa did not develop further in vitro when they were cultured in the presence of inhibitors to DNA repair (Ahmadi and Ng, 1999cGo). Oocyte DNA repair capacity is limited and is related to the degree of sperm DNA damage. Damage beyond the capacity to repair will result in fragmentation, which finally leads to degeneration of the embryo. This may be one of the causes of fragmentation in human embryos seen in clinical IVF programmes.

It is known that injection of previously immotile spermatozoa results in either poor or no fertilization (Dozortsev et al., 1995Go; Hoshi et al., 1995Go; Nagy et al., 1995Go; Nijs et al., 1996Go). However, actual necrozoospermia is associated with markedly decreased frequency of embryo formation (Sherins et al., 1995Go). Poor fertilization could be due to the inability of spermatozoa to trigger the activation process of the oocyte. It has been suggested that many fertilization failures and probably implantation failure and early spontaneous abortion after ICSI could be avoided by activating the oocyte when deficient spermatozoa are used (Tesarik, 1998Go). Our results show that the key point in fertilization by damaged spermatozoa is activation of the injected oocytes. This is easily overcome by the injection of cytosolic sperm factor, a natural activator in fertilization, together with the damaged spermatozoa. Spermatozoa introduce substances inside the oocyte following gamete fusion and thereby induce Ca2+ transients without necessarily utilizing signals generated at the egg surface. Dale et al. (1985) were the first to demonstrate that intracellular injection of sperm extract causes the activation of oocyte similar to that seen in the normal fertilization process (Dale et al., 1985Go). Since then overwhelming evidence has suggested that the internally acting factor might be: (i) calcium ions (Creton and Jaffe, 1995Go); (ii) other small molecules such as inositol 1,4,5-triphosphate (IP3) (Iwasa et al., 1990Go); or (iii) a soluble protein (Swann and Lawrence, 1996Go). The CSF is a heat-sensitive protein, and is only active after introduction into the oocyte (Dozortsev et al., 1995Go). Recently it has been shown that cytosolic sperm factor contains a 33 kDa protein called oscillin which is related to a prokaryote hexose phosphate binding protein and responsible for causing the Ca2+ oscillations that trigger egg activation at fertilization in mammals (Parrington et al., 1996Go). The oscillin is located in the equatorial segment of mammalian spermatozoa, the region where the spermatozoon is fused with the oocyte in mammals, and is readily extractable by simple freezing and thawing. It has been demonstrated that oscillin is the mammalian form of glucosamine-6-phosphate deaminase (GNPDA) as cloned oscillin has a robust GNPDA activity and can account for all such activity in mammalian tissue extracts (Wolosker et al., 1998Go). Recombinant GNPDA and GNPDA purified from hamster spermatozoa failed to trigger Ca2+ oscillations when injected into the oocyte. Therefore the calcium-releasing or oscillin activity of sperm extract is due to a substance other than GNPDA. The oscillin reported (Parrington et al., 1996Go) was, in fact, a major contaminant of the active fractions following column chromatography (Swann et al., 1998Go).

However, evidence in non-mammalian species like sea urchin and amphibians suggest that the putative sperm factor is a relatively small, heat-resistant molecule, and active when applied outside the oolemma (Osawa, 1994Go; Iwao et al., 1995Go). It is also incapable of triggering the entire response in unfertilized oocytes. In fact, the sperm extract is heat- or protease-labile, and its biological activity is fully retained in >10 kDa fractions, but not in <10 kDa fractions. Recent evidence revealed that tr-kit, a truncated form of the c-kit tyrosine kinase receptor corresponding to the phosphotransferase portion of the cytoplasmic catalytic domain and the carboxyterminus, could also be a sperm factor inducing the early events of natural fertilization (Sette et al., 1997Go). It is localized mainly in the residual cytoplasm of spermatozoa with maximal accumulation in the midpiece of the flagellum, and may be partially associated with triton-insoluble components of the spermatozoa. Microinjection of recombinant tr-kit protein or synthetic tr-kit RNA is sufficient to trigger the complete set of events in the oocyte activation process including cortical granule exocytosis, pronuclear formation and cleavage stages. Therefore one may conclude that sperm cytosolic factor is likely to contain more than one natural Ca2+ oscillation-inducing molecule.

By activating oocytes with CSF, the damaged human spermatozoa could decondense and develop to pronuclei within hamster oocytes at the same rate as live intact spermatozoa. Fertilization is not related to the integrity of either the cellular component (except CSF) or the genetic material. The next question as to whether development of resultant embryos in vitro and in vivo is possible, was tested in the mouse homologous model. Fertilization was similar when damaged spermatozoa at the various stages of disintegration were injected into the mouse oocyte. Furthermore, activated oocytes with CSF had the same blastocyst development rate as live intact spermatozoa, suggesting that CSF is not only responsible for the activation of oocyte in the fertilization process but also supports in-vitro embryonic development up to blastocyst stage. However, the ability to develop to the blastocyst was reduced when there was DNA damage in the spermatozoa. Furthermore, development of live fetuses was lower when there was damage to protamine and DNA although the implantation rate was not significantly different. Hence when there is genetic damage, though some embryos may reach to the blastocyst stage, natural selection ensures that most of them would abort and not go to term. Currently, we are unable to select individual spermatozoa with intact DNA that can be injected to the ooplasm, though this may be possible in the future. Infrared technology may be useful for this purpose; this technology was first proposed by Lua and Ng as a possible method of selecting viable and genetically intact embryos (Lua and Ng, 1987Go). More research should be done into the selection of genetically intact cells.


    Acknowledgments
 
This investigation was funded by Research Grant No: RP3940336 of the National University of Singapore. We are grateful to Professor Alan Trounson of the Centre for Early Human Development, Monash Medical Centre for his support. We also thank the staff of the micromanipulation, IVF and Andrology laboratories for their help. We also would like to extend our thanks to staff of the Gamma Radiation Unit of the Physics Department and Flow Cytometry Laboratory of the National University Medical Institutes for their assistance.


    Notes
 
1 To whom correspondence should be addressed Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
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Submitted on December 7, 1998; accepted on May 24, 1999.