Comparison of oocyte activation and Ca2+ oscillation-inducing abilities of round/elongated spermatids of mouse, hamster, rat, rabbit and human assessed by mouse oocyte activation assay

Hiroyuki Yazawa1, Kaoru Yanagida, Haruo Katayose, Syoutaro Hayashi and Akira Sato

1 Department of Obstetrics and Gynecology, Fukushima Medical University, Hikarigaoka Fukushima, Fukushima 960-1295, Japan


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Oocyte activation and Ca2+ oscillation-inducing abilities of round spermatid (ROS) and elongated spermatid (ELS) of some rodents and human were assessed by their injection into mouse (B6D2F1) oocytes (mouse test). With mice (B6D2F1, ICR) and rat, ROS displayed no oocyte activation or Ca2+ oscillation-inducing abilities. Although ELS could induce activation at 87, 86 and 31% of injected oocytes respectively, most of the intracellular calcium concentration ([Ca2+]i) responses of ELS-injected oocytes did not show oscillation patterns; only several transient [Ca2+]i rises (transient pattern) were seen. Similarly, with hamster, rabbit and human, while ROS could induce oocyte activation efficiently (70, 71 and 52% respectively), most of the [Ca2+]i patterns of injected oocytes were transient patterns, and not oscillation patterns. When ROS nuclei only from these latter species were injected into mouse oocytes, most of the oocytes could not be activated. [Ca2+]i patterns of oocytes injected with immature sperm cells changed from transient pattern to oscillation pattern while the cells were maturing into spermatozoa. With hamster ROS, oocyte-activating factor was found to be distributed mainly in the cytoplasm. It was interesting that there is a dissociation between the timings of appearance of oocyte activation and that of Ca2+ oscillation of oocytes injected with developing immature sperm cells.

Key words: human/intracellular calcium oscillation/intracytoplasmic spermatid injection/oocyte activation/rodents


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Since the time that pregnancies were first reported after intracytoplasmic sperm injection (ICSI) (Palermo et al., 1992Go), this technique has become increasingly popular in clinical therapies for infertile couples affected by severe male factors (Van Steirteghem et al., 1993Go; Nagy et al., 1995Go; Palermo et al., 1995Go). In cases of obstructive or non-obstructive azoospermia, epididymal or testicular spermatozoa have been used for ICSI with successful results (Craft et al., 1993Go, 1995Go; Schoysman et al., 1993Go; Devroey et al., 1994Go; Tournaye et al., 1994Go; Tournaye, 1999Go). When no spermatozoa can be found in the testicular biopsy specimens, use of immature sperm cells is the only alternative for pregnancy, except for artificial insemination with donor spermatozoa. Recently, immature sperm cells have been used in the clinical treatment of men with spermatogenesis failure. While several pregnancies using spermatids have been reported, reported fertilization and pregnancy rates for these immature sperm cells are lower than for spermatozoa (Tesarik et al., 1996Go; Fishel et al., 1997Go; Vanderzwalmen et al., 1997Go; Sousa et al., 1999Go). Functional immaturity may be present in spermatids despite the fact that their nuclei have completed meiosis, just like those of mature spermatozoa. Moreover, the functional differences of spermatids and spermatozoa are not obvious. There are no reports investigating both oocyte activation and Ca2+ oscillation in spermatid fertilization without any treatment of artificial oocyte activation. It is also not clear whether fertilization using spermatids of any stage brings oocyte activation with Ca2+ oscillation.

The present study was initiated to determine four factors: (i) the oocyte activation and Ca2+ oscillation-inducing abilities of immature sperm cells in some rodents and human; (ii) the timing at which these abilities are established during spermiogenesis; (iii) the nature of changing intracellular calcium concentration ([Ca2+]i) response in injected oocytes during spermiogenesis; and (iv) the main locations of oocyte-activating factor distribution within round spermatids.

A mouse oocyte activation assay (mouse test) was used to compare the above abilities for the subject species, since the oocytes of B6D2F1 mice provide a good assessment of oocyte activation by spermatogenic cells, especially in human, due to the difficulties of using human oocytes in experiments.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Preparation of oocytes
Female mice (B6D2F1) aged 6–8 weeks were superovulated by consecutive i.p. injections (48 h apart) of 8 IU pregnant mare's serum gonadotrophin (Teikokuzoki Co., Tokyo, Japan) and 8 IU human chorionic gonadotrophin (HCG; Mochida Pharmaceutical Co., Tokyo, Japan). Oocyte–cumulus complexes were obtained from the oviducts at ~16 h after HCG injection and treated with HEPES-buffered human tubal fluid medium (mHTF; Irvine Scientific, Santa Ana, CA, USA) containing 0.1% hyaluronidase (from bovine testis; 825 IU/mg; Sigma, St Louis, MO, USA) to disperse cumulus cells. Cumulus-free oocytes were kept in mHTF (Irvine Scientific) with 10% synthetic serum substitute (SSS; Irvine Scientific) at 37°C under 5% CO2, 5% O2 and 90% N2 for up to 2 h before ICSI.

Preparation of round spermatids (ROS) and elongated spermatids (ELS)
Mature male mice (B6D2F1 and ICR), hamster (golden), rat (Wistar) and rabbit (Japanese White) were killed by cervical dislocation or i.v. injection of pentobarbital sodium (Nembutal; Abbott Laboratories, North Chicago, IL, USA) and their testes were isolated. After removal of the tunica, seminiferous tubules were placed in 1 ml of mHTF and cut into small pieces with a pair of scissors. One part of suspension containing seminiferous tubules was mixed thoroughly with one part of 0.9% NaCl containing 10% polyvinylpyrrolidone (PVP-360; Sigma). A 3 µl aliquot of this suspension, which contained spermatozoa as well as spermatogenic cells at various stages of development, was placed in a plastic Petri dish, covered with mineral oil, and retained for up to 2 h before injection into oocytes.

Human spermatids were obtained from patients with non-obstructive azoospermia. Samples were drawn during ICSI treatment using testicular sperm extraction (TESE). Informed consent was obtained from all participating patients before the experiment was started; in addition, the agreement of the ethical committee at Fukushima Medical University was obtained.

Mature spermatozoa from rodent species were obtained from the cauda epididymis of mature male animals. Spermatozoa were incubated in 3 ml of mHTF for 10–15 min. Immediately before injection, these sperm suspensions were lightly sonicated (Kuretake et al., 1996Go) for 5 s at 5 W power output, using an ultrasonic sonicator (Microson; Misonic Inc., Farmingdale, NY, USA). A 3 µl aliquot of the suspension was then mixed with an equal volume of 0.9% NaCl containing 10% PVP.

Human spermatozoa were obtained from consenting volunteers of proven fertility as a control. Semen samples were allowed to liquefy for 30 min at room temperature. Motile spermatozoa were collected by the swim-up method using HTF medium. In the case of human spermatozoa, no sonication was applied; instead, motile spermatozoa were immobilized by the application of piezo pulses to their tails immediately before injection.

Microinjection of spermatids and spermatozoa into mouse oocytes (mouse test)
Intracytoplasmic spermatids/spermatozoa injection was performed using a micromanipulator with piezo-electric elements (Kimura and Yanagimachi, 1995aGo; Yanagimachi, 1998Go; Yanagida et al., 1999aGo). ROS from the subject rodents (stage 1–7, Gorgi-phase to early cap phase) and humans (Sa1) have largely common morphological characteristics and can be easily distinguished from other spermatogenic and somatic cells by their small size (for rodents, ~10 µm in diameter; for humans, ~7–10 µm) and round nucleus with a centrally located nucleolar structure (Oakberk, 1956Go; Ogura and Yanagimachi, 1993Go; Sousa et al., 1999Go) (Figure 1Go). ELS (stage 9–12) and mature spermatozoa (stage 15–16) of the subject rodents can also be easily identified by their distinctive morphology (Oakberk, 1956Go).



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Figure 1. Round spermatids (ROS) and elongated spermatids (ELS) used in this study. (A) Mouse (B6D2F1); (B) hamster; (C) rabbit; (D) rat. (E) Intact ROS of human obtained from testis of azoospermic patient (upper picture). In some experiments, the nucleus (N) of ROS was separated from the bulk of the cytoplasm (C) by drawing ROS in and out of the injection pipette repeatedly (lower picture). Most of the flagella of ELS were lost during the treatment for isolation of these cells.

 
During injection, a single spermatid/spermatozoon was drawn into the injection pipette (5–6 µm inner diameter at the tip) and its plasma membrane was damaged by application of a piezo pulse. A B6D2F1 mouse oocyte was fixed to the holding pipette using a metaphase II spindle at the 12 or 6 o'clock position, and spermatid/spermatozoa injection was then performed from the 3 o'clock position. Initially, the filled injection pipette was allowed to penetrate only the zona pellucida while piezo pulses were applied. When the tip of the pipette had been injected deeply into the ooplasm, the plasma membrane was ruptured by a single piezo pulse and the spermatid/spermatozoon was expelled into the oocyte. In some species (hamster and rabbit) and some human cases, the nucleus of ROS was separated from the bulk of the cytoplasm by drawing ROS in and out of the injection pipette repeatedly, and then injected into oocytes separately to examine the distribution of the oocyte activation factor.

During spermatid/spermatozoon injection, no additional special procedures capable of inducing oocyte activation were performed, including vigorous cytoplasmic aspiration. All the procedures of intracytoplasmic injection were performed in 3 µl of mHTF on the stage cooled to 17°C. After injection, oocytes were kept at 17°C for 10 min and at room temperature for a further 10 min, then washed three times in HTF and incubated under 5% CO2, 5% O2 and 90% N2 at 37°C (Kimura and Yanagimachi, 1995aGo). The reason for cooling oocytes to 17°C after microinjection was because the oolemma of mouse oocytes is highly elastic and the wound-healing capacity after microinjection is inferior to that of other animals. When mouse oocytes were inseminated by microinjection at room temperature or at 37°C, most were degenerated. At a lower temperature (17–18°C), closure of the pipette-made channel was promoted and the oolemma wound healed completely, resulting in an intact injected oocyte (Kimura and Yanagimachi, 1995aGo). After 4–5 h incubation, oocytes were fixed and stained by acetocarmine to allow cytological examination by phase-contrast microscope (Yanagida et al., 1991Go), and considered to be activated when they possessed a second polar body and more than one pronucleus.

Measurement of [Ca2+]i of injected oocytes
The Ca2+ response of spermatids/spermatozoa-injected oocytes was investigated using a Ca2+-imaging method and a confocal laser scanning microscope system (Bio-Rad MRC-600, Nippon Bio-Rad Laboratories, Tokyo, Japan). Oocytes were loaded with a Ca2+-sensitive fluorescent dye, fluo-3 acetoxymethyl ester (Fluo-3/AM, Molecular Probes Inc, Eugene, OR, USA) dissolved in dimethyl sulphoxide, final concentration 44 µmol/l in HTF with 0.02% Pluronic F-127, for 30 min under 5% CO2, 5% O2 and 90% N2 at 37°C. Loaded oocytes were washed thoroughly and placed in a 3 µl droplet of mHTF on a chambered coverglass (Lab-Tek, Nunc Inc., Naperville, USA) covered with mineral oil. The chamber was mounted on the stage of a phase-contrast inverted microscope equipped with an image processor and [Ca2+]i responses of injected oocytes were measured at room temperature. Most measurements were initiated at 15–20 min after the injection procedure because the injected oocytes were kept on the stage of a microscope cooled at 17°C for 10 min, and the measurements continued for about 60 min at 20 s intervals. Some oocytes were measured starting before injection, while others were measured for up to 180 min.

Statistical significance was assessed using a {chi}2-test; P < 0.05 was considered to be statistically significant.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
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 References
 
Oocyte activation and [Ca2+]i response following injection of ROS/ELS
The [Ca2+]i response patterns of injected oocytes were classified into four groups, as shown in Figure 2Go. Normal oscillation pattern (type A) consisted of repetitive spike-shaped Ca2+ rises at intervals of 2–10 min. Abnormal oscillation pattern (type B) consisted of continuous Ca2+ elevation for several minutes (20–30 min) before the normal oscillation pattern. This pattern was very rare in intact (i.e. not degenerated after injection procedure) oocytes and in this experiment, only four type B patterns were observed out of 223 [Ca2+]i measured oocytes. The transient pattern (type C) consisted of only several (one to four) transient [Ca2+]i rises. Finally, the no-response pattern (type D) consisted of no [Ca2+]i rises during observation periods.



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Figure 2. Intracellular calcium concentration ([Ca2+]i) pattern of spermatid/spermatozoa-injected oocytes. The [Ca2+]i patterns were classified into four groups (types A–D). Type A: normal oscillation pattern, consists of repetitive spike-shaped [Ca2+]i rises for which the intervals are 2–10 min. Type B: abnormal oscillation pattern, consists of a combination of a continuous [Ca2+]i rise and oscillation pattern. Type C: transient pattern, consists of several (one to four) transient [Ca2+]i rises. Type D: no-response pattern, consists of no [Ca2+]i rises during observation periods. Because most of the measurements were initiated at 15–20 min after injection, the large initial [Ca2+]i rise due to the puncture of the oocyte plasma membrane with the pipette was not recorded. x-axis = time from starting measurement; y-axis = intensity.

 
Because most [Ca2+]i measurements were initiated at 15–20 min after injection, the large initial [Ca2+]i rise due to the puncture of the oocyte plasma membrane with the injection needle was not recorded.

Mouse spermatids/spermatozoa injection
The results of experiments in which spermatids/spermatozoa of B6D2F1 mice were injected into oocytes are summarized in Table IGo. As a control, when each oocyte was injected with a bolus (~5 pl) of mHTF without spermatids/spermatozoa, no oocytes were activated, and all the oocytes showed no-response patterns of [Ca2+]i.


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Table I. Activation and intracellular calcium concentration ([Ca2+]i) patterns of mouse (B6D2F1) round spermatid (ROS)/elongated spermatid (ELS) and spermatozoa-injected oocytes
 
When ROS were injected, none of the oocytes was activated, and almost all of the oocytes were arrested at metaphase II of the second meiotic division. [Ca2+]i rises were not observed in all oocytes. When ELS were injected, most of the oocytes were activated (89%), and [Ca2+]i responses of almost all of the oocytes (94%) showed transient patterns (type C). When spermatozoa were injected, 92% of the oocytes were activated normally, and all of the examined oocytes showed normal oscillation patterns (type A).

The results of experiments in which spermatids/spermatozoa of ICR mice were injected into oocytes of B6D2F1 mice are summarized in Table IIGo. When ROS were injected, only 6.2% of oocytes were activated. None of the oocytes showed normal oscillation patterns and only two out of 16 (13%) showed transient patterns (type C); others showed type D patterns. When ELS were injected, 86% of oocytes were normally activated. Normal oscillation patterns were observed in only three out of 13 oocytes examined (23%); most oocytes (69%) showed transient patterns (type C). When spermatozoa were injected, all of the oocytes were activated, and all showed normal oscillation patterns (type A).


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Table II. Activation and intracellular calcium concentration ([Ca2+]i) patterns of mouse (ICR) round spermatid (ROS)/elongated spermatid (ELS) and spermatozoa-injected oocytes
 
In the case of mice, oocyte activation-inducing ability appears during transformation to ELS (Kimura et al., 1998Go), and Ca2+ oscillation-inducing ability appears during transformation to spermatozoon.

Hamster spermatids/spermatozoa injection
The results of experiments in which spermatids/spermatozoa of hamsters were injected into oocytes of B6D2F1 mice are summarized in Table IIIGo. When ROS were injected, most of the oocytes (70%) were normally activated; about 30% of examined oocytes showed transient patterns (type C). When ELS were injected, 74% of oocytes were activated and six of nine oocytes (67%) showed normal oscillation patterns (type A). When spermatozoa were injected, almost all of the oocytes were activated and showed normal oscillation patterns.


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Table III. Activation and intracellular calcium concentration ([Ca2+]i) patterns of hamster round spermatid (ROS)/elongated spermatid (ELS) and spermatozoa-injected oocytes
 
When ROS were separated into nucleus and cytoplasm and injected into oocytes separately, only 3.6% of oocytes were activated by injection of nucleus, whereas 71% of oocytes were activated by injection of cytoplasm.

Rabbit spermatids/spermatozoa injection
The results of experiments in which spermatids/spermatozoa of rabbit were injected into oocytes are summarized in Table IVGo. When ROS were injected, 71% of oocytes were normally activated. Some 33% of examined oocytes showed normal oscillation patterns, and 33% showed transient patterns (type C). When ELS and mature spermatozoa were injected, over 75% of oocytes were activated and all of the examined oocytes showed normal oscillation patterns (type A).


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Table IV. Activation and intracellular calcium concentration ([Ca2+]i) patterns of rabbit round spermatid (ROS)/elongated spermatid (ELS) and spermatozoa-injected oocytes
 
When ROS nucleus and cytoplasm were injected separately into oocytes, only 16 and 4.5% of oocytes were activated respectively.

Rat spermatids/spermatozoa injection
The results of experiments in which spermatids/spermatozoa of rat were injected into oocytes are summarized in Table VGo. When ROS were injected, none of the oocytes was activated, and 88% showed a type D response. When ELS were injected, 31% of the oocytes were activated, and 55% showed a type C response. When mature spermatozoa were injected, 53% of oocytes were activated and 55% of the examined oocytes showed some types of oscillation pattern (type A, B).


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Table V. Activation and intracellular calcium concentration ([Ca2+]i) patterns of rat round spermatid (ROS)/elongated spermatid (ELS) and spermatozoa-injected oocytes
 
Human spermatids/spermatozoa injection
Six patients with non-obstructive azoospermia and receiving TESE–ICSI as treatment were examined. All six produced spermatozoa upon testicular extraction and underwent successful TESE–ICSI.

The results of experiments in which spermatids/spermatozoa from these subjects were injected into oocytes of B6D2F1 mice are summarized in Table VIGo. When ROS were injected, 52% of oocytes were normally activated and 51% of examined oocytes showed normal oscillation patterns, while 22% showed transient patterns (type C). In this study the ELS-injected oocytes were not examined because it was not possible to obtain sufficient ELS from these patients for examination. However, it is easily estimated that ELS can induce oocyte activation and Ca2+ oscillation more efficiently than ROS (>50%). When mature spermatozoa from fertile donors were injected as a control, 97% of oocytes were activated normally, and nine out of 10 injected oocytes (90%) showed normal oscillation patterns.


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Table VI. Activation and intracellular calcium concentration ([Ca2+]i) patterns of human round spermatid (ROS)/elongated spermatid (ELS) and spermatozoa-injected oocytes
 
When the nucleus and cytoplasm of human ROS were injected separately into oocytes, 29 and 26% of oocytes were normally activated respectively.


    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In the present study, immature sperm cells (ROS/ELS) and mature spermatozoa from selected rodent species and humans were injected into oocytes from B6D2F1 mice in order to examine the oocyte activation and Ca2+ oscillation-inducing abilities of these cells. This experimental situation was termed a `mouse oocyte activation assay', or `mouse test'. Mouse (B6D2F1) oocytes are activated with certainty by injecting mature spermatozoa of many species, yet are largely immune to activation by sham alternatives (e.g. injection medium only). Consequently, oocytes from B6D2F1 mice constitute an excellent material for estimating the oocyte activation-inducing abilities of immature sperm cells injected without any artificial stimulation (such as vigorous cytoplasmic aspirations, Ca-ionophore or electrical stimulation). This approach has been used in recent reports (Rybouchkin et al., 1995Go; Yanagida et al., 1999bGo), and was also used in the present study, not only to observe oocyte activation but also to investigate the Ca2+ oscillation-inducing abilities of immature sperm cells. This report is the first describing that Ca2+ oscillation was induced by heterologous gametes.

Spermatids, as spermatogenic cells that have just completed meiosis, possess a haploid set of chromosomes, as do spermatozoa. Fertilization with ROS nuclei has been reported previously (Ogura and Yanagimachi, 1993Go). These authors confirmed that the ROS nuclei of hamster microsurgically injected into mature oocytes were capable of participating in syngamy. Subsequently, numerous reports have described fertilization efforts using immature spermatogenic cells, either through animal experiments or clinical treatments of men with non-obstructive azoospermia. Normal offspring have been obtained by electrofusion or intracytoplasmic injection of ROS both in mice (Ogura et al., 1994Go; Kimura and Yanagimachi, 1995bGo) and in rabbit (Sofikitis et al., 1996Go). Recently, clinical pregnancies and births resulting from intracytoplasmic ROS/ELS injection have also been reported (Fishel et al., 1995Go, 1997Go; Tesarik et al., 1995Go, 1996Go; Vanderzwalmen et al., 1997Go; Sousa et al., 1999Go). However, fertilization and pregnancy rates after injection of ROS are significantly lower than those of ELS or testicular spermatozoa. Possible partial explanations for these lower rates include the absence of a normal centrosome (Fishel et al., 1996Go), incomplete maturation of nuclear protein (Tesarik et al., 1996Go; Sousa et al., 1998Go), lack of active oocyte-activation factor and lack of Ca2+ oscillation response caused by the cytoplasmic immaturity of ROS (Vanderzwalmen et al., 1998Go) relative to ELS and mature spermatozoa. In the present study particular attention was paid to oocyte activation and Ca2+ oscillation during the process of fertilization, and to establishing differences in the fertilization mechanisms of spermatids and spermatozoa.

The results of these experiments are summarized in Table VIIGo. In general, oocyte activation-inducing abilities of ROS differed by species, with ROS of mice and rats failing to activate mouse (B6D2F1) oocytes and ROS of hamster, rabbit and humans achieving efficient activation. For all species, ELS achieved clear activation of mouse oocytes. It was confirmed that the oocyte-activating factor of spermatogenic cells appeared (or became active) while the cells were developing into ROS or ELS.


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Table VII. Summary of oocyte activation and intracellular calcium concentration [Ca2+]i response pattern of spermatid/spermatozoa-injected oocytes
 
[Ca2+]i responses of oocytes in which immature sperm cells were injected changed from transient pattern (type C) to normal oscillation pattern (type A) in all species examined, while the cells were developing to mature spermatozoa. The timing of the transition from type C to type A differed among species, occurring in ELS in hamsters and rabbits, and in spermatozoa in mice and rats. In the case of humans, more than half of ROS-injected oocytes showed normal oscillation patterns, while the transition from type C to type A seemed to occur partly at the ROS stage. These findings suggest that human ROS may be more mature than those of the rodent species examined in this experiment.

It was interesting to note the dissociation between the timing of acquisition of oocyte activation-inducing ability and Ca2+ oscillation-inducing ability of spermatogenic cells. For example, hamster ROS could activate oocytes but could not induce Ca2+ oscillation, while hamster ELS could induce Ca2+ oscillation of injected oocytes. Similar results were obtained for rabbit ROS. In mice and rats, ELS could activate oocytes, but could not induce Ca2+ oscillation, while spermatozoa could induce Ca2+ oscillation. Based on these findings, it is presumed that oocyte activating factor and Ca2+ oscillation-inducing factor (COIF) appeared (or became active) gradually, while the maturation of spermatogenic cells and the thresholds required to induce oocyte activation and Ca2+ oscillation might differ (i.e. the threshold of activation is lower than that of Ca2+ oscillation).

It was possible to confirm from another experiment that Ca2+ oscillation is not essential for normal embryo development, because some normal offspring could be obtained after the transfer of embryos from ELS injection of B6D2F1 mice (activation occurred, but Ca2+ oscillation did not; unpublished data).

Typical [Ca2+]i response patterns could be easily distinguished from type A (oscillation pattern) to type C (transient pattern), not only because of frequency but also because the duration of [Ca2+]i elevation differs between the two types (duration of [Ca2+]i elevation was 3–5 min for type C and <=1 min for type A) (Figure 1Go). This suggests that the Ca2+ released from the endoplasmic reticulum persists in the cytoplasm, because re-uptake into the endoplasmic reticulum was delayed when more immature sperm cells were injected into oocytes. Such behaviour might be due to an insufficiency of COIF.

ROS of hamster, rabbit and humans were able to activate mouse oocytes efficiently. In these cases, the ROS nucleus was separated from the bulk of cytoplasm by repetitive pipetting and injected into each oocyte separately to examine the distribution of oocyte activating factor. With hamsters, ROS nuclei (ROS-N) could not activate oocytes, whereas ROS cytoplasm (ROS-C) could activate oocytes efficiently—which indicated that oocyte activating factor was distributed mainly in the cytoplasm of hamster ROS. With rabbits, both ROS-N and ROS-C were largely unable to activate oocytes, which might be due to oocyte activating factor being distributed in both the nucleus (or perinucleus) and cytoplasm evenly so that neither the nucleus (or perinucleus) nor cytoplasm had enough oocyte activating factor to activate the oocyte efficiently. It was observed by confocal laser scanning microscopy that when rabbit ROS were treated with hypotonic solution and gentle pipettings for separation of the nucleus from the cytoplasm, a significant amount of cytoplasm remained around the nucleus of ROS (Yamamoto et al., 1999Go). The cytoplasm of rabbit ROS may have a lower concentration of oocyte activating factor than hamster ROS. Thus, injection of the same amount of cytoplasm could not activate the oocyte efficiently, although both the cytoplasm of hamster and whole ROS (including whole cytoplasm) of rabbit could provide efficient activation. In humans, the activation rates of ROS-N or ROS-C were both ~30%, whereas the activation rate of whole ROS injection was ~50%. Moreover, the oocyte activating factor seems to be distributed evenly in the nucleus (or perinucleus) and cytoplasm in human ROS.

In this experiment, the ROS used was obtained from patients who had azoospermia and were able to receive TESE–ICSI (incomplete spermiogenesis failure; Amer et al., 1997). Previous reports indicate that fertilization outcomes, embryo quality and conception by ROS injection can all be affected by the severity of testicular pathology (Vanderzwalmen et al., 1997Go) or damage of the testis (Sofikitis et al., 1996Go). The study included one human subject with complete spermiogenesis failure (maturation arrest at the spermatid stage; ROS could be identified upon testicular biopsy, but spermatozoa could not); his ROS were largely unable to induce oocyte activation and Ca2+ oscillation (data not shown). Additional data involving cases of complete spermiogenesis failure are required because ROS injection may be indicated in such cases, rather than the cases of incomplete spermiogenesis failure. In addition, it must be determined without delay whether the clinical use of ROS injection is appropriate.

In order to determine the exact activity of oocyte activating factor-COIF in immature spermatogenic cells, it appears necessary to perform the current experiments using homologous oocytes and ROS/ELS. Such experiments, using experimental animals, are planned.


    Notes
 
To whom correspondence should be addressed. E-mail: h-yazawa{at}fmu.ac.jp


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
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Submitted on March 13, 2000; accepted on August 16, 2000.