Abstract
The efficiency of intracytoplasmic sperm injection (ICSI) in the bovine is low compared to other species. It is unknown whether defective oocyte activation and/or sperm head decondensation limit the success of this technique in this species. To elucidate where the main obstacle lies, we used homologous and heterologous ICSI and parthenogenetic activation procedures. We also evaluated whether in vitro maturation negatively impacted the early stages of activation after ICSI. Here we showed that injected bovine sperm are resistant to nuclear decondensation by bovine oocytes and this is only partly overcome by exogenous activation. Remarkably, when we used heterologous ICSI, in vivo-matured mouse eggs were capable of mounting calcium oscillations and displaying normal PN formation following injection of bovine sperm, although in vitro-matured mouse oocytes were unable to do so. Together, our data demonstrate that bovine sperm are especially resistant to nuclear decondensation by in vitro-matured oocytes and this deficiency cannot be simply overcome by exogenous activation protocols, even by inducing physiological calcium oscillations. Therefore, the inability of a suboptimal ooplasmic environment to induce sperm head decondensation limits the success of ICSI in the bovine. Studies aimed to improve the cytoplasmic milieu of in vitro-matured oocytes and to replicate the molecular changes associated with in vivo capacitation and acrosome reaction will deepen our understanding of the mechanism of fertilization and improve the success of ICSI in this species.
Introduction
Intracytoplasmic sperm injection (ICSI) has become an indispensable tool to overcome infertility in humans (Palermo et al. 1992). In addition, ICSI has become an important method to study the mechanisms of fertilization and early events of egg activation in mammals (Yanagimachi 2005). ICSI involves the fertilization of oocytes in metaphase II stage (MII) by direct injection of a spermatozoon (Goto 1997). This procedure is nearly as effective as in vitro fertilization (IVF) in producing offspring in humans (Palermo et al. 1992) and mice (Kimura & Yanagimachi 1995), and it has been a valuable tool for conservation purposes in species where other assisted reproductive technologies are not available or are not optimized (Perry et al. 1999). Nevertheless, the success of ICSI in the bovine is poor with rates of embryo development well below those obtained by IVF (Rho et al. 2004, Arias et al. 2014). Specifically, in this species, the majority of eggs fail to activate following ICSI (Catt & Rhodes 1995, Malcuit et al. 2006) and display delayed and/or incomplete sperm head decondensation (Chen & Seidel 1997, Suttner et al. 2000). These defects together conspire to limit success of ICSI in the bovine.
During fertilization the sperm delivers into the ooplasm a sperm-specific phospholipase C (PLCZeta1), PLCζ (Saunders et al. 2002, Knott et al. 2005), which is responsible for initiating the intracellular calcium ([Ca2+]i) oscillations that are a hallmark of mammalian fertilization (Wakai & Fissore 2013). The [Ca2+]i oscillations induce all downstream events of egg activation, which is the first stage of embryo development (Schultz & Kopf 1995). There are early events of activation such as exocytosis of the cortical granules, prevention of polyspermy and exit from the MII stage, which are initiated soon after sperm entry. Late events of egg activation such as sperm head decondensation, progression to the pronuclear (PN) stage and DNA synthesis and recruitment of maternal RNAs unfold over a period of 10 h (h). Remarkably, in species where ICSI is successful, the events of egg activation are closely recapitulated following sperm injection (Kimura et al. 1998, Sato et al. 1999, Yanagida et al. 2001). Nevertheless, this is not the case in the bovine, as whereas bovine eggs are capable of undergoing normal activation following IVF including formation of male PNs with high efficiency, the same cohort of eggs are incapable of supporting similar rates of activation and formation of male PN following fertilization by injection of a spermatozoon.
As noted above, a necessary step for egg activation is the initiation of [Ca2+]i oscillations. This step is defective in the bovine following ICSI, as oscillations fail to initiate and/or are short lived (Malcuit et al. 2006). Another step of egg activation that is defective is the formation of the male PN. Decondensation of the sperm head requires the sperm’s nuclear and cytoplasmic contents and its surrounding membranes to mingle with the ooplasm (Sutovsky & Schatten 2000). Further, the transformation into a male PN requires among other things the swapping of the DNA-associated sperm’s protamines with maternal histones as well as the incorporation of other chaperone proteins (Florman & Fissore 2014). It is known that IVM oocytes display reduced developmental potential (Rizos et al. 2002, Kim et al. 2004, Virant-Klun et al. 2013), and it is possible that IVM bovine eggs might be unable to process sperm that have not undergone in vivo capacitation and the acrosome reaction. Therefore, it is unknown if the defective activation with delayed and incomplete sperm head decondensation observed after ICSI in the bovine is caused by a deficient stimulus of egg activation or if it is due to ‘suboptimal ooplasm’ of IVM eggs that are unable to convert the sperm into a male PN.
Thus, to elucidate where the main obstacle for successful bovine ICSI lies, we used homologous and heterologous ICSI and artificial activation procedures to evaluate the ability of sperm exposed to these different conditions to induce [Ca2+]i responses, undergo sperm head decondensation and PN formation. Our results show that bovine sperm injected into IVM bovine eggs are highly stable and resistant to nuclear decondensation, and exogenous activation by parthenogenetic procedures only partly overcomes this defect. Remarkably, in vivo- but not in vitro-matured mouse oocytes supported normal [Ca2+]i oscillations, sperm head decondensation and PN formation. Together, the results suggest that less than optimal ooplasmic conditions present in IVM eggs undermine ICSI in the bovine. Protocols to improve the ooplasmic environment as well as to replicate in vitro the molecular changes associated with capacitation and the acrosome reaction will improve ICSI in this species.
Materials and methods
Reagents
Unless stated otherwise, all chemicals were purchased from Sigma Aldrich.
Collection of gametes
All animal procedures were performed in accordance with the Animal Care and Use Committee (IACUC) guidelines of UMass-Amherst. Mouse GV oocytes were collected from the ovaries of 6- to 10-week-old CD-1 female mice 44–46 h after injection of 5 IU of pregnant mare serum gonadotropin (PMSG; Sigma). Cumulus intact GV oocytes were recovered in a HEPES-buffered Tyrode’s lactate solution (TL-HEPES) containing 5% heat-treated fetal calf serum (FCS; Gibco) and were matured in vitro for 12–14 h in IBMX-free Chatot, Ziomek and Bavister (CZB) (Chatot et al. 1989) media supplemented with 3 mg/mL bovine serum albumin (BSA, Sigma) under mineral oil at 37°C in a humidified atmosphere of 5% CO2, as described before (Lee et al. 2016). MII mouse oocytes were collected from the oviducts into a HEPES-buffered solution (TL-Hepes) supplemented with 10% heat-treated calf serum (FCS; Gibco), 12–14 h after administration of 5 IU of human chorionic gonadotropin (hCG), which was administered 46–48 h after PMSG. All animal procedures were performed according to research animal protocols approved by the University of Massachusetts Institutional Animal Care and Use Committee. Bovine MII oocytes were purchased from Boviteq USA (Wisconsin, Madison). Germinal vesicle-stage bovine oocytes were isolated from cow ovaries collected from an abattoir located in the mid-western region of the USA. Oocytes were isolated from follicles 2–8 mm in diameter. Following isolation, oocytes were placed in vials in 2 mL TCM-199 medium (Gibco), supplemented with 10% FCS, 0.1 m/L luteinizing hormone (LH, Sioux Biochemical, Sioux Center, IA, USA) and 1 µg/mL oestradiol for 20 h. Oocytes were shipped overnight at 39°C in a portable incubator (MiniTube of America, Verona, WI, USA). Oocytes for ICSI experiments were denuded of their cumulus cells by vortexing in the presence of 1 mg/mL hyaluronidase and selected for the presence of the first polar body. In all cases, mouse and bovine MII oocytes were rinsed thoroughly and held in KSOM (Specialty Media, Phillipsburg, NJ, USA) supplemented with 0.1% fraction V BSA until initiation of the ICSI procedure.
Sperm preparation
Cauda epididymal sperm from CD1 male mice were collected in 500 µL of a modified Krebs-Ringer medium (Whitten’s HEPES-buffered medium) (Navarrete et al. 2015) and allowed to swim out for 10 min at 37°C. The sperm suspension was washed three times and resuspended in 0.5 mL injection buffer. These samples were then washed three times in injection buffer prior to injection. Bovine sperm were obtained from frozen semen samples kindly donated by American Breeder Services (DeForest, WI, USA). After thawing the straws, the sperm were separated using a Percoll gradient. These spermatozoa (5 × 106 spermatozoa/mL) were then washed in Sp-TALP medium (Parrish et al. 1988) for 5 min and then used for ICSI.
Microinjection of PLCζ cRNA
Microinjections were performed as described previously (Kurokawa et al. 2005). PLCζ cRNA was prepared as previously described by us and stored at −20°C (Ross et al. 2009). After thawing, the samples were centrifuged, and the top 1–2 µL was used to prepare microdrops from which glass micropipettes were loaded by aspiration. cRNA was delivered into oocytes by pneumatic pressure (PLI-100 picoinjector, Harvard Apparatus, Cambridge, MA). Each oocyte received 5–10 pL, which is approximately 1–3% of the total volume of the egg.
[Ca2+]i measurements
[Ca2+]i monitoring was performed as previously described (Kurokawa et al. 2005). [Ca2+]i changes in mouse oocytes were measured using the Ca2+-sensitive dye Fura-2-acetoxymethyl ester (Fura 2-AM, Molecular Probes; Invitrogen). Mouse oocytes were incubated and loaded with 1.25 µM Fura-2AM supplemented with 0.02% pluronic acid (Molecular Probes) for 20 min at room temperature. For bovine oocytes, [Ca2+]i was measured using Fura-2-dextran (Fura 2-Dextran, Molecular Probes; Invitrogen), which was delivered by injection into the oocytes. Following loading with the dye, oocytes were placed in microdrops of TL-HEPES on a monitoring glass bottom dish (Mat-Tek Corp.) under mineral oil. Eggs were monitored simultaneously using inverted microscopes (Nikon) outfitted for fluorescence measurements. Fura 2-AM and Fura2-Dextran were excited by UV light alternating wavelengths of 340 nm and 380 nm by a filter wheel (Ludl Electronic Products Ltd.), and fluorescence was captured every 20 s. A 75 W Xenon lamp provided the excitation light. The excitation wavelength was alternated between 340 and 380 nm by a filter wheel (Ludl Electronic Products, Hawthorne, NY, USA), and fluorescence ratios were obtained every 20 or 30 s. The emitted light was passed through a 510 nm barrier filter and collected with either a cooled Photometrics SenSys CCD or a cool SNAP ES digital camera (Roper Scientific, Tucson, AZ, USA). SimplePCI software (Compix Imaging Inc., Cranberry, PA, USA) was used to monitor [Ca2+]i and synchronize the rotation of the filter wheel. [Ca2+]i values are reported as the ratio of 340/380 nm fluorescence in the whole egg.
Bovine in vitro fertilization (IVF) and embryo culture
Matured bovine oocytes and sperm were co-incubated for 18–20 h in IVF-TL supplemented with 0.2 mM sodium pyruvate, 3 mg fatty acid-free BSA and 0.025 mg gentamicin sulfate/mL (Parrish et al. 1986, Felmer et al. 2011). Final IVF-TALP contained PHE (2 mM penicillamine, 1 mM hypotaurine and 0.25 mM epinephrine), 2 μg heparin and 1 × 106 Percoll separated frozen-thawed sperm/mL. Presumptive zygotes were stripped of cumulus cells via vortex and randomly assigned to the different culture systems. In vitro maturation and fertilization were conducted in 400 μl drops (50 COCs and/or eggs per well) at 38.5°C and 5% CO2 in humidified atmosphere, while embryo culture was carried out in 100 μL drops (∼45 embryos per drop) under mineral oil of KSOM–0.4% BSA medium (EmbryoMax, Chemicom International, USA) at 38.5°C and 5% CO2, 5% O2 and 90% N2, in a humidified atmosphere.
Intracytoplasmic sperm injection
ICSI was carried out as previously described (Kurokawa & Fissore 2003) using Narishige manipulators (Medical System Corp., Great Neck, NY) mounted on a Nikon diaphot microscope (Nikon). ICSI was performed in CZB medium (Chatot et al. 1989) at room temperature using CD1 eggs. One part sperm suspension was mixed with one part injection buffer containing 12% polyvinylpyrrolidone (PVP, M.W. 360 kDa; Sigma). Sperm were delivered into the eggs’ cytosol using a piezo micropipette-driving unit (Piezodrill; Burleigh Instruments Inc., Rochester, NY). In certain experiments, the bovine sperm were immobilized by applying a few piezo pulses to the sperm tail prior to ICSI; the immotile sperm still was connected to the tail, and the whole sperm was injected. When sperm heads were injected, sonication was used to decapitate the heads (Kimura et al. 1998); sonication (XL2020, Heat Systems Inc., Farmingdale, NY) was carried out for 5 s at 4°C. In certain bovine ICSI experiments, oocytes were washed at least three times in TL-HEPES and then activated as described below.
Collection of mating-derived mouse zygotes
Ovulation was induced in mice by intraperitoneal injection of 10 IU of Pregnant Mare’s Serum Gonadotropin Folligon (PMSG; Sigma) followed by 10 IU of human chorionic gonadotropin (hCG; Sigma) after 48 h. The female mouse was mated with a male mouse overnight. The following morning, the mating was confirmed by inspection of the vaginal plug. Mouse with visible copulation plug was killed by cervical dislocation, and then ovary burses were surgically removed and collected in a HEPES-buffered Tyrode’s lactate solution (TL-HEPES) containing 5% heat-treated fetal calf serum (FCS; Gibco) (Lee et al. 2016). Under stereomicroscope, zygotes were dissected out from the swollen ampulla and treated with 0.1% bovine testes hyaluronidase (Sigma). The collected zygotes were transferred into 100 μL of TL-HEPES and fixed to determine pronuclear stage.
Parthenogenetic oocyte activation
Bovine oocytes were activated by injection of PLCZ1 cRNA into the ooplasm (concentration as indicated) or chemically. After cRNA microinjection, oocytes were cultured in potassium simplex optimized medium (KSOM; EmbryoMax; Millipore).
Chemical activation was performed with 5 µM ionomycin (Calbiochem) for 5 min, followed by incubation in KSOM containing 10 mg/mL cycloheximide (CHX) for 5 h (Io + CHX). After activation, oocytes were allocated to 50 µL culture drops (maximum 20–25 embryos per drop) consisting of KSOM medium. For all experiments, embryo culture was performed at 38.5°C with a gas mixture of 5% CO2, 5% O2, 90% N2 and saturation humidity. Cleavage and Blastocyst rates were assessed at 72 and 192 h, respectively. Pronuclear formation was assessed between 14–18 h post activation (hpa), depending on the experiment.
Determination of Pronuclear (PN) stage
After ICSI or egg activation, oocytes were fixed in paraformaldehyde (4%) at two different times after activation, 14 and 18 h post activation (hpa) and then stained with Hoechst 33342 (10 mg/mL) for 25 min to determine the presence of pronucleus and/or male sperm decondensation. In case of IVF, zygotes were fixed at 14 and 18 h post insemination (hpi). The PN stage and its size were recorded using a Nikon Optiphot microscope equipped with epifluorescence optics (×200). Image analysis was conducted using the free software ImageJ v1.48, downloaded from the NIH website (https://imagej.nih.gov/ij/download.html).
Statistical analysis
Data were analyzed by descriptive statistics based on the mean plus/minus the standard deviation (s.d.) calculated for each of the variables. Differences among treatments were analyzed using one-way ANOVA. Post hoc analysis to identify differences between groups was performed using Scheffe’s test. Pronuclear formation was analyzed by a Chi-squared test with Bonferroni correction. Significant differences were considered if P < 0.05.
Results
Sperm head decondensation and PN formation after ICSI using bovine gametes
A common defect of bovine ICSI is the inability of eggs to timely decondense the sperm nucleus and form a male PN. Therefore, we first examined if the eggs used in our study were capable of undergoing normal rates of PN formation following in vitro fertilization or common parthenogenetic activation procedures. PN formation was evaluated at 14 and 18 h after insemination or activation with ionomycin (Io) + cycloheximide (CHX). Over 80% of the eggs showed 2 PNs by 14 h post IVF, and if observed a few hours later, all fertilized zygotes had cleaved to the 2-cell stage (Table 1 and Fig. 1; also, data not shown). Similar results were observed with the parthenogenetic zygotes. A final control group was used in which the same parthenogenetic activation procedure was performed but instead of a sperm a comparable volume of PVP (5%) was injected. As observed in Table 1 and Fig. 1, the injection procedure did not affect the ability of eggs to undergo PN formation. Therefore, we conclude that the eggs used in our study are capable of undergoing high rates of male and female PN formation and cleavage to the 2-cell stage following IVF. Moreover, the injection procedure does not disturb the capacity of eggs to undergo activation.
Representative images of PN formation in bovine oocytes fertilized by conventional in vitro fertilization (IVF), chemically activated oocytes without and with sham ICSI injection, or fertilized by ICSI. Upper images were captured after Hoechst staining, whereas the corresponding phase-contrast images are at the bottom. A and B: In vitro fertilization: zygotes examined 14 h post insemination. C and D: Parthenogenetically activated group with Io + CHX (14 h post activation). E and F: Sham-injected group, where injection of comparable volume of PVP (5%) was performed followed by chemical activation with Io + CHX (14 h post activation). G and H: ICSI, bovine zygote generated by ICSI without exogenous activation. White circle in G denotes injected, intact sperm head. I and J: ICSI – Io + CHX 14 hpa, ICSI bovine zygote 14 h post-chemical activation. White circle in I surrounds a partially decondensed sperm head. K and L: ICSI – Io + CHX 18 hpa, ICSI bovine zygote 18 h post-chemical activation. M and N: ICSI + bPLCζ: ICSI bovine zygote activated with bovine PLCζ 18 hpa. Arrows indicate PBs and arrowheads PNs. Magnification: 200×, scale bar in microns.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Representative images of PN formation in bovine oocytes fertilized by conventional in vitro fertilization (IVF), chemically activated oocytes without and with sham ICSI injection, or fertilized by ICSI. Upper images were captured after Hoechst staining, whereas the corresponding phase-contrast images are at the bottom. A and B: In vitro fertilization: zygotes examined 14 h post insemination. C and D: Parthenogenetically activated group with Io + CHX (14 h post activation). E and F: Sham-injected group, where injection of comparable volume of PVP (5%) was performed followed by chemical activation with Io + CHX (14 h post activation). G and H: ICSI, bovine zygote generated by ICSI without exogenous activation. White circle in G denotes injected, intact sperm head. I and J: ICSI – Io + CHX 14 hpa, ICSI bovine zygote 14 h post-chemical activation. White circle in I surrounds a partially decondensed sperm head. K and L: ICSI – Io + CHX 18 hpa, ICSI bovine zygote 18 h post-chemical activation. M and N: ICSI + bPLCζ: ICSI bovine zygote activated with bovine PLCζ 18 hpa. Arrows indicate PBs and arrowheads PNs. Magnification: 200×, scale bar in microns.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Representative images of PN formation in bovine oocytes fertilized by conventional in vitro fertilization (IVF), chemically activated oocytes without and with sham ICSI injection, or fertilized by ICSI. Upper images were captured after Hoechst staining, whereas the corresponding phase-contrast images are at the bottom. A and B: In vitro fertilization: zygotes examined 14 h post insemination. C and D: Parthenogenetically activated group with Io + CHX (14 h post activation). E and F: Sham-injected group, where injection of comparable volume of PVP (5%) was performed followed by chemical activation with Io + CHX (14 h post activation). G and H: ICSI, bovine zygote generated by ICSI without exogenous activation. White circle in G denotes injected, intact sperm head. I and J: ICSI – Io + CHX 14 hpa, ICSI bovine zygote 14 h post-chemical activation. White circle in I surrounds a partially decondensed sperm head. K and L: ICSI – Io + CHX 18 hpa, ICSI bovine zygote 18 h post-chemical activation. M and N: ICSI + bPLCζ: ICSI bovine zygote activated with bovine PLCζ 18 hpa. Arrows indicate PBs and arrowheads PNs. Magnification: 200×, scale bar in microns.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
PN formation in bovine oocytes following in vitro fertilization, common parthenogenetic activation procedures, ICSI, ICSI + chemical activation or PLC cRNA injection.
| Group | Io + CHX | n | Pronuclear Formation 14 hpa | UF/NA | n | Pronuclear Formation 18 hpa | UF/NA | ||
|---|---|---|---|---|---|---|---|---|---|
| PDSH (%) | PN (%) | PDSH (%) | PN (%) | ||||||
| Bovine IVF | – | 22 | 0 (0%)a | 18 (82%)a | 4 (18%)a | 23 | 0 (0%)a | 20 (87%)a | 3 (13%)a |
| Parthenothes | + | 23 | 0 (0%)a | 17 (74%)a | 6 (26%)a | 25 | 0 (0%)a | 20 (80%)a | 5 (20%)a |
| Sham injection | + | 11 | 0 (0%)a | 9 (82%)a | 2 (18%)a | 18 | 0 (0%)a | 13 (72%)*a | 5 (28%)a |
| ICSI | – | 19 | 1 (5%)a | 0 (0%)b | 18 (95%)b | 14 | 1 (7%)a | 1 (7%)b | 12 (86%)b |
| ICSI | + | 16 | 8 (50%)ab | 4 (25%)b | 4 (25%)a | 17 | 5 (30%)a | 7 (40%)b | 5 (30%)a |
| ICSI + bPLCz | – | 15 | 12 (80%)b | 0 (0%)b | 3 (20%)a | 17 | 5 (30%)a | 9 (53%)ab | 3 (17%)a |
Data within the same column with different superscripts are significantly different (P < 0.05).
These oocytes had already cleaved and were 2 cells.
ICSI, Injection of bovine sperm; ICSI + bPLCζ, Injection of sperm followed by injection with bovine bPLCζ cRNA (0.5 µg/µL); Io + CHX, Ionomycin plus cycloheximide; PDSH, Partially decondensed sperm head; PN, Pronucleus; 2PB, Two polar bodies; UF/NA, unfertilized or non-activated oocytes.
We next examined if the same cohort of eggs was capable of undergoing activation and formation of a male PN after injection of a sperm (Table 1; Fig. 1). As expected, without additional parthenogenetic stimulation, ICSI resulted in low rate of PN formation and only 1 of 19 eggs showed a partially decondensed sperm head; there were not marked differences between 14 and 18 h post ICSI. We repeated these studies but accompanied the injection of sperm with exogenous stimulation. In the ICSI-Io + CHX group, 25% (4/16) of the fertilized eggs displayed a fully formed male PN by 14 h, whereas the majority of eggs only showed partially decondensed sperm heads (8/16). Although a slight improvement was observed by 18 h, as 40% (7/17) of the zygotes showed a normal male PN, the rest of the injected zygotes were still unable to form a male PN by this time. We also activated eggs with bovine PLCζ cRNA, ICSI + bPLCζ, but this treatment was even less effective, as none of the zygotes formed a male PN by 14 h post injection (0/15), although nearly 53% of the zygotes formed a male PN by 18 h (9/17). Together, these results show that whereas IVM bovine eggs are capable of responding to common parthenogenetic treatments with formation of female PNs, they are unable to efficiently and timely decondense injected bovine sperm.
[Ca2+]i responses after ICSI and supplementary activation treatments
Given the protracted formation of the male PN after bovine ICSI in general and after ICSI combined with bPLCζ cRNA injection in particular, we examined whether these oocytes/zygotes were capable of mounting [Ca2+]i responses. We also examined whether the delayed male PN formation prevented cleavage to the 2-cell stage. Injection of sperm alone into bovine oocytes induced [Ca2+]i oscillations in only 4/20 oocytes, which were spaced by 80 min intervals (Fig. 2A). These results are consistent with previous studies that showed that in response to sperm injection only a few bovine oocytes can mount persistent oscillations (Malcuit et al. 2006). Nevertheless, injection of bPLCζ cRNA after ICSI induced [Ca2+]i oscillations in nearly all oocytes that were spaced by shorter intervals, which is similar to the responses observed after normal fertilization (Fig. 2B). We also confirmed that all oocytes activated by Io + CHX showed the expected single [Ca2+]i rise of ~5 min duration (Fig. 2C). Together, the results demonstrate that despite replicating the activation stimulus, or bypassing it using Io + CHX, bovine eggs are unable to timely decondense the injected sperm heads and form a male PN. The data therefore suggest that the initial inability to decondense the homologous sperm head is not due to failure of the activation stimulus per se. Remarkably, eggs undergoing ICSI aided with artificial activation were capable of experiencing cleavage at rates similar to those of IVF zygotes and parthenogenetically activated eggs (Table 2), although development to the blastocyst stage was lower for these treatments compared to IVF. Thus, bovine zygotes generated by ICSI are capable of initiating development, although the asynchronous PN formation might undermine the long-term developmental competence of these embryos.
[Ca2+]i responses induced by injection of bovine sperm with and without injection of PLCζ cRNA (0.5 µg/µL) or by the addition of ionomycin. (A) Injection of sperm-induced [Ca2+]i responses in the minority of injected oocytes. (B) Co-injection of bull sperm with bovine PLCzeta1 cRNA (0.5 μg/μL) caused [Ca2+]i responses in all injected oocytes. (C) Addition of Io caused a single [Ca2+]i rise in all exposed oocytes and caused the baseline remain high as long as Io was in the media.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
[Ca2+]i responses induced by injection of bovine sperm with and without injection of PLCζ cRNA (0.5 µg/µL) or by the addition of ionomycin. (A) Injection of sperm-induced [Ca2+]i responses in the minority of injected oocytes. (B) Co-injection of bull sperm with bovine PLCzeta1 cRNA (0.5 μg/μL) caused [Ca2+]i responses in all injected oocytes. (C) Addition of Io caused a single [Ca2+]i rise in all exposed oocytes and caused the baseline remain high as long as Io was in the media.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
[Ca2+]i responses induced by injection of bovine sperm with and without injection of PLCζ cRNA (0.5 µg/µL) or by the addition of ionomycin. (A) Injection of sperm-induced [Ca2+]i responses in the minority of injected oocytes. (B) Co-injection of bull sperm with bovine PLCzeta1 cRNA (0.5 μg/μL) caused [Ca2+]i responses in all injected oocytes. (C) Addition of Io caused a single [Ca2+]i rise in all exposed oocytes and caused the baseline remain high as long as Io was in the media.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
ICSI-generated bovine zygotes initiate embryo development.
| Group | n | Cleavage at 72 h | Blastocysts at 192 h |
|---|---|---|---|
| IVF | 29 | 23 (79%)a | 9 (31%)a |
| ICSI | 28 | 3 (11%)b | 0 (0%)b |
| ICSI – Io + CHX | 27 | 20 (74%)a | 4 (15%)c |
| ICSI + bPLCZeta1 | 28 | 19 (68%)a | 3 (11%)c |
| Parthenogenetic activation | 32 | 24 (75%)a | 3 (9%)c |
Parthenogenetic activation: oocytes chemically activated with Io + CHX. Data followed by different superscripts are significantly different (P < 0.05).
PN formation after heterologous ICSI using mouse oocytes
The inability of IVM bovine eggs to promote male PN formation in a timely fashion after ICSI even after exogenous activation treatments, as shown by us and by many other research groups (Perreault et al. 1988, Goto et al. 1990, Rho et al. 1998), suggests that in vitro maturation might impair the ability of eggs to process intact sperm heads such are those delivered by ICSI. To test our hypothesis the logical next step was to perform the same experiments in in vivo-matured bovine oocytes. Nevertheless, it is prohibitively expensive, and technically challenging, to obtain enough in vivo-matured bovine eggs to carry out meaningful experiments. We therefore resorted to heterologous ICSI, as in vivo-matured mouse eggs are easy to collect. Further, in a previous study we had shown that in vivo-matured mouse eggs were capable of responding to injection of bovine sperm with persistent oscillations (Malcuit et al. 2006). We therefore examined if these eggs, and also in vitro-matured mouse eggs, were capable of inducing PN formation following injection of bovine sperm. PN formation was assessed 5–6 h post sperm injection. Injection of mouse sperm heads into in vivo matured, ovulated, eggs (MIIOV), as expected, induced PN formation in all injected eggs (Table 3, Figs. 3 and 4), which is consistent with results by many other groups and our own previous data (Kimura & Yanagimachi 1995, Kimura et al. 1998, Mizuno et al. 2002). In vitro-matured mouse eggs (MIIIVM) were also efficient at inducing male PN formation following injection of mouse sperm heads (Figs. 3 and 4), although an increasing number of sperm heads remained condensed by the time of observation (Table 3). Remarkably, MIIOV eggs were very efficient at inducing male PN formation following injection of bovine sperm heads, although approximately 25% of the sperm heads remained condensed (Table 3). Importantly, MIIIVM eggs were largely devoid of capacity to render a bovine sperm into a male PN, as less than 10% of the injected sperm formed a male PN (Table 3; Figs. 3 and 4). Similarly, bovine IVM eggs were largely incompetent to process mouse sperm head regardless of whether or not they received supplementary parthenogenetic activation (Table 3). Altogether, these results indicate that the source of oocytes and eggs and the maturation conditions greatly impact the ability of eggs to process and transform sperm heads into male PNs, especially for sperm that bypass the physiological processes associated with natural fertilization.
Representative fluorescent (upper) and phase-contrast (bottom) images of mouse oocytes following IVF or ICSI using mouse or a bovine sperm. A and B: IVF. C and D: Homologous ICSI using ovulated oocytes. E and F: Homologous ICSI using IVM oocytes. G and H: Heterologous ICSI using ovulated oocytes. I and J: Heterologous ICSI using IVM oocytes. M: Male pronuclei. FM: Female pronuclei. MP: Metaphase plate. Arrows indicate PBs and red circles denote PNs. Magnification 200×. Scale bar in microns.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Representative fluorescent (upper) and phase-contrast (bottom) images of mouse oocytes following IVF or ICSI using mouse or a bovine sperm. A and B: IVF. C and D: Homologous ICSI using ovulated oocytes. E and F: Homologous ICSI using IVM oocytes. G and H: Heterologous ICSI using ovulated oocytes. I and J: Heterologous ICSI using IVM oocytes. M: Male pronuclei. FM: Female pronuclei. MP: Metaphase plate. Arrows indicate PBs and red circles denote PNs. Magnification 200×. Scale bar in microns.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Representative fluorescent (upper) and phase-contrast (bottom) images of mouse oocytes following IVF or ICSI using mouse or a bovine sperm. A and B: IVF. C and D: Homologous ICSI using ovulated oocytes. E and F: Homologous ICSI using IVM oocytes. G and H: Heterologous ICSI using ovulated oocytes. I and J: Heterologous ICSI using IVM oocytes. M: Male pronuclei. FM: Female pronuclei. MP: Metaphase plate. Arrows indicate PBs and red circles denote PNs. Magnification 200×. Scale bar in microns.
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Source of sperm and oocytes affect the size of PNs following homologous and heterologous ICSI using ovulated and IVM mouse oocytes. (A) Female PN. (B) Male PN. Values represent the mean + s.d. Bars with different letters are significantly different (P < 0.05).
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Source of sperm and oocytes affect the size of PNs following homologous and heterologous ICSI using ovulated and IVM mouse oocytes. (A) Female PN. (B) Male PN. Values represent the mean + s.d. Bars with different letters are significantly different (P < 0.05).
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Source of sperm and oocytes affect the size of PNs following homologous and heterologous ICSI using ovulated and IVM mouse oocytes. (A) Female PN. (B) Male PN. Values represent the mean + s.d. Bars with different letters are significantly different (P < 0.05).
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
PN formation in mouse and bovine oocytes after homologous and heterologous ICSI.
| Pronuclear Formation | ||||||
|---|---|---|---|---|---|---|
| 1 FPN + 2PB | ||||||
| Group | Io + CHX | n | PDSH (%) | MPN (%) | ISH (%) | ISH + MII (%) |
| MIIOV + mouse sperm | – | 11 | 0 (0%)a | 11 (100%)a | 0 (0%)a | 0 (0%)a |
| MIIIVM + mouse sperm | – | 10 | 0 (0%)a | 7 (70%)b | 2 (20%)b | 1 (10%)a |
| MIIOV + bovine sperm | – | 23 | 5 (22)c | 17 (74%)b | 1 (4%)b | 0 (0%)a |
| MIIIVM + bovine sperm | – | 24 | 2 (8%)d | 0 (0%)c | 0 (0%)c | 22 (92%)a |
| bMIIIVM + mouse sperm | – | 25 | 0 (0%)a | 0 (0%)c | 0 (0%)c | 25 (100%)c |
| bMIIIVM + mouse sperm | + | 24 | 0 (0%)a | 0 (0%)c | 24 (100%)c | 0 (0%)c |
PN formation in mouse oocytes was evaluated at 6 h post sperm injection, and in bovine oocytes at 18 hpi. Data within the same column followed by different superscripts are significantly different (P < 0.05).
bMIIIVM, in vitro-matured bovine oocyte; FPN, female pronucleus; ISH, intact sperm head; MII, oocyte arrested at metaphase II; MIIOV, mouse oocytes collected from oviduct; MIIIVM, in vitro-matured mouse oocytes; MPN, male pronucleus.
The formation of male and female PNs requires several steps the last of which is ‘nuclear swelling,’ which is when the pronuclei attain their final size (Jenkins & Carrell 2012). Given the diminished ability of in vitro-matured mouse eggs to convert injected sperm into males PNs, we wondered if differences could be detected between the sizes of the PNs formed following injection of mouse and bovine sperm into in vivo- and in vitro-matured mouse eggs. We used as a reference point the size attained by the PNs of in vivo-fertilized zygotes. Our observations for mouse fertilization revealed, as expected, that the area of the male PN was larger than that of the female PN, 191 ± 40 µm2 and 116 ± 38 µm2, respectively (Figs. 3 and 4). Remarkably, these differences in sizes between males and female mouse PNs were maintained when heterologous ICSI was performed using either ovulated or IVM oocytes, although the average size of the PNs was smaller when IVM eggs were used (Fig. 3) (P > 0.05). Injection of bovine sperm into mouse eggs, surprisingly, induced an overall increase in the size of both PNs when it was performed in MIIOV eggs, but as it was the case for mouse eggs, the area of the PNs was smaller in IVM oocytes (Fig. 3). For example, the average area of male and female PNs in MIIOV eggs was of 338 ± 78 µm2 and 236 ± 75 µm2, respectively, but it was only of 38 ± 19 µm2 and 151 ± 58 µm2 in MIIIVM eggs, respectively (P < 0.05). It is worth noting that the area of PNs is even greater when the gametes are both bovine. For example, after IVF the PN areas were 1322 ± 411 and 945 ± 292 µm2 and after ICSI the dimensions were 708 ± 269 and 667 ± 329 µm2. For homologous bovine studies, PN formation was always evaluated at 18 hpi and it was not possible to distinguish between male and female PNs. Altogether, our results demonstrate the source of the female gamete and maturation conditions impact the ability to transform sperm heads into male PNs.
[Ca2+]i responses after bovine ICSI into mouse oocytes
The findings that in vivo- and in vitro-matured mouse eggs display different capacity to transform injected sperm into male PNs led us to examine if these two different source of eggs also influenced the [Ca2+]i responses induced by the injection of sperm. Homologous ICSI using mouse gametes and ovulated oocytes triggered [Ca2+]i oscillations with mean intervals ranging between 15 and 25 min (Fig. 5A), which is in agreement with previous data in the literature indicating that in the mouse the [Ca2+]i responses are influenced by the preparation of the sperm and the strains (Sato et al. 1999, Kurokawa & Fissore 2003). Further, these values are similar to those induced by conventional IVF (Knott et al. 2003). Remarkably, the responses were less uniform when ICSI was performed into in vitro-matured eggs, with 11/18 showing slower oscillations (Fig. 5B) and the rest of the oocytes roughly equally divided between high-frequency responders or non-responders (Fig. 5B and inset, respectively). When heterologous ICSI was performed, bovine sperm caused high-frequency [Ca2+]i responses in 11/16 MIIOV eggs, although 5/16 eggs displayed low-frequency oscillations (Fig. 5C and inset, respectively). On the other hand, in vitro-matured mouse eggs displayed highly variable responses, and only 7/23 eggs mounted the expected high-frequency [Ca2+]i oscillations (Fig. 5C) , whereas 7/23 showed fewer and spaced out responses and 9/23 eggs failed to oscillate (Fig. 5D and inset). Collectively, these results indicate that ICSI failure in in vitro-matured eggs is associated with multiple defects in the processing of the sperm heads, including the very early steps of sperm head decondensation and the release of the sperm factor responsible for the [Ca2+]i oscillations that drive egg activation.
[Ca2+]i responses after homologous and heterologous ICSI in mouse oocytes. (A and B) Injection of mouse sperm into MIIOV induced consistent and expected [Ca2+]i responses, although injection into IVM oocytes induced variable responses [Ca2+]i (B). (C) Injection of bull sperm into MIIOV induced mostly high-frequency oscillations, although a few oocytes showed low-frequency oscillations (inset). (D) Injection of bull sperm into IVM induced high-frequency oscillations in half of the oocytes, although in the other half, the oscillations were less frequent or absent (inset).
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
[Ca2+]i responses after homologous and heterologous ICSI in mouse oocytes. (A and B) Injection of mouse sperm into MIIOV induced consistent and expected [Ca2+]i responses, although injection into IVM oocytes induced variable responses [Ca2+]i (B). (C) Injection of bull sperm into MIIOV induced mostly high-frequency oscillations, although a few oocytes showed low-frequency oscillations (inset). (D) Injection of bull sperm into IVM induced high-frequency oscillations in half of the oocytes, although in the other half, the oscillations were less frequent or absent (inset).
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
[Ca2+]i responses after homologous and heterologous ICSI in mouse oocytes. (A and B) Injection of mouse sperm into MIIOV induced consistent and expected [Ca2+]i responses, although injection into IVM oocytes induced variable responses [Ca2+]i (B). (C) Injection of bull sperm into MIIOV induced mostly high-frequency oscillations, although a few oocytes showed low-frequency oscillations (inset). (D) Injection of bull sperm into IVM induced high-frequency oscillations in half of the oocytes, although in the other half, the oscillations were less frequent or absent (inset).
Citation: Reproduction 154, 3; 10.1530/REP-17-0270
Discussion
In this study we examined factors that undermine the success of ICSI in the bovine. We posited that either the inability of bovine sperm to induce [Ca2+]i oscillations and egg activation or the incompetence of the ooplasm to process non-capacitated sperm or a combination of both might explain the low activation rates, asynchronous PN formation and low rates of pre-implantation development observed after ICSI in the bovine. Our results show that although bovine eggs in this study can undergo normal activation and cleavage following IVF or parthenogenetic treatment, they are unable to consistently and timely form male PNs despite supplementing the sperm’s egg-activating signal with physiological activation stimuli. Nevertheless, following heterologous ICSI we observed that in vivo-matured mouse eggs rendered bovine sperm heads into male PNs with high frequency, whereas this was not the case with in vitro-matured eggs. We therefore interpret our results to mean that activation failure is not the main factor limiting the success of ICSI in the bovine. Instead, we propose that in vitro-matured eggs are developmentally compromised and when confronted with a ‘trespassing sperm,’ i.e., a sperm that has not undergone ‘admission clearance’ normally associated with capacitation, acrosome reaction and fusion with the oolema, is unable to timely convert it into a male PN. Improvement of oocyte maturation conditions and better understanding of the molecular mechanisms and cellular changes associated with sperm preparation for fertilization will improve the success of ICSI in the bovine and in other mammalian species.
To ascertain the underlying reasons that limit the success of ICSI in the bovine, we first examined if the gametes used to study this process were competent under optimal conditions. We examined egg activation and embryo development after IVF or parthenogenetic activation. Our results show that the gametes in our study were capable of initiating development, as both procedures induced high rates of cleavage by 72 h along with acceptable rates of development to the blastocyst stage. These results show the bovine gametes used for ICSI are not intrinsically defective. Nevertheless, when performing ICSI with the same gametes, the formation of male PN formation is temporally delayed and greatly reduced. Furthermore, supplementation of ICSI with exogenous activation partially overcomes the deficit, as the rate of male PN formation increases to reach values of near 50% of the injected oocytes, although the formation of the male PN is still delayed. Therefore, as presently performed, the decondensation of the male nucleus appears to be main obstacle to the success of ICSI in the bovine. The difficulty in remodeling the male chromatin following sperm injection is not unique to the bovine, as earlier studies noted impaired and asynchronous chromatin remodeling following ICSI in the mouse (Shoukir et al. 1998, Ajduk et al. 2006), pig (Kren et al. 2003, Lee et al. 2003, Nakai et al. 2011) and humans (Dozortsev et al. 1994, Sousa & Tesarik 1994, Flaherty et al. 1995). Several common factors have been suggested to hinder the remodeling of injected sperm, such as the lack of capacitation and acrosome reaction (Sekhavati et al. 2012, Zambrano et al. 2016, Aguila et al. 2017). Nevertheless, unique features of the bovine spermatozoa may further complicate PN formation in this species. For instance, the plasma membrane of bovine spermatozoa is reportedly more stable than the plasma membrane of other mammalian spermatozoa, which may slow down its disintegration and delay the mingling of the sperm’s content with the ooplasm (Perreault et al. 1988). Bovine spermatozoa also only contain protamine I, unlike the spermatozoa of primates, horses and rodents that have different proportions of protamine II (Balhorn 2007); protamine I contains greater number of disulfide bonds that could hinder sperm nucleus decondensation. In support of this possibility, comparisons of the rates of decondensation of sperm from mammals with different inherent protamine P1/P2 ratios using hamster eggs have shown that the sperm content of protamine P2 appears to regulate the rate with which sperm chromatin decondenses (Perreault et al. 1988). In other words, sperm containing a higher proportion of P2 decondense more quickly than sperm containing lower proportions of it (Perreault et al. 1988). Therefore, the absence of protamine II in bull and boar spermatozoa may greatly undermine their ability to undergo timely decondensation and PN formation following the injection of sperm, which bypasses all the physiological milestones and checkpoints aimed to ensure fertilization by the fittest sperm.
Questions have persisted as to whether the injection procedure or the abnormal [Ca2+]i responses contribute to the low success of ICSI in the bovine. Our results here show that the injection procedure, at least using the piezoelectric-actuated methodology, is not detrimental, as sham-injected groups showed similar PN formation and cleavage rates than parthenogenetic derived-zygotes. Regarding [Ca2+]i responses and despite the highly abnormal oscillations induced by the fertilizing bovine spermatozoon delivered by injection (Malcuit et al. 2006), the results of our studies rule this out as an important factor responsible for undermining the success of this technique. First, even though we supplemented the injection of sperm with injection of PLCzeta1 cRNA, which was capable of inducing nearly identical [Ca2+]i responses to those induced by fertilization, the rates and timing of male PN formation remained low. It is worth noting that female PNs formed in a timely manner in nearly all of the PLCzeta1 cRNA injected eggs, indicating that MPF activity have decreased as expected following activation induced by this stimulus. These data are also consistent with previous results demonstrating that PLCzeta1 cRNA injected is a very efficient activation stimulus to generate SCNT-derived bovine embryos (Ross et al. 2009). Second, another method of activation that is widely used for SCNT, Io + CHX, also was inefficient in our hands to timely decondense and remodel the male chromatin and form a male PN despite inducing 100% female PN formation. Thus, even though we generated optimal [Ca2+]i responses or bypassed them with efficient, exogenous parthenogenetic activation procedures, male nuclear decondensation was largely defective following bovine ICSI. Here, we therefore propose that the defective [Ca2+]i responses are not responsible for the poor success of ICSI, but rather are one of the earliest manifestations of flawed male chromatin remodeling, as the male factor, PLCzeta1, cannot be appropriately released/activated following sperm injection, and it is therefore unable to initiate normal [Ca2+]i oscillations.
It is well established that the female gamete acquires developmental capacity during oocyte maturation (Ferreira et al. 2009, Cheon et al. 2013, Oqani et al. 2016), and that in vitro-matured eggs have reduced developmental competence (Kim et al. 2004, Gioia et al. 2005). In this regard, the great majority of cattle eggs used for experimental purposes are in vitro matured, which, as pointed out, have inherently lower developmental capacity (Kim et al. 2004, Gilchrist 2011, Ptak et al. 2013). Therefore, given that bovine eggs resulting from in vitro maturation are unable to timely remodel the male chromatin of sperm delivered by ICSI, even after exogenous activation, we performed experiments using in vivo-maturated mouse eggs. We have shown that mouse eggs mount normal [Ca2+]i oscillations when injected with sperm of other species (Knott et al. 2003). Therefore, we examined if ovulated mouse eggs were also able to convert the bovine spermatozoa into male PNs. We found that nearly 70% of the bovine sperm were transformed into a male PN within 6 h post injection, which is only slightly lower than what is induced by injection of homologous sperm in the mouse, but considerably higher than what is accomplished by bovine eggs. Furthermore, consistent with the above results, ovulated mouse eggs displayed high-frequency oscillations in response to injection of bovine sperm (Knott et al. 2003). Remarkably, in vitro-matured mouse eggs were largely unable to decondense bovine spermatozoa and were less efficient at remodeling the mouse sperm head. In agreement with these results, IVM mouse eggs were less able to mount [Ca2+]i responses to injection of either sperm. Altogether, these results demonstrate that in vitro-matured eggs have significantly lower capabilities to reprogram sperm nucleus delivered by ICSI. These results suggest that despite significant progress in improving IVM conditions, IVM eggs do not attain the same developmental competence of in vivo-matured eggs. This is agreement with a recent study using in vivo- and in vitro-matured porcine eggs, where it was shown that the use of in vitro-matured eggs compromised the rates of activation and embryo development regardless of the pattern of [Ca2+]i oscillations (Nakai et al. 2016). Further, there is extensive evidence in the literature of multiple methods used by researchers to supplement the ‘compromised’ remodeling ability of eggs obtained after in vitro maturation or to treat sperm to render them more prone to undergo PN formation, including methods of sperm immobilization (Wei & Fukui 1999, Horiuchi et al. 2002), damage to sperm membranes (Arias et al. 2014, Zambrano et al. 2016), or addition of agents to enhance decondensation of spermatozoa DNA such as dithiothreitol (DTT) (Rho et al. 1998, Galli et al. 2003) and heparin-glutathione (Sekhavati et al. 2012). However, thus far, none of these methods has consistently improved the success of ICSI in this species.
Altogether, our results show that bovine sperm is especially resistant to nuclear decondensation, which is possibly associated with the exclusive expression of protamine I to condense its DNA. Further, the near exclusive reliance on in vitro-matured eggs, which contain a suboptimal ooplasm, does not aid in the success of this technique. Considering these results, additional studies are necessary to elucidate the mechanism(s) that underlie the sperm’s preparation for fertilization and devise effective methods that render bovine spermatozoa ‘remodeling-ready’ so that post-fertilizing events after injection can be initiated in a timely and more efficient manner. Such a progress will increase the efficiency of this technique in large domestic species, and will also help in the design of better in vitro fecundation methods to be applied to humans and animal models.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This study was funded in part by a Hatch-NIFA project (grant MAS439) to R Fissore L Aguila received the Doctoral Scholarship (Grant 21120581) from CONICYT (Comisión Nacional de Investigación Científica y Tecnológica), Chile. D Martin-Hidalgo is recipient of a post-doctoral Grant from the Government of Extremadura (Spain) and by Fondo Social Europeo (PO14005).
Acknowledgements
The authors would like to acknowledge Alexandra Flores for her constant support to Chang Lee, Goli Ardestani, Aujan Mehregan and María Gracia Gervasi.
References
Aguila L, Zambrano F, Arias ME & Felmer R 2017 Sperm capacitation pretreatment positively impacts bovine intracytoplasmic sperm injection. Molecular Reproduction and Development. 84 649–659. (doi:10.1002/mrd.22834)
Ajduk A, Yamauchi Y & Ward MA 2006 Sperm chromatin remodeling after intracytoplasmic sperm injection differs from that of in vitro fertilization. Biology of Reproduction 75 442–451. (doi:10.1095/biolreprod.106.053223)
Arias ME, Sanchez R, Risopatron J, Perez L & Felmer R 2014 Effect of sperm pretreatment with sodium hydroxide and dithiothreitol on the efficiency of bovine intracytoplasmic sperm injection. Reproduction, Fertility and Development 26 847–854.
Balhorn R 2007 The protamine family of sperm nuclear proteins. Genome Biology 8 227. (doi:10.1186/gb-2007-8-9-227)
Catt JW & Rhodes SL 1995 Comparative intracytoplasmic sperm injection (ICSI) in human and domestic species. Reproduction, Fertility, and Development 7 161–166; discussion 167. (doi:10.1071/RD9950161)
Chatot CL, Ziomek CA, Bavister BD, Lewis JL & Torres I 1989 An improved culture medium supports development of random-bred 1-cell mouse embryos in vitro. Journal of Reproduction and Fertility 86 679–688. (doi:10.1530/jrf.0.0860679)
Chen SH & Seidel GE 1997 Effects of oocyte activation and treatment of spermatozoa on embryonic development following intracytoplasmic sperm injection in cattle. Theriogenology 48 1265–1273. (doi:10.1016/S0093-691X(97)00369-5)
Cheon B, Lee HC, Wakai T & Fissore RA 2013 Ca2+ influx and the store-operated Ca2+ entry pathway undergo regulation during mouse oocyte maturation. Molecular Biology of the Cell 24 1396–1410. (doi:10.1091/mbc.E13-01-0065)
Dozortsev D, De Sutter P & Dhont M 1994 Behaviour of spermatozoa in human oocytes displaying no or one pronucleus after intracytoplasmic sperm injection. Human Reproduction 9 2139–2144. (doi:10.1093/oxfordjournals.humrep.a138406)
Felmer RN, Arias ME, Munoz GA & Rio JH 2011 Effect of different sequential and two-step culture systems on the development, quality, and RNA expression profile of bovine blastocysts produced in vitro. Molecular Reproduction and Development 78 403–414. (doi:10.1002/mrd.21317)
Ferreira EM, Vireque AA, Adona PR, Meirelles FV, Ferriani RA & Navarro PA 2009 Cytoplasmic maturation of bovine oocytes: structural and biochemical modifications and acquisition of developmental competence. Theriogenology 71 836–848. (doi:10.1016/j.theriogenology.2008.10.023)
Flaherty SP, Payne D, Swann NJ & Mattews CD 1995 Aetiology of failed and abnormal fertilization after intracytoplasmic sperm injection. Human Reproduction 10 2623–2629. (doi:10.1093/oxfordjournals.humrep.a135757)
Florman HM & Fissore R 2014 Fertilization in mammals. In Plant T M, Zeleznik A (eds) Knobil and Neill’s Physiology of Reproduction, vol 1, 4th edn. Academic Press, New York, pp 149–196.
Galli C, Vassiliev I, Lagutina I, Galli A & Lazzari G 2003 Bovine embryo development following ICSI: effect of activation, sperm capacitation and pre-treatment with dithiothreitol. Theriogenology 60 1467–1480. (doi:10.1016/S0093-691X(03)00133-X)
Gilchrist RB 2011 Recent insights into oocyte-follicle cell interactions provide opportunities for the development of new approaches to in vitro maturation. Reproduction, Fertility, and Development 23 23–31. (doi:10.1071/RD10225)
Gioia L, Barboni B, Turriani M, Capacchietti G, Pistilli MG, Berardinelli P & Mattioli M 2005 The capability of reprogramming the male chromatin after fertilization is dependent on the quality of oocyte maturation. Reproduction 130 29–39. (doi:10.1530/rep.1.00550)
Goto K 1997 Current status and future of micromanipulation-assisted fertilization in animals and human. Journal of Reproduction and Development 43 107–119. (doi:10.1262/jrd.43.107)
Goto K, Kinoshita A, Takuma Y & Ogawa K 1990 Fertilisation of bovine oocytes by the injection of immobilised, killed spermatozoa. Veterinary Record 127 517–520.
Horiuchi T, Emuta C, Yamauchi Y, Oikawa T, Numabe T & Yanagimachi R 2002 Birth of normal calves after intracytoplasmic sperm injection of bovine oocytes: a methodological approach. Theriogenology 57 1013–1024. (doi:10.1016/S0093-691X(01)00701-4)
Jenkins TG & Carrell DT 2012 Dynamic alterations in the paternal epigenetic landscape following fertilization. Frontiers in Genetics 3 143.
Kim DH, Ko DS, Lee HC, Lee HJ, Park WI, Kim SS, Park JK, Yang BC, Park SB & Chang WK et al. 2004 Comparison of maturation, fertilization, development, and gene expression of mouse oocytes grown in vitro and in vivo. Journal of Assisted Reproduction and Genetics 21 233–240. (doi:10.1023/B:JARG.0000042008.83699.cc)
Kimura Y & Yanagimachi R 1995 Intracytoplasmic sperm injection in the mouse. Biology of Reproduction 52 709–720. (doi:10.1095/biolreprod52.4.709)
Kimura Y, Yanagimachi R, Kuretake S, Bortkiewicz H, Perry AC & Yanagimachi H 1998 Analysis of mouse oocyte activation suggests the involvement of sperm perinuclear material. Biology of Reproduction 58 1407–1415. (doi:10.1095/biolreprod58.6.1407)
Knott JG, Kurokawa M & Fissore RA 2003 Release of the Ca(2+) oscillation-inducing sperm factor during mouse fertilization. Developmental Biology 260 536–547. (doi:10.1016/S0012-1606(03)00251-3)
Knott JG, Kurokawa M, Fissore RA, Schultz RM & Williams CJ 2005 Transgenic RNA interference reveals role for mouse sperm phospholipase Czeta in triggering Ca2+ oscillations during fertilization. Biology of Reproduction 72 992–996. (doi:10.1095/biolreprod.104.036244)
Kren R, Kikuchi K, Nakai M, Miyano T, Ogushi S, Nagai T, Suzuki S, Fulka J, Fulka J Jr. 2003 Intracytoplasmic sperm injection in the pig: where is the problem? Journal of Reproduction and Development 49 271–273. (doi:10.1262/jrd.49.271)
Kurokawa M & Fissore RA 2003 ICSI-generated mouse zygotes exhibit altered calcium oscillations, inositol 1,4,5-trisphosphate receptor-1 down-regulation, and embryo development. Molecular Human Reproduction 9 523–533. (doi:10.1093/molehr/gag072)
Kurokawa M, Sato K, Wu H, He C, Malcuit C, Black SJ, Fukami K & Fissore RA 2005 Functional, biochemical, and chromatographic characterization of the complete [Ca2+]i oscillation-inducing activity of porcine sperm. Developmental Biology 285 376–392. (doi:10.1016/j.ydbio.2005.06.029)
Lee JW, Tian XC & Yang X 2003 Failure of male pronucleus formation is the major cause of lack of fertilization and embryo development in pig oocytes subjected to intracytoplasmic sperm injection. Biology of Reproduction 68 1341–1347. (doi:10.1095/biolreprod.102.009001)
Lee HC, Yoon SY, Lykke-Hartmann K, Fissore RA & Carvacho I 2016 TRPV3 channels mediate Ca(2)(+) influx induced by 2-APB in mouse eggs. Cell Calcium 59 21–31. (doi:10.1016/j.ceca.2015.12.001)
Malcuit C, Maserati M, Takahashi Y, Page R & Fissore RA 2006 Intracytoplasmic sperm injection in the bovine induces abnormal [Ca2+]i responses and oocyte activation. Reproduction, Fertility, and Development 18 39–51. (doi:10.1071/RD05131)
Mizuno K, Hoshi K & Huang T 2002 Fertilization and embryo development in a mouse ICSI model using human and mouse sperm after immobilization in polyvinylpyrrolidone. Human Reproduction 17 2350–2355. (doi:10.1093/humrep/17.9.2350)
Nakai M, Kashiwazaki N, Ito J, Maedomari N, Ozawa M, Shino M, Noguchi J, Kaneko H & Kikuchi K 2011 Factors affecting fertilization and embryonic development during intracytoplasmic sperm injection in pigs. Journal of Reproduction and Development 57 183–187. (doi:10.1262/jrd.10-200E)
Nakai M, Ito J, Suzuki SI, Fuchimoto DI, Sembon S, Suzuki M, Noguchi J, Kaneko H, Onishi A & Kashiwazaki N et al. 2016 Lack of calcium oscillation causes failure of oocyte activation after intracytoplasmic sperm injection in pigs. Journal of Reproduction and Development 62 615–621. (doi:10.1262/jrd.2016-113)
Navarrete FA, Garcia-Vazquez FA, Alvau A, Escoffier J, Krapf D, Sanchez-Cardenas C, Salicioni AM, Darszon A & Visconti PE 2015 Biphasic role of calcium in mouse sperm capacitation signaling pathways. Journal of Cellular Physiology 230 1758–1769. (doi:10.1002/jcp.24873)
Oqani RK, Lin T, Lee JE, Kim SY, Sa SJ, Woo JS & Jin DI 2016 Inhibition of P-TEFb disrupts global transcription, oocyte maturation, and embryo development in the mouse. Genesis 54 470–482. (doi:10.1002/dvg.22961)
Palermo G, Joris H, Devroey P & Van Steirteghem AC 1992 Pregnancies after intracytoplasmic injection of single spermatozoon into an oocyte. Lancet 340 17–18. (doi:10.1016/0140-6736(92)92425-F)
Parrish JJ, Susko-Parrish JL, Leibfried-Rutledge ML, Critser ES, Eyestone WH & First NL 1986 Bovine in vitro fertilization with frozen-thawed semen. Theriogenology 25 591–600. (doi:10.1016/0093-691X(86)90143-3)
Parrish JJ, Susko-Parrish J, Winer MA & First NL 1988 Capacitation of bovine sperm by heparin. Biology of Reproduction 38 1171–1180. (doi:10.1095/biolreprod38.5.1171)
Perreault SD, Barbee RR, Elstein KH, Zucker RM & Keefer CL 1988 Interspecies differences in the stability of mammalian sperm nuclei assessed in vivo by sperm microinjection and in vitro by flow cytometry. Biology of Reproduction 39 157–167. (doi:10.1095/biolreprod39.1.157)
Perry AC, Wakayama T, Kishikawa H, Kasai T, Okabe M, Toyoda Y & Yanagimachi R 1999 Mammalian transgenesis by intracytoplasmic sperm injection. Science 284 1180–1183. (doi:10.1126/science.284.5417.1180)
Ptak GE, D’Agostino A, Toschi P, Fidanza A, Zacchini F, Czernik M, Monaco F & Loi P 2013 Post-implantation mortality of in vitro produced embryos is associated with DNA methyltransferase 1 dysfunction in sheep placenta. Human Reproduction 28 298–305. (doi:10.1093/humrep/des397)
Rho GJ, Kawarsky S, Johnson WH, Kochhar K & Betteridge KJ 1998 Sperm and oocyte treatments to improve the formation of male and female pronuclei and subsequent development following intracytoplasmic sperm injection into bovine oocytes. Biology of Reproduction 59 918–924. (doi:10.1095/biolreprod59.4.918)
Rho GJ, Lee SL, Kim YS, Yeo HJ, Ock SA, Balasubramanian S & Choe SY 2004 Intracytoplasmic sperm injection of frozen-thawed bovine oocytes and subsequent embryo development. Molecular Reproduction and Development 68 449–455. (doi:10.1002/mrd.20110)
Rizos D, Fair T, Papadopoulos S, Boland MP & Lonergan P 2002 Developmental, qualitative, and ultrastructural differences between ovine and bovine embryos produced in vivo or in vitro. Molecular Reproduction and Development 62 320–327. (doi:10.1002/mrd.10138)
Ross PJ, Rodriguez RM, Iager AE, Beyhan Z, Wang K, Ragina NP, Yoon SY, Fissore RA & Cibelli JB 2009 Activation of bovine somatic cell nuclear transfer embryos by PLCZ cRNA injection. Reproduction 137 427–437. (doi:10.1530/REP-08-0419)
Sato MS, Yoshitomo M, Mohri T & Miyazaki S 1999 Spatiotemporal analysis of [Ca2+]i rises in mouse eggs after intracytoplasmic sperm injection (ICSI). Cell Calcium 26 49–58. (doi:10.1054/ceca.1999.0053)
otherSaunders CM, Larman MG, Parrington J, Cox LJ, Royse J, Blayney LM, Swann K & Lai FA 2002 PLC zeta: a sperm-specific trigger of Ca(2+) oscillations in eggs and embryo development. Development 129 3533–3544.
Schultz RM & Kopf GS 1995 Molecular basis of mammalian egg activation. Current Topics in Developmental Biology 30 21–62. (doi:10.1016/s0070-2153(08)60563-3)
Sekhavati MH, Shadanloo F, Hosseini MS, Tahmoorespur M, Nasiri MR, Hajian M & Nasr-Esfahani MH 2012 Improved bovine ICSI outcomes by sperm selected after combined heparin-glutathione treatment. Cell Reprogram 14 295–304.
Shoukir Y, Chardonnens D, Campana A & Sakkas D 1998 Blastocyst development from supernumerary embryos after intracytoplasmic sperm injection: a paternal influence? Human Reproduction 13 1632–1637. (doi:10.1093/humrep/13.6.1632)
Sousa M & Tesarik J 1994 Ultrastructural analysis of fertilization failure after intracytoplasmic sperm injection. Human Reproduction 9 2374–2380. (doi:10.1093/oxfordjournals.humrep.a138455)
Sutovsky P & Schatten G 2000 Paternal contributions to the mammalian zygote: fertilization after sperm-egg fusion. International Review of Cytology 195 1–65. (doi:10.1016/s0074-7696(08)62703-5)
Suttner R, Zakhartchenko V, Stojkovic P, Muller S, Alberio R, Medjugorac I, Brem G, Wolf E & Stojkovic M 2000 Intracytoplasmic sperm injection in bovine: effects of oocyte activation, sperm pretreatment and injection technique. Theriogenology 54 935–948. (doi:10.1016/S0093-691X(00)00403-9)
Virant-Klun I, Knez K, Tomazevic T & Skutella T 2013 Gene expression profiling of human oocytes developed and matured in vivo or in vitro. BioMed Research Internationa 2013 article ID 879489 23 1138–1144. (doi:10.1155/2013/879489)
Wakai T & Fissore RA 2013 Ca(2+) homeostasis and regulation of ER Ca(2+) in mammalian oocytes/eggs. Cell Calcium 53 63–67. (doi:10.1016/j.ceca.2012.11.010)
Wei H & Fukui Y 1999 Effects of bull, sperm type and sperm pretreatment on male pronuclear formation after intracytoplasmic sperm injection in cattle. Reproduction, Fertility, and Development 11 59–65. (doi:10.1071/RD98106)
Yanagida K,, Katayose H, Hirata S, Yazawa H, Hayashi S & Sato A 2001 Influence of sperm immobilization on onset of Ca(2+) oscillations after ICSI. Human Reproduction 16 148–152. (doi:10.1093/humrep/16.1.148)
Yanagimachi R 2005 Intracytoplasmic injection of spermatozoa and spermatogenic cells: its biology and applications in humans and animals. Reproductive BioMedicine Online 10 247–288. (doi:10.1016/S1472-6483(10)60947-9)
Zambrano F, Aguila L, Arias ME, Sanchez R & Felmer R 2016 Improved preimplantation development of bovine ICSI embryos generated with spermatozoa pretreated with membrane-destabilizing agents lysolecithin and Triton X-100. Theriogenology 86 1489–1497. 10.1016/j.theriogenology.2016.05.007
