Postovulatory ageing modifies sperm-induced Ca²⁺ oscillations in mouse oocytes through a conditions-dependent, multi-pathway mechanism

Abstract

Postovulatory ageing of mammalian oocytes occurs between their ovulation and fertilization and has been shown to decrease their developmental capabilities. Aged oocytes display numerous abnormalities, including altered Ca²⁺ signalling. Fertilization-induced Ca²⁺ oscillations are essential for activation of the embryonic development, therefore maintaining proper Ca²⁺ homeostasis is crucial for the oocyte quality. In the present paper, we show that the mechanism underlying age-dependent alterations in the pattern of sperm-triggered Ca²⁺ oscillations is more complex and multifaceted than previously believed. Using time-lapse imaging accompanied by immunostaining and molecular analyses, we found that postovulatory ageing affects the amount of Ca²⁺ stored in the cell, expression of Ca²⁺ pump SERCA2, amount of available ATP and distribution of endoplasmic reticulum and mitochondria in a manner often strongly depending on ageing conditions (in vitro vs. in vivo). Importantly, those changes do not have to be caused by oxidative stress, usually linked with the ageing process, as they occur even if the amount of reactive oxygen species remains low. Instead, our results suggest that aberrations in Ca²⁺ signalling may be a synergistic result of ageing-related alterations of the cell cycle, cytoskeleton, and mitochondrial functionality.

Introduction

Postovulatory ageing in mammals occurs between ovulation of mature oocytes and their fertilization and has been shown to decrease oocytes’ ability to react properly to sperm penetration and to develop further into the embryo^1–8. Aged oocytes display a wide range of various abnormalities, such as dysfunction of actomyosin and microtubular cytoskeleton^8–13, decreased activity of M-phase promoting kinases^11,14,15, deregulation of energy metabolism^2,16–18 and epigenetic alterations^19,20. The extent of these abnormalities depends on the length of ageing period and, most likely, the genetic background of the cell. It has been also reported that postovulatory ageing disturbs Ca²⁺ signalling in oocytes and alters the pattern of sperm-induced Ca²⁺ oscillations^4,21–27. The exact mechanism of this particular disturbance is not known, although it has been suggested that it may be caused by ageing-dependent oxidative stress²⁵.

Fertilization-induced Ca²⁺ oscillations are essential for activation of the embryo development. They trigger processes such as completion of meiosis, establishment of the block to polyspermy and recruitment of maternal mRNAs required for activation of the embryonic genome^28,29. In addition, Ca²⁺ oscillations regulate functioning of mitochondria^30–32 and influence gene expression during further development of the embryo^33,34. It has been shown that Ca²⁺ must be elevated for different time to induce properly each of these processes. If the number of Ca²⁺ transients is too low (i.e. the total duration of Ca²⁺ elevation is too short), the embryo implantation is impaired; on the other hand, an excessive number of Ca²⁺ transients (i.e. Ca²⁺ is elevated for too long) affects postimplantation development of the embryo^33–35.

Ca²⁺ oscillations are triggered in oocytes by phospholipase C zeta (PLC zeta), introduced by a fertilizing sperm³⁶. PLC zeta cleaves phosphatidylinositol 4,5-bisphosphate (PIP₂) to 1,4,5-inositol triphosphate (IP₃) and diacylglycerol. IP₃ binds to its receptor (IP₃ receptor type 1, IP3R1, dominates in oocytes³⁷) located in the endoplasmic reticulum (ER) membrane. IP3R1 acts as a Ca²⁺ channel and opens upon IP₃ binding, enabling the release of Ca²⁺ ions from the ER lumen into the cytoplasm. When the cytoplasmic Ca²⁺ concentration increases above a threshold level (low micromolar concentrations), the Ca²⁺ channel closes; the release of Ca²⁺ stops and the uptake of Ca²⁺ ions to the ER lumen is facilitated. The uptake of Ca²⁺ ions is mediated by ATP-dependent Ca²⁺ pumps (the sarco/endoplasmic reticulum Ca²⁺ ATPases, SERCA). Restoration of the Ca²⁺ level in the ER is required for the next Ca²⁺ release^38,39. This interplay between IP3R1-mediated Ca²⁺ release and SERCA-mediated Ca²⁺ uptake constitutes the main mechanism of cytoplasmic Ca²⁺ oscillations observed in fertilized oocytes. Ca²⁺ oscillations cease after few hours when oocytes enter interphase^40–42.

In the present paper, we show that postovulatory ageing in in vivo conditions has significantly more detrimental effect on Ca²⁺ homeostasis in mouse oocytes than in vitro ageing. In both conditions, postovulatory ageing affects several elements of the mechanism generating fertilization-induced Ca²⁺ oscillations, such as the amount of Ca²⁺ stored in the cell, expression of IP3R1 and SERCA2, the amount of available ATP and distribution of ER and mitochondria, but in a different way or to a different extent. We also argue that those changes are not necessarily caused by oxidative stress, as they occur even if the reactive oxygen species (ROS) level is not increased. Instead, our data suggest that aberrations in Ca²⁺ oscillatory pattern are a synergistic result of many ageing-related modifications, including alterations of the cell cycle, cytoskeleton, and mitochondrial functionality.

Results and Discussion

In vitro and in vivo postovulatory ageing differently alters the pattern of Ca²⁺ oscillations

In order to examine the effect of different postovulatory ageing conditions (a short and a long duration, in vitro and in vivo) on the pattern of Ca²⁺ oscillations generated in fertilized mouse oocytes, we subjected mature, metaphase II oocytes to 9- or 25-hour-long aging, either in in vitro culture or in female oviducts. Then, we labelled oocytes for Ca²⁺ ions using a fluorescent dye Oregon Green 488 BAPTA-1AM (Oregon Green BAPTA), fertilized them in vitro and subjected to time-lapse imaging. Representative Ca²⁺ traces are presented in Supplementary Fig. S1. 9 hours of ageing, both in in vitro and in vivo conditions, increased the frequency of Ca²⁺ oscillations, as compared to freshly ovulated counterparts. However, only in vivo ageing shortened their total duration as well (Fig. 1A,B, Table 1). 25 hours of ageing led to more versatile reactions. In both in vitro and in vivo experimental variants, some oocytes displayed very frequent Ca²⁺ oscillations, while others generated Ca²⁺ transients with a frequency similar to that in freshly ovulated oocytes (on average, the mean interval between Ca²⁺ transients in in vivo aged oocytes was significantly shorter than in fresh ones). The same rule applied to the total length of Ca²⁺ oscillations: in some aged oocytes they were shorter and in some significantly longer than in freshly ovulated ones (Fig. 1A,B, Table 1). Although the average duration of Ca²⁺ oscillations in oocytes aged for 25 hrs did not differ from that observed in the fresh counterparts, in over 25% of aged oocytes Ca²⁺ oscillations were still in progress when the recording ended (even though they started on average at the same time after the imaging onset as in the fresh oocytes), suggesting that the 25-hour-long ageing may prolong Ca²⁺ oscillations. It has been shown that in mouse Ca²⁺ oscillations cease when fertilized oocytes enter interphase of the 1^st embryonic division. This decline may be caused by PLC zeta sequestration into the newly formed pronuclei^40–42 (although this process seems not to be universal across species⁴³) or/and decrease in IP3R1 phosphorylation by M-phase kinases, IP3R1 degradation, and change in ER distribution^44–47. Oocytes aged for 25 hrs often did not form pronuclei during 7 hrs of the recording (data not shown), and instead they remained arrested in M-phase with disarrayed spindles, and, in consequence, displayed most likely high activity of M-phase kinases. It has been reported before that oocytes arrested in metaphase II by microtubule depolymerizing drugs, such as nocodazole or colcemid, generated significantly prolonged Ca²⁺ oscillations (lasting through the whole recording period, i.e. even up to 22 hrs) of low frequency and constant amplitude^48,49. It seems that the inability of oocytes aged for 25 hrs to enter interphase is related to the spindle abnormality present already at the time of fertilization that inhibits M-phase/interphase transition through the spindle assembly checkpoint (SAC)⁵⁰ (Fig. 1C,D). It has been shown previously that postovulatory ageing negatively affects microtubular cytoskeleton and spindle structure in oocytes^9,10. Difficulties with the pronuclear formation in oocytes aged in vitro for 24 hrs have been also reported by Lacham-Kaplan and Trounson³.

Figure 1.

Fertilization-induced Ca²⁺ oscillations in postovulatory aged oocytes. (A,B) Point charts presenting the total duration of Ca²⁺ oscillations and the mean interval between Ca²⁺ transients in freshly ovulated oocytes, oocytes aged for 9 and 25 hrs in vitro (A) and in vivo (B). (C,D) Representative immunostainings (β-tubulin in green, DNA in red) of nuclear apparatus in unfertilized freshly ovulated and aged oocytes (C), and 6 hrs after fertilization of freshly ovulated and aged oocytes (D). Scale bar 50 µm. (E–F) Point charts presenting the amplitude and the duration of the 1^st Ca²⁺ transient in freshly ovulated oocytes and oocytes aged for 9 and 25 hrs in vitro (E) and in vivo (F). (G–H) Point charts presenting the rates of Ca²⁺ increase and decrease during the 1^st Ca²⁺ transient in freshly ovulated oocytes and oocytes aged for 9 and 25 hrs in vitro (G) and in vivo (H). (A,B, E–H) Each dot/triangle represents one oocyte, the number of analysed oocytes is included in Table 1. Triangles (in (A,B)) mark oocytes that did not finish Ca²⁺ oscillations during the filming.

Table 1.

Effect of postovulatory aging on the pattern of Ca²⁺ oscillations and Ca²⁺ ER store in oocytes.

	fresh	9 hrs in vitro	25 hrs in vitro	9 hrs in vivo	25 hrs in vivo
	Median (Q1; Q3)
No. of Ca2+ transients	7.0a (6.0; 8.0) n = 64	8.0f (7.0; 9.0) n = 57	8.0 (6.0; 10.0) n = 30	6.0f,b (5.0; 7.5) n = 51	12.0a,b (7.0; 19.0) n = 43
Duration of Ca2+ oscillations (min)	158.0a (128.3; 182.2) n = 64	159.9b (141.7; 171.8) n = 57	165.3c (107,1; 209.0) n = 30	88.0a,b,c,d (64.1; 114.0) n = 47	183.5d (117.9; 279.5) n = 42
Mean interval between Ca2+ transients during 1st 2 hrs (min)	23.4a,b,f (20.5; 25.3) n = 64	19.6f (16.2; 21.3) n = 57	20.7 (15.5; 26.0) n = 30	16.8a (14.6; 19.0) n = 47	14 b (10.1; 19.9) n = 42
Amplitude of the 1st Ca2+ transient (ΔF/F0)	0.9a,b,c (0.8; 1.0) n = 64	0.8d,e (0.7; 0.9) n = 57	0.7a (0.6; 0.8) n = 30	0.6b,d (0.5; 0.7) n = 50	0.6c,e (0.5; 0.6) n = 43
Amplitude of the 3rd Ca2+ transient (ΔF/F0)	0.8a,b,c (0.8; 0.9) n = 64	0.8d,h (0.7; 0.8) n = 57	0.7a,h (0.6; 0.7) n = 30	0.7b (0.6; 0.7) n = 45	0.6c,d (0.5; 0.6) n = 41
Duration of the 1st Ca2+ transient (min)	3.2a,b,f (2.8; 3.5) n = 64	2.2a,g (2.0; 2.5) n = 57	1.7b,c,h (1.4; 2.0) n = 30	3.3c,g (2.5; 4.7) n = 50	2.2f,h (1.8; 3.0) n = 43
Duration of the 3rd Ca2+ transient (min)	1.1 (1.0; 1.2) n = 64	1.2 (1.0; 1.2) n = 57	1.0 (0.8; 1.2) n = 30	1.2 (1.0; 1.2) n = 45	1.0 (0.8; 1.2) n = 41
Rate of the 1st Ca2+ increase (min−1)	3.1a,b (2.4; 3.7) n = 64	3.9c,d (3.1; 4.6) n = 57	3.0h,f (2.3; 3.5) n = 30	2.2a,c,h (1.4; 2.7) n = 49	2.0b,d,f (1.6; 2.3) n = 43
Rate of the 3rd Ca2+ increase (min−1)	4.7a,b,c (4.4; 5.2) n = 64	4.3d,h (4.0; 4.6) n = 57	3.7a,h (3.4; 4.0) n = 30	3.8b (3.3; 4.2) n = 45	3.1c,d (2.9; 3.3) n = 41
Rate of the 1st Ca2+ decrease (min−1)	−4.1a,b,c (−4.4; −3.6) n = 64	−3.5d,f (−3.9; −3.2) n = 57	−2.4a,f (−3.4; −2.0) n = 30	−3.1b (−3.4; −2.5) n = 49	−2.7c,d (−3.1; −2.1) n = 43
Rate of the 3rd Ca2+ decrease (min−1)	−4.7a,b,c (−5.2; −4.4) n = 64	−4.2f,h (−4.5; −3.9) n = 57	−3.4a,h (−3.8; −2.9) n = 30	−3.9b (−4.2; −3.4) n = 45	−3.2c,f (−3.7; −2.9) n = 41
Amplitude of the TG-induced Ca2+ release (ΔF/F0)	1.6a,b,c, (1.4; 1.7) n = 65	1.3a,f (1.3; 1.4) n = 75	1.3b,g (1.2; 1.4) n = 53	1.3c,f,g (1.2; 1.3) n = 63	1.4c (1.3; 1.4) n = 56

Values marked with the same letter are significantly different: ^a,b,c,d,ep < 0.001, ^f,gp < 0.01, ^hp < 0.05.

Postovulatory ageing in vivo decreased the amplitude of Ca²⁺ oscillations (we analysed in detail the 1^st and the 3^rd Ca²⁺ transient), both in the 9- and the 25-hour-long variants. In case of in vitro ageing, Ca²⁺ transients had lower amplitudes only after 25 hrs. We also noticed that the duration of the 1^st Ca²⁺ peak in oocytes aged for 9 hrs in vitro, but not in vivo, was shorter than in freshly ovulated ones, whereas in oocytes aged for 25 hrs the shortening occurred in both conditions (Fig. 1E,F, Table 1, Supplementary Fig. S2A,B). Ageing altered also dynamics of Ca²⁺ transients. In case of in vitro ageing (but only after 25 hrs), the concentration of cytoplasmic Ca²⁺ increased at a slower rate during the 1^st Ca²⁺ transient and decreased at a slower rate during the 1^st and the 3^rd Ca²⁺ transients while compared to fresh oocytes. In in vivo conditions, the increase and decrease of Ca²⁺ cytoplasmic levels during the 1^st and the 3^rd Ca²⁺ spikes slowed down already after 9 hrs and remained like this after 25 hrs (Fig. 1G,H, Table 1, Supplementary Figs S1B, S2C,D). Noteworthy, changes in the dynamics of Ca²⁺ oscillations observed in oocytes aged for 25 hrs (apart from the duration of Ca²⁺ oscillations in the in vivo group and the duration of the 1^st Ca²⁺ transient in the in vitro group) do not seem to be related to their M-phase kinase activity: they are comparable in oocytes that progressed to interphase after fertilization and in those that remained in M-phase (Supplementary Table S1). It is important to note that our measurements of the amplitudes and, in consequence, also the rates of Ca²⁺ increase/decrease, are only estimations, as Oregon Green BAPTA is not a ratiometric dye and its response to increasing Ca²⁺ concentration is not linear (its output flattens for higher Ca²⁺ concentrations)⁵¹. Nevertheless, the tendencies we observed in the analysed groups seem to be real, as they accord with the previously reported data^{4,17,21,22,25}. Due to the characteristics of Oregon Green BAPTA susceptibility to Ca²⁺, the actual differences in the Ca²⁺ concentrations between the groups may be just more pronounced than what we have shown.

In summary, in vivo postovulatory ageing alters the Ca²⁺ oscillation pattern faster than in vitro ageing: it changed 8 out of 11 analysed parameters after 9 hrs, comparing to 2/11 in the in vitro variant. After 25 hrs in both variants more than half of the parameters were changed: 9/11 in in vivo and 6/11 in in vitro ageing. Ageing affected the frequency of Ca²⁺ transients, their amplitude, duration and dynamics of Ca²⁺ increase/decrease, and most likely, the total duration of Ca²⁺ oscillations. Interestingly, in vivo and in vitro aging affects differently not only Ca²⁺ oscillations, but also activities of M-phase kinases (MPF and MAPK) and profile of protein synthesis; in this case, the same as in our experiments, the effect of in vivo ageing is much more severe than that of in vitro aging^15,52.

Postovulatory ageing decreases the ER Ca²⁺ store and SERCA2 expression

Ageing-related changes in the Ca²⁺ oscillation pattern may be caused by the modified amount of Ca²⁺ stored in oocytes or/and altered expression of proteins involved in generation of the oscillations. To assess the amount of Ca²⁺ stored in the ER, we labelled oocytes with Oregon Green BAPTA and treated with thapsigargin (TG), a SERCA inhibitor, which induces a release of Ca²⁺ from the ER cisterns to the cytoplasm. Increase in the Oregon Green BAPTA fluorescence observed afterwards reflects the size of the ER Ca²⁺ store. Both in vitro and in vivo ageing, for 9 and 25 hrs, led to a decrease in the amount of Ca²⁺ available from the ER (Fig. 2A,B, Table 1). We also examined mRNA and protein levels of two main regulators of Ca²⁺ oscillations: IP3R1 and SERCA2. Postovulatory ageing decreased both mRNA and protein levels of SERCA2, although the decline in the protein level appeared only after 25 hrs. In case of IP3R1, both mRNA and protein expression remained relatively constant during postovulatory ageing (Fig. 2C–E, Supplementary Fig. S2E).

Figure 2.

Effect of postovulatory ageing on Ca²⁺ ER store, expression of IP3R1 and SERCA2 and amount of ROS. (A,B) Mean Ca²⁺ release triggered by thapsigargin (TG), calculated for freshly ovulated oocytes and oocytes aged for 9 and 25 hrs in vitro (A) and in vivo (B). The number of analysed oocytes is included in Table 1. Mean values +/− SD are shown. Time-point “0” was set as a moment when the cytoplasmic Ca²⁺ concentration in oocytes started to rise. (C-D) Relative expression of mRNA for Ip3r1 (Itpr1) (C) and Serca2 (Atp2a2) (D) genes in freshly ovulated and aged oocytes. (E) Western blot analysis of protein expression for IP3R1 and SERCA2. Results of the densitometric analysis are presented as numbers below the blots. Ponceau S staining was used to confirm an equal sample loading. (F) Mean intensity of CellROX Orange staining of reactive oxygen species (ROS) calculated for 65 freshly ovulated oocytes, 65 and 85 oocytes aged in vitro (for 9 and 25 hrs, respectively), 85 and 83 oocytes aged in vivo (for 9 and 25 hrs, respectively), 23 fresh oocytes treated with H₂O₂ and 28 oocytes aged for 25 hrs in vitro in medium supplemented with NAC. All intensity values were normalized with the mean fluorescence intensity calculated in the particular experiment for the control, freshly ovulated oocytes. ^ap < 0.001 vs. fresh oocytes, ^bp < 0.001 vs. 25hrs in vitro, ^cp < 0.001 vs. 9 hrs in vitro. (G) CellROX Orange staining in representative oocytes from the experimental variants analysed in (F). The dashed white line marks oocytes’ circumferences. Scale bar 50 µm. (C-D, F) Graphs present medians and the 1^st and the 3^rd quartile values. The ends of the whiskers are set at 1.5*IQR above the third quartile and 1.5*IQR below the first quartile. Dots show the minimum and maximum values if they are outside the range (outliers).

Importantly, the decreased expression of SERCA2 protein, especially in in vivo aged oocytes, correlates with the slower rate of Ca²⁺ decrease during Ca²⁺ transients. Depleted ER Ca²⁺ stores explain, on the other hand, the lower amplitude of Ca²⁺ transients in postovulatory aged oocytes, which accords with the previous reports^4,53,54. It has been suggested that the decreased amount of Ca²⁺ stored in the ER of ageing oocytes may be caused by a Ca²⁺ leak resulting from expression of a truncated caspase 3-cleaved form of IP3R1^27,55,56, likely triggered by increased expression of miR-98⁵⁷. The low amplitude of Ca²⁺ oscillations can be also related to the deregulation of other posttranslational modifications of IP3R1. It has been shown that IP3R1 functionality, particularly its sensitivity to IP₃, depends on the phosphorylation status of the receptor^44,58,59. MPF and MAPK are key kinases phosphorylating IP3R1 and stimulating its action^44,58,60. As postovulatory ageing leads to a decrease in their activity^11,14,15,20, IP3R1 phosphorylation, and, in result, its functionality, is inhibited²⁷. Decreased functionality of IP3R1 could also explain the slower Ca²⁺ release rate during Ca²⁺ transients. However, the slower rate of Ca²⁺ rise could be caused also simply by the lower cellular Ca²⁺ stores^4,53,54. Moreover, it is possible that postovulatory ageing affects plasma membrane channels responsible for Ca²⁺ entry. It has been shown recently that mouse oocytes lacking both TRPM7 and Ca_V3.2 channels stop oscillating prematurely⁶¹. Inhibition of Ca²⁺ oscillations, in terms of the peak number and amplitude, has been also observed in porcine oocytes with downregulated expression of ORAI1 or STIM1 proteins, the key elements in the store-operated Ca²⁺ entry (SOCE) pathway, and in porcine oocytes overexpressing ORAI1 (but not STIM1)^62,63.

Cumulus cells are not sufficient to trigger the in vivo ageing phenotype

It has been suggested that cumulus cells extrude factors that accelerate ageing of oocytes^14,64–66. We noticed that indeed, Ca²⁺ oscillations in oocytes aged for 9 hrs in vivo, i.e. in the company of cumulus cells, were shorter, displayed less Ca²⁺ spikes, and the 1^st Ca²⁺ transient had on average lower amplitude and slower rate of Ca²⁺ release than oocytes aged for the same time in vitro. Those differences between the ageing conditions diminished or disappeared after 25 hrs. We, therefore, examined whether the presence of cumulus cells during the 9-hour-long in vitro culture enhanced the effect of ageing on Ca²⁺ oscillations. Interestingly, we did not notice such a tendency. Although the amount of Ca²⁺ stored in the ER was depleted in oocytes cultured with cumulus cells to the level typical for oocytes aged in vivo (Supplementary Fig. S3A, Table 1, Supplementary Table S2), Ca²⁺ oscillations in oocytes aged for 9 hrs in vitro in presence of cumulus cells resembled – regarding all the above-mentioned parameters - oscillations in the in vitro, not in vivo, aged oocytes (Supplementary Fig. S3B–G, Table 1, Supplementary Table S2). Thus, cumulus cells are not sufficient to generate changes in Ca²⁺ oscillations typical for in vivo ageing. In this respect, our results accord with the data reported by Takahashi et al.⁴, who have also shown no pronounced negative effect of cumulus cells on fertilization-induced Ca²⁺ response in in vitro aged oocytes. Nonetheless, our data suggest that there are some factors present in an oviduct and absent in in vitro culture that accelerate oocyte ageing.

Oxidative stress is not requisite for the age-related modifications of Ca²⁺ oscillations

It has been suggested in the literature that ageing-related changes in oocyte physiology, including Ca²⁺ homeostasis, can be attributed to the negative impact of oxidative stress^67–69. To investigate this hypothesis, we first analysed the amount of reactive oxygen species (ROS) in oocytes aged in vitro and in vivo for 9 or 25 hrs. Measurements of fluorescence intensity of CellROX Orange, a ROS-sensitive dye, indicated that 9 hrs of in vitro ageing did not affect the abundance of ROS in oocytes, and a significant increase in ROS amount was observed only after 25 hrs. In contrast, both 9 and 25 hrs of in vivo ageing led to a decrease in ROS level, as compared to freshly ovulated oocytes (Fig. 2F,G).

Although oxidative stress did not seem to be related to altered Ca²⁺ homeostasis in in vivo aged oocytes or in oocytes aged in vitro for 9 hrs, it still may have been responsible for changes in Ca²⁺ signalling recorded in oocytes aged in vitro for 25 hrs. As 15 min incubation in 100 µM H₂O₂ induces a similar increase in the ROS level as 25-hour-long in vitro ageing (Fig. 2F,G), we examined whether it could also mimic the ageing impact on Ca²⁺ oscillations. Experiments with TG and A23187 ionophore showed that H₂O₂ led to severe depletion of the ER Ca²⁺ stores, but not to depletion of Ca²⁺ stored in other cellular compartments (Supplementary Fig. S4A). Moreover, oocytes pre-treated with H₂O₂ and then fertilized generated Ca²⁺ transients of lower amplitude and slower dynamics of Ca²⁺ increase/decrease than fresh untreated oocytes, the same as oocytes aged for 25 hrs in vitro. On the other hand, H₂O₂ treatment did not shorten the duration of the 1^st Ca²⁺ spike, as 25-hour-long in vitro ageing did. Instead, it shortened the total duration of Ca²⁺ oscillations, the mean interval between subsequent Ca²⁺ spikes and the duration of the 3^rd Ca²⁺ spike – alterations that were not recorded in oocytes aged in vitro for 25 hrs (Fig. 3A–C, Tables 1, 2, Supplementary Fig. S4B–D).

Figure 3.

Impact of oxidative stress on Ca²⁺ homeostasis in oocytes. (A–C) Point charts presenting the total duration of Ca²⁺ oscillations and the mean interval between Ca²⁺ transients (A), the amplitude and the duration of the 1^st Ca²⁺ transient (B) and the rates of Ca²⁺ increase and decrease during the 1^st Ca²⁺ transient (C) in fresh oocytes, fresh oocytes treated with H₂O₂ or oocytes aged for 25 hrs in vitro. (D–F) Point charts presenting the total duration of Ca²⁺ oscillations and the mean interval between Ca²⁺ transients (D), the amplitude and the duration of the 1^st Ca²⁺ transient (E) and the rates of Ca²⁺ increase and decrease during the 1^st Ca²⁺ transient (F) in freshly ovulated oocytes and oocytes aged for 25 hrs in vitro with and without NAC. (A–F) Each dot/triangle represents one oocyte, the number of analysed oocytes is included in Tables 1 and 2. Triangles in (A) and (D) mark oocytes that did not finish Ca²⁺ oscillations during the filming.

Table 2.

Effect of oxidative stress, antioxidants, actin depolymerisation and parthenogenetic activation on the pattern of Ca²⁺ oscillations and Ca²⁺ ER store in oocytes.

	fresh + H2O2	25 hrs in vitro + NAC	fresh + CCD	fresh + EtOH
	Median (Q1; Q3)
No. of Ca2+ transients	8.0 (5.3; 10.0) n = 38	14.0aaa,bb,cc,ddd (11.0; 19.8) n = 54	9.5aaa,ddd (7.0; 11.0) n = 98	7.0e (5.0; 12.0) n = 59
Duration of Ca2+ oscillations (min)	71.4aaa,bbb,cc,eee (45.5; 113.9) n = 38	315.6aaa,bbb,ccc,ddd,e (225.0; 360.9) n = 54	196.1a,b,ddd (174.9; 217.8) n = 97	105.8a,bb,ccc,e (86.5; 126.6) n = 59
Mean interval between Ca2+ transients during 1st 2 hrs (min)	12.9aaa, c (7.2; 19.3) n = 38	18.8aa (14.4; 23.3) n = 54	18.0aaa (13.8; 22.9) n = 97	18.8aaa (10.1; 26.5) n = 59
Amplitude of the 1st Ca2+ transient (ΔF/F0)	0.5aaa,bbb (0.4; 0.7) n = 38	0.6aaa,bbb (0.5; 0.6) n = 53	0.9cc,ddd,eee (0.8; 1.0) n = 95	0.6aaa,bbb (0.4; 0.7) n = 59
Amplitude of the 3rd Ca2+ transient (ΔF/F0)	0.5aaa,bbb,d (0.3; 0.6) n = 38	0.6aaa,bbb,d (0.5; 0.6) n = 52	0.9ccc,ddd,eee (0.8; 0.9) n = 95	0.6aaa,bbb (0.5; 0.7) n = 59
Duration of the 1st Ca2+ transient (min)	4.7ccc (0.7; 7.0) n = 37	1.3aaa,bbb,ddd,eee (1.3; 1.5) n = 53	3.3bbb,ccc,ee (3.0; 3.7) n = 94	2.8ccc (2.3; 3.5) n = 59
Duration of the 3rd Ca2+ transient (min)	1.3a,cc,eee (1.2; 1.6) n = 38	1.0 (0.8; 1.2) n = 52	1.0 (0.8; 1.2) n = 38	1.0b (0.7; 1.2) n = 59
Rate of the 1st Ca2+ increase (min−1)	2.0aaa,bbb,cc (1.5; 2.4) n = 38	2.5bbb (2.1; 2.8) n = 53	2.9bb,ddd,eee (2.3; 3.4) n = 87	1.8aaa,bbb,ccc (1.3; 2.4) n = 58
Rate of the 3rd Ca2+ increase (min−1)	2.7aaa,bbb,dd (1.4; 3.5) n = 36	3.1aaa,bbb,d (2.7; 3.4) n = 52	4.7ccc,ddd,eee (4.2; 5.2) n = 91	3.3aaa,bbb (2.6; 4.0) n = 59
Rate of the 1st Ca2+ decrease (min−1)	−2.2aaa,bbb,d (−2.8; −1.6) n = 38	−2.0aaa,bbb,ddd (−2.3; −1.7) n = 53	−4.1ccc,ddd,eee (−4.5; −3.6) n = 87	−3.0aaa,b (−3.6; −2.2) n = 58
Rate of the 3rd Ca2+ decrease (min−1)	−2.8aaa,bbb,dd (−3.4; −1.7) n = 36	−2.5aaa,bbb,ddd,e (−2.8; −2.1) n = 52	−4.8ccc,ddd,eee (−5.3; −4.3) n = 91	−3.5aaa,b (−4.1; −2.8) n = 59
Amplitude of the TG-induced Ca2+ release (ΔF/F0)	n/a	1.3a,bbb,ccc,eee (1.2; 1.3) n = 42	n/a	n/a

^aaap < 0.001, ^aap < 0.01, ^ap < 0.05 vs. fresh oocytes, ^bbbp < 0.001, ^bbp < 0.01, ^bp < 0.05 vs. 9 h in vitro,

^cccp < 0.001, ^ccp < 0.01, ^cp < 0.05 vs. 25 h in vitro, ^dddp < 0.001, ^ddp < 0.01, ^dp < 0.05 vs. 9 h in vivo, ^eeep < 0.001, ^eep < 0.01, ^ep < 0.05 vs. 25 h in vivo.

We also investigated whether changes in Ca²⁺ homeostasis caused by 25-hour-long in vitro ageing could be rescued with antioxidant treatment. To this end, oocytes were cultured for 25 hrs in medium supplemented with 5 mM N-acetylcysteine (NAC)⁷⁰. The amount of ROS in such oocytes was significantly lower than in oocytes aged without NAC and similar to the ROS level in fresh oocytes (Fig. 2F,G). Interestingly, this decrease in the ROS amount prevented neither the age-dependent decrease in the ER Ca²⁺ store (Supplementary Fig. S4E, Table 2) nor the change in the pattern of Ca²⁺ oscillations. The amplitudes of Ca²⁺ transients, the duration of the 1^st Ca²⁺ spike and the rates of increases and decreases in Ca²⁺ during Ca²⁺ spikes were the same in oocytes aged with and without NAC. Moreover, Ca²⁺ oscillations lasted for even longer and were even more frequent in oocytes aged with NAC than in those aged in a pure medium (Fig. 3D–F, Tables 1, 2, Supplementary Fig. S4F–H).

Taken together, our data indicate that oxidative stress may not be responsible for the alterations in Ca²⁺ homeostasis observed in postovulatory aged oocytes. First, the increase in ROS was not observed in the majority of our ageing variants, even though the Ca²⁺ oscillations were disturbed in all of them. Secondly, although some of the changes in Ca²⁺ oscillations recorded in oocytes treated with H₂O₂ resembled those in aged ones, this resemblance may be at least partially incidental, as treatment with the antioxidant did not reverse the ageing phenotype.

Ageing affects the functionality of mitochondria

Since oxidative stress does not seem to be the main cause of defective Ca²⁺ homeostasis in postovulatory aged oocytes, we wished to investigate other potential mechanisms. The activity of mitochondria and Ca²⁺ oscillations are intertwined^71,72, so disrupted functionality of these organelles may be a reason for altered Ca²⁺ homeostasis in postovulatory aged oocytes. To investigate this possibility, we examined how postovulatory ageing affected the functionality of mitochondria. To this end, we used TMRE, a fluorescent probe of mitochondrial membrane potential⁵¹. TMRE staining revealed that 25-hour-long ageing, both in vitro and in vivo, caused severe aggregation of active mitochondria (Fig. 4A). Additionally, we also observed a decrease in the TMRE fluorescence intensity in those two experimental groups (Fig. 4B). Even though TMRE has certain limitations as a potentiometric dye⁷³, it suggests that the prolonged ageing leads to a decrease in the mitochondrial membrane potential.

Figure 4.

Functionality of mitochondria and distribution of organelles in postovulatory aged oocytes. (A) TMRE staining of active mitochondria in representative freshly ovulated oocytes and oocytes aged in vitro and in vivo for 9 and 25 hrs. The dashed white line marks oocytes’ circumferences. Scale bar 50 µm. (B) Mean intensity of TMRE staining (indicative of mitochondrial membrane potential) for the experimental variants depicted in (A) calculated for 43 freshly ovulated oocytes, 47 and 44 oocytes aged in vitro (for 9 and 25 hrs, respectively) and 58 and 53 oocytes aged in vivo (for 9 and 25 hrs, respectively). All intensity values were normalized with the mean fluorescence intensity calculated in the particular experiment for the control, freshly ovulated oocytes. ^ap < 0.05, ^bp < 0.001 vs. fresh oocytes, ^cp < 0.001 vs. 9 hrs in vitro, ^dp < 0.001 vs. 9 hrs in vivo. (C) Ca²⁺ (in orange; Rhod-2 fluorescence) and Mg²⁺ (in blue; MgGreen fluorescence) oscillations in a representative freshly ovulated oocyte. The cytoplasmic concentration of free Mg²⁺ ions is inversely proportional to the ATP concentration. Letters a-e indicate values used for further analysis of the fertilization-induced ATP production and presented in the graph (D) and Supplementary Fig. S6E,F. (D) General decrease in the Mg²⁺ concentration (value ‘e’ in (C)), indicative of general increase in the ATP production, calculated for 31 freshly ovulated oocytes, 24 and 10 oocytes aged in vitro (for 9 and 25 hrs, respectively), and 17 and 30 oocytes aged in vivo (for 9 and 25 hrs, respectively). ^ap < 0.01, ^bp < 0.05, ^cp = 0.05 vs. fresh oocytes, ^dp < 0.05 vs. 9 hrs in vitro. (B,D) Graphs present medians and the 1^st and the 3^rd quartile values. The ends of the whiskers are set at 1.5*IQR above the third quartile and 1.5*IQR below the first quartile. Dots show the minimum and maximum values if they are outside the range (outliers). (E-F) Immunofluorescence staining of calnexin, an ER marker, in representative freshly ovulated oocytes and oocytes aged in vitro and in vivo for 9 and 25 hrs (E), and in control and cytochalasin D (CCD)-treated oocytes (F). The dashed white line marks the zoomed regions. Scale bar 50 µm. (G) TMRE staining of active mitochondria in representative control and CCD-treated oocytes. The dashed white line marks the zoomed regions. Scale bar 50 µm.

Next, we wished to examine whether postovulatory ageing impaired Ca²⁺-induced ATP production, typical for fertilized oocytes. To this end we labelled oocytes with fluorescent dyes: Magnesium Green (MgGreen) for Mg²⁺ and Rhod-2 for Ca²⁺ ions (concentration of Mg²⁺ reflects inversely ATP level, i.e. it decreases when ATP concentration increases and vice versa⁷⁴; see Methods section), fertilized them and subjected to time-lapse imaging. The specificity of MgGreen dye towards Mg²⁺ vs. Ca²⁺ ions was tested by challenging the labelled oocytes with FCCP or TG. As expected, FCCP led to an abrupt increase in the MgGreen fluorescence reflecting a decrease in the ATP amount caused by uncoupling oxidation from phosphorylation in mitochondria⁷⁵, whereas TG did not alter significantly the MgGreen fluorescence (Supplementary Fig. S5A,B). We also did not observe MgGreen sequestration into the ER cisterns (Supplementary Fig. S5C).

As shown before², in fresh oocytes fertilization induced oscillatory changes in the MgGreen fluorescence reflecting readjustments of the ATP concentration during the sperm-triggered Ca²⁺ transients. Those oscillations were accompanied by a general decrease in the MgGreen fluorescence, indicative of general increase in the ATP production observed after fertilization (Fig. 4C,D). In in vivo aged (for 9 and 25 hrs) oocytes, the general decrease in the MgGreen signal was lower than in freshly ovulated oocytes. A similar decrease in the ATP production was also observed in oocytes aged in vitro for 25 hrs (Fig. 4D, Supplementary Fig. S6A–D). Additionally, aged oocytes (25 hrs in vitro and 9 and 25 hrs in vivo) displayed difficulties in readjusting ATP levels after fertilization-induced Ca²⁺ transients (Supplementary Fig. S6E,F): a decrease in the MgGreen fluorescence (i.e. increase in the ATP level) during the 1^st and the 3^rd Ca²⁺ transients was significantly lower than in freshly ovulated oocytes. Therefore, it seems that ageing may decrease the functionality of mitochondria that in consequence may affect ATP-dependent generation of Ca²⁺ transients and explain some of the alterations in the pattern of Ca²⁺ oscillations. This is, however, a two-way process, as hindering Ca²⁺ oscillations may further affect the ATP production^31,32. Interestingly, despite the fact that mitochondrial dysfunction is often linked to oxidative stress^76,77 (we noticed that it accompanies the alleviation of ROS concentration in oocytes aged in vitro for 25 hrs), we observed it also in oocytes aged in vivo, in which ROS levels were actually decreased in comparison to the fresh oocytes.

Changes in actin cytoskeleton and organelle distribution modify the pattern of Ca²⁺ oscillations

Postovulatory ageing often leads to aggregation of organelles, including mitochondria and ER cisterns^10,17,18,78. Indeed, we showed that ageing alters the structure of ER and mitochondria network, facilitating formation of the cistern aggregates (Fig. 4A,E). As ER and mitochondria localization depends on actin filaments^79,80, such redistribution may be related to a decreased functionality of actin cytoskeleton observed in aged oocytes^12,13. Therefore, we wished to examine whether dysfunctional actin cytoskeleton and, in consequence, redistribution of ER and mitochondrial network, altered Ca²⁺ signalling. To hinder actin cytoskeleton and trigger aggregation of organelles, we treated fresh oocytes with cytochalasin D (CCD), an actin-depolymerizing agent. As expected, we observed aggregates of ER cisterns and mitochondria similar to those in postovulatory aged oocytes, although never as pronounced as in oocytes aged for 25 hrs in vivo (Fig. 4F,G). In fertilized oocytes, CCD treatment did not change the characteristics of the single Ca²⁺ transients, but it altered the pattern of Ca²⁺ oscillations as a whole: increased the duration and the frequency of oscillations as well as the number of Ca²⁺ transients. Similar trends were observed in postovulatory aged oocytes, especially in 25-hour-long variants (Fig. 5A–C, Tables 1, 2, Supplementary Fig. S7A–C). Indeed, the data published previously indicate that generation of proper Ca²⁺ oscillations in response to fertilization depends on distribution of the ER cisterns^46,79,81. Interestingly, it has been also shown in somatic cells that intact cortical actin cytoskeleton is required for maintaining ER-plasma membrane junctions that participate in Ca²⁺ signalling and lipid (including phosphatidylinositol) metabolism^82,83. Moreover, it seems that actin cytoskeleton regulates also ER-mitochondria contact sites that are crucial for Ca²⁺ signalling in the cell^84–86.

Figure 5.

Impact of actin depolymerisation and parthenogenetic activation on Ca²⁺ homeostasis. (A–C) Point charts presenting the total duration of Ca²⁺ oscillations and the mean interval between Ca²⁺ transients (A), the amplitude and the duration of the 1^st Ca²⁺ transient (B) and the rates of Ca²⁺ increase and decrease during the 1^st Ca²⁺ transient (C) in freshly ovulated oocytes treated with CCD, and oocytes aged for 25 hrs in vitro and in vivo. Blue ovals indicate the region, where dots representing freshly ovulated oocytes would have been located. (D-F) Point charts presenting the total duration of Ca²⁺ oscillations and the mean interval between Ca²⁺ transients (D), the amplitude and the duration of the 1^st Ca²⁺ transient (E) and the rates of Ca²⁺ increase and decrease during the 1^st Ca²⁺ transient (F) in freshly ovulated oocytes, oocytes pre-activated parthenogenetically with EtOH and oocytes aged for 9 hrs in vivo. Each dot/triangle represents one oocyte, the number of analysed oocytes is included in Tables 1 and 2. Triangles (in (A) and (D)) mark oocytes that did not finish Ca²⁺ oscillations during the filming.

Premature activation modifies the pattern of Ca²⁺ oscillations

Last but not least, altered Ca²⁺ homeostasis in postovulatory aged oocytes may be caused by deregulation of the cell cycle. As we described before, prolonged Ca²⁺ oscillations in oocytes aged for 25 hrs are likely related to the inability of those oocytes to enter the 1^st embryonic interphase. In oocytes aged for 9 hrs in vivo, we observed another cell cycle abnormality: they tend to undergo spontaneous activation before fertilization, which was manifested as formation of the 2^nd polar bodies (Supplementary Fig. S8A). It has been reported that postovulatory ageing decreases the activity of M-phase kinases in mouse oocytes, which in turn could induce such a phenotype^{11,14,15,20,67}. To test whether modifications of Ca²⁺ oscillations typical for 9-hour-long in vivo ageing were related to this phenomenon, we analysed the pattern of Ca²⁺ oscillations in oocytes pre-treated with ethanol (EtOH), a parthenogenetic agent. EtOH induces in oocytes a single large Ca²⁺ transient⁸⁷. It activated fresh oocytes very effectively, so within approx. 1.5 hrs they reached the same stage of activation as seen in oocytes aged for 9 hrs in vivo (Supplementary Fig. S8A). Consequently, it is likely that the activity of M-phase kinases important for regulation of the Ca²⁺ signalling (see previous paragraphs) decreased in the EtOH-treated oocytes to the same level as in the aged oocytes. We showed that in the pre-activated oocytes the duration of Ca²⁺ oscillations, the frequency and the amplitude of Ca²⁺ transients, as well as the rate of Ca²⁺ increase/decrease during the transients were similar to those in oocytes aged for 9 hrs in vivo and significantly decreased as compared to the fresh oocytes. Moreover, parameters such as the duration of the 1^st and the 3^rd Ca²⁺ transients or the number of Ca²⁺ transients remained unchanged compared to fresh oocytes, the same as in oocytes aged for 9 hrs in vivo (Fig. 5D–F, Tables 1, 2, Supplementary Fig. S8B–D).

As ageing-related premature activation of oocytes does not necessarily involve an increase in the cytoplasmic Ca²⁺ concentration, we decided to examine additionally how Ca²⁺-independent parthenogenetic activation affected the pattern of Ca²⁺ oscillations. To this end, we incubated fresh oocytes with 6-DMAP, an inhibitor of protein kinases, and cycloheximide (CHX), an inhibitor of protein synthesis. This procedure activated oocytes less uniformly and efficiently than EtOH and at the time of fertilization, usually only a subset of cells formed anaphase bulges (as opposed to the 2^nd polar bodies visible in the majority of EtOH-treated oocytes and the oocytes aged for 9 hrs in vivo) (Supplementary Fig. S8A). 6-DMAP/CHX treatment seemed to have a more diverse effect on the pattern of Ca²⁺ oscillations in oocytes than EtOH pre-activation or 9-hour-long in vivo ageing: e.g. in some oocytes the duration of Ca²⁺ oscillations or the rates of Ca²⁺ rise/decline were lower than in fresh oocytes, whereas in others such decrease did not occur. Nevertheless, on average its impact on Ca²⁺ oscillations resembled in many ways that of the above-mentioned experimental conditions: it decreased the duration of Ca²⁺ oscillations and the rate of Ca²⁺ increase/decrease during the transients, and did not affect the duration of the 3^rd Ca²⁺ transient and the number of Ca²⁺ spikes. On the other hand, it did not increase the frequency nor decrease the amplitudes of Ca²⁺ transients (Supplementary Table S2, Supplementary Fig. S9).

Alterations in the pattern of Ca²⁺ oscillations caused by EtOH and 6-DMAP/CHX treatments varied slightly, possibly due to a different extent of the cell cycle progression (i.e. decrease in the activity of M-phase kinases) or to the different mechanism of activation. Ca²⁺-induced activation leads for example to degradation of IP3R1 receptor⁸⁸ that most likely is not observed in Ca²⁺-independent mode of oocyte stimulation. Taken together, these results suggest, however, that spontaneous activation may be at least partially responsible for the modification of the Ca²⁺ oscillation pattern in in vivo aged oocytes.

The pattern of Ca²⁺ oscillations and embryo fragmentation

It has been suggested that change in the pattern of Ca²⁺ oscillations caused by postovulatory ageing may lead to oocyte apoptosis and fragmentation^23,24,89. Pro-apoptotic stimuli provided by postovulatory ageing, such as the decreased amount of BCL-2, ATP or increased oxidative stress^{4,6,10,17,24,90,91}, may push aged oocytes upon fertilization to cell death. Indeed, majority of oocytes aged for 25 hrs (63.3% (19/30) in in vitro and 53.5% (23/43) in in vivo conditions) underwent fragmentation (believed to be usually an effect of apoptosis^92,93) within few hours after fertilization, while fragmentation occurred neither in freshly ovulated oocytes, nor oocytes aged for 9 hrs (Fig. 6A). Interestingly, fragmentation in aged oocytes was observed only in those activated (i.e. in oocytes that completed meiosis and entered the 1^st embryonic interphase); fertilized oocytes that had remained arrested in metaphase II did not undergo this process (Figs 1D, 6B,C). Therefore, we analysed which alterations in the pattern of Ca²⁺ oscillations correlate with this age-related susceptibility to fragmentation. In in vitro aged oocytes, fragmentation was more likely if the 1^st Ca²⁺ transient lasted for longer (p < 0.01, Fig. 6D). It seems plausible, that too long increase in cytoplasmic Ca²⁺ concentration translates to Ca²⁺ overload in mitochondria, where it can cause a persistent increase in permeability of the mitochondrial membrane, and in result, trigger release of proapoptotic factors, such as cytochrome c⁹⁴. On the other hand, in in vivo aged group, fragmentation correlated with the shorter total duration of Ca²⁺ oscillations (p < 0.001, Fig. 6E). It could be related to the fact that short Ca²⁺ oscillations reflect the depleted Ca²⁺ ER store, which in turn may lead to ER stress and activate the unfolded protein response that triggers cell death⁹⁵. A similar relationship has been observed before, but for in vitro aged oocytes^23,24.

Figure 6.

Developmental potential of postovulatory aged oocytes. (A) Frequency of fragmentation in fertilized freshly ovulated oocytes (n = 64) and oocytes aged for 9 and 25 hrs in vitro and in vivo (n = 57 and 30 and n = 51 and 43, respectively). (B) Percentage of oocytes arrested in metaphase II or activated (in interphase) among fertilized freshly ovulated oocytes and oocytes aged for 9 and 25 hrs in vitro and in vivo, and frequency, with which they undergo fragmentation. (C) Immunostaining (β-tubulin in green, DNA in red) or representative fragmented embryos derived from oocytes aged for 25 hrs in vitro and in vivo. Scale bar 50 µm. (D) The duration of the 1^st Ca²⁺ transient in normal and fragmented embryos derived from oocytes aged for 25 hrs in vitro. (E) The total duration of Ca²⁺ oscillations in normal and fragmented embryos derived from oocytes aged for 25 hrs in vivo. (D,E) Graphs present medians and the 1^st and the 3^rd quartile values. The ends of the whiskers are set at 1.5*IQR above the third quartile and 1.5*IQR below the first quartile. Dots show the minimum and maximum values if they are outside the range (outliers). ^ap < 0.01 vs. normal, non-fragmented embryos.

In summary, we show that the effect of postovulatory ageing on Ca²⁺ homeostasis in oocytes is much more complex and multifaceted than it has been reported before. Our results indicate that in order to understand it properly many various factors have to be taken into consideration, oxidative stress, often associated with age-related aberrations, being only one of them. Ageing-induced alterations in mitochondrial and cytoskeletal functionality, organelle distribution and cell cycle regulation, seem to be of special importance here. As postovulatory ageing affects oocytes of all mammalian species studied so far, including livestock and humans^{5,9,10,17,18,78}, further research on this process has a practical meaning, as it may help to optimize in vitro fertilization procedure.

Moreover, our observations lead to an exciting question regarding the mechanism, in which the oviductal environment accelerates oocyte ageing. It seems plausible that the accelerated deterioration of oocytes in in vivo conditions, preventing their development after fertilization, is evolutionarily justified. Embryonic development in mammals is very costly for a female organism and it may be beneficial to enable it only when fertilization occurs on time, ascertaining good quality of the fertilizing sperm. Indeed, it has been reported that in humans probability of successful conception is the highest when insemination takes place just before or on the day of ovulation⁹⁶.

Methods

Animals and reagents

F1 (C57Bl6/Tar × CBA/Tar) mice were maintained in the Animal Facility of the Faculty of Biology, University of Warsaw at 14:10 light/dark cycle and provided with food and water ad libitum. Animals were sacrificed by cervical dislocation. All experiments were approved by the Local Ethics Committee for Experimentation on Animals no. 1 (Warsaw, Poland) and were performed in compliance with the national regulations. All reagents were purchased from Sigma-Aldrich unless stated otherwise.

Oocyte collection

Female mice were superovulated with an intraperitoneal injection of 10 IU of pregnant mare serum gonadotrophin (PMSG, Intervet) followed 48 hrs later by 10 IU of human chorionic gonadotrophin (hCG, Intervet). Ovulated oocytes were recovered from oviducts: (i) 15 hrs after hCG (fresh and in vitro aged oocytes); (ii) 24 hrs after hCG (oocytes aged in vivo for 9 hrs); (iii) 40 hrs after hCG (oocytes aged in vivo for 25 hrs). Except for some experiments (ageing with cumulus cells), recovered oocytes were placed in hyaluronidase solution (150 IU/ml in PBS) to remove the cumulus cells. Denuded oocytes were washed in M2 medium (M16 buffered with Hepes⁹⁷) and either subjected to further experimental procedures or cultured in vitro for 9 or 25 hrs (in vitro ageing for 9 and 25 hrs) in M2 medium and 37.5 °C and 5% CO₂ in the air. In some experiments, freshly ovulated oocytes were incubated for 4 hrs with or without cytochalasin D (CCD; 2 µg/ml in M2 medium). Some fresh oocytes were also parthenogenetically activated with 8% ethanol solution (in M2 medium, for 8 min) or with 2.5 mM 6-DMAP and 10 μg/ml cycloheximide (CHX) (in M2 medium for the whole period of preparatory manipulations (isolation, staining, etc.) i.e. approx. 1.5 hrs), and others – treated for 15 min with 100 µM H₂O₂ (in M2 medium). In some experiments, oocytes were aged in vitro for 25 hrs in M2 medium supplemented with 5 mM N-acetylcysteine⁷⁰.

Sperm collection

Epididymal sperm was isolated from male mice and capacitated in 0.5 ml of fertilization medium with 5 mg/ml BSA⁹⁸ for 1.5–2 hrs in 37.5 °C and 5% CO₂ in the air.

Imaging of Ca²⁺ oscillations

Oocytes loaded with 5 μM fluorescent Ca²⁺ indicator Oregon Green 488 BAPTA-1AM (Oregon Green BAPTA; Molecular Probes, Thermo Fisher Scientific; in pure M2 medium or M2 medium supplemented with 2 µg/ml CCD or with 2.5 mM 6-DMAP and 10 μg/ml CHX, 30 min) were subjected to acidic Tyrode’s solution (pH 2.5)⁹⁹ in order to remove zonae pellucidae. Oocytes without zonae were transferred to a glass-bottom dish (MatTek Corporations) with M2 medium without BSA and allowed to stick to the glass bottom (in some experiments M2 medium without BSA but with 2 µg/ml CCD was used). Next, the dish was placed on the time-lapse imaging system (Zeiss Axiovert microscope with AxioCam HRm camera) equipped with an environmental chamber sustaining a temperature of 37.5 °C. 1-2 μl of capacitated sperm suspension was added to the oocytes. The experiment was repeated 3-5 times for each variant with consistent results (Supplementary Dataset 1).

Time-lapse imaging with single-plane images taken every 10 s was performed for approx. 7 hrs. Oocytes were illuminated with light passing through a 450–490 nm excitation filter, and the emitted light was collected with a 500–550 nm emission filter (exposure time 50 ms, 4 × 4 binning). Changes in cytoplasmic Ca²⁺ concentration were assessed by measuring the mean intensity of Oregon Green BAPTA fluorescence in time. To avoid any additional variability between the experiments caused by the different extent of dye loading, the initial (pre-fertilization) mean intensity of fluorescence was calculated for each oocyte and then used to normalize the measurements in this oocyte. The resulting values are ratios: measured fluorescence intensity (F) /initial fluorescence intensity (F₀). The rates of increase and decrease of Ca²⁺ concentration during the Ca²⁺ transients were calculated as tangents of the rising/decreasing slopes of the Ca²⁺ transients (‘a’ parameter in y = ax + b linear function fitted into these slopes; Supplementary Fig. S1B).

Ca²⁺ store measurement

Oocytes were loaded with 5 μM Oregon Green BAPTA (in M2 medium, 30 min) and transferred to a glass-bottom dish with M2 medium devoid of Ca²⁺ and Mg²⁺ ions. After 2 minutes of time-lapse imaging (conditions the same as described in ‘Imaging of Ca²⁺ oscillations’ section) thapsigargin (TG) or A23187 ionophore were added to the medium (to the final concentration of 10 μM and 1 μM, respectively). Single-plane images were taken every 10 s for approximately 30 min (the settings were the same as described for imaging of Ca²⁺ oscillations). Changes in the Oregon Green BAPTA fluorescence were analysed as described for the imaging of Ca²⁺ oscillations.

Assessment of ATP level

To assess changes in the ATP concentration during fertilization, we visualised Mg²⁺ ions, as described by Igarashi et al.². The intracellular Mg²⁺ pool is predominantly present as MgATP²⁻ and the affinity of ATP⁴⁻ for Mg²⁺ is about 10-fold greater than that of ADP at physiological pH. Therefore, Mg²⁺ increases when ATP hydrolysis exceeds the rephosphorylation of ADP to ATP and vice versa⁷⁴. To simultaneously image changes in the cytoplasmic Ca²⁺ and Mg²⁺ concentrations, oocytes were incubated for 30 min with 5 μM Rhod-2 AM (Molecular Probes, Thermo Fisher Scientific) and 5 μM Magnesium Green AM (MgGreen; Molecular Probes, Thermo Fisher Scientific), respectively. Then, their zonae pellucidae were removed with acidic Tyrode’s solution and oocytes were transferred to a glass-bottom dish with M2 medium without BSA and allowed to stick to the glass bottom. Next, they were placed on the time-lapse imaging system (Zeiss Axiovert microscope with AxioCam HRm camera) equipped with an environmental chamber sustaining a temperature of 37.5 °C, and fertilized as described in ‘Imaging of Ca²⁺ oscillations’ section. To test the specificity of MgGreen towards Mg²⁺ ions vs. Ca²⁺ ions, oocytes, labelled as described above, were subjected in separate experiments to 1 μM FCCP or 10 μM TG while imaged. Imaging was performed in normal M2 medium or M2 medium devoid of Ca²⁺ and Mg²⁺ ions, respectively.

Time-lapse imaging with single-plane images taken every 10 s was performed for approx. 7 hrs for the fertilization experiments and 30 min for the experiments testing MgGreen specificity. Oocytes were illuminated with light passing through 450–490 nm and 538–562 nm excitation filters, and the emitted light was collected with 500–550 nm and 570–640 nm emission filters, for MgGreen and Rhod-2, respectively (exposure times 20 and 50 ms, 4 × 4 binning). Changes in the cytoplasmic Ca²⁺ concentration were assessed by measuring the mean intensity of Rhod-2 fluorescence in time, whereas changes in the Mg²⁺ concentration – the mean intensity of MgGreen fluorescence. To avoid any additional variability between the experiments caused by the different extent of dye loading, the initial (pre-fertilization) mean intensity of fluorescence was calculated for each oocyte and then used to normalize the measurements in this oocyte. The resulting values were ratios: measured fluorescence intensity (F) /initial fluorescence intensity (F₀). Additionally, a drift caused by the photobleaching of MgGreen was calculated for each experiment (we used traces obtained for unfertilized oocytes) and then subtracted from the experimental measurements for fertilized oocytes.

Visualisation of reactive oxygen species

Oocytes were loaded with 5 µM CellROX Orange (Molecular Probes, Thermo Fisher Scientific; in M2 medium, 30 min) and transferred to a glass-bottom dish with M2 medium. Single equatorial plane images of the oocytes were taken on a fluorescence microscope (Zeiss Axiovert microscope with AxioCam HRm camera) equipped with an environmental chamber sustaining a temperature of 37.5 °C. Oocytes were illuminated with light passing through a 538–562 nm excitation filter, and the emitted light was collected with a 570–640 nm emission filter (exposure time 100 ms, 4 × 4 binning). ROS concentration was assessed by measuring the mean intensity of CellROX Orange fluorescence. In each experiment the mean fluorescence intensity in aged/treated oocytes (F_exp) was normalized with the mean fluorescence intensity in control, freshly ovulated oocytes dyed and imaged simultaneously (F_contr).

Visualization of mitochondria

Oocytes were loaded with 100 nM TMRE (in M2 medium) for 30 min and transferred to a glass-bottom dish with M2 medium. Single equatorial plane images of the oocytes were taken on a fluorescence microscope (Zeiss Axiovert microscope with AxioCam HRm camera) equipped with an environmental chamber sustaining a temperature of 37.5 °C. Oocytes were illuminated with light passing through a 538–562 nm excitation filter, and the emitted light was collected with 570–640 nm emission filter (exposure time 50 ms, 1 × 1 binning). As TMRE is a potential-sensitive mitochondrial dye⁵¹, we assessed the mitochondrial membrane potential in oocytes by measuring the mean intensity of TMRE fluorescence. In each experiment the mean fluorescence intensity in aged/treated oocytes (F_exp) was normalized with the mean fluorescence intensity in control, freshly ovulated oocytes dyed and imaged simultaneously (F_contr).

Immunofluorescence staining

Oocytes were fixed in 4% PFA (30 min, RT), permeabilized with 0.5% Triton-X100 (30 min, RT) and blocked with 3% BSA. Calnexin, an ER marker, was labelled with a rabbit polyclonal antibody (1:200, Abcam) followed by an Alexa Fluor 633-conjugated goat anti-rabbit IgG (1:200; Invitrogen, Thermo Fisher Scientific) and β-tubulin – with a mouse monoclonal antibody conjugated with FITC (1:50). Embryos were incubated in the primary antibodies overnight at 4 °C, washed in PBS and 3% BSA and then, if required, incubated with the secondary antibody for 2 hrs in RT. DNA was stained with propidium iodide (0.01 mg/ml in PBS; 30 min, RT or overnight, 4 °C). Oocytes were analysed on an inverted confocal microscope (Zeiss and Olympus).

Western blotting

Expression of IP3R1 and SERCA proteins was examined in samples of 50–80 oocytes, depending on the experiment. Cell lysates were mixed with 4× NuPage LDS sample Buffer and 10× NuPage Sample Reducing Agent (Invitrogen, Thermo Fisher Scientific) and heated for 10 min in 70 °C. The samples were subjected to NuPage Novex 3–8% Tris-Acetate gels (Invitrogen, Thermo Fisher Scientific) and separated proteins were transferred onto PVDF membranes (Hyperbond-P, Amersham Biosciences). The blots were then stained with Ponceau S to confirm the equal sample load, blocked in 5% non-fat milk in TTBS and probed for 1 h with a rabbit polyclonal antibody (Rbt03) raised against a 15 amino acid peptide sequence of the C-terminal end of the IP3R1¹⁰⁰ and with a goat anti-SERCA2 antibody, diluted 1:350 and 1:200, respectively, in 5% non-fat milk in TTBS. A donkey anti-goat antibody and a goat anti-rabbit antibody (Bio-Rad), respectively, conjugated with horseradish peroxidase diluted 1:5000 were used as secondary antibodies in 1-hour-long incubation. Detection was performed by the enhanced chemiluminescence technique using SuperSignal West Dura Extended Duration Substrate reagents (Pierce, Thermo Fisher Scientific) according to the manufacturer’s instruction. Western blot analysis was repeated 3 times for IP3R1 and 2 times for SERCA2.

Real-time RT-PCR

Oocytes were transferred in groups of 30 into 20 μl of Lysis/Binding Buffer (Dynabeads mRNA DIRECT Micro Kit, Thermo Fisher Scientific) and stored in −80 °C until further analysis. mRNA was isolated from the samples using the Dynabeads mRNA DIRECT Micro Kit (Thermo Fisher Scientific) according to the manufacturer’s protocol. In short, thawed samples were rotated with 10 μl of paramagnetic oligo-(dT)₂₅ bead suspension for 30 min at RT. mRNA was eluted from the beads by adding 10 μl of DEPC-treated water and heated for 10 min at 70 °C with 0.5 μg oligo-(dT)₂₅. The reverse transcription was performed in a total volume of 20 μl using 200 U of Superscript II Reverse Transcriptase, 0.5 mM dNTPs and 40 IU RNase inhibitor (Invitrogen, Thermo Fisher Scientific) at 42 °C for 50 min. Synthesized cDNA was diluted twice with nuclease-free water (Thermo Fisher Scientific) and subjected to preamplification procedure (T100 thermocycler, Bio-Rad; 10 cycles: 95 °C/15 s, 60 °C/4 min) with TaqMan PreAmp Mix (Thermo Fisher) and pooled TaqMan Gene Expression Assay probes (Itpr1/IP3R1: cat. no. Mm00439907_m1; Atp2a2/SERCA2: Mm01275320_m1; Actb/Actin B: Mm01205647_g1; Applied Biosystems, Thermo Fisher Scientific). The preamplification product was diluted twice and subjected to a real-time PCR using TaqMan Gene Expression MasterMix in StepOne Real-Time PCR System thermocycler (Applied Biosystems, Thermo Fisher Scientific; 50 °C/2 min; 60 °C/10 min; 50 cycles: 95 °C/15 sec, 60 °C/1 min). The relative level of expression was calculated using 2^−ΔCt method¹⁰¹ with actin B expression used for normalization. PCR analysis was repeated 3–9 times depending on the experimental variant.

Statistical analysis

Kruskal-Wallis test with the post-hoc analysis and Mann–Whitney U test were applied to analyse the data. The differences between groups were considered statistically significant for p < 0.05.

Author Contributions

M.S., A.W., D.S.P. and P.S. conducted experiments and data analysis. T.I. conducted experiments. R.M. conducted statistical analysis. K.S. consulted PCR experiments. A.A. secured funding, designed and conducted experiments, analysed data and wrote the manuscript.

Competing Interests

The authors declare no competing interests.

Supplementary information

Supplementary information accompanies this paper at 10.1038/s41598-019-48281-3.