Separation and Loss of Centrioles From Primordidal Germ Cells To Mature Oocytes In The Mouse

Simerly, Calvin; Manil-Ségalen, Marion; Castro, Carlos; Hartnett, Carrie; Kong, Dong; Verlhac, Marie-Hélène; Loncarek, Jadranka; Schatten, Gerald

doi:10.1038/s41598-018-31222-x

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Published: 24 August 2018

Separation and Loss of Centrioles From Primordidal Germ Cells To Mature Oocytes In The Mouse

Scientific Reports volume 8, Article number: 12791 (2018) Cite this article

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Abstract

Oocytes, including from mammals, lack centrioles, but neither the mechanism by which mature eggs lose their centrioles nor the exact stage at which centrioles are destroyed during oogenesis is known. To answer questions raised by centriole disappearance during oogenesis, using a transgenic mouse expressing GFP-centrin-2 (GFP CETN2), we traced their presence from e11.5 primordial germ cells (PGCs) through oogenesis and their ultimate dissolution in mature oocytes. We show tightly coupled CETN2 doublets in PGCs, oogonia, and pre-pubertal oocytes. Beginning with follicular recruitment of incompetent germinal vesicle (GV) oocytes, through full oocyte maturation, the CETN2 doublets separate within the pericentriolar material (PCM) and a rise in single CETN2 pairs is identified, mostly at meiotic metaphase-I and -II spindle poles. Partial CETN2 foci dissolution occurs even as other centriole markers, like Cep135, a protein necessary for centriole duplication, are maintained at the PCM. Furthermore, live imaging demonstrates that the link between the two centrioles breaks as meiosis resumes and that centriole association with the PCM is progressively lost. Microtubule inhibition shows that centriole dissolution is uncoupled from microtubule dynamics. Thus, centriole doublets, present in early G2-arrested meiotic prophase oocytes, begin partial reduction during follicular recruitment and meiotic resumption, later than previously thought.

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Introduction

Centrioles, found at the poles of mitotic spindles, are vital for reproduction and development. Long thought to be contributed by the sperm during fertilization and lost during fetal oogenesis, they are essential in innumerable processes¹. Indeed, centriole defects appear as the root causes of a broad set of diseases, ranging from blindness and cancers through microcephaly and ciliopathies^2,3. Centrioles are often surrounded by the pericentriolar material (PCM), and together, the two structures define the canonical centrosome, the cell’s major microtubule organizing center (MTOC)³.

In most mammals, haploid female gametes produced during oogenesis lose their centrosomes, although the mechanism of when and how remains elusive^4,5,6. Most studies on centrosome reduction in gametes involve ultrastructural observations^4,7,8. In humans, centrioles have been detected in fetal oogonia at 13–15 weeks post-gestation and within early growing oocytes⁹. However, centrioles have not been found in fully grown germinal vesicle (GV)-stage oocytes, and the metaphase-I and -II spindles formed after meiotic resumption are anastral, barrel-shaped structures with spindle poles devoid of centrioles or PCM⁸. In mice, ultrastructural and marker tracing have identified intact centriole pairs in fetal oogonia and early post-natal stage (P4) mouse primordial oocytes^10,11,12. In later, preovulatory stages, growing mouse oocytes apparently lose centrioles¹³ while maintaining dispersed acentriolar PCM throughout the cytoplasm.

As the oocyte reaches maturity and competency to enter meiosis, a perinuclear MTOC, composed of PCM constituents such as γ-tubulin and pericentrin, gradually enlarges near the GV nucleus^14,15,16. Upon meiotic resumption, the acentriolar PCM fragments along the GV nucleus, mediated by PLK1, which releases the centriole adhesion protein cNAP1 (centrosomal Nek2-associated protein-1)^17,18 and then is stretched and fragmented by BicD2-anchored dynein in a microtubule-dependent manner¹⁸. Finally, KIF11 mediates further MTOC fragmentation to allow segregation of PCM material to opposing spindle poles¹⁸. The kinases Aurora A and PLK4 also enhance microtubule growth and first meiotic spindle assembly as chromosomal divisions ensue¹⁹. The arrested mouse metaphase-II spindle is anastral and acentriolar but maintains assembled PCM material at the spindle poles and within distinct cytoplasmic foci^1,20,21,22. Interestingly, the mouse sperm does not contribute a centriole at fertilization^23,24,25, and zygotes rely on convergent cytoplasmic PCM and kinesin-5 to progress through mitotic divisions during early development until the blastocyst stage, when centrioles reappear at the spindle poles^26,27,28,29.

The most prominent permanent core components found, nearly universally, in the centriole and within the centrosomes are centrin, pericentrin, and γ-tubulin. Centrin is an EF-hand calcium-binding protein found in the lumen of assembled centrioles³⁰. Centrins are required for basal body formation and positioning of the spindle pole body in yeast, algae, and ciliates^31,32. Mammals express four centrin genes (CETN1-4), but their cellular functions are not known^33,34. γ-tubulin is the tubulin isoform responsible for serving as the MTOC³⁵ and is a component of the γ-tubulin ring complex (γ-TuRC)³⁶. Pericentrin is a conserved coiled-coil PCM scaffolding protein that complexes with γ-tubulin and other proteins to initiate microtubule nucleating activity and cell cycle regulation^37,38.

Centrioles have been reliably traced dynamically with transgenic reporter green fluorescent protein (GFP)-labeled centrin, including GFP-centrin-2 (GFP CETN2)^{39,40,41,42,43,44}. A stable transgenic mouse strain that constitutively expresses GFP CETN2 in every cell of the body, including gametes, has been generated and shown to be a reliable probe for tracing centriole behavior in a variety of organs in the mouse^25,45. These transgenic mice, expressing GFP CETN2, appear nearly normal physiologically when compared to wild-type mice^29,45.

Here, using this transgenic mouse constitutively expressing GFP CETN2 to trace centrioles and γ-tubulin to track centrosomes, along with microtubule and DNA probes, we found centriole pairs in somatic cells (e.g., cumulus and stromal cells) as well as in oogonia from fetal ovaries persisting in immature oocytes in the adult ovaries. As maturing oocytes transition into full competency to resume meiotic maturation and chromosome-reductional divisions, these pairs begin to disassemble, although they maintain the ability to nucleate microtubules throughout meiosis. Surprisingly, GFP-centrin-tagged structures are visible at meiotic spindle pole PCM at metaphase-I and -II spindles. Yet, following both GFP-centrin and PCM in living oocytes shows that their association is weak and that, in oocytes, the CETN2 sites do not retain the capacity to organize the PCM as meiosis I progresses as they do in mitotic cells. These foci comprise the largest, best organized PCM in the cytoplasm, and drug recovery from nocodazole microtubule disassembly shows prominent microtubule nucleation from these GFP CETN2-containing PCM foci after drug reversal. Collectively, this study suggests that centriole remnants persist in fully grown oocytes and that the dogma regarding the destruction of the maternal centrioles in early oogenesis may need to be reconsidered.

Results

Primordial germ cells and oogonia in fetal ovaries contain GFP CETN2-labeled centrioles

Since centriole elimination is reported to occur in fetal stages⁴, we first investigated primordial germ cells (PGCs) in the nascent gonads in GFP CETN2-expressing CB6F1 females, with sex identified by y chromosome PCR (see Methods and Materials), and traced their history through fetal oogonia formation in developing ovaries prior to birth. Isolated PGCs from e11.5 post-coitus gonadal tissues were identified from undifferentiated or somatic cells by the expression of the germ line cell surface marker Stage-Specific Embryonic Antigen-1 (SSEA-1; Fig. 1)⁴⁶. Initial findings on this transgenic mouse reported GFP CETN2 expression from e14.5 forward (Jackson Laboratory Mouse Database Description at https://www.jax.org/jax-mice-and-services#)⁴⁵, so we were initially surprised to find that all e11.5 PGCs also contained GFP CETN2-labeled centrioles within microtubule arrays, whether mitotic or interphase (Fig. 1A–D). When PGCs stopped mitotic divisions, and entered pre-meiotic G2 arrest around e12.5 post-coitus, twin pairs of GFP CETN2 foci, in association with microtubules, were identified in SSEA-1-positive e14.5 and e18.5 oogonia, as well as in surrounding somatic cells (Fig. 1E–H). We conclude that GFP CETN2 centrin detects centriole pairs throughout fetal development, including their presence in female primordial germ cells.

Diminished GFP CETN2 punctate foci, perhaps centriole remnants, are typically observed in metaphase-I and -II spindle poles

We traced the fate of the GFP CETN2-labeled structures during meiotic maturation. At meiotic resumption, separated GFP CETN2-labeled pairs embedded in γ-tubulin ribbons were observed at the assembling spindle poles (Fig. 2A–G). Cumulus cells, serving as an internal somatic cell control, also showed GFP CETN2-labeled centriole pairs within γ-tubulin PCM, including at the spindle poles of a rare bipolar mitotic cell (Fig. 2A,C: arrowheads). Using correlative light and electron microscopy (CLEM), we investigated whether classical 9-triplet microtubule organization could be identified in oocytes at this stage (Suppl Fig. S1). Analysis revealed linear MTOC-like structures with associated GFP CETN2 foci (Suppl Fig. S1B, inset) in live oocytes. EM sectioning through the imaged site, however, showed only linear MTOC structures and multi-vesicular PCM aggregates (13; Suppl Fig. S1B–F), but no classical centrioles with microtubule walls, despite careful examination of hundreds of 80-nm-thick sequential serial sections. Typical canonical centrioles were identified in adhering cumulus cells attached to the same oocyte (Fig. 2E, inset; Suppl Fig. S1G–I). Collectively, the data suggest that significant oocyte centriole dissolution begins with meiosis onset. Altered oocyte centrioles still display GFP CETN2 labeling but are either too fragile to survive EM processing protocols or have undergone partial disassembly. This observation is supported by finding classical centrioles with 9 + 0 triplet microtubules in the surrounding somatic cumulus cells.

Support for oocyte centriole alterations occurring during meiotic resumption was also derived from measuring changes in PCM and GFP CETN2 foci area sizes (in μm²) during maturation (Suppl Fig. S2D,E). After germinal vesicle breakdown (GVBD) and through complete maturation to metaphase-II arrest, we measured increased γ-tubulin area as the GV-associated MTOC foci were stretched into expanded ribbons and fragmented, as previously reported^16,18 (Suppl Fig. S2D). However, significantly decreased GFP CETN2 areas were measured after GVBD (Suppl Fig. S2E), suggesting structural changes in oocyte centrioles with diminishing GFP CETN2 detection. Interestingly, areas for GFP CETN2 centrioles in somatic cumulus cells were consistent in primordial follicles, growing antral follicles, and surrounding mature GVs after follicular release (0.47 ± 0.17, 0.42 ± 0.23, and 0.43 ± 0.19 µm², respectively). Despite reduced areas in GFP CETN2 foci after meiotic resumption, most spindle poles in metaphase-I and -II oocytes still contained GFP CETN2 foci at the ends of γ-tubulin short segments or foci on the outer edges of the barrel-shaped spindles (Fig. 2H–W, Suppl Fig. S2F).

Tracking GFP CETN2 foci in first and second meiotic spindles showed many localization patterns. A minority of oocytes at metaphase-I or -II were found with GFP CETN2 foci at both poles (Fig. 2H–O), even after first polar body extrusion (Fig. 2P–W; panels enhanced in Suppl Fig. S8). Most metaphase-I spindles, however, contained single GFP CETN2 foci at the extreme edges of the barrel-shaped spindle (Suppl Fig. S3A–G), unlike centrosomes in somatic cell spindles that reside more centrally in the fusiform spindle pole. Diminished GFP CETN2 foci association with the γ-tubulin PCM was also observed at this stage, sometimes with foci residing within the spindle lattice near the aligned bivalents (Suppl Fig. S3H–V). In metaphase-II-arrested oocytes, a minority of GFP CETN2 foci were observed in the extruded first polar body (Pb1) or in the microtubule-based asters (cytasters)^20,21,22 in the cytoplasm (Suppl Fig. S4). However, the majority of GFP CETN2 foci were found in association with meiotic spindles throughout maturation to metaphase-II arrest (Suppl Fig. S2F). Taken together, our analyses suggest that centrioles show significant reduction during meiotic progression, looser association with the PCM, and could not be identified as traditional centrioles with 9-fold symmetry in EM serial sections, consistent with centriole dissolution prior to the onset of meiosis. However, GFP CETN2 foci identified in fetal stages survive in mature oocytes and are found in γ-tubulin-containing meiotic spindle poles through metaphase-II arrest following meiotic resumption. GFP CETN2 foci are found in atypical spindle pole positions relative to traditional localization of centrioles in somatic cells.

Our analysis of the fate of GFP CETN2 centriole doublets, traced from e11.5 PGCs isolated from the genital ridge through mature metaphase-II-arrested oocytes is summarized in Fig. 3 and Supplemental Fig. S5. GFP CETN2 foci associated with γ-tubulin or pericentrin PCM in PGCs, fetal oogonia, and early oocytes from P4 neonate ovaries showed double pairs of GFP CETN2 centrioles (Fig. 3a–c, graph; Fig. 3A–C, lower panel) closely apposed in the cytoplasm (Suppl Fig. S5a–c). Incompetent GVs (<66 µm diameter) from adult ovaries, as well as mature GV’s collected after hormonal simulation (>75 µm diameter), showed the average distance between double pairs of GFP CETN2 foci dramatically increased (Fig. 3D,E, lower panel; Suppl Fig. S5d,e; *p < 0.01). Interestingly, around 3–10% of incompetent or competent GV oocytes showed more than two doublet pairs (Fig. 3d–g, graph, green bars). No evidence of centriole duplication in these quiescent oocytes was found, suggesting these aberrant oocytes were produced by fetal-stage mitotic errors or, perhaps, by early oocyte cyst breakdown inconsistencies. Often, oocytes with abnormal doublets showed widely dispersed GFP CETN2 foci in the cytoplasm or cortex or along the GV surface, with distances >12 µm generally observed. Regardless, centriole dissolution appears to accelerate following meiotic resumption, with an increase in the number of single GFP CETN2 pairs at the spindle poles of after GVBD (Fig. 3f–h, graph; Fig. 3F–H, lower panel). Collectively, the data show that GFP CETN2 centrioles embedded within PCM are maintained as two doublets until follicular recruitment in post-adolescent females, when GV-arrested oocytes within growing follicles first separate into GFP CETN2 pairs and, later, single-pair dissolution occurs. Following meiotic resumption, single-centriole-pair dissolution accelerates. No evidence of centriole duplication was found during meiotic maturation. Of the metaphase-II-arrested oocytes with detectable GFP CETN2 foci at the spindle poles (~38%; Suppl Fig. S2A), nearly all showed diminished single pairs at their spindle poles.

Immature, growing, and mature oocytes contain GFP CETN2 foci in γ-tubulin PCM

Fixed neonatal Day-4 post-birth female ovaries (P4), a stage at which centrioles have been identified by EM¹⁰, showed two tightly apposed doublets of GFP CETN2 foci within γ-tubulin PCM, adjacent to the GV (Fig. 4A). Pre-antral and early antral follicles in adult ovaries maintained GFP CETN2 foci, with some displaying doublet pair separation, beginning with antral formation (Fig. 4B,C). Some cytoplasmic GFP CETN2 aggregates appeared in early follicular stages and could be distinguished from GFP CETN2 foci in centrosomes by their exclusion from co-localization with γ-tubulin.

Tracking GFP CETN2 doublet fate in ovarian sections is challenging and provides the rationale for isolating immature oocytes of assorted sizes during follicular growth (Suppl Fig. S6). Incompetent GV oocytes, which could not initiate meiotic maturation, showed GFP CETN2 foci embedded within γ-tubulin PCM in all oocytes, mostly without microtubule aster assembly (Fig. 5A). Only 70% of fully competent mature GV oocytes maintained GFP CETN2 foci with γ-tubulin (Suppl Fig. S2A). Although numerous GFP protein aggregates in growing GV oocytes were present, these supernumerary foci did not interact with γ-tubulin or pericentrin, nor assemble microtubules. The GFP CETN2 foci in association with γ-tubulin or pericentrin maintained consistent area sizes during oocyte growth, although their distribution within the oocyte changed from the GV nuclear surface in small follicle GVs to the cytoplasm during mid-follicular growth, before returning to the GV surface just before meiotic resumption (Fig. 5A7–A8; enhanced image in Suppl Fig. S8; graphic analysis, Suppl Fig. S2B,C).

Mature GVs, preparing to enter first meiosis, had decondensed, stretching PCM along the GV surface, vastly increasing the surface area (Fig. 5B and graph)^16,18. Oocytes with two GFP CETN2 pairs typically separated to opposing ends of the enlarging, fragmenting MTOC (Fig. 5B3; see enhance image Suppl Fig. S8). Other nuclear and cytoplasmic PCM foci lacking GFP CETN2 foci did not immediately undergo expansion along the GV surface (Fig. 5B3, short arrows; Suppl Fig. S8N–R). However, PCM stretching along the GV surface does not require GFP CETN2 foci (Fig. 5C). Collectively, GFP CETN2 foci, present from early pre-antral follicle oocytes, are maintained through follicular growth to full maturity. These structures remain within the γ-tubulin or pericentrin PCM as it undergoes decondensation, stretching, and fragmentation along the GV surface in preparation for meiotic spindle assembly.

Since GFP CETN2 doublets appear to separate during GV growth, we investigated whether the expression of Cep135, a centrosomal scaffolding protein that anchors cNAP1 in somatic cell centrioles and whose disruption or overexpression causes premature centrosome separation⁴⁷, might be the underlying mechanism (Fig. 6). Adhering cumulus cells, as well as incompetent GVs isolated from small, medium, and large growing follicles, showed bright GFP CETN2 foci co-localized with pericentrin and Cep135 (Fig. 6A–F). As oocytes transitioned toward meiotic resumption, GV oocytes expanded and the MTOC fragmented along the GV surface as Cep135 detection was significantly reduced in the PCM and the GFP CETN2 doublet pairs separated (Fig. 6G–J). cNAP1, anchored by Cep135 at somatic cell centrioles, has been identified in mouse oocyte MTOCs⁴⁸, and PLK1 has been implicated in release of cNAP1 to permit MTOC expansion in the mouse GV oocyte¹⁸. Therefore, we utilized the PLK1 inhibitor BI 2536 to investigate whether the inhibitor impacts Cep135 localization at MTOCs and subsequent GFP CETN2 doublet separation. A 2-hr treatment of mature GV oocytes with BI 2536 inhibitor blocked Cep135 reduction from MTOCs at the GV surface, which still slightly expanded (Fig. 6K). Remarkably, despite Cep135 retention in the MTOC, the GFP CETN2 doublet pairs separated to opposite ends of the PCM (Fig. 6K–M). Even the combined treatment of BI 2536 with the microtubule inhibitor nocodazole, which blocks MTOC expansion and fragmentation, showed slightly elongated MTOCs on the GV surface, with retained Cep135 in the PCM and GFP CETN2 foci separated into four pairs (Fig. 6N–P). Collectively, these results suggest that GFP CETN2 doublet separation does not require the loss of Cep135 centrosomal core protein at the MTOC. Centriole doublet separation and dissolution may involve other unknown mechanisms occurring upstream of meiotic resumption.

GFP CETN2 foci present a lose association with the PCM

To verify that the GFP CETN2 foci observed in fully grown mouse oocytes were not an artifact due to fixation, we also followed them in living oocytes. These investigations, which were performed in France on similar and identical mouse strains, obtained independently, provide confirmation of the reliability of both the research resources and the experimental strategies. For these experiments, oocytes expressing mCherry-Plk4 (either from transgenic mice or from cRNA injection)⁴⁹, a PCM marker¹⁹, together with GFP or Venus-tagged CETN2 (also from either transgenic mice or cRNA injection) were utilized. No differences in behavior or localization were observed between cRNA and transgenic expression of these constructs (Fig. 7, compare A and D). Live imaging also revealed the presence of GFP CETN2 doublets in follicular cells surrounding the oocyte coming from transgenic GFP CETN2 mice (Fig. 7A upper panels) as well as GFP CETN2 foci co-localizing with the cortical PCM in incompetent oocytes (Fig. 7A lower panel). The distance between the two doublets increased while the major PCM foci moved toward the nuclear envelope (Fig. 7B), arguing that anchoring of the major MTOC to the nuclear envelope favored the building of forces able to perturb its internal architecture, as observed by Luksza et al.¹⁶. This distance increased even further in competent oocytes able to resume meiosis (Fig. 7C). This is fully consistent with observations made in fixed oocytes (Suppl Fig. S5). Importantly, as meiosis I progressed, live imaging showed that the association between GFP CETN2 foci and the PCM, labeled with mCherry-Plk4, was weak (Fig. 7D and Suppl Movie S1) at all stages of meiosis I spindle assembly (Suppl Movies S2 and S3). Foci of GFP CETN2 could be extruded into the first polar body (Fig. 7D and Suppl Movie S1) as also observed in fixed oocytes (Fig. S2F). Collectively, live imaging demonstrated GFP CETN2 foci in association with organize PCM, although co-localization appears to be progressively lost as the oocytes resumed meiosis.

GFP CETN2 foci that function in meiotic spindle reassembly after microtubule inhibition are present in PCM

Since meiotic spindle assembly is initiated by multiple MTOCs, we explored whether GFP CETN2 foci within fragmented MTOCs would reassemble microtubules following disruption with the microtubule inhibitors nocodazole and paclitaxel (Fig. 8). In prometaphase-I oocytes, 3.5-hr nocodazole exposure increased MTOC volumes near the condensed bivalents with embedded GFP CETN2 foci but no microtubule assembly (Fig. 8A–F). Upon a 5-min rescue from nocodazole, the MTOC with GFP CETN2 foci expanded and assembled a large, well-arrayed microtubule aster, suggesting that GFP CETN2-containing centrosomes can function as MTOCs that could potentially participate in spindle assembly (Fig. 8G–L).

Paclitaxel enhances microtubule assembly. After GVBD, paclitaxel initiated microtubule aster assembly around the GV nucleus from fragmented, dispersed MTOCs. The largest MTOC retained two embedded GFP CETN2 foci from which the best organized microtubule aster assembled next to the condensing bivalents (Fig. 8M–R). All other identified cytoplasmic and cortical microtubule asters assembled from MTOCs but without detectable GFP CETN2 foci (Fig. 8R). In later prometaphase-I, GFP CETN2 foci within many small fragments of γ-tubulin PCM were observed around the circular bivalents and extensive bundles of microtubules assembled outward toward the cortex in a large ring pattern (Fig. 8S–X). Thus, paclitaxel-induced microtubule enhancement prevents PCM assembly at opposing spindle poles. GFP-expressing CETN2 foci remain with the scattered MTOCs around the condensing bivalents, but these centrosomes neither nucleate microtubules nor assemble bipolar spindles.

Analysis of changes to GFP CETN2 foci areas during exposure to nocodazole and paclitaxel is presented in Supplemental Fig. S7. GV oocytes exposed continuously to nocodazole or paclitaxel for 3.5 hrs in the presences of dbcAMP to retard meiotic resumption showed significant decreases in overall GFP CETN2 foci areas, perhaps indicating GFP CETN2 foci instability in the face of altered microtubule dynamics (Suppl Fig. S7A, green bars). Similar observations were seen when dbcAMP was removed to permit GV oocytes to progress to prometaphase-I in the presence of nocodazole but not after paclitaxel exposure (Suppl Fig. S7A, blue bars). Regardless, GFP CETN2 foci areas decreased following resumption of meiotic maturation in all microtubule inhibitor treatments when analyzed with their corresponding GV controls (Suppl Fig. S7B). Collectively, disruption of microtubule assembly or disassembly reduces GFP CETN2 areas, even in arrested GV oocytes, suggesting that structural alterations have already occurred prior to oocyte full maturation that render the oocyte centrioles susceptible to changes in microtubule dynamics before meiotic resumption.

Discussion

The transgenerational contribution of centrosomes and centrioles was thought to have been solved by Boveri^50,51 who wrote that: “The ripe egg possesses all of the elements necessary for development save an active division-center (centriole). The sperm, on the other hand, possesses such a center but lacks the protoplasmic substratum in which to operate”. Later, the pioneering ultrastructural imaging of centrioles by Fawcett and Phillips⁵², previously depicted as mere dots, recognized that centrioles and basal bodies were composed of 9-triplet microtubules. However, predictions that molecular genetic approaches would answer definitively inheritance questions, following on the discovery of DNA in mitochondria^53,54, failed. Without a permanent tracer molecule, as DNA in mitochondria, it has not been possible to answer this question conclusively, until recently. Now, breakthroughs in understanding the nature and dynamism of the core protein components of both centrosomes and centrioles⁵⁵, and the realization that their assembly is tightly regulated and orchestrated provide the tools to address inheritance. With considerable simplification, centrioles assemble upon a SAS-6 cartwheel⁵⁶, upon which the other components of the 9-triplet structures assemble and onto which centrosomal PCM later concentrates³⁶. In consequence, the question as to whether centrosomes and centrioles are inherited, strictly speaking, may need to be examined more stringently. Findings demonstrating that they are elaborate, dynamic protein assemblies support the concept of “transmission”, as thoughtfully proposed by Ross and Normark⁵⁷. Without the DNA of chloroplasts, mitochondria and other plastids, perhaps the term “inheritance” should not be applied casually to organelles, assembled at times de novo, which lack their own genomic material.

Centriole disassembly, studied here in oocytes during the last two meiotic divisions, remains challenging, though elimination mechanisms have been discovered in flies⁵⁸ and starfish⁵⁹. In mice, two centriole pairs persist from fetal stages¹¹ and in oocytes within adolescent ovaries¹⁰. During maturation, the centriole pairs first separate within the surrounding PCM, with the disassembly of one of the two doublets prior to full oocyte growth. Centriole adhesion does not appear to depend on the Cep135 centriolar core protein that anchors cNAP1 at the centrioles. Centriole dissolution accelerates upon meiotic resumption, as the number and area significantly decrease following GVBD (Fig. 3; Suppl Fig. S2E). Perhaps these centriole-like remnants anchor the stretching PCM to the GV surface^16,18. Later, the metaphase-I and -II spindles typically display at least a single pair near or at a spindle pole. Overall, this suggests that the maternal centriole is not lost until the completion of oocyte maturation, as in starfish oocytes^59,60. Centrioles were found in first or second mouse meiotic spindles neither by Szollosi et al.⁷, nor in our CLEM investigations here. Locating centrioles within an oocyte is akin to looking for a needle in a haystack, what with the enormous cytoplasmic volume and the lack of any fiduciary marks to help locate the centriole precisely. Here however, the CLEM approach defined the site of the GFP CETN2 pairs, and still, only osmiophilic PCM was observed (Suppl Fig. S1). This suggests that the canonical 9-triplet centriole structure disassembles prior to meiotic maturation, while the adjacent cumulus cells with intact centrioles provide ideal controls (Fig. 2E, inset and Suppl Fig. S1).

The GFP CETN2 doublets identified here likely fall into the category of centrioles described as “advertisements,” “remnants”, “passenger” or “zombie” structures^61,62,63,64; i.e., vestigial, non-functional, and non-replicating centrioles. Perhaps centriole alterations mirror events seen in sperm distal centriole dissolution, where proximal centriole-like structures identified in the sperm of flies and humans lose their 3-dimensional 9-triplet microtubule structure, but retain PCM antigens and the ability to organize into functional MTOCs under the right conditions, such as after fertilization in the activated zygotic cytoplasm^64,65. Indeed, the destruction of first one, and then the second, centriole in mouse sperm as these transit through the epididymis²⁵ underscores the complex behavior of centrosomes and centrioles in gametes even after they depart from the gonad. While these GFP CETN2 doublets in maturing oocytes appear non-functional, their microtubule nucleating activity can be revived after recovery from nocodazole microtubule disassembly (Fig. 8). Upon this revival, robust microtubule arrays reassemble around the now reactivated GFP CETN2 doublets, which themselves, are surrounded by PCM foci.

The precise mechanism of CETN2 doublet separation is not currently known. Cep135, important in centriole biogenesis, is present in the MTOCs of incompetent oocytes and is lost when the PCM begins to expand and fragment prior to resumption of meiosis (Fig. 6). In somatic cells, Cep135 disruption leads to centrosome splitting and abnormal mitotic spindle phenotypes through its interaction with cNAP1, a centrosomal linker protein implicated in centriole-centriole cohesion^17,66,67. cNAP1 is also in mouse GV MTOCs, although disruption of cNAP1 by microinjection of specific double-stranded RNAs did not affect first or second meiotic spindle assembly⁴⁸. Recently, the initiation of MTOC stretching on the GV surface before nuclear envelope breakdown¹⁶, critical for PCM fragmentation and first meiotic spindle assembly, was shown to be dependent on PLK1 phosphorylation of cNAP1, although the presence and functional role of cNAP1 in these MTOCs was unclear since they were not thought to have centrioles¹⁸. Our results suggest that maturing oocytes split the two CETN2 doublets early in follicular growth before attaining full maturation (Fig. 3), with Cep135 and cNAP1 continuing to be expressed until just before meiotic resumption. Thus, perhaps other proteins play a key role in mouse centriole-centriole cohesion upstream of Cep135 and cNAP1. Investigations on the precise molecules essential for centriole persistence and PCM functionality in the doublets and their fates during oocyte growth and maturation to metaphase-II arrest await future studies.

Centriole destruction may be prerequisite for terminal differentiation, and the retention of centrioles is associated with proliferation, regeneration, pluripotency, perhaps even totipotency. Skeletal muscle differentiation results in centriole loss^68,69 as during neuron differentiation⁷⁰. Cardiomyocyte differentiation is particularly instructive, since centrioles are lost during terminal differentiation in murine hearts, which do not regenerate^71,72. Remarkably, centrioles are retained in the regenerating hearts of fish and newts⁷³. Here, centrioles are lost upon the onset of meiotic maturation, with immature oocytes retaining their double paired centrioles (Fig. 3, graph). Perhaps the last phase of oogenesis (i.e., oocyte maturation) is another example of terminal differentiation, including with the elimination of centrioles and centrosomes. Interestingly, while it appears necessary to undergo two mitotic cycles to generate centrioles, perhaps during both spermatogenesis and oogenesis, the last two meiotic cycles also are prerequisite for centriole destruction.

Methods and Materials

Mouse husbandry, handling, and Institutional oversight

All research protocols complied with the National Institute of Health’s Office of Laboratory Animal Welfare Guide for the Care and Use of Laboratory Animals regulations and by the DGRI in France (Direction Generale de la Recherche et de l′Innovation: Agrément OGM; DUO-1783) and approved by the University of Pittsburgh’s and Magee-Womens Research Institute’s Institutional Animal Care and Use Committees (IACUCs; protocol #16027448). CB6-Tg (CAG-EGFP/CETN2)3-4Jgg/J mice (Stock number: 008234)⁴⁵ were obtained from the Jackson Laboratory (Bar Harbor, ME) as juveniles, bred and analyzed as described previously²⁵. All mice were housed in an Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC)-accredited mouse facility or in the CIRB (French agreement number C75-05-12). We investigated isolated germ cells from 14 fetal gonads, follicle and mature GVs from >36 females, and produced more than 500 oocytes for analysis.

Fetal stage sex determination by PCR

Genomic DNA was isolated for Y-chromosome detection in GFP CETN2-expressing e11.5, e14.5 and e18.5 mouse tail tip tissues and PCR performed using MyTaq Extract-PCR Kit (Bioline, Taunton, MA). A 331-bp gene segment was amplified using the Jarid primers (forward: 5′ CTG AAG CTT TTG GCT TTG AG; reverse: 3′ CCG CTG CCA AAT TCT TTG G; Life Technologies, Carlsbad, CA). Β-actin loading control used the primers: 5′ GAT GAC GAT ATC GCT GCG CTG GTC G 3′ (forward), and 5′ GCC TGT GGT ACG ACC AGA GGC ATA CAG 3′ (reverse). PCR conditions were: 95 °C for 3 min, followed by 35 cycles of 95 °C for 15 seconds, 60 °C for 15 seconds, and 72 °C for 20 seconds. PCR product was analyzed in 2% agarose gel and visualized using ethidium bromide. Lanes expressing two bands were classified as males, the remainder female. Direct immunofluorescence of GFP CETN2 expression from non-gonadal cells at e11.5, e14.5, and e18.5 stages was accomplished as previously described²⁵.

Fetal germ cell collection

GFP CETN2-expressing female fetal gonads were collected following mating with GFP males (day of copulation plug = 0.5-day post-coitus [dpc]). Pregnant GFP CETN2 females were sacrificed at e11.5, e14.5, e15.5, and e18.5 dpc and gonads harvested as described in DeMiguel and Donovan⁷⁴. Gonads were digested in 200 µl of 0.05% trypsin:0.02% EDTA (ATCC, Manassas, VA) at 37 °C for 8–10 min, then inactivated with Trypsin Neutralization Buffer (ATCC) for 10 min at 37 °C. Digested gonads in 200 µl of warm modified human tubal fluid (HTF) medium without calcium or protein supplementation [mHTF-Hepes; formulation modified from Irvine Scientific, Santa Ana, CA] were mechanically pipetted 25–50 strokes with a P-200 sterile micropipette tip to release germ cells.

Follicular and mature germinal vesicle (GV)-stage oocyte collection

GFP CETN2 CB6F1 ovaries in EmbryoMax M2 (EMD Millipore, Billerica, MA) were minced mechanically to collect follicles of various sizes (Suppl Fig. S6) and GV’s released using sterile needles. Cumulus cells were removed with a 75-µm tip (Stripper pipet; Origio Mid-Atlantic Diagnostics, Trumbull, CT). Oocyte diameters were determined with Elements software (Nikon USA, Melville, NY) after image capture on a Nikon digital sight camera (DS Fi1). Oocytes were maintained in KSOM until fixation. Mature oocytes were harvested after stimulation with an intraperitoneal injection of 7.5 IU PMSG (Sigma-Aldrich, St. Louis, MO) for 48 hrs and kept in M2 supplemented with 100 µg/ml dibutyryl adenosine 3′,5′-cyclic monophosphate (dbcAMP; Sigma-Aldrich) to prevent meiosis resumption. To initiate meiotic maturation, mature GVs were rinsed 3x in M2 and placed in KSOM in a 37 °C, 5% CO₂ humidified incubator.

Cytoskeletal and PLK1 inhibitors

10 mM stocks of nocodazole, paclitaxel and BI 2536 PLK1 inhibitors (Sigma Aldrich) were prepared in DMSO and stored at −80 °C. Inhibitors were diluted to final concentrations in KSOM. Rescue experiments from inhibitors were performed by rinsing treated oocytes 3x in M2 before returning to inhibitor-free KSOM for recovery at the times specified.

Immunocytochemistry

Isolated primordial germ cells, oogonia from fetal gonads, and zona-denuded (EmbryoMax acid Tyrode’s, Millipore) GVs were attached to polylysine-coated (2 mg/ml; Sigma-Aldrich) 22 mm² coverslips and fixed in either 2% paraformaldehyde (pFA; Electron Microscopy Services, Hatfield, PA) or 0.5% pFA (5 min), followed by postfixing for 5 min in absolute ethanol (−20 °C). Primary antibodies included: mouse anti-SSEA-1 (Developmental Studies Hybridoma Bank, Iowa City, IA; 1:50), rabbit anti-Ak15 anti-γ-tubulin (Sigma-Aldrich; 1:500), mouse anti-pericentrin (clone 30; BD Transduction Laboratories, San Jose, CA; 1:200), rabbit anti-Cep135 (Sigma Aldrich; 1:800), rabbit anti-Cep250/cNAP1 (Proteintech, Rosemount, IL; 1:100), and rat anti-tubulin YOL 1/34 (EMD Millipore; 1:200), all diluted in PBS and applied overnight at 4 °C. Primary antibodies were detected with appropriate fluorescently-tagged secondary antibodies (1:500; Life Technologies) for 2 hr at room temperature in the dark after PBS rinses. DNA was labeled with Hoechst 33342 (10 µg/ml; 10 min) before mounting in ProLong Diamond antifade (Life Technologies).

Immunohistochemistry of intact ovaries for primary oocyte detection

P4 neonatal and adult excised ovaries were fixed in toto in 4% pFA (Electron Microscopy Services) in mHTF-Hepes overnight at 37 °C, rinsed 3x in sterile PBS, air dried at −20 °C for 2–4 hr, and then embedded in optimal cutting temperature compound (OTC; Tissue-Tek; VWR, Bridgeport, NJ) prior to storage at −80 °C. 7-µm sections were cut on a cryostat at −20 °C (CM1850 UV cryostat; Leica, Buffalo Grove, IL), floated onto clean glass slides (MidSci, Valley Park, MO), and stored at −80 °C. For immunostaining, slides were warmed at room temperature for 5 min, treated for 10 secs with Surgipath O-Fix (Thermo Fisher), and washed briefly in distilled H₂O. Immunostaining was performed as described above.

Equipment, Imaging, analysis, and settings

Fixed slide imaging was accomplished with a Nikon A1 four-laser line confocal microscope²⁵. Images were collected at 1024 × 1024 size at ¼ frames per second, a 79.2 μm pinhole, and a z-depth of 0.25 μm through the entire oocyte, with a differential interference contrast (DIC) Plan Fluor ×100 (1.3 NA) objective. The same laser photomultiplier tube settings for each channel across specimens helped facilitate comparison between oocytes. Fluorescent intensity ratios, mean intensities, area or volume measurements were performed on binarized images, using Elements software threshold tool and region-of-interest statistical menu, and downloaded to Microsoft Excel for statistical analysis. For image panels, a subtracted background image from outside of the oocyte was performed on generated confocal nd2 files. Deconvolution software (Landweber; supplied by Nikon USA), using the point scan confocal command and same filter (clear) at eight iterations was applied and selected deconvolved.tiff images prepared in Adobe Photoshop (Adobe Systems, San Jose, CA) for image panels.

Correlative light and electron microscopy

CLEM on GFP CETN2 mouse oocytes was adapted from a published protocol⁷⁵. GFP CETN2 foci were imaged on an inverted microscope (Eclipse Ti; Nikon, Tokyo, Japan) with a spinning-disk confocal head (CSUX Spinning Disk; Yokogawa Electric Corporation, Tokyo, Japan) and 200-nm-thick Z-sections spanning the entire oocyte recorded to register GFP CETN2 doublets positions. Adhering oocytes were then perfused with 2.5% glutaraldehyde, marked with a diamond scribe, and rinsed extensively in PBS. Next, samples were stained with 2% osmium tetroxide and 1% uranyl acetate, dehydrated, and then embedded in EMbed 812resin. Oocytes identified by light microscopy were then serially sectioned (200-nm-thick), transferred onto copper slot grids, stained with uranyl acetate and lead citrate, and imaged by transmission electron microscope (H-7650; Hitachi, Tokyo, Japan).

Mouse strains and genotyping

Both CB6-Tg (CAG-EGFP/CETN2)3-4Jgg/J and ZP3 Cre+; mCherry-Plk4+ female mice were used. The [C57BL/6-Tg(Zp3-cre)93Knw/J] breeding pairs were obtained from Jackson Laboratories. mCherry-Plk4^flox/wt mice were generated by random insertion of a pCAG-loxCATloxmCherryPlk4SV40pA construct in the genome of C57BL/6 N mice⁴⁹.

Microinjection

Injection of in vitro-transcribed cRNAs into the cytoplasm of Prophase I-arrested oocytes was performed using an Eppendorf FemtoJet microinjector as described⁷⁶, and oocytes kept for 1–3 hrs in milrinone to allow fusion protein expression. Oocytes were released from prophase I arrest by washing and transferring into milrinone-free M2.

Plasmid construction and in vitro transcription of cRNA

The mCherry-Plk4 construct produced the transgenic lines⁴⁹ and the Venus-Centrin 2 construct was subcloned into the pRN3 vector. cRNAs were synthesized with the T3 mMessage mMachine Kit (Ambion) and resuspended in RNase-free water⁷⁶.

Confocal spinning disk imaging of living oocytes

Spinning disk movies were acquired using a Plan APO 40 Å~/1.25 N.A. objective on a Leica DMI6000B microscope enclosed in a thermostatic chamber at 37 °C (Life Imaging Services), and equipped with a CoolSnap HQ2/CCD-camera (Princeton Instruments) or EMCCD camera (Evolve) coupled to a Sutter filter wheel (Roper Scientific) and a Yokogawa CSU-X1-M1 confocal scanner. MetaMorph software (Universal Imaging) collected the data.

Statistics

Means ± standard deviations were determined by EasyCalculations.com. We used Microsoft Excel to prepare graphs and box plots, which show median (horizontal lines), means (black squares), 25th and 75th percentiles (small boxes), and 5th and 95th percentiles (whiskers). Statistical significance was determined by Student’s t test (two-tailed), with actual p valves expressed, and was performed with GraphPad software (La Jolla, CA). Significance was determined at p < 0.05. Graphical analyses shown are indicative of average values ± standard deviation for more than three trials for most experiments.

Data availability

Datasets generated and/or analyzed during this study are available from the corresponding author on reasonable request as set forth in the journal guidelines.

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Acknowledgements

We thank Bruce Campbell and Angela Palermo Lauff for expert editorial assistance and manuscript submission. We are grateful for the sponsorship of the National Institutes of Health and Pennsylvania Department of Health (#4100077065) for project funding. We also acknowledge the technical help of the core lab facility at Magee-Womens Research Institution for assistance with immunohistochemistry, as well as the important contributions of Dr. Alice Meunier (IBENS, Paris, France) for the gift of GFP-CENT2 mice and Dr. Renata Basto (Curie Institute, France) for the gift of the Venus-Centrin 2 construct and the mCherry-Plk4 mice. MHV acknowledges funding from the FRM (FRM Label-DEQ20150331758 to MHV) and the Inca (grant PLBIO 2016-270-TRAN) in France.

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Authors and Affiliations

Departments of Cell Biology; Obstetrics, Gynecology and Reproductive Sciences; and Bioengineering, University of Pittsburgh School of Medicine, Pittsburgh, PA, 15213, USA
Calvin Simerly, Carlos Castro, Carrie Hartnett & Gerald Schatten
Center for Interdisciplinary Research in Biology (CIRB) Collège de France, CNRS, INSERM, PSL Research University, Equipe labellisée FRM, Paris, France
Marion Manil-Ségalen & Marie-Hélène Verlhac
Laboratory of Protein Dynamics and Signaling, National Institutes of Health/Center for Cancer Research/National Cancer Institute-Frederick, Frederick, MD, 21702, USA
Dong Kong & Jadranka Loncarek

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Calvin Simerly
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Marion Manil-Ségalen
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Carlos Castro
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Carrie Hartnett
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Dong Kong
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Marie-Hélène Verlhac
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Contributions

C.S. and G.S. designed the experiments and analyzed the data. C.S. and C.H. conducted the experiments, archived data, and assisted in manuscript preparation. M.M.S and M.-H.V performed live oocyte time-lapse imaging and analyzed the data. C.C. and C.H. prepared immunohistochemistry data with data analysis. D.K. and J.L. performed and analyzed the CLEM data. G.S. wrote the manuscript. All authors reviewed and approved the final manuscript.

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Correspondence to Gerald Schatten.

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Simerly, C., Manil-Ségalen, M., Castro, C. et al. Separation and Loss of Centrioles From Primordidal Germ Cells To Mature Oocytes In The Mouse. Sci Rep 8, 12791 (2018). https://doi.org/10.1038/s41598-018-31222-x

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Received: 02 July 2018
Accepted: 09 August 2018
Published: 24 August 2018
DOI: https://doi.org/10.1038/s41598-018-31222-x

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