Spatial dynamics of centromeres and telomeres during chromosome territory formation

To study the dynamics of chromosome territory formation, we performed time-lapse imaging of spermatocytes expressing His2Av-mRFP and Cenp-A/Cid-EGFP. His2Av-mRFP resulted in weak diffuse signals throughout the nucleus and in stronger signals in regions with chromatin (Fig 1A). Based on the diffuse signals, the nuclear diameter (d_n) was determined, which correlates with the spermatocyte developmental stage [9,10]. In spermatocytes with d_n around 9 μm, i.e. during late S2 stage, our time-lapse imaging revealed the conversion of the His2Av-mRFP chromatin signals from a state without to a state with spatially separated chromosome territories (Fig 1A), as expected [9]. The impossibility to delineate individual bivalents before territory formation precluded a precise scoring of the start of the relatively gradual process. However, to a first approximation, the spatial re-organization of chromatin appeared to take about three hours in the fastest cases and up to about six hours, during which nuclear diameters grew minimally (1–4%).

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Fig 1. Disruption of centromere clusters during chromosome territory formation.

Spermatocytes expressing His2Av-mRFP and Cenp-A/Cid-EGFP were analyzed by time-lapse imaging. (A) Labeling of Cid-EGFP dots indicates their association with distinct chromosomes. “Aa” and “Ab” designate centromere clusters associated with the large autosomes (chr2 and chr3) with “1” and “2” denoting the two homologs. “X”, “Y” and “4” designate the clustered centromeres of the additional chromosomes (chrX, chrY and chr4). Complete separation of chromosome territories (dashed lines) is evident at the last time point. Scale bar = 5 μm. (B) Signal intensity of the Cid-EGFP dots displayed by the spermatocyte shown in (A). Dashed grey lines indicate dot-splitting events. (C) Duration of the stages with three, four and five Cid-EGFP dots per nucleus, respectively. Grey values represent actual stage durations determined for spermatocytes that progressed through both dot-splitting events delimiting a particular stage. Mean ± s.d. of these values is displayed as well. Values plotted in color are from cells, in which only one of the two delimiting dot-splitting events was observed during the imaging period, thus indicating minimal durations. (D) Relative Cid-EGFP signal intensity of the XY4 dot at the four-dot stage compared to the Aa1 dot (formed by the two tightly associated, unresolved sister centromeres of a large autosome). Mean ± s.d. is displayed as well (n = 10). (E) Cid-EGFP dots are far from the nuclear periphery during dot-splitting events. A representative event (Aa into Aa1 and Aa2) with a graph indicating the shortest distances between dots and nuclear periphery is shown. t = 0: start of stretching of the separating Cid-EGFP dot. (F) Position of Cid-EGFP dots relative to the nuclear periphery were tracked (after time-lapse imaging at five-second intervals over 7.5 min). Nine spermatocytes with three dots (Aa, Ab and XY4) were analyzed. Separation distance of a given Cid-EGFP dot at each time point (n = 89) is plotted, as well as mean ± s.d. (G) Cid-EGFP dots move faster during the S2 stage when chromosome territories form. Cid-EGFP dots were tracked (after time-lapse imaging at five-second intervals). Bars represent mean dot velocity (± s.d.) obtained by averaging over all dots present in a cell (i.e., 3–5 dots depending on stage) over all 99 time intervals and over all cells analyzed for a given stage (3–5 spermatocytes, see S3 Fig). *** indicates p < .001 (t test). Scale bars = 5 μm.

https://doi.org/10.1371/journal.pgen.1009870.g001

Time-lapse imaging of the centromeric Cenp-A/Cid-EGFP dots indicated that chromosome territory formation was accompanied by centromere de-clustering (Fig 1A and 1B and S1 Movie), as implied previously by live imaging at single time points [10] and by microscopy with fixed samples [54]. The overwhelming majority of S1/2 spermatocytes, which did not yet have well separated territories according to the His2Av-mRFP signals, displayed 2–3 Cid-EGFP dots (97%, n = 184). Spermatocytes with either one or more than three dots at these early stages were rare (2 and 1%, respectively). A series of dot splitting events was observed during chromosome territory formation and thereafter, raising dot number up to six or seven (Fig 1A and 1B). More rarely, dot fusions were observed as well, which reduced the total dot number but only transiently. The temporal dynamics of centromere de-clustering varied considerably (Figs 1C and S1). The four-dot stage had the lowest variability and the shortest mean duration compared to the three- and five-dot stages (Fig 1C). Based on the spermatocytes progressing from the three- to the five-dot stage during the 12 hour imaging period, the mean duration of the four-dot stage was 72 min (± 50 s.d., n = 15) (Fig 1C). This is a slight underestimate, because there were a few among the imaged cells with a more extended four-dot stage. The actual duration of the four-dot stage could not be determined in these exceptional cells, because only one of the two dot-splitting events delimiting the four-dot stage was observed within the imaging period (12 hours). In case of the three-dot stage, only three among the analyzed cells displayed both delimiting dot-splitting events during the imaging period. However, cells progressing through only one of the dot-splitting events provided additional confirmation that the three-dot stage lasts longer than the four-dot stage.

Centromere de-clustering was paralleled by changes in signal intensities of the centromeric Cid-EGFP dots. Intensity quantification in combination with the tracking of centromeric signals over time indicated that the four-dot stage rarely represented a state where each bivalent is associated with a single centromere cluster. During entry into the first meiotic division, when each of the four bivalents can be identified, centromeres display highly comparable Cid-EGFP intensities except for the chrY centromere, which exhibits a twofold higher intensity [54,55]. Accordingly, if each dot at the four-dot stage represents a cluster of all the centromeres of a bivalent, the dot on the sex chromosome bivalent is expected to have a 1.5 fold greater intensity than the remaining three autosomal dots of equal intensity. However, the intensities at the four-dot stage did not conform to this prediction. Compared to the weakest centromere dot, the strongest was on average 6.4 fold stronger rather than just 1.5 fold more intense (Fig 1D). Moreover, dot tracking over time revealed that the dots present at the four-dot stage were not partitioned each into a distinct chromosome territory. In the majority of the analyzed spermatocytes (85%, n = 20), the two weakest dots of the four-dot stage were partitioned together into one of the two large autosomal chromosome territories (Fig 1A). These large autosomal territories were characterized by more uniform His2Av-mRFP signals compared to the territories of chr4 and chrXY, which were usually in close spatial association [54,55].

Back-tracking centromeric dot signals from stages after territory formation to earlier stages revealed a most frequent centromere de-clustering program. When a single centromere cluster was present at the earliest stage, a first dissociation event liberated a smaller centromere cluster (Aa), containing all centromeres of one of the large autosomal bivalents (i.e., a cluster with the two homologous pairs of sister centromeres). A subsequent event dissociated an analogous cluster with all centromeres of the other large autosome (Ab) from the most intense centromere cluster. This most intense cluster thereafter still contained all centromeres of chrX, Y and 4. Accordingly, centromere clusters Aa, Ab and XY4 were present at the three-dot stage (Fig 1A). Importantly, the subsequent four-dot stage did not result from XY4 cluster splitting into a chrXY and a chr4 cluster. Rather, the Aa centromere cluster was separated into two dots (Aa1 and Aa2), each containing an unresolvable sister centromere pair (Fig 1A). Typically, chromosome territory formation paralleled the transition from the three—to the four-dot stage. The subsequent splitting of the Ab cluster into two dots (Ab1 and Ab2) resulted in the five-dot stage (Fig 1A). The six- and seven-dot stages arose by release of dots from the most intense XY4 cluster. At the seven-dot stage, each of the large autosome territories Aa and Ab contained two dots, and the XY4 territory comprised three additional dots: the paired sister centromeres of chrX (dot 1) and of chrY (dot 2), and a dot with all the chr4 centromeres (dot 3). A final de-clustering step dissociated the single chr4 dot into two, each containing a pair of sister centromeres. This final transition to the eight-dot stage occurred usually after a long delay just around the time of NEBD I at the end of the S6 stage [55]. A permanent separation of sister centromeres before M I was never observed, but occasional transient breathing of sister centromeres in large autosomal territories could be detected, as described [55].

The de-clustering program described above was observed in 65% of the spermatocytes analyzed by dot tracking (n = 20). The remaining 35% displayed deviations from the canonical program. In 20%, the variation concerned progression from the four to the five-dot stage. In these cases, the five-dot stage was reached by the release of a dot from the most intense XY4 dot before Ab splitting. The newly released dot contained all centromeres of either chrY or chr4, because it was about twofold more intense than Aa1 or Aa2. The final 15% of the tracked spermatocytes varied in the progression from the three to the four-dot stage. In these cells, the release of a dot from the most intense XY4 dot occurred even earlier, already at the three-dot stage before Aa splitting. Again, the dot released early represented all centromeres of either chrY or chr4 based on its intensity. Of relevance for Cid-EGFP dot intensity quantification, the Aa dot splitting into Aa1 and Aa2, as well as the analogous Ab dot splitting and the occasional transient sister centromere breathing events were all accompanied by an approximate 1:1 partitioning of the precursor dot intensity (Fig 1B), indicating that Cid-EGFP quantification permits a reliable detection of twofold differences, in particular when positions along the z axis remained comparable.

Overall, these results disprove the presence of an obligate four-dot stage, where each bivalent displays a single centromere cluster, as previously claimed [10]. Such a centromere pattern is present in at most 15% of the spermatocytes.

The absence of an obligate four-dot stage with a single centromere cluster per bivalent does not necessarily rule out an involvement of centromeres in territory formation. Cytoskeletal forces might still act via centromeres to increase chromosome mobility in scenarios more complicated than a straightforward separation of bivalents via unique handles. However, if territory formation were to involve centromere movements by cytoskeletal forces acting via LINC complexes (or comparable linkages), centromeres would be expected to move at close distance along the NE. Therefore, we analyzed the intranuclear positions of the Cid-EGFP dots. First, we focused on dot de-clustering events (two-to-three and three-to-four dot transitions). In six out of eight analyzed events, dot splitting occurred far from the nuclear periphery. The splitting dot was between 0.4 and 2.3 μm away from an isosurface delineating the nuclear His2Av-mRFP signal (Fig 1E). Cid-EGFP dots present in addition to the splitting dot were also localized far from the nuclear periphery (Fig 1E).

For further analysis of Cid-EGFP dot positions, spermatocytes were imaged at 5 sec intervals for error-free dot tracking at high resolution. The spatial separation of Cid-EGFP dots from an isosurface delineating the nuclear His2Av-mRFP signal was determined in spermatocytes at the S2 stage when chromosome territories form. As described above, three Cid-EGFP dots were usually present at this stage. Cid-EGFP dots at positions close to the nuclear periphery (d < 0.4 μm) were almost never observed (Fig 1F), even in case of the XY4 dot, which tended to be closer to the nuclear periphery than the Aa and Ab dots (Fig 1F). Finally, we labeled Cid-EGFP whole mount testis preparations with anti-Lamin and a DNA stain. Centromere positions were found to be at least 0.5 μm and up to 3.5 μm away from the nuclear lamina during the stages S1 to S5 (S2 Fig). Moreover, at the stages before complete separation of the territories, DNA signals were separated from the nuclear lamina by an unstained gap (S2 Fig). In conclusion, given the considerable distance separating centromeres from the nuclear envelope, a direct mechanical coupling between cytoskeletal forces and centromeres for the purpose of territory formation appears improbable.

The Cid-EGFP dot tracking was also used for the characterization of centromere movements. After correcting lateral nuclear drift (using an isosurface around the His2Av-mRFP signal), centromere dot velocities were determined. Similar analyses with Drosophila ovaries have revealed rapid nuclear rotations in gonial cells during the stage where the cysts consist of eight cells [3,56]. In testes, the average speed of centromeres was around 0.03 μm/sec during interphase in gonial eight-cell cysts, as well as in S1-S5 spermatocytes (Figs 1G and S3). This is tenfold slower than during the nuclear rotations in ovarian eight-cell cysts [3,56], and also considerably slower than the RPMs during C. elegans meiosis [57]. Interestingly, however, in Drosophila spermatocytes at the S2 stage, transient phases of more rapid movements in particular of the most intense XY4 centromere cluster tended to be more frequent than in the preceding and following stages (S3 Fig). Averaging across all Cid-EGFP dots, all time points and all cells confirmed that centromere movements were significantly faster during the S2 stage compared to the stages before and thereafter (S1, S3, S4/5) (Fig 1G). Therefore, territory formation, which occurs during the S2 stage, is accompanied by phases with increased chromosome mobility. Similar phases with comparatively fast movements of at least some centromeres were observed in early round spermatids (S3 Fig), in which up to four Cid-EGFP dots were detectable. The increased centromere mobility in spermatids coincides with the known, striking relocalization of LINC and nuclear pore complexes from a spherical to a hemispherical distribution within the NE of these cells [58,59].

In mammals and yeast, RPMs during meiosis are led by telomeres linked via LINC complexes to cytoskeletal forces [3,4]. In Drosophila, electron microscopy has failed to reveal a classical bouquet stage with telomeres attached or close to the NE in pachytene oocytes [60]. To characterize telomere behavior in spermatocytes, we exploited an EGFP knockin at caravaggio (cav) [61]. This gene codes for the telomere protein HOAP. If each telomere were resolved as a dot, the number of EGFP-HOAP dots per nucleus would be 16 in G1 and 32 in G2. However, only four to six EGFP-HOAP dots were detected in early embryos and larval brain cells, as a result of telomere clustering [62]. Based on limited data from fixed spermatocytes, partial telomere clustering occurs in these cells also [61,63]. For a detailed analysis, we applied time-lapse imaging. Analysis of M I confirmed that EGFP-HOAP is a faithful telomere marker in spermatocytes (Fig 2A). EGFP-HOAP dots were localized at the trailing ends of the large chromosomes during anaphase I. Counts of the EGFP-HOAP dots provided further support that they represent telomeres. At the late S6 stage, after partial condensation of bivalents into well-isolated territories just before NEBD I, 25 EGFP-HOAP dots were present in the cell shown for illustration (Fig 2A). Eight dots were present in one of the large autosome territories (Fig 2A), corresponding to the expected number in case of separated sister telomeres. Only seven EGFP-HOAP dots were resolved in the other large autosome territory (Fig 2A), because two were presumably too close. Finally, ten EGFP-HOAP dots were associated with the XY4 territory, and two of those were clearly brighter than the rest (Fig 2A). Each of these two bright dots contained the four clustered telomeres of a chr4 homolog, as indicated by their subsequent segregation during M I. Numbers and distributions of EGFP-HOAP dots were comparable in the analyzed M I spermatocytes (n = 7 from five distinct cysts). In conclusion, close to all telomeres can be resolved individually in late spermatocytes except for those of the small chr4. Moreover, the amount of EGFP-HOAP per telomere does not vary extensively.

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Fig 2. Telomeres display limited movements far from the nuclear periphery during territory formation.

Spermatocytes expressing His2Av-mRFP and the telomere protein EGFP-HOAP were analyzed by time-lapse imaging. (A) Progression through M I. Time (min:sec) is given relative to the onset of NEBD I. Chromosome territories labeled as in Fig 1A. The two most intense EGFP-HOAP dots (arrows) represent telomere clusters of the two homologs of the small dot-like chr4. During anaphase, EGFP-HOAP dots were clearly at the trailing ends of long chromosome arms (see arrowheads for example). (B) Telomere de-clustering at the end of the S6 stage. A large autosome territory is shown with the number of EGFP-HOAP dots indicated (top). Time (min:sec) is given relative to the onset of NEBD I. (C) During the stage of chromosome territory formation, telomeres are partially clustered. EGFP-HOAP dots vary in numbers and intensity, as illustrated with two neighboring spermatocyte nuclei from an S2 cyst with the number of EGFP-HOAP dots per nucleus indicated. (D,E) Intranuclear positions and velocities of telomeres during territory formation. After time-lapse imaging (at five-second intervals), all EGFP-HOAP dots sufficiently strong for reliable tracking over time were analyzed in five distinct cells (c1-c5) during the S2 stage. All distances (D) and all velocities (E) observed for a given EGFP-HOAP dot during an 8.5 min period were plotted (n = 102), as well as the mean ± s.d. Scale bars = 3 μm (A), 1 μm (B) and 3 μm (C).

https://doi.org/10.1371/journal.pgen.1009870.g002

Time-lapse imaging at earlier stages revealed fewer EGFP-HOAP dots of more variable intensities, indicating partial telomere clustering. Indeed, de-clustering of EGFP-HOAP dots could be observed readily during the S6 stage (Fig 2B). In early spermatocytes during the stages of chromosome territory formation, the number of telomere clusters was variable, as well as the intensities of the associated EGFP-HOAP signals. Even neighboring cells within the same cyst were observed to differ with regard to the extent of telomere clustering (Fig 2C). The EGFP-HOAP dots in S2 spermatocytes were tracked. As for the Cid-EGFP signals, the distance between telomere dots and an isosurface delineating the nuclear His2AvD-mRFP signal was determined, as well as their speed. Only the stronger EGFP-HOAP dots that could be tracked reliably were considered. Our results revealed that telomere clusters are neither consistently close to the NE (Fig 2D) nor moving fast (Fig 2E) during the stages of chromosome territory formation. Therefore, telomeres are unlikely used for the mobilization of chromosomes by mechanisms comparable to those acting during the bouquet stage in yeast, mammals, or C. elegans [3,4].

Disruption of centromere clusters and chromocenters during chromosome territory formation in spermatocytes requires Cap-D3 and Cap-H2

While centromeres and telomeres do not appear to have a prominent role during chromosome territory formation in Drosophila spermatocytes, condensin II (Fig 3A) was suggested to be crucial for this process [23]. Previous analyses of Cap-D3 and Cap-H2 function were completed with mutant alleles that might not necessarily cause a complete loss of function. While the presence of several genes within the first Cap-D3 intron complicates the isolation of null mutations, this appeared feasible in case of Cap-H2. Applying CRISPR/Cas9 in combination with a pair of single guide RNAs for the generation of intragenic deletions of central Cap-H2 regions led to three distinct alleles (Fig 3B). Cap-H2^cc1 and Cap-H2^cc2 are intragenic, in-frame deletions of 918 and 1488 bp, respectively. Cap-H2^cc3 carries a 5 bp, frame-shifting deletion in the gRNA1 target region, as well as a 918 bp deletion further downstream. These newly induced alleles are expected to affect all four annotated Cap-H2 isoforms (D-G) (Fig 3B). In wild type, the isoforms E, F and G are identical except for a short internal stretch of 13 amino acids (aa) that is either present (E), absent (F), or shortened to 12 aa (G), as a result of differential splicing of the fourth intron. The D isoform is most unusual. It is a variant F isoform without the N terminal-most region and with a distinct C terminal region, resulting from the use of a downstream transcriptional start site and differential splicing. The newly isolated Cap-H2^cc3 mutation might still permit expression of N-terminal Cap-H2 fragments (aa 1–186 of E- G; aa 1–8 of D) followed by a few extra amino acids after the frame shift until the premature stop. These truncated Cap-H2^cc3 products lack the regions predicted to bind Cap-D3, Cap-G2 and SMC4/Gluon (Fig 3B) [64–66], but the N-terminal SMC2-binding region is still present. Thus, the putative truncated E-G isoforms encoded by Cap-H2^cc3 might exert a dominant-negative effect in principle, although mitigated by non-sense mediated RNA decay presumably. The protein products encoded by Cap-H2^cc1 and Cap-H2^cc2 include both the N- and the C-terminal domain for binding to SMC2 and SMC4, respectively (Fig 3B). However, these internally truncated β-kleisins are predicted to be defective in Hawk recruitment. In case of Cap-H2^cc2, the regions predicted to bind Cap-D3 and Cap-G2 are both absent, while only the former is missing in Cap-H2^cc1 (Fig 3B).

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Fig 3. Intragenic Cap-H2 deletions abolish disruption of centromere clusters and chromocenters in spermatocytes.

(A) Condensin II complex organization. (B) Drosophila Cap-H2 codes for four isoforms annotated as E, F, G and D. Predicted binding sites for condensin II subunits are indicated, as well as a degron [48]. Drosophila Cap-G2 existence is uncertain (see discussion). Three intragenic deletions, cc1 –cc3, were generated by CRISPR/Cas9. Regions deleted in case of the in-frame deletions cc1 and cc2 are indicated (grey shading with dashed borders). Two deletions are present in cc3, a small 5 bp deletion resulting in a premature stop (grey arrow) and a downstream in-frame deletion identical as in cc1. (C) Fertility of adult flies with the indicated genotypes. The deficiency Df(3R)Exel6159 (Df) eliminates Cap-H2. Bars report male fertility relative to +/+ males (100%), and female fertility relative to cc3/+ (100%). (D) Failure of centromere cluster disruption. Squash preparations of testes from control (+ / +) and the indicated Cap-D3 and Cap-H2 mutants were labeled with anti-Cid and a DNA stain. High magnification views of spermatocyte nuclei at the indicated stages are displayed. The dot plot presents the number of anti-Cid dots per nucleus, as well as the mean dot number (red cross) at the indicated stages. (E) Failure of chromocenter disruption. Whole mount preparations of testes from the indicated genotypes were labeled with anti-Prod and anti-Lamin Dm0. High magnification views from regions with spermatogonial cells (sg) and spermatocytes at the indicated stages are presented. Scale bars = 5 μm.

https://doi.org/10.1371/journal.pgen.1009870.g003

Zygotes with either one of the three newly isolated Cap-H2 alleles in trans over a deficiency deleting Cap-H2 developed to the adult stage with Mendelian frequencies. Morphologically, the hemizygous adults were apparently normal and females were fully fertile (Fig 3C). In contrast, male fertility was severely reduced in case of Cap-H2^cc1, almost completely eliminated by Cap-H2^cc2 (one escaping progeny fly), and totally abolished by Cap-H2^cc3 (Fig 3C). Cap-H2^cc1, therefore, and perhaps even Cap-H2^cc2 have residual function. Moreover, we conclude that Cap-H2 function is essential exclusively for male fertility, and it appears to contribute in a non-essential manner to cellular processes like chromosome condensation during M phase [67], interphase chromosomal organization and gene expression [17,22,24,49]. In contrast to Drosophila, the mouse ortholog Ncaph2 is required for embryonic development [68].

For an initial analysis of the effects of the newly isolated Cap-H2 alleles on chromosome territory formation, we analyzed squash preparations of testes after DNA staining. The characteristic compartmentalization of nuclear DNA staining that reveals chromosome territories in control spermatocytes at stage S2b and later, was never observed in Cap-H2^cc3/ Df(3R)Exel6159 mutants (S4 Fig), as previously reported for other strong Cap-H2 and Cap-D3 alleles [23]. The phenotypes observed in Cap-H2^cc3/ Df(3R)Exel6159 and Cap-D3^EY00456/ Df(2L)Exel7023 were indistinguishable (S4 Fig). As Cap-H2^cc3, Cap-D3^EY00456 is very likely an amorphic allele [23]. The other alleles, Cap-H2^cc1 and Cap-H2^cc2, also resulted in severe defects in chromosome territory formation (S4 Fig).

To clarify whether Cap-D3 and Cap-H2 are required for the disruption of centromere clusters during chromosome territory formation and subsequent spermatocyte maturation, we applied anti-Cenp-A/Cid labeling (Fig 3D). In control spermatocytes at the S2/3 stage, when chromosome territories are formed, an average of 3.3 (± 0.8 s.d.) dots were detected by anti-Cid (Fig 3D). At stages S4 and S5/6, the number of dots increased to 5.2 (± 1.2 s.d.) and 6.7 (± 0.6 s.d.), respectively (Fig 3D). However, in spermatocytes of Cap-D3 and Cap-H2 mutant males (Cap-D3^EY00456/ Df(2L)Exel7023 and Cap-H2^cc3/ Df(3R)Exel6159) centromere de-clustering was not observed. The mean number of anti-Cid dots per nucleus remained essentially constant at around two (Fig 3D). This result strongly argues against the notion that chromosome territories might still be formed in condensin II protein mutants, remaining cryptic however when analyzed by standard DNA staining because of extensive territory overlap resulting from a chromatin condensation failure. Evidently, the elimination of non-homologous chromosome associations is largely blocked in the absence of condensin II proteins.

Recently, the heterochromatin proteins D1 and Prod were shown to be crucial for chromocenter formation in Drosophila cells [69,70]. These proteins bind to pericentromeric satellites. D1 binds primarily to the {AATAT}_n satellite, which is abundant on chrX, Y and 4 but scarce on chr2 and 3. However, chr2 and 3 contain extensive arrays of the {AATAACATAG}_n satellite, which recruits Prod. Dynamic binding of D1 and Prod to their target satellites, in combination with dynamic interactions between these two proteins, was proposed to drive intranuclear clustering of pericentromeric heterochromatin of all chromosomes into a chromocenter [69,70]. Conversely, chromosome territory formation in spermatocytes, which disrupts the non-homologous chromosome associations in the chromocenter, might depend on control of D1 and Prod behavior. A re-distribution of D1 and a disappearance of Prod was reported to accompany chromosome territory formation during normal spermatogenesis [70]. Moreover, experimental prolongation of Prod expression was shown to result in incompletely separated, bridged chromosome territories [70], suggesting that the normal disappearance of Prod during spermatogenesis is crucial for normal territory formation. However, it remains to be explained why D1 is re-distributed and why Prod disappears from spermatocytes around the stages of chromosome territory formation. In principle, these processes might be driven by condensin II.

To evaluate whether condensin II proteins are required for displacing D1 and Prod from their pericentromeric target satellites, we analyzed the behavior of these proteins in Cap-H2^cc3/ Df(3R)Exel6159 mutants and controls (Figs 3E and S5). In wild-type testes, D1-sfGFP expressed under control of the endogenous D1 cis-regulatory region was readily detected in the previously reported pattern [70] (S5 Fig). In spermatogonial cells, D1-sfGFP was strongly enriched in 1–3 dots per nucleus. In contrast, in S1/2 spermatocytes, D1-sfGFP signals were reduced and far more diffuse throughout the nucleus. This drastic change in distribution was also observed in Cap-H2 mutant testis (S5 Fig), indicating that the re-organization of D1 before chromosome territory formation does not depend on Cap-H2.

Prod expression in wild-type testis detected with anti-Prod [71] was also observed essentially in the previously reported pattern [70], although with a clear asynchrony in the re-organization of D1 and Prod during spermatogenesis (S5 Fig). In spermatogonial cells, anti-Prod signals were strongly enriched in 1–2 intranuclear dots. Early spermatocytes during the stages S2b/S3, when chromosome territories are formed, displayed primarily two Prod dots per nucleus (Fig 3E). During S4, anti-Prod signals decreased in intensity and in S5/6 spermatocytes, they were no longer detectable (Fig 3E). In Cap-H2 mutants, spermatogonial cells displayed anti-Prod signals indistinguishable from wild-type (Fig 3E). However, most S2b/S3 spermatocytes in Cap-H2 mutants displayed one rather than two anti-Prod dots (Fig 3E). At later stages, Prod disappeared also from Cap-H2 mutant spermatocytes as in wild-type (Figs 3E and S5). Interestingly, during Prod disappearance, the anti-Prod signals in both control and Cap-H2 mutants were frequently re-organized in space into a series of finer dots.

In conclusion, these results indicate that condensin II does not promote chromosome territory formation by enforcing the release of D1 and Prod from the chromocenter. Moreover, the disappearance of Prod might be of limited importance for chromosome territory formation. In wild-type, this disappearance occurred primarily after the partitioning of DNA into territories. Anti-Prod dots of maximal intensity were still present during the stages of chromosome territory formation in wild-type S2b/S3 spermatocytes and the disintegration of the anti-Prod dots during disappearance suggested that the separation forces acting on chromatin during territory formation might be greater than a putative Prod-mediated heterochromatin cohesion. However, the persistence of a single anti-Prod focus in Cap-H2 mutants during territory formation indicated that successful chromocenter disruption requires Cap-H2.

Failure of bivalent individualization in Cap-D3 and Cap-H2 mutants precludes regular bi-orientation and segregation during M I

Meiotic chromosome missegregation in Cap-D3 and Cap-H2 mutant males has been described previously based on segregation analysis of genetic markers and cytology with fixed samples [23]. For further characterization, we performed time-lapse imaging with condensin II mutant spermatocytes expressing Cenp-A/Cid-EGFP and His2Av-mRFP. Analogous analyses of control spermatocytes were reported previously [55]. In control spermatocytes before NEBD I, His2Av-mRFP reveals the three major chromosome territories containing chrXY4, chr2 and chr3, respectively (Fig 4A). Each of the two large autosome territories (Fig 4A, Aa and Ab) almost invariably displays two Cid-EGFP dots. In contrast, the number of Cid-EGFP dots associated with the chrXY4 territory is more variable. In spermatocytes with fewer than four Cid-EGFP dots within the chrXY4 territory, residual centromere clusters are disrupted during NEBD I, when the final rapid chromosome condensation converts the territories into compact chromosome blobs. The His2Av-mRFP marker reveals the large bivalents clearly (chrXY, 2 and 3), but the dot chr4 contains very little His2Av-mRFP (Fig 4A). Rapid centromere-led, saltatory movements of bivalents during prometaphase I precede their eventual bi-polar integration into a compact metaphase plate before bivalents split in anaphase I (Fig 4A).

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Fig 4. Centromere and chromatin dynamics during M I in condensin II protein mutants.

Time-lapse imaging of His2Av-mRFP and Cenp-A/Cid-EGFP was applied for analysis of progression through M I in (A) control (+ / +), (B-E) Cap-H2 mutants (Cap-H2^cc3/ Df), and (F) Cap-D3 mutants (Cap-D3^EY/ Df). Time (min:sec) is indicated relative to the onset of NEBD I. (C) Arrowheads point to a chr4 bivalent that is separated out of the single clump of chromatin with clustered centromeres. (D) Arrowheads point to micronucleus resulting after bridge formation during anaphase. (E) Single clump of chromatin with clustered centromeres perduring into late anaphase and telophase after a complete failure of chromosome segregation. (G) Bars indicate the percentage of spermatocytes with chromosome bridges during anaphase I in the indicated genotypes. n = 48 from 5 cysts (+ / +), 62 from 10 cysts (Cap-H2^cc3/ Df), 48 from 6 cysts (Cap-D3^EY00456/ Df) and 28 from 6 cysts (Cap-D3^EY00456/ Df; mnm^z3-5578/ mnm^z3-3298). (H) Bars indicate the percentage of spermatocytes with 4:4 segregation of centromeres based on Cid-EGFP dot tracking during exit from M I. n = 48 from 5 cysts (+ / +), 45 from 8 cysts (Cap-H2^cc1/ Df), 8 from 2 cysts (Cap-H2^cc2/ Df), 63 from 10 cysts (Cap-H2^cc3/ Df), 46 from 6 cysts (Cap-H^EY00456/ Df) and 28 from 6 distinct cysts (Cap-D3^EY00456/ Df; mnm^z3-5578/ mnm^z3-3298). Scale bars = 3 μm.

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During the S6 stage, Cap-H2 mutant spermatocytes (Cap-H2^cc3/ Df(3R)Exel6159) did not display the characteristic chromatin compartmentalization into three major territories. While chromatin was still enriched at the nuclear periphery, it was in a contiguous reticular pattern. Moreover, the large majority displayed only one or two Cid-EGFP dots. These centromere clusters were dissociated into multiple dots (usually around six per nucleus) during the final two hours before NEBD I, in parallel with the initial slow condensation and release of chromatin from the NE that accompanied nuclear rounding (Fig 4B). However, the individualized Cid-EGFP dots remained in close proximity during NEBD I, when chromatin was rapidly compacted into a single large clump (Fig 4B). Remarkably, this final rapid chromosome condensation during NEBD I, proceeded with apparently normal efficiency, presumably driven by condensin I. During prometaphase I, the single chromatin clump was partly broken up, when centromeres along with associated chromatin were pulled away by spindle forces. The chr4 bivalent was most often separated away (Fig 4C), indicating that this small chromosome was less stably detained within the chromatin clump compared to other bivalents. Released bivalents were usually bi-oriented eventually, moving into a metaphase plate that also contained the residual chromosome clump. During anaphase, centromeres were pulled towards the spindle poles (Fig 4B) and chromosomes were separated with some lagging and with variable success. 44% of the spermatocytes (n = 63) displayed chromatin bridges during anaphase (Fig 4G). These chromatin bridges were eventually resolved during telophase in some cells, but also persistence of bridges and micronuclei formation was observed in 22% of the spermatocytes (n = 63) (Fig 4D) or a complete failure of chromatin clump separation into two portions in 11% of the spermatocytes (n = 63) (Fig 4E).

Bi-orientation of centromere pairs during prometaphase I was severely compromised in Cap-H2 mutants. As an estimate for the upper limit of the bi-orientation success, the fraction of spermatocytes with a 4:4 separation of centromeres during anaphase I was determined. The chromatin clumping in the mutants precluded an assignment of Cid-EGFP dots to specific bivalents, and thus an unknown number of the 4:4 separation events might reflect balanced syntelic rather than regular amphitelic chromosome attachment. In Cap-H2^cc3/ Df(3R)Exel6159 mutants, 4:4 segregation during anaphase was detected in only 14% of the cells (Fig 4H). Accordingly, 86% or more of the mutant spermatocytes failed at regular bi-orientation of bivalent centromeres. The Cap-H2^cc1 allele caused fewer mistakes in centromere bi-orientation (Fig 4H). In case of Cap-H2^cc2, which was less extensively analyzed, bivalent mis-orientation appeared to be intermediate (Fig 4H).

For comparison, we performed analogous time-lapse analyses with Cap-D3^EY00456/ Df(2L)Exel7023 mutants. The abnormal phenotype observed during progression through M I in these mutants (Fig 4F) was highly similar to that in Cap-H2^cc3/ Df(3R)Exel6159. Chromatin bridges were displayed during anaphase I also in 44% of the spermatocytes and the fraction of cells with 4:4 segregation of centromeres during anaphase I was only 6% (Fig 4G and 4H).

In conclusion, our results confirm that the condensin II proteins Cap-D3 and Cap-H2 are essential for chromosome territory formation and regular chromosome segregation during M I [23]. Moreover, they provide additional insights into the role of chromosome territory formation. Previous studies have led to the proposal that chromosome missegregation during M I in Cap-D3 and Cap-H2 mutants arises from heterologous chromosome entanglements that exert their damaging effect during anaphase I by inhibiting chromosome separation [23]. Our analysis of centromere behavior revealed that meiotic chromosome segregation in Cap-D3 and Cap-H2 mutants is irregular for an additional reason. The failure of chromocenter break-up in the mutants has highly detrimental effects already before anaphase I. It generates a cluster of mechanically coupled centromeres that precludes regular bi-orientation of homologous centromeres during prometaphase I.

We add that our time-lapse imaging also revealed the unexpected finding that chromosome territories do not only form in spermatocytes but also in cyst cells in a condensin II-dependent manner (S6 Fig).

Limited contribution of alternative homolog conjunction to meiotic chromosome missegregation in condensin II protein mutants

In principle, the chromosome bridges during exit from M I in Cap-D3 and Cap-H2 mutants, might result from a failure in elimination of alternative homolog conjunction (AHC). Normally, AHC is eliminated by separase-mediated cleavage of the AHC protein UNO during the metaphase-to-anaphase I transition [53]. To address whether condensin II proteins are required to remove AHC during M I, we performed time-lapse imaging with Cap-D3^EY00456/ Df(2L)Exel7023 spermatocytes expressing UNO-EGFP.

Normally, UNO-EGFP is co-localized with other AHC proteins throughout spermatocyte maturation [53]. At the start of M I, it most strongly enriched on the chrXY pairing region, where it forms a very prominent dot that disappears within minutes at the onset of anaphase I (Fig 5A and 5C) [53]. UNO-EGFP signals on autosomes, which disappear concomitantly, are far lower and difficult to detect above background [53]. These weak autosomal UNO-EGFP signals are below detection limit with the chosen display settings (Fig 5A).

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Fig 5. Limited contribution of alternative homolog conjunction to meiotic chromosome missegregation in condensin II protein mutants.

(A-C) The disappearance of the alternative homolog conjunction protein UNO during M I was analyzed by time-lapse imaging in (A) control (+ / +) and (B) Cap-D3 mutants (Cap-D3^EY00456/ Df). Beyond UNO-EGFP, spermatocytes expressed His2Av-mRFP, and in case of the Cap-D3 mutants also Cenp-A/Cid-EGFP. The prominent UNO-EGFP dot on the chrXY pairing site (arrowheads) disappeared during anaphase I with comparable rapid kinetics in both genotypes, as confirmed (C) by quantification of the UNO-EGFP dot intensity over time. (D) Progression into and through M I was analyzed by time-lapse imaging in Cap-D3 mutants (Cap-D3^EY00456/ Df) (top) and Cap-D3 mnm double mutants (Cap-D3^EY00456/ Df; mnm^z3-5578/ mnm^z3-3298) (bottom). Chromosomes condense into a single clump irrespective of presence or absence of AHC. Time (min:sec) indicated relative to the onset of NEBD I. Scale bars = 3 μm.

https://doi.org/10.1371/journal.pgen.1009870.g005

To detect the metaphase-to-anaphase I transition more clearly in our time-lapse imaging with Cap-D3 mutant spermatocytes, they expressed not just UNO-EGFP and His2Av-mRFP but also Cenp-A/Cid-EGFP (Fig 5B). As shown previously [53], the strong UNO-EGFP dot on the chrXY pairing region can be distinguished readily from centromeric Cid-EGFP dots based on intensity and shape. As in normal spermatocytes, a single prominent UNO-EGFP dot was present in Cap-D3 mutant spermatocytes just prior to NEBD I (Fig 5B), indicating that AHC protein localization is not affected, at least on the sex chromosome bivalent. However, we were unable to resolve whether autosomal UNO-EGFP dots are present or absent in Cap-D3 mutant spermatocytes. Strong enhancement of the EGFP display settings, which reveals the weak autosomal UNO-EGFP dots in normal spermatocytes [53], did not allow a reliable identification of green signals with an unequivocal association with autosomal chromatin. Importantly, the highly prominent chromosomal UNO-EGFP dot that was present in Cap-D3 mutant spermatocytes at the start of M I, disappeared rapidly around the metaphase-to-anaphase I transition, with kinetics indistinguishable from that in normal spermatocytes (Fig 5B and 5C). In conclusion, Cap-D3 is not required for the elimination of AHC during M I.

Although AHC is eliminated on time during exit from M I in Cap-D3 mutants, it might still contribute to the meiotic chromosome segregation defects observed in these mutants, as previously suggested [23]. In condensin II protein mutants, conjunction might be established not just between homologous chromosomes but also between non-homologous chromosomes as a consequence of the failure of chromosome territory formation. Such ectopic AHC between non-homologous chromosomes might preclude timely disentangling of bivalents during spermatocyte maturation; the temporal window for disentangling remaining after AHC elimination in M I might be too short for a proper anaphase I without chromosome bridges. Addressing potential ectopic AHC by direct localization of AHC proteins is problematic, as barely detectable levels of AHC proteins are sufficient for stable conjunction of autosomal homologs during normal meiosis [52,53]. However, this issue has previously been addressed by phenotypic analysis of Cap-H2 teflon double mutants [23]. In contrast to loss of teflon, mutations in mnm results in a complete loss of AHC [52]. Therefore, we studied Cap-D3 mnm double mutants (Cap-D3^EY00456/ Df(2L)Exel7023; mnm^z3-5578/ mnm^z3-3298) by time-lapse imaging to resolve whether a complete AHC loss might result in an equally complete suppression of anaphase bridging (Fig 5D). Compared to Cap-D3 single mutants, the percentage of spermatocytes with chromatin bridges during anaphase I was reduced in the Cap-D3 mnm double mutants, from 44% to 8% (Fig 4G). The fraction of cells with anaphase bridges remaining in the Cap-D3 mnm double mutants was still higher than in mnm single mutants, where anaphase bridge frequency is around 2% [72]. The incomplete suppression of bridges in Cap-D3 mnm double mutants indicates that some type of linkages other than those mediated directly by AHC contribute to the failure of chromosome segregation in condensin II protein mutants.

Interestingly, while anaphase bridging was substantially suppressed in Cap-D3 mnm double mutants compared to Cap-D3 single mutants, centromere mis-orientation during M I was not. The fraction of cells with 4:4 centromere segregation, i.e., the upper bound for correct segregation, was comparably low in both Cap-D3 mnm double and Cap-D3 single mutants (Fig 4H). In fact, time-lapse imaging revealed very similar abnormalities at the start of M I in the double mutants as in the single mutants (Fig 5D). NEBD I was accompanied by formation of a single chromatin clump with clustered centromeres in both Cap-D3 single and Cap-D3 mnm double mutants (Fig 5D). In contrast, in mnm single mutants (mnm^z3-5578/ mnm^z3-3298), bivalents were separated prematurely into univalents, as previously reported [52,55]. Moreover, our time-lapse analysis of centromere behavior during anaphase I in mnm mutants revealed a distribution of spermatocytes (n = 44) with either 4:4 (25.0%), 5:3 (43.2%), 6:2 (25.0%), 7:1 (6.8%) and 8:0 (0%) segregation, in excellent agreement with the mathematical prediction for the case of random segregation.

In summary, the importance of chromosome territory formation for regular chromosome segregation during M I goes beyond an avoidance of ectopic AHC between non-homologous chromosomes. Even in complete absence of AHC, non-homologous chromosome associations remain so prominent in Cap-D3 mutants that a single clump of chromatin with clustered centromeres unfit for bi-orientation is generated at the start of M I.

A TALE light system for analysis of chromocenter disruption dynamics during chromosome territory formation

While centromere clusters were disrupted in condensin II mutant spermatocytes just before NEBD I, the individualized centromeres nevertheless remained in close vicinity, apparently anchored by an intact chromocenter that also bundled the distinct emanating arm regions together (Fig 5D). The failure of chromocenter disruption in the mutants appeared to be largely responsible for the missegregation of chromosomes during M I. For specific analyses of chromocenter disruption, we adapted TALE lights for use in spermatocytes. TALE lights [73,74] are fusions of a fluorescent protein with a TALE domain, an engineered sequence-specific DNA binding domain. The DNA target sequences of the chosen TALE lights corresponded to those of two distinct satellites. A first red-fluorescent TALE light targeted the 359-bp satellite in the centromere-proximal heterochromatin of chrX (Fig 6A). A second green-fluorescent TALE light targeted the two 1.686 satellite blocks, which bind Prod [71] and are present in pericentromeric heterochromatin of chr2 and chr3 (Fig 6A). For use in spermatocytes, we generated Drosophila lines with UASt transgenes, designated as UASt-RedX and UASt-Green2/3, respectively. For validation, these transgenes were expressed in testes using bamP-GAL4-VP16. Immuno-FISH with antibodies recognizing the TALE lights and FISH probes for the targeted satellite loci confirmed that the TALE lights localized to their target sites (S7 Fig). Moreover, direct observation of TALE lights fluorescence revealed intranuclear dot signals in spermatocytes that were clearly above background (S7 Fig). The dot signals generated by bam>RedX were relatively weak and no longer detectable after the S4 stage. When detectable, there was a single RedX dot per nucleus, which was localized within the XY4 territory (S7 Fig). The dot signals resulting with bam>Green2/3 were stronger and still detectable in early postmeiotic spermatids. The number of Green2/3 dots per nucleus was variable. One or two dots were present in early spermatocytes, and usually four dots in late spermatocytes, i.e., one dot pair per major autosome territory (S7 Fig). These dot numbers observed in spermatocytes were in agreement with previous FISH analyses [11]. In addition to the dot signals, bam>RedX and bam>Green2/3 spermatocytes also displayed weaker diffuse nuclear signals.

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Fig 6. Time-lapse imaging of chromocenter disruption in spermatocytes.

(A) The fluorescent TALE light proteins RedX and Green2/3 were used for analysis of chromocenter disruption by time-lapse imaging. The karyogram indicates the location of the 359 bp satellite locus (red) and the 1.686 satellite loci (green) that recruit RedX and Green2/3, respectively. Heterochromatic regions represented by grey shading and centromeres by circles. (B) Spermatocytes expressing RedX and Green2/3 (bam>RedX Green2/3) were analyzed by live imaging. The diameter (d) of the cell nucleus (dashed circle), as revealed by the weak diffuse red and green signals, was used as a proxy for developmental stage [9,10]. While a single RedX dot was present invariably during the stages S2 and S3, the number of Green2/3 dots was variable. Bar diagram for comparison of the number of Green2/3 dots in spermatocytes before (d < 9 μm) and after (d = 9–12 μm) chromosome territory formation. n = 270 (d < 9 μm) and 20 (d = 9–12 μm). (C) Tracking of RedX and Green2/3 dots over time. Still frames (top) illustrate the splitting of a Green2/3 dot into two widely separated dots, indicating chromocenter disruption. Time (h:min) indicated relative to the start of imaging. The distances separating RedX and Green2/3 dots in this spermatocyte were plotted over time (bottom). (D) Radial positions of RedX and Green2/3 dots during chromocenter disruption. Radial positions over time (left) after dot tracking in a representative spermatocyte. Bar diagram (middle) displays radial positions in five cells with a single Green2/3 dot (cells 1–5) and in five cells with two Green2/3 dots (cells 6–10). Bar diagram (right) provides the mean (± s.d.) after averaging the radial dot positions observed in cells 1–5 and 6–10, respectively. ** indicates p < .01 (t test). Scale bars = 2 μm.

https://doi.org/10.1371/journal.pgen.1009870.g006

For time-lapse imaging, UASt-RedX and UASt-Green2/3 were co-expressed in spermatocytes with bamP-GAL4-VP16. Staging of spermatocytes was based on the nuclear diameter (d) revealed by the diffuse nuclear signals. Early spermatocytes with d < 9 μm, i.e., during S1 and S2 [9,10], displayed a single RedX dot (Fig 6B), as in the analyses with fixed samples. The number of Green2/3 dots was variable in early spermatocytes (Fig 6B), also as in fixed cells. In early spermatocytes (d < 9 μm), the number of Green2/3 dots per nucleus was one, two or three in 31%, 67% and 1% of the cells, respectively (Fig 6B). Therefore, in almost a third of the early spermatocytes, the 1.686 satellite blocks of chr2 and chr3 were in intimate association, as indicated by only a single Green2/3 dot per nucleus. This non-homologous association of pericentromeric heterochromatin regions of chr2 and chr3 reflects chromocenter formation. However, the 359-bp satellite block on chrX was not recruited into the region occupied by the 1.686 satellite loci of chr2 and chr3. Substantial overlap of Green2/3 and RedX dot signals was never observed. Nucleolus formation might separate the 359-bp satellite away from the main chromocenter because the 359-bp satellite is close to the rDNA locus of chrX.

Importantly, at later stages, in S3/S4 spermatocytes (d = 9–12 μm), non-homologous associations between the 1.686 satellite blocks of chr2 and chr3 were no longer apparent. None of the analyzed spermatocytes nuclei contained a single Green2/3 dot (Fig 6B). The majority (60%) displayed four Green2/3 dots, indicating that not just non-homologous associations were disrupted at this stage but also tight homolog pairing. These observations are in agreement with the conclusions of Vazquez et al. [10] that homologs are separated during stage S3 immediately after chromosome territory formation during stage S2b.

To analyze the temporal dynamics of chromocenter disruption, we tracked the RedX and Green2/3 dot signals in early spermatocytes (d = 7.1–8.3 μm). Spermatocytes with a single Green2/3 dot that was split into two during the imaging period were analyzed in detail (n = 20 from eight distinct cysts). Distances between the dots were determined, as well as their separation from the nuclear periphery (Fig 6C). After splitting of the Green2/3 dot, the distance between the two progeny dots increased up to 7.8 μm within one to two hours, with a dot speed around 0.03 μm/s. The single RedX dot was always relatively close to the nuclear periphery at the start of the chromocenter disruption phase and it remained peripheral over time (Fig 6D). In contrast, the single Green2/3 dot had a central position and the two progeny dots moved towards the nuclear periphery after splitting (Fig 6D).

In conclusion, fluorescent dot tracking in bam>RedX and Green2/3 spermatocytes revealed the dynamics of chromocenter disruption.

Stretching of 1.686 chromatin instead of efficient disruption of non-homologous 1.686 associations in condensin II protein mutants

For characterization of the consequences of the loss of condensin II proteins for chromocenter dynamics in spermatocytes, we analyzed Cap-D3 and Cap-H2 mutant spermatocytes with bam>RedX Green2/3. Beyond the presumed null alleles Cap-D3^EY00456 and Cap-H2^cc3, the hypomorphic mutation Cap-H2^cc1 was analyzed as well. As in controls, the earliest spermatocytes (d < 9 μm, S1-S2) invariably displayed a single RedX and one or two Green2/3 dot signals in all three hemizygous mutant genotypes (Fig 7A). However, early spermatocytes with only one Green2/3 dot were more than twofold more frequent in the mutants (Fig 7A). This increase was less pronounced with the hypomorphic Cap-H2 allele compared to the strong Cap-H2 and Cap-D3 alleles (Fig 7A). In the early Cap-H2^cc3 mutant spermatocytes with a single RedX and a single Green2/3 dot, their separation was significantly reduced compared to controls (Fig 7B) and the single Green2/3 dot was more peripheral (Fig 7C).

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Fig 7. Prolonged stretching of 1.686 satellite chromatin in the absence of condensin II proteins.

Spermatocytes with bam>RedX Green2/3 in different genetic backgrounds were analyzed by live imaging. (A) The diameter (d) of the cell nucleus was used as a proxy for developmental stage. Bars indicate the number of Green2/3 dots in spermatocytes before (d < 9 μm) and after (d = 9–12 μm) chromosome territory formation in the indicated genotypes. Control data (+/+) is identical to that in Fig 6B. n = 270, 20, 119, 72, 155, 71, 212, 58 (from left to right). (B) Significantly shorter average separation between RedX and Green2/3 dots in Cap-H2^cc3 mutant spermatocytes compared to controls. Spermatocytes with one RedX and one Green2/3 dot at the S1/2 stage were analyzed. Mean ± s.d. is shown; n = 18 (+/+) and 22 (Cap-H2^cc3/Df). (C) Average radial positions of RedX and Green2/3 dots (± s.d.; n = 6) in Cap-H2^cc3 mutant spermatocytes are not significantly different, in contrast to controls (Fig 6D). (D-F) Instead of the normal rapid splitting of single Green2/3 dots into two, condensin II protein mutants display episodes with prolonged Green2/3 dot stretching and occasional splitting. Time (h:min) is given relative to the start of displayed image sequence. Graphs display the extent of stretching over time. Spot detection (using IMARIS) resulted in the identification of two green spots after sufficient stretching even if a connection was still detectable in between. A distance value of zero indicates that only a single Green2/3 spot was detected. The plots start just before the stretching/splitting of the Green2/3 dot. Scale bars = 3 μm. (G) Percentage of S1-S4 spermatocytes that displayed at least one episode of Green2/3 dot stretching (as in panel E) or Green2/3 dot splitting (as in panel F) within the first 2.5 hours of imaging. n = 126 from 14 cysts (Cap-H2^cc1/Df) and 45 from 4 cysts (Cap-H2^cc3/Df).

https://doi.org/10.1371/journal.pgen.1009870.g007

During spermatocyte maturation, differences between controls and condensin II protein mutants increased dramatically. In case of the strong alleles, the fraction of spermatocytes with non-homologous associations of chr2 and 3 (i.e., only a single Green2/3 dot) was largely maintained at around 80% (Fig 7A). In contrast, non-homologous associations were completely absent in controls at the S3/S4 stage (d = 9–12 μm) and even homolog pairing was already disrupted in the majority of cells (Fig 7A). In the hypomorphic Cap-H2^cc1 hemizygotes, an increase of cells with two or even three Green2/3 dots at the expense of those with only one was evident when comparing S1/S2 with S3/S4 spermatocytes (Fig 7A). Thus, non-homologous associations were eliminated in Cap-H2^cc1 mutants, although to a far lower extent than in controls.

Although chromocenter disruption did not succeed without condensin II proteins, the TALE lights time-lapse imaging exposed that some chromatin-separating forces were still at work in the mutants. In mutant spermatocytes with a single Green2/3 dot, a striking dynamic stretching of this signal occurred during the stages of chromosome territory formation and the subsequent stages S3 and S4 (Fig 7D–7G). Although transient stretching of the single Green2/3 dot was also observed occasionally in the controls just before definitive dot splitting, the stretching episodes lasted much longer in the mutants. To characterize the stretching episodes in the mutants, we applied software-based spot detection after time-lapse imaging (n = 10 spermatocytes for Cap-D3^EY/Df, 23 for Cap-H2^cc3/Df, and 21 for Cap-H2^cc1/Df). With the chosen parameters, two spots were identified in the green channel not only after complete splitting of the Green2/3 dot but also after sufficient stretching of this dot. Once two spots were identified during a stretching episode, they were usually separated to about 2 μm (Fig 7D–7F). Occasionally separation reached up to 4.8 μm. Moreover, in a minority of the mutant spermatocytes, even a definitive splitting of the single Green2/3 dot into two could be observed. In about half of the analyzed Cap-H2^cc3/Df spermatocytes during S2b-S4, we observed at least one Green2/3 dot stretching episode (Fig 7G). Definitive splitting was detected in only 2.2% of the Cap-H2^cc3/Df spermatocytes (Fig 7G). Stretching episodes and definitive dot splitting were no longer detectable during S5 and S6. Compared to Cap-H2^cc3, Green2/3 dot stretching, as well as its definitive splitting, were more frequent in the weaker Cap-H2^cc1 allele (Fig 7G).

In conclusion, instead of the efficient disruption of non-homologous associations between the 1.686 satellite loci on chr2 and chr3 that accompanies normal territory formation, prolonged stretching of 1.686 chromatin rarely succeeding in definitive disruption of the non-homologous associations occurred in condensin II protein mutants. Thus, force generators other than condensin II appear to be at work during chromosome territory formation. However, in the absence of condensin II, they fail to achieve bivalent individualization. As documented (S8 Fig), we have addressed whether these unidentified force generators depend on cytoskeletal dynamics using inhibitors of F actin and microtubules. The inhibitors failed to affect both territory formation in control and Green2/3 dot stretching in condensin II protein mutants.

The control of Cap-H2 expression levels in spermatocytes is crucial for bivalent formation

Analyses with Drosophila ovaries, larval salivary glands and cultured cells have demonstrated that Cap-H2 levels determine the extent of somatic homolog pairing, chromocenter formation and centromere clustering [17,22,24,25,49]. In testis, the isoform F of Cap-H2 is most abundant according to RNA-Seq data [75]. Therefore, for an analysis of overexpression effects, we generated a line with an UASt transgene encoding this isoform with EGFP at the N terminus. UASt-EGFP-Cap-H2 expression driven by bamP-GAL4-VP16 was readily detected in early spermatocytes (Fig 8A). However, the EGFP-Cap-H2 signals were clearly above background only until about the S3 stage. In contrast, other EGFP fusion proteins, including those of the AHC proteins MNM, SNM and UNO [53,76], remain detectable throughout spermatocyte maturation when expressed analogously (i.e., using bam> UASt). Thus, EGFP-Cap-H2 appears to be a relatively unstable protein.

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Fig 8. Increased Cap-H2 levels inhibit the pairing of autosomal homologs.

(A-D) UASt-EGFP-Cap-H2 (encoding isoform F) and UASt-EGFP-Cap-H2^D (encoding isoform D) were expressed in spermatocytes using bamP-GAL4-VP16. (A) EGFP signals and DNA staining in testis tip regions (left) and high magnification views with S3 spermatocytes (right). EGFP is present only transiently in early spermatocytes on chromatin (EGFP-Cap-H2) and cytoplasm (EGFP-Cap-H^D). (B) Expression of UASt-EGFP-Cap-H2 results in too many chromosome territories (arrowheads) and meiotic chromosome missegregation in contrast to UASt-EGFP-Cap-H2^D. Spermatocytes at the stages S3 and S5 are displayed, as well as early round spermatids (sp), after labeling testis squash preparations with anti-Lamin Dm0 and a DNA stain. (C) Fertility of males with the indicated genotypes. Four test crosses, each with a single male parent, were set up and the resulting number of F1 progeny was counted. Mean fertility of bam-GAL4-VP16 males without a UAS transgene (bam>) was set to 100%. *** indicates p < .001 (t test) and whiskers s.d. (D) Progression through MI was analyzed by time-lapse imaging of spermatocytes with bam>EGFP-Cap-H2, His2Av-mRFP and Cenp-A/Cid-EGFP. Time (min:sec) is indicated relative to the onset of NEBD I. Arrowheads indicate the centromeres of chr4 (green), chrX (dark red) and chrY (light red). After stable bi-orientation of the sex chromosome bivalent (t = 15:45), chrX and chrY are separated apart regularly during anaphase I (last three still frames). In contrast, autosomes are present as univalents at the start of M I, which segregate randomly after failure of congression into a metaphase plate (as indicated for example by chr4). (E) Live imaging of spermatocytes with bam>EGFP-Cap-H2, RedX and Green2/3. Four Green2/3 dots (arrowheads) are present already in early S1/2 spermatocytes. Scale bars = 40 μm (A, left), 10 μm (A, right), 5 μm (B, D), 3 μm (E).

https://doi.org/10.1371/journal.pgen.1009870.g008

In early spermatocytes, EGFP-Cap-H2 was enriched on chromatin. Interestingly, the DNA staining pattern was abnormal in these spermatocytes. Already during the S1 stage, the DNA appeared to be more compact. At the S3 stage, more than the normal number of three territories were present (Fig 8B). The increased territory number was even more apparent at later stages, after nuclear growth and wider separation of the NE-associated territories (Fig 8B). Post-meiotic spermatids were also abnormal in bam> EGFP-Cap-H2 testes. The size of spermatid nuclei was highly variable (Fig 8B), while normal testes display far less variation [53,76]. Finally, the fertility of bam> EGFP-Cap-H2 males was reduced to about 40% of controls (Fig 8C).

For further characterization of the effects of EGFP-Cap-H2 overexpression on territories and meiotic chromosome segregation, we applied time-lapse imaging. First, we analyzed bam> EGFP-Cap-H2 spermatocytes expressing also His2Av-mRFP and Cenp-A/Cid-EGFP. After NEBD I, instead of four normal bivalents, seven chromosomal entities were evident in bam> EGFP-Cap-H2 spermatocytes (Fig 8D). Six of these had only one associated Cid-EGFP dot. These six displayed independent saltatory movements and failed to congress into the metaphase plate, indicating that they were univalents. The seventh chromosome with two associated Cid-EGFP dots displayed the asymmetries characteristic of the chrXY bivalent [55]. This apparently normal sex chromosome bivalent was regularly bi-oriented and segregated during anaphase I (Fig 8D). The same phenotype was observed consistently in all of the analyzed bam> EGFP-Cap-H2 spermatocytes (n = 28, from four cysts).

To characterize the effects of bam> EGFP-Cap-H2 in early spermatocytes, we analyzed the behavior of RedX and Green2/3 signals by time-lapse imaging (Fig 8E). The relatively weak nuclear EGFP-Cap-H2 signals in these spermatocytes during S1/2 did not preclude detection and tracking of the Green2/3 dots. In all the analyzed spermatocytes with bam> EGFP-Cap-H2, RedX, Green2/3, four completely resolved and independent Green2/3 dots were present already in S1/2 spermatocytes (n = 160) (Fig 8E). Tracking of these Green2/3 dots over time did not reveal permanent splitting, although some transient breathing of presumably sister chromatids was readily detectable. We conclude that increased levels of Cap-H2 appear to block the pairing of autosomal homologs, rather than inducing a premature separation of bivalents.

In conclusion, the effects of Cap-H2 overexpression emphasize the importance of control over Cap-H2 levels in early spermatocytes. Not only loss of Cap-H2 but also its excess is clearly detrimental for the success of male meiosis.

The importance of precise control of Cap-H2 levels in spermatocytes made the annotated D isoform interesting (Fig 3B). The D isoform of Cap-H2 is predicted to lack the N-terminal region required for binding to SMC2, whereas the conserved winged helix domain required for binding to SMC4 is present in the C-terminal region. While incapable of connecting the two core SMC subunits, the D isoform might act in an inhibitory manner by titrating Hawks away from condensin II and by blocking the binding of SMC4 to canonical Cap-H2 isoforms (if isoform D is still capable of binding to SMC4). The D isoform is also distinct from the canonical isoforms E–G in lacking a C terminal Slmb/β-TRCP binding site (Fig 3B), which keeps protein levels of canonical isoforms low [48]. Stabilization of the D isoform might boost its dominant-negative effect on condensin II activity. Moreover, this D isoform might be testis-specific according to RNA-Seq and EST data. Overall, it appeared conceivable, that the D isoform might down-regulate condensin II activity in spermatocytes. To evaluate the role of the D isoform, we generated lines with UASt-EGFP-Cap-H2^D integrated into the same chromosomal landing site as UASt-EGFP-Cap-H2. In bam> EGFP-Cap-H2^D spermatocyte, EGFP signals accumulated during the initial S1/2 stages predominantly in the cytoplasm with some enrichment on the fusome (Fig 8A). However, in contrast to expectations, EGFP-CapH2^D was similarly unstable as EGFP-CapH2. EGFP signals were absent from late spermatocytes. Moreover, no obvious abnormalities with regard to chromosome territories were detected in bam> EGFP-Cap-H2^D spermatocytes (Fig 8B) and male fertility was normal (Fig 8C). In conclusion, these results do not support the speculation that the D isoform is a negative regulator of condensin II activity in spermatocytes. The function of the D isoform remains to be clarified.

Drosophila lines

The lines with the following mutations or transgenes have been described before: Df(3R)Exel6159 (Bloomington Drosophila stock Center (BDSC) # 7638), Df(2L)Exel7023 (BDSC # 7797), Cap-D3^EY00456 (BDSC # 7797), mnm^Z3-3298 and mnm^Z3-5578 [52], cav^EGFP (EGFP-HOAP) [61], g-uno-EGFP III.1 [53], His2Av-mRFP and gCid-EGFP-Cid [87], UbiP{GFP(S65T)-βTub56D}17–1 (DGRC Kyoto # 109603), P{y[+t7.7] w[+mC] = D1-GFP.FPTB}attP40 (BDSC # 66454), bamP-GAL4-VP16 [88].

Lines with the following transgenes were generated by microinjection (BestGene Inc., Chino Hills, CA, USA) with the plasmid constructs described further below: UASt-EGFP-Cap-H2, UASt-EGFP-Cap-H2^D, UASt-RedX, and UASt-Green2/3. The first two pUASt-attB constructs were integrated into the landing site P{CaryP}attP40. The UASt-RedX transgene in the established Drosophila lines was observed to be unstable. Initial analyses of testis from bam> RedX males clearly resulted in a single nuclear red dot in early spermatocytes. Subsequent analogous analyses after standard maintenance of the UASt-RedX lines for about 3 years in the laboratory did not reveal these red dots as previously in the initial experiments. Depending on the subline, dots were either substantially weaker or no longer detectable above the diffuse red nuclear signal that was still present. The UASt-RedX coding sequence encompasses 20 TALE repeats of 102 bp, which are identical in sequence except for 2–6 central base pairs in some repeats. Therefore, recombination between repeats might occur occasionally and result in altered protein products with weakened or abolished DNA binding activity. Such instabilities were not encountered in case of UASt-Green2/3 even though it contains the same number of TALE repeats.

The Cap-H2 alleles (Cap-H2^cc1, Cap-H2^cc2, and Cap-H2^cc3) were generated using CRISPR/Cas9. Embryos from y¹ M{w[+mC] = Act5C-Cas9.P}ZH-2A w* (Bloomington Drosophila stock Center #54590) were injected (BestGene Inc., Chino Hills, CA, USA) with pCFD5 derived plasmids described further below. Each of these pCFD5 derivatives allowed expression of a gRNA pair targeting two distinct gene regions. Adults obtained from the injected eggs were crossed singly with w*;; Sb/TM3, Ser. A fraction of the F1 progeny was used for preparation of genomic DNA and analysis by PCR using the primers LV048 and LV049 (see S1 Table for sequences of all synthetic DNA oligonucleotides). These primers annealed 337 bp upstream and 490 bp downstream of the two gRNA targets sites, respectively. If the PCR analysis indicated the presence of CRISPR/Cas9-induced Cap-H2 deletion alleles among the tested progeny of a particular founder animal, additional F1 progeny males derived from the same founder were crossed singly with w*;; Sb/TM3, Ser to establish balanced lines. These lines were tested again by PCR with LV048 and LV049 for the presence of Cap-H2 deletion alleles. Moreover, the amplified fragments were sequenced. The sequences around the intragenic deletion breakpoints (-II-) were: 5’-AGCGAAGTCGAG-II-CCCACATTTGAC-3’) (Cap-H2^cc1), 5’-CGACAAGCGCTTCAACGC-II-TCGACCCACATTTG-3’ (Cap-H2^cc2) and 5’-ATGTCGGACGACAAGCGCTTCAACG-II-GGCTTTGTAAAGT-3’ (Cap-H2^cc3). In case of Cap-H2^cc3, a second intragenic deletion, which is identical to that in Cap-H2^cc1, was detected further downstream. The chromosomes carrying Cap-H2^cc1 and Cap-H2^cc3 are homozygous lethal but hemizygous viable, suggesting the presence of second-site recessive lethal mutations on these chromosomes. The chromosome carrying Cap-H2^cc2 is homozygous viable but male sterile.

X0 males were generated by crossing virgin females isolated from the stock C(1;Y)2, y B/0 & C(1)RM, y v/0) (BDSC #2487) with X/Y males (w¹¹¹⁸).

Standard crossing was used for the generation of the various strains used for experimental analyses. The genotypes of the flies analyzed are described in detail in the supporting information (S2 Table). All flies analyzed were raised at 25°C.

Plasmids

For generation pCFD5-Cap-H2_gRNAs1-2, a synthetic double stranded DNA fragment (LV046) (Integrated DNA Technologies, Coralville, IA, USA) was inserted into of pCFD5 [89] after digestion of vector and insert with BbsI.

For generation of pUASt-RedX, a modified version of pUASt was generated first. After digestion of pUASt with EcoRI and XbaI, a linker generated by annealing oligonucleotides CL305 and CL306 was inserted. This linker insertion resulted in the elimination of the EcoRI and XbaI sites and in the introduction of a NheI and an XhoI site. pET28-359_2-mCherry [73] was digested with NheI and XhoI to release the region coding for 359 TALE Light-mCherry, which was inserted into the corresponding newly introduced restriction sites of the modified pUASt vector.

For the generation of pUASt-Green2/3, we modified pUASt-mcs-EGFP [90] in a first step by digestion with EcoRI and XhoI, followed by insertion of a linker generated by annealing the oligonucleotides CL318 and CL319. This linker insertion resulted in the introduction of a NheI and an XhoI site. pET28_CM3_EGFP [73] was digested with NheI and SalI to release the 1.686-TALE coding region, which was inserted into the newly introduced restriction sites of the modified pUASt-mcs-EGFP vector.

For the generation of our UASt-Cap-H2 transgene constructs, we modified pUASp1-EGFP-Cap-H2 [79] (kindly provided by S. Heidmann, Unversity of Bayreuth). This starting plasmid had been constructed with a cDNA isolated from an EST clone (SD18322), which contained the coding sequence for the isoform G with a few genetic polymorphisms. This coding region was mutated into a version that codes for the isoform F. Mutagenesis was performed as described [91] with the primer CL316. The region coding for EGFP-Cap-H2 (isoform F) was then isolated after digestion with NotI and XhoI and transferred into the corresponding sites of a pUASt-attB variant. This variant was obtained by digestion of pUASTattB [92] with EcoRI and BglII, followed by insertion of a linker generated by annealing oligonucleotides CL312 and CL313. With a final step, the Cap-H2 3’UTR sequences were deleted by digestion with EcoRI and XhoI, followed by insertion of a linker obtained by annealing the oligonucleotides CL314 and CL315. The final construct pUASt-attB-EGFP-Cap-H2 (isoform F) contains the SV40 3’UTR sequences, as present in pUASTattB, downstream of the EGFP-Cap-H2 coding sequence. For generation of pUASt-attB-EGFP-Cap-H2 (isoform D), we first deleted the sequences coding for the isoform F-specific N-terminal region from pUASt-EGFP-Cap-H2 (isoform F) as described [91] using the primer CL317. A NotI-AatII fragment was isolated from the mutagenized plasmid and used for replacement of the NotI-AatII fragment in pUASt-attB-EGFP-Cap-H2 (isoform F) resulting in the last intermediate plasmid. In a final step, we converted the C-terminal coding sequences to those present in isoform D. This was achieved with a synthetic DNA fragment (CL320) (GenScript Biotech, Leiden, Netherlands) of about 550 bp flanked by AatII and EcoRI restriction sites. This synthetic DNA fragment was used for replacement of the AatII-EcoRI fragment in the last intermediate, yielding the final construct pUASt-attB-EGFP-Cap-H2 (isoform D). The presence of the correct insert sequences in all of our constructs was verified by DNA sequencing.

Fertility tests

For analysis of male fertility, we crossed single males with three w virgin females. Five to ten replicate crosses were started. For analysis of female fertility, three virgin females of the genotype to be tested were pooled and crossed with three w males. Ten replicate crosses were set up. After two days of mating, crosses were transferred into a fresh vial. After an additional two days, all adult flies were discarded, followed by counting all of the adult progeny that developed subsequently at 25°C.

Fixation and labeling of testis preparations

For whole mount testis preparations, dissection was performed in testis buffer (183 mM KCl, 47 mM NaCl, 10 mM Tris-HCl, pH 6.8). Testes were fixed in phosphate buffered saline (PBS) containing 4% formaldehyde and 0.1% Triton X-100 in 0.2 ml Eppendorf tubes for 20 minutes on a rotating wheel. For immunolabeling, mouse monoclonal anti-Lamin Dm0 antibody ADL67.10 (Developmental Studies Hybridoma Bank, University of Iowa, Iowa City, IA, USA) was used at 1:50 and the rabbit antibody against Prod [71] at 1:5000. Secondary antibodies were Alexa568- or Alexa488-conjugated goat antibodies against mouse IgG (Invitrogen, A11004 and A10631) or Alexa568-conjugated goat antibodies against rabbit IgG (Invitrogen, A11011) diluted 1:500. For DNA staining, testes were incubated for 10 minutes in PBS, 0.1% Triton X-100 (PBTx) containing Hoechst 33258 (1 μg/ml). After three washes with PBS, testes were transferred into a drop of mounting medium (70% glycerol, 1% n-propyl gallate, 0.05% p-phenylenediamine, 50 mM Tris-HCl pH 8.5) on a slide before adding a cover slip.

Testis squash preparations were made and stained essentially as described previously [93], according to protocol 3.3.2, except that the mounting medium described above was used. For immunolabeling, mouse monoclonal anti-Lamin Dm0 antibody ADL67.10 (Developmental Studies Hybridoma Bank) was used also at 1:50 and rabbit anti-Cid IS1 [94] was used 1:1000. Secondary antibodies were used as described above.

For immuno-FISH, testes were dissected and fixed with 4% formaldehyde in PBS, followed by permeabilization with PBS containing 0.3% Triton X-100 and 0.3% sodium deoxycholate. Immunolabeling was done as described above with the following antibodies: rabbit anti-EGFP (IS28) [90] diluted 1:3000 or rabbit anti-mCherry (IS751) diluted 1:3000. Alexa488-conjugated goat anti-rabbit IgG (Invitrogen, A11008) diluted 1:1000 was used as secondary antibody. Ethanol incubations and dehydration with a formamide series were done as described (immuno-FISH protocol 3.2, steps 10–26) [95]. An oligonucleotide (5’-TTTTCCAAATTTCGGTCATCAAATAATCAT-3’) with Atto-565 on both the 5’- and the 3’ end (Integrated DNA Technologies, Leuven, Belgium) was used for detection of the X-specific 359 bp satellite sequences at a concentration of 1 ng/μl in hybridization buffer. An oligonucleotide (5’- ATAACATAGAATAACATAGAATAACATAGA -3’) with Atto-565 on both the 5’- and the 3’ end (Integrated DNA Technologies, Leuven, Belgium) was used for detection of the 1.686 satellite sequences in pericentric heterochromatin of chr2 and chr3 at a concentration of 1 ng/μl in hybridization buffer. The denaturation step was performed at 98°C for 6 min, and hybridization over night at 18°C. Slides were washed twice for ten minutes for each wash with 50% formamide, 2x SSCT at 18°C. Thereafter, additional washes of ten minutes were performed at room temperature, first once in 25% formamide, 2x SSCT and then three times in 2x SSCT. DNA was stained with Hoechst 33258 (1 μg/ml) for 10 minutes. Before mounting, slides were washed twice in PBS for 5 minutes.

Generally, about 20 dissected testes were mounted per slide. Images (z stacks) were acquired using a Zeiss Cell Observer HS wide-field microscope using 40×/0.75 or 63×/1.4 objectives, except for the images presented in S2 Fig. These latter images were acquired with an Olympus FluoView 1000 laser-scanning confocal microscope using a 60×/1.42 objective.

Testis preparations for live imaging

Time-lapse imaging of progression through meiosis was performed as recently described [55]. In brief, testes from pupal or young adult males were dissected in Schneider’s Drosophila Medium (Invitrogen, #21720) supplemented with 10% fetal bovine serum (Invitrogen) and 1% penicillin/streptomycin (Invitrogen, #15140). The dissected testes were transferred into 45 μl of medium in a 35 mm glass bottom dish (MatTek Corporation, #P35G-1.5-14-C) and opened with fine tungsten needles to release the cysts. To reduce sample movements, 15 μl of 1% w/v methylcellulose (Sigma, #M0387) was added. A wet filter paper was placed inside along the dish wall before sealing the lid with parafilm. For long-term time-lapse imaging of territory formation over up to six hours, 1.5 ml of medium and 0.5 ml of methylcellulose were used. No wetted filter papers were used in these experiments.

Drugs were administered by pipetting small volumes of stock solution directly to the final testis preparation in the 35 mm dishes. Colcemid (demecolcine, Sigma, #D6165) was dissolved in DMSO (10 mM) before further dilution in tissue culture medium and used at a final concentration of 10 μM. Latrunculin B (Sigma-Aldrich L5288) was also dissolved in DMSO (10 mM) before further dilution in tissue culture medium and used at a final concentration of 4 μM. Cytochalasin D (Sigma-Aldrich C2618) was further diluted in DMSO (1 mM) and used at a final concentration of 10 μM.

Imaging was performed at 25°C in a room with temperature control using a spinning disc confocal microscope (VisiScope with a Yokogawa CSU-X1 unit combined with an Olympus IX83 inverted stand and a Photometrics evolve EM 512 EMCCD camera, equipped for red/green dual channel fluorescence observation; Visitron systems, Puchheim, Germany). A 60×/1.42 oil immersion objective was used for acquisition of z stacks. The z stacks acquired for analysis of progression through M I comprised 46 focal planes spaced by 500 nm and the stacks were acquired at 45 seconds intervals (Figs 2A and 2B, 4, 5, S6, S8 panel A and B). The z stacks acquired for the analysis of chromosome territory formation comprised 29 focal planes spaced by 800 nm. The time intervals between stack acquisitions were 10 minutes (Fig 1A–1D), one minute (Figs 6, 7, S8 panel C—E), 30 seconds (Fig 1E), 10 seconds (Fig 2), or five seconds (Figs 1F and S3).

Image processing and analysis

Maximum intensity projections were generated using ImageJ for wide-field images and IMARIS (Bitplane; versions 8.4.0, 9.2.0, 9.7.2) for confocal images. For measuring the distance of Cid-EGFP dots from the nuclear lamina, an isosurface was created based on the anti-Lamin Dm0 signal using IMARIS for image segmentation. The same software was used for spot detection based on the Cid-EGFP signals, setting the parameter “estimated xy diameter” to 500 nm with background subtraction as described [55]. Distance values were obtained by using the function “distance transformation” of the IMARIS software. The resulting values were exported as a Microsoft Excel file.

For the analysis of chromosome segregation during M I, centromeric Cid-EGFP signals were tracked after spot detection as described above and the algorithm “Autoregressive Motion” of IMARIS software. The same procedure was also applied for the analysis of centromere behavior during chromosome territory formation. Telomeric EGFP-HOAP dots were tracked analogously using an estimated xy diameter of 400 nm. The software identified the majority of the centromeric Cid-EGFP and telomeric EGFP-HOAP dots correctly with the chosen parameter settings. Tracking of the spots over time resulted in tracks that contained at most a few incorrect linkages without significant effects on the results, if the image stacks sequences had been acquired at maximal spatial and temporal resolution (Figs 1F, 2D and 2E, and S3). For time-lapse data acquired at a lower frame rate, tracks were corrected as follows. Spots for centromeric or telomeric signals that were not recognized automatically were added manually. Similarly, spots assigned to background signals were deleted. Manual correction of tracks was readily possible because the distinct features of the His2Av-mRFP signals associated with centromeric or telomeric dot signals could also be taken into account, while these were ignored during the automatic detection by the IMARIS software. Centromeric and telomeric signals observed during progression through M I were assigned to specific chromosomes using criteria as previously described [55]. Moreover, NEBD and later transitions during progression through M I were also scored as described [55].

For the quantification of centromeric Cid-EGFP signal intensities as well as UNO-EGFP signals associated with the XY pairing center (Figs 2B and 2E and 5C), IMARIS software was used for image segmentation. The parameters used for the generation of isosurfaces were iteratively modified until signals of interest were completely surrounded by the isosurface. Signal intensities of voxels within a given isosurface were summed and exported as a Microsoft Excel file.

To measure the spatial separation of centromeric or telomeric dot signals from the nuclear periphery (Figs 1E and 1F and 2D), we used an analogous procedure as described above for images acquired with Cid-EGFP and anti-Lamin Dm0 signals (S2 Fig). However, instead of using the anti-Lamin Dm0 signal the His2Av-mRFP signal was used for the generation of an isosurface around the nuclear periphery. The resulting separation distances were exported as a Microsoft Excel file.

To measure the velocity of the movements of tracked centromeric and telomeric dot signals using IMARIS software (Figs 1G, 2E, and S3), we also exploited the His2Av-mRFP signals for the generation of an isosurface around the periphery of the nucleus of interest. After tracking the isosurface over time, its translational drift was corrected with the IMARIS software. The velocity of the residual movements of the tracked centromeric and telomeric dot signals at each time point were then exported as a Microsoft Excel file.

RedX and Green2/3 dots signals were tracked during the stages of chromosome territory formation using the spot detection function of IMARIS. In this case, the parameter “estimated xy diameter” was set to 1 μm. To determine radial positions of these dots, an isosurface was created around the nucleus based on the weak diffuse nucleoplasmic RedX signals. Thereafter, the separation of the dots from the nuclear periphery was determined as described above. The weak diffuse nucleoplasmic RedX signal was also used to determine the radius of the nucleus. The distances of each dot from the nuclear periphery at the first 10 time points of imaging were exported as a Microsoft Excel file, which was used for calculation of the radial positions and their mean. To measure the distance between two dots (Figs 6C and 7E and 7F; d(RG_a), d(RG_b) and d(G_aG_b)), spot positions at each time point of the track were exported as a Microsoft Excel file. Python scripts were used for further analysis and calculation of the separation distances.

Figures display maximum intensity projections unless stated otherwise. The projections were generated with either ImageJ or IMARIS. Export of projections from IMARIS as movies or still frames after live imaging was made with interpolated image display. Moreover, display parameters for the His2Av-mRFP signals were adjusted manually over time to reveal chromosomes clearly throughout the movies, thereby correcting photobleaching and partially also the changes in the extent of chromosome condensation during M I. Graphs were generated with Microsoft Excel or GraphPad Prism. P values were calculated using a two tailed student t test (* = p < 0.05; ** = p < 0.01; *** = p < 0.001). Adobe Photoshop, Adobe Illustrator and Inkscape were used for production of figures.