Before a cell divides, it duplicates all its genetic information, which is stored on chromosomes. Then, each chromosome evenly divides into two new cells so that each cell ends up with identical copies of the genetic information. Because the cellular machinery that evenly divides chromosomes is built to recognize chromosomes that were duplicated exactly once, it is important to maintain this pattern of alternating one round of duplication with one round of division. Cells that instead duplicate their chromosomes more than once can make mistakes during division that are associated with diseases such as cancer.

Chromosomes with extra duplications are present in normal tissues such as the placenta of mammals. They can also occur in human diseases and may even result from chemotherapy treatment. However, we know almost nothing about how cells respond to these problematic chromosomes when dividing.

By studying cells from the Drosophila melanogaster species of fruit fly, Stormo and Fox discovered two distinct ways in which cells respond to extra chromosome duplications. One response occurs in cells that were experimentally engineered to undergo an extra chromosome duplication. These cells delay division so that the chromosome separation machinery can somehow adapt to the challenge of separating more than two chromosome copies at once. The second response occurs in cells that naturally undergo extra chromosome duplications before division. In these cells, Stormo and Fox discovered a new type of chromosome separation, whereby the extra chromosome copies move apart from each other before cell division. In doing so the chromosomes can better interact with the chromosome separation machinery during division.

Stormo and Fox also found that a protein named Mad2 is important in both responses, and gives the cell enough time to respond to extra chromosome copies. Without Mad2, the separation of chromosomes with extra duplications is too hasty, and can lead to severe cell division errors and cause organs to form incorrectly.

Having uncovered two new responses that cells use to adapt to extra chromosomes, it will now be important to find other proteins like Mad2 that are important in these events. Understanding these processes and the proteins involved in more detail could help to prevent diseases that are associated with extra chromosomes.

Regulating mitotic chromosome structure is critical to preventing genomic instability (Gordon et al., 2012; Pfau and Amon, 2012). During mitosis, chromatids associate in sister pairs, which facilitates their bi-orientation and subsequent segregation to opposite spindle poles. A frequently occurring and long-recognized departure from this paired chromosome structure occurs when the genome reduplicates without chromatid separation (hereafter: genome reduplication). Following a single extra S-phase, cells frequently form diplochromosomes: four sister chromatids conjoined at centromeres (White, 1935). A more general term for chromosomes formed by any degree of genome reduplication without chromatid separation is 'polytene' (Painter, 1934; Zhimulev et al., 2004).

While incompletely understood, it is appreciated that multiple layers of physical connections tightly intertwine the multiple sister chromatids of polytene chromosomes. These connections likely include cohesins (Cunningham et al., 2012; Pauli et al., 2010) as well as topological entanglements that can be removed by Condensin II activity (Bauer et al., 2012; Smith et al., 2013; Wallace et al., 2015). Additionally, recurring regions of DNA under-replication occur between chromatids in some polytene cells (Beliaeva et al., 1998; Gall et al., 1971; Hannibal et al., 2014; Nordman et al., 2011; Yarosh and Spradling, 2014) whereas DNA replication is more complete in others (Dej and Spradling, 1999; Fox et al., 2010). In addition to connections between sister chromatids, another layer of chromosome association - pairing between homologs - also occurs in some polytene cells. This pairing results in polyploid/polytene cells that exhibit only the haploid number of distinct chromosomes (Metz, 1916; White, 1954). Given these multiple physical connections between polytene chromatids, mitosis in polytene cells is considered 'ill-advised for mechanical reasons' (Edgar and Orr-Weaver, 2001). Indeed, separation of polytene diplochromosomes at anaphase causes chromosome mis-segregation (Vidwans et al., 2002).

Given the association of polytene chromosomes with mitotic errors, it is not surprising that these structures are often associated with aberrant development and disease. Polytene chromosomes have been observed in cells from spontaneous human abortions (Therman et al., 1978), in muscular dystrophy patients (Schmidt et al., 2011), in a variety of tumors (Biesele and Poyner, 1943; Erenpreisa et al., 2009; Therman et al., 1983) and can also precede tumor formation in mice (Davoli and de Lange, 2012). Polytene chromosomes also occur after treatment with currently used anti-mitotic chemotherapeutics such as those that inhibit Topoisomerase II (Cantero et al., 2006; Sumner, 1998). Disruption of numerous other processes crucial for mitosis, including spindle formation (Goyanes and Schvartzman, 1981; Takanari et al., 1985) sister chromatid cohesion (Wirth et al., 2006) or genome integrity control (Davoli et al., 2010) also cause genome reduplication and polyteny. Thus, polytene chromosomes, a source of mitotic instability, are a conserved and common outcome of ectopic genome reduplication.

To understand how cells adapt the cell cycle machinery to the challenge of segregating the intertwined polytene chromatids found in genome-reduplicated cells, naturally occurring models of this problem can prove useful. Programmed genome reduplication cycles of successive S-phase without M-phase (endocycles, Edgar et al., 2014; Fox and Duronio, 2013 see nomenclature) produce polytene chromosomes in many plant, insect, and mammalian species, including humans (Zhimulev et al., 2004; Zybina et al., 1996). However, many cells with programmed genome reduplication do not subsequently divide, preventing study of how nature has circumvented the issue of segregating polytene chromosomes. In contrast, we previously demonstrated that rectal papilla (hereafter: papillar cells), ion-absorbing structures in the Drosophila hindgut, are built entirely by mitosis of endocycled cells (Fox et al., 2010; Schoenfelder et al., 2014). Surprisingly, we never observed polytene chromosomes in hundreds of papillar metaphases (Fox et al., 2010; Schoenfelder et al., 2014), suggesting papillar cells are programmed to either avoid or eliminate polyteny and its associated mitotic defects. Interestingly, previous studies suggest that polyteny can be at least partially undone without anaphase in both normal and tumorous tissue (Dej and Spradling, 1999; Grell, 1946; Levan and Hauschka, 1953). Thus, in some cases, polyteny may be actively regulated or eliminated.

Taken together, the potential negative impact of genome reduplication on mitotic chromosome structure is clear. However, the responses that enable either developing or tumorous cells to continue dividing after reduplication, despite profound chromosome structure changes, remain unclear. Here, using Drosophila tissue models of both ectopic and naturally occurring genome reduplication, we uncover two distinct cellular responses to reduplicated chromosomes. Both reduplication responses require the conserved spindle assembly checkpoint (SAC) protein Mad2, which inhibits the Anaphase Promoting Complex to both delay anaphase in response to unattached or tensionless kinetochores and to also regulate overall mitotic timing from nuclear envelope breakdown (NEBD) to anaphase onset (London and Biggins, 2014; Musacchio, 2015). In reduplicated cells that retain polytene chromosomes at metaphase, we show Mad2 is involved in a SAC wait-anaphase response. This anaphase delay does not fully prevent the mitotic errors and the resulting aneuploidy associated with mitosis of polytene chromosomes, but it substantially reduces apoptosis, tissue malformation, and organismal death. In contrast to this wait-anaphase response, we also define a second response in reduplicated cells that actively eliminates polyteny before anaphase. In this response, polytene chromosomes undergo a dynamic, spindle-independent process we term Separation Into Recent Sister chromatid pairs (SIRS), which eliminates any trace of polyteny before anaphase. Unlike mitosis with metaphase polytene chromosomes, mitosis with SIRS does not trigger a Mad2-dependent anaphase delay. Yet, we find Mad2 promotes efficient SIRS by allowing sufficient time between nuclear envelope breakdown and anaphase, which allows polytene chromosomes to separate into conventional mitotic sister chromatid pairs. Our results therefore define two distinct responses to reduplicated chromosomes, each of which depends on a distinct Mad2 response.

To understand the mechanisms employed by cells with reduplicated chromosomes, we took advantage of accessible developmental models and in vivo genetic tools in Drosophila. While ectopic genome reduplication was previously established to generate polytene diplochromosomes and subsequent mitotic errors in Drosophila, the effects were examined within the time-frame of the terminal embryonic mitotic cell cycle (Vidwans et al., 2002). Thus, the long-term effects of ectopic genome reduplication on cell viability and tissue development, and key molecular regulation of reduplicated chromosomes has remained unexplored. In parallel, our development of rectal papillae as a non-ectopic, naturally occurring model of mitosis after genome reduplication enabled us to also study how cells programmed to undergo genome reduplication can regulate polytene chromosome structure during mitosis.

Ectopic genome reduplication yields polyteny and continued aneuploid cell division

We first ectopically induced genome reduplication in proliferating tissues of developing larvae by transiently re-programming mitotic cycles to endocycles. fizzy-related (fzr, mammalian Cdh1) plays a conserved role in endocycles by targeting the anaphase promoting complex to destroy the mitotic Cyclins A, B, and B3 (Larson-Rabin et al., 2009; Sigrist and Lehner, 1997). fzr overexpression was previously shown to transform mitotic cycles into endocycles (Sigrist and Lehner, 1997). To transiently induce endocycles, we used a brief heat shock (HS) pulse to express ectopic fzr (HS>fzr, Figure 1A). Using the cell cycle marker system Fly-FUCCI (Zielke et al., 2014; Figure 1B) we find that pulsed fzr overexpression temporarily eliminates expression of the S/G2/M mRFP-CyclinB reporter in wing imaginal disc cells (Figure 1C vs. C’, D). This same population of mRFP-CyclinB-negative cells continues to express the G2/M/G1 GFP-E2F1 reporter, but in greater proportion (Figure 1C vs. C’, D). Together, these data suggest HS>fzr promotes G1 accumulation (91% of cells compared to 36% in controls, Figure 1B–D). To test whether this G1 accumulation is due to direct conversion of G2 cells to G1, as opposed to an acceleration of the cell cycle through G2/M, we stained for the mitotic marker Phospho-Histone H3 at several time points after pulsed fzr expression. For up to 7 hr after fzr overexpression, there is essentially no mitosis in the wing imaginal disc, whereas wing cells in heat shocked wild type flies continue to divide after heat shock (Figure 1—figure supplement 1A,B). Based on previous studies of fzr function and our FUCCI and Phospho-Histone H3 data, we conclude that pulsed fzr expression converts G2 cells to a G1 state by eliminating mitotic cyclins (Figure 1A).

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To determine if the G2 cells re-programmed to G1 proceed through a second genome duplication, we examined mitotic chromosome number when mitosis of HS>fzr tissue first resumes (ten hours after heat shock). At this time-point, we observe frequent tetraploidy (41% of mitotic cells, equivalent to 93% of all G2 cells prior to heat shock, Figure 1E v. F,F’,H). We obtained similar results when examining the results of fzr overexpression in diploid brain progenitors (Figure 1—figure supplement 1C). These results confirm our ability to induce genome reduplication in normally diploid tissues. We also examined chromosome structure in our induced tetraploid cells. When HS>fzr-induced tetraploid DNA first condenses post-heat shock, all chromatids of each chromosome type are closely aligned in a polytene configuration, as evidenced by having the haploid number of distinguishable chromosomes (four in females, Figure 1F). Frequently, we observe un-pairing of the homologous groups of centromeres within each polytene chromosome (Figure 1F, asterisk, also see discussion). Later, at the first metaphase, homologous chromosomes of each polytene are now completely separated, but the four centromeres of each group of sister chromatids remain conjoined within diplochromosomes (observed for 96% of tetraploid cells, Figure 1F’ inset, H). Thus, HS>fzr induces ectopic genome reduplication, resulting in tetraploid cells with metaphase polytene diplochromosomes.

We next examined the mitotic fidelity of cells with diplochromosomes by two independent means: chromosome karyotype analysis and live imaging. By examining the metaphase chromosomes of the division immediately following diplochromosome division, we could detect whether aneuploidy results from diplochromosome segregation. Following the division of cells with diplochromosomes, we observe tetraploid-aneuploid cells with one or two extra or missing chromosomes (8.6% of mitotic cells). In these cells, diplochromosomes are no longer present and instead chromatids are found in distinct sister pairs (Figure 1G). This suggests that during or after anaphase of the first post-reduplication division, diplochromosomes can separate into individual chromatids. Further, these diplochromosome divisions can produce aneuploid daughter cells, many of which continue to divide (Figure 1G,H,K, Figure 1—figure supplement 1C).

We also live imaged mitosis of wing imaginal disc and brain progenitor (neuroblasts and ganglion mother) cells, both with and without ectopic genome reduplication. In addition to using a histone marker to observe chromosomes, we used the Cenp-C-Tomato marker to observe kinetochores. Control diploid cells divide without errors (Figure 1I No HS, J, Video 1). In contrast, most (80%) tetraploid divisions with diplochromosomes exhibit lagging chromosomes, DNA bridges, or both (Figure 1I HS+10 hr, J, Video 2). In our live imaging, diplochromosomes were identifiable as quartets of centromeres and their associated chromosome arms in very close proximity. In some of these divisions we clearly observe four chromatids of a diplochromosome quartet segregating 3:1 (in agreement with prior work in embryos by Vidwans et al., 2002, suggesting incomplete or imprecise sister chromatid disjunction is the cause of chromosome gains and losses (Figure 1I, Figure 1—figure supplement 1D, Video 2). Mitotic errors in the first division of HS>fzr cells appear to result primarily from diplochromosomes and not tetraploidy itself, as tetraploid cells in the subsequent divisions (which lack diplochromosomes) do not exhibit obvious chromosome quartets and have a substantially reduced error rate (Figure 1J 4N, Figure 1—figure supplement 1E, Video 3).

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We previously reported centrosome amplification to contribute to polyploid mitotic errors in Drosophila (Schoenfelder et al., 2014). However, centrosome amplification does not appear to be a major contributor to mitotic errors in the first (or subsequent) polyploid division of HS>fzr animals, as few tetraploid cells amplify centrosomes, and multipolar division is very rare (Figure 1—figure supplement 1F,G, Video 4). In spite of the high initial error rate caused by separation of diplochromosomes, tetraploid-aneuploid cell divisions continue to occur for at least 5 days after genome reduplication, as determined by cytology (Figure 1H HS+120 hr). We conclude that division of diplochromosomes in the mitotically expanding diploid progenitor tissues that we surveyed can lead to the generation of aneuploid cells, which can continue to divide (Figure 1K).

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To determine the long-term effect of tetraploid-aneuploid divisions on tissue development, we took advantage of the fact that expression of HS>fzr occurs in adult progenitor tissues. We thus examined the survival of these animals to adulthood. Survival is only subtly affected in animals with mild (23.0% tetraploid [S.E.M. 4.9%]) levels of induced error-prone tetraploid progenitor division (Figure 2A), and resulting adult tissues appear normal (Figure 2B). In contrast, when tetraploidy is further increased by increasing the duration of heat shock, organism survival decreases in a tetraploid-dependent fashion (Figure 2A, Figure 2—figure supplement 1A). Together, these results show that ectopic genome reduplication in multiple progenitor tissues yields tetraploid cells with polytene metaphase diplochromosomes, which are aneuploid-prone (Figure 1K). These conclusions are in agreement with a previous study in the terminal embryonic division of embryos (Vidwans et al., 2002). We further show that such aneuploid-prone cells can continue to propagate, and that only at high frequencies are these error-prone tetraploid mitotic events lethal to the organism.

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Polyteny response 1: Spindle assembly checkpoint (SAC)-mediated anaphase delay

Our data (Figure 1H, Figure 1—figure supplement 1C) suggest that many polytene diplochromosome divisions in a variety of tissues do not lead to aneuploidy. Little is known about aneuploidy prevention mechanisms in cells with polytene chromosomes, despite the numerous mechanisms that can generate these aberrations. Through live imaging, we uncovered one such aneuploidy-prevention mechanism in cells with metaphase diplochromosomes. Because our heat shock protocol only affects cells in G2, a single HS>fzr pulse creates a mixed population of unaltered diploid cells and diplochromosome-containing tetraploid cells, allowing us to simultaneously live image both cell types in the same tissue. Metaphase in cells with diplochromosomes (Figure 2C, yellow dotted outline) is significantly longer than in diploid cells (Figure 2C blue dashed outline, Video 5, Figure 2D), consistent with previous work on diplochromosomes formed in Drosophila Securin mutants (Pandey et al., 2005). We thus hypothesized that diplochromosomes trigger the SAC, which activates a wait-anaphase signal until all kinetochores attach to microtubules and are under tension (London and Biggins, 2014; Musacchio, 2015). To test this model, we examined SAC-defective mad2 null animals (Buffin et al., 2007; Emre et al., 2011) Figure 2—figure supplement 1B). Using live imaging of wing imaginal discs before and after heat shock, we find that loss of mad2 eliminates the lengthened period of metaphase caused by diplochromosomes (Figure 2D). When the checkpoint is active unattached or misattached kinetochores generate a wait-anaphase signal by localizing SAC proteins such as BubR1 to those kinetochores (Musacchio, 2015). To confirm that diplochromosomes have localized SAC proteins we co-imaged kinetochores and BubR1 in wing disc cells after fzr expression (Royou et al., 2010). We find that in diploid cells BubR1-GFP is clearly evident on kinetochores following nuclear envelope break down and remains there until anaphase. This signal is relatively evenly spread across all the kinetochores (Figure 2—figure supplement 1C, diploid, Video 6). In contrast BubR1-GFP remains localized for much longer in cells with diplochromosomes and is often localized strongly to a specific kinetochore group rather than evenly distributed, suggesting that a subset of diplochromosomes may have trouble forming attachments that satisfy the checkpoint (Figure 2—figure supplement 1C, diplochromosomes, Video 7) From these data, we conclude that diplochromosomes trigger a SAC wait-anaphase response.

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Although important for mitosis in cultured Drosophila S2 cells (Orr et al., 2007), the Mad2-directed SAC is reported to be dispensable in Drosophila tissue mitosis (Buffin et al., 2007; Emre et al., 2011). mad2 null animals are viable with no obvious tissue defects (Figure 2B; Buffin et al., 2007) due in part to an apoptotic response (Morais da Silva et al., 2013). In contrast, the Mad2-dependent wait-anaphase response is essential during development of HS>fzr animals. Even at low levels of tetraploidy, which affect the survival of HS>fzr animals only slightly, few HS>fzr, mad2 animals survive to adulthood (15.4%, Figure 2A, Figure 2—figure supplement 1A). To understand why HS>fzr, mad2 animals have survival defects, we analyzed the surviving animals. In these animals, we find a variety of developmental defects in normally diploid tissues, including smaller eyes, ectopic wing veins, and melanotic abdominal masses (Figure 2B). Increased apoptosis is associated with these tissue malformation phenotypes, as progenitor tissue from HS>fzr, mad2 animals have much higher rates of apoptotic cell death as shown by both TUNEL labeling (Figure 2E,F), and cleaved caspase staining (Figure 2—figure supplement 1D,E). Further, 100% of mad2 diplochromosome divisions exhibit lagging chromosomes, or DNA bridges (Figure 2—figure supplement 1F, Video 8), compared with 80% of divisions in HS>fzr cells. These data may suggest that diplochromosomes are likely susceptible to at least two classes of mitotic errors, one that can be corrected by the SAC, and one that cannot. It is also likely that the mad2 diplochromosome divisions are qualitatively more erroneous than in WT, which may account for differences in survival and tissue phenotype between these genotypes. Taken together, our data identify an important role for the Mad2-dependent SAC in delaying anaphase in the presence of metaphase polytene diplochromosomes.

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Polyteny response 2: Separation Into Recent Sisters (SIRS)

Having defined a response to mitosis after ectopic genome reduplication, we next asked if this same mechanism operates in a tissue that we previously found to divide after programmed genome reduplication. In earlier work we found Drosophila rectal papillar cells (hereafter: papillar cells) naturally undergo two fzr-dependent endocycles during the 2^nd larval instar to generate octoploid cells and then divide, on average, two times during pupal development. An intervening S-phase accompanies these polyploid divisions, and cells at the papillar base undergo one additional S-phase after the final polyploid mitosis (Figure 3A; Fox et al., 2010; Schoenfelder et al., 2014). Thus, as with HS>fzr induction in diploid tissues, papillar development naturally involves genome reduplication followed by mitosis. Our previous work established that papillar mitoses can be error prone, so the same problems with dissociating polytene chromosomes in HS>fzr tissues could be responsible for a portion of these errors during papillar divisions. However, we previously did not see, in hundreds of observed cells, any instances of metaphases with persistent polyteny in papillar cells, suggesting that papillar cells somehow avoid mitosis of polytene chromosomes. Through careful re-examination of the first octoploid metaphase, we confirmed that papillar chromosomes in these octoploid cells are arranged in individual sister chromatid pairs (Figure 3B’’ inset, C). This suggested two possibilities: 1) papillar cells never form polytenes, or 2) papillar cells form polytenes, but somehow separate into recent sister pairs prior to the first metaphase.

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To distinguish these two possibilities, we examined papillar karyotypes from the moment chromosome condensation could be detected. Papillar cells re-enter mitosis from a G2-like state, as evidenced by expression of the G2/M regulator Cdc25/string just before the onset of pupal cell cycles (Fox et al., 2010) (Figure 3—figure supplement 1A). At time points early in the first mitosis, we indeed find that papillar chromosomes are polytene (Figure 3B, Polytene). In these polytenes, we again see examples of cells where the centromeric regions are no longer tightly associated, as we did in our studies of HS>fzr-induced polyteny. However, unlike in cells with induced polyteny, the centromeres in papillar polytene cells are able to not only separate into groups of homologs, but to further separate into individual sister chromatid pairs (asterisks in Figure 3B and Figure 3—figure supplement 1B vs. Figure 1F, also see discussion). In flies heterozygous for inversion-containing balancer chromosomes, papillar polytene structure is locally perturbed, likely due to the disruption of somatic homolog pairing (Figure 3—figure supplement 1B). At this early mitotic time point, we also observe cells where polytenes are absent. Instead, in these cells, sister chromatid pairs and homologs of each chromosome type are separated but remain clumped closely together, as if the polytene chromosome recently separated into pairs containing only the most recent sister chromatids (Figure 3B’, D, Clumped). Neither the polytene nor the clumped configurations remain during the second division (Figure 3C,D), suggesting a specific chromosome structure is present early in the first division of papillar cells. Similarly, by Fluorescent In Situ Hybridization (FISH) we find examples of both closely associated (polytene) and dispersed (separated/non-polytene) signals during the period of the first papillar mitosis (Figure 3—figure supplement 1C,C’). Thus, a key difference between the response to genome reduplication between papillar and HS>fzr cells is the elimination of polyteny before anaphase in papillar cells.

To examine if a majority of (if not all) papillar cells transition from polytene to separate/non-polytene chromosomes during the first mitosis, we used drug treatment to isolate specific chromosome structures during the transition into the first papillar division. To enrich for early mitotic and pre-mitotic chromosomes, we induced Premature Chromosome Compaction (PCC) in papillar cells at a time point just prior to the first mitosis (Methods). PCC causes interphase chromosomes to condense and makes it possible to visualize interphase chromosome structure by standard cytological methods (Figure 3—figure supplement 1B). Using this technique, we find that in pre-mitotic papillar tissue, clear polytene chromosomes are present in nearly every cell (Figure 3D Pre 1^st Div, Figure 3—figure supplement 1B). If we instead enrich for cells in metaphase of the first mitosis by treating with the spindle poison colcemid, we find zero examples where chromosomes are still polytene. In these metaphase-enriched samples, all chromosomes are separated into recent-sister pairs, and even cells with clumped chromosomes are rare (Figure 3D, 1^st Div +colc). Thus, our pharmacological studies further suggest that essentially all genome-reduplicated papillar cells are programmed to completely eliminate polytene chromosomes as cells progress into the first metaphase.

To observe the temporal dynamics of the pre-anaphase elimination of papillar polyteny, we used live imaging, using the same markers used to image diplochromosome division. Prior to the first papillar mitosis, the kinetochores from each homolog are closely associated into an average of 4.1 large foci, close to the haploid number of distinct chromosomes (4 for females and 5 for males due to X/Y un-pairing, Figure 3E). As time progresses in the first division, it is possible to watch these large kinetochore foci disperse into many smaller foci prior to metaphase (Figure 3F, inset, Video 9). In contrast, prior to the second division kinetochores are already separated into many more foci (an average of 15.1 observably distinct foci per cell) before entry into mitosis (Figure 3E). During the second division, the number of resolvable foci remains essentially constant (Figure 3G, Video 10). Additionally, the histone marker reveals that polytene chromosomes are visible when chromosomes first condense and can then be seen to disperse during the first but not the second division (Figure 3F v. 3G, -18:00 min). This result confirms the model that genome-reduplicated papillar cells eliminate polyteny during the first mitosis, then undergo an intervening S-phase before the next division (Figure 3A). We also confirmed that each clump of 4 or 5 pre-first division centromeres only contains a single chromosome type. To do so, we took advantage of the fact that dosage compensation in flies relies on upregulation of transcription on the male X chromosome via the Dosage Compensation Complex (Conrad and Akhtar, 2011). By live imaging papillar cells expressing the DCC complex protein MSL3 tagged with GFP, which localizes only to the male X-chromosome (Strukov et al., 2011), we find that indeed only a single Cenp-C-Tomato focus is MSL3-GFP positive prior to polytene dissociation (Figure 3—figure supplement 1D, Video 11). Taken together, we find papillar cells avoid mitosis of polytene chromosomes in part by undergoing a pre-anaphase chromosome separation process we term Separation Into Recent Sisters (SIRS, Figure 3H).

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Previously, we confirmed that a similar polyploid mitotic program occurs in the developing hindgut of the mosquito Culex pipiens (Fox et al., 2010). Interestingly, classical descriptions of mitosis in this part of the Culex hindgut seem to suggest a polytene organization is present only early in the first polyploid mitosis (Grell, 1946). In agreement with this observations, we find Phospho-Histone H3 positive polytene chromosomes during the period of the first polyploid mitosis (Figure 3—figure supplement 1E, Video 12). From our Drosophila and Culex studies, we conclude that unlike cells that enter metaphase with polytene chromosomes, a separate mechanism, SIRS, can eliminate polyteny as cells enter metaphase.

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Spindle-independent mitotic timing by Mad2 promotes efficient SIRS

Our dual genome-reduplication systems identified two distinct cellular responses to polytene chromosomes: a Mad2-dependent response that delays anaphase when polytenes remain at metaphase, and a SIRS response that eliminates polyteny as cells enter metaphase. Despite the lack of metaphase polytene chromosomes in papillar cells, we also identified an important role for Mad2 during SIRS. Because mad2 loss has no reported mitotic defects in Drosophila animals (Buffin et al., 2007), we were surprised to find that first division mad2 papillar cells exhibit a substantial increase in DNA bridges (Figure 4A,B, Video 13, Video 14). We did not detect similar defects during papillar mitosis of animals null for mad1, another SAC component (Figure 4A,B, Video 15). Thus, a Mad1 independent function of Mad2 is important in cells during SIRS.

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In Drosophila, Mad2 plays a conserved, cell type-dependent role in regulating NEBD-to-anaphase onset timing (Buffin et al., 2007; Meraldi et al., 2004; Rodriguez-Bravo et al., 2014; Yuan and O'Farrell, 2015). As for the wait-anaphase response, this mitotic timing role involves Mad2 inhibition of the Anaphase Promoting Complex. However, Mad2’s control of overall mitotic timing is irrespective of SAC kinetochore attachment surveillance (Meraldi et al., 2004; Rodriguez-Bravo et al., 2014). Interestingly, Drosophila Mad1 is reported to be dispensable for regulation of NEBD-to-anaphase timing (Emre et al., 2011). Given the lack of mad1 phenotypes with respect to the first papillar division, we thus hypothesized that SIRS enables papillar cells to bypass the SAC-mediated anaphase delay, and that Mad2 control of overall mitotic timing is important during SIRS. If so, one would predict cells undergoing SIRS to not trigger an anaphase delay, but to still depend on mitotic timing.

To first test if papillar cells employ the SAC wait-anaphase in response to polytene chromosomes, we treated animals with colcemid, a known SAC wait-anaphase trigger. This treatment increases the mitotic index of wild type papillar cells, whereas the mitotic index of mad2 null animals is unaffected (Figure 4C,D) Thus, spindle defects trigger the SAC wait-anaphase response in papillar cells. We next asked if the SAC wait-anaphase responds to polytene chromosomes during SIRS. If so, the first divisions (polytenes present) should have a longer metaphase than the second division (polytenes absent). However we find that metaphase is not any longer in the first papillar division than in the second papillar division, while in contrast metaphase is almost twice as long in wing cells with diplochromosomes than in those that are polyploid but lack diplochromosomes (Figure 4E). We then tested if triggering the SAC wait-anaphase response can prevent or delay SIRS completion. We find SIRS occurs on schedule even in the presence of colcemid concentrations that are sufficient to eliminate a detectable spindle and inhibit anaphase (Figure 4F,G, Video 16, Video 17). We conclude that: a) SIRS is not regulated by the SAC wait-anaphase response, and b) chromosome separation during SIRS does not require a mitotic spindle.

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We next tested whether Mad2-dependent control of overall mitotic timing is crucial for efficient SIRS. Using an NEBD marker, we first confirmed that Mad2 regulates NEBD-to-anaphase timing and that mad2 cells spend significantly less time in mitosis than wild type cells (Figure 5A,B, Video 18, Video 19). We also find that Nuclear Envelope Breakdown and the onset of SIRS are synchronous in wild type cells and that SIRS generally continues until up to the onset of anaphase (Figure 5C). This suggests that the rapid mitosis in mad2 cells might lead to a failure of complete SIRS, which could cause the resulting DNA bridges. In our live imaging, we saw evidence that a pre-SIRS group of homologs would often fail to completely disperse prior to anaphase (Figure 5A, mad2, yellow arrowhead). To quantify this, we generated heat maps and line profiles of centromere signals at the metaphase plate. We performed this analysis just prior to the onset of mitosis in wild type cells, mad2 cells that did not generate bridges, and mad2 cells that did generate DNA bridges (Figure 5D). From these measurements, we found that SIRS fails to complete before anaphase in mad2 animals, leading to a high variance of centromere intensity signal across the metaphase plate (reflecting failure of centromere dissociation/SIRS completion). This high variance disrupts the bilateral symmetry of the metaphase plate in mad2 cells that form DNA bridges (Figure 5D’). Taken together, our data show mitotic fidelity after genome reduplication is improved by one of two Mad2-dependent functions: 1) in the presence of metaphase polytene chromosomes a Mad2-dependent wait-anaphase signal is generated, 2) the efficient elimination of polytenes as cells enter metaphase by SIRS requires a Mad2 (but SAC wait-anaphase-independent) NEBD-anaphase timer (Figure 5E).

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Author details

1. Benjamin M Stormo

   Department of Cell Biology, Duke University Medical Center, Durham, United States

   Contribution

   BMS, Contributed to conception and design, Data acquisition, Analysis of data, Drafting/revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Donald T Fox

   1. Department of Cell Biology, Duke University Medical Center, Durham, United States
   2. Department of Pharamacology and Cancer biology, Duke University Medical Center, Durham, United States

   Contribution

   DTF, Contributed to conception and design, Data acquisition, Analysis of data, Drafting/revising the article

   For correspondence

   don.fox@duke.edu

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-0436-179X

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