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. 2018 Mar 26;7:e34122. doi: 10.7554/eLife.34122

A conserved function for pericentromeric satellite DNA

Madhav Jagannathan 1, Ryan Cummings 1,3, Yukiko M Yamashita 1,2,3,✉
Editor: K VijayRaghavan4
PMCID: PMC5957525  PMID: 29578410

Abstract

A universal and unquestioned characteristic of eukaryotic cells is that the genome is divided into multiple chromosomes and encapsulated in a single nucleus. However, the underlying mechanism to ensure such a configuration is unknown. Here, we provide evidence that pericentromeric satellite DNA, which is often regarded as junk, is a critical constituent of the chromosome, allowing the packaging of all chromosomes into a single nucleus. We show that the multi-AT-hook satellite DNA-binding proteins, Drosophila melanogaster D1 and mouse HMGA1, play an evolutionarily conserved role in bundling pericentromeric satellite DNA from heterologous chromosomes into ‘chromocenters’, a cytological association of pericentromeric heterochromatin. Defective chromocenter formation leads to micronuclei formation due to budding from the interphase nucleus, DNA damage and cell death. We propose that chromocenter and satellite DNA serve a fundamental role in encapsulating the full complement of the genome within a single nucleus, the universal characteristic of eukaryotic cells.

Research organism: D. melanogaster

eLife digest

On Earth, life is divided into three domains. The smallest of these domains includes all the creatures, from sunflowers to yeasts to humans, that have the genetic information within their cells encased in a structure known as the nucleus. The genomes of these organisms are formed of long pieces of DNA, called chromosomes, which are packaged tightly and then unpackaged every time the cell divides. When a cell is not dividing, the chromosomes in the nucleus are loosely bundled up together.

It is well known that DNA is the blueprint for the building blocks of life, but actually most of the genetic information in a cell codes for nothing, and has unknown roles. An example of such ‘junk DNA’ is pericentrometric satellite DNA, small repetitive sequences found on all chromosomes.

However, new experiments by Jagannathan et al. show that, in the nucleus of animal cells, certain DNA binding proteins make chromosomes huddle together by attaching to multiple pericentrometric satellite DNA sequences on different chromosomes. In fact, if these proteins are removed from mice and fruit flies cells grown in the laboratory, the chromosomes cannot be clustered together. Instead, they ‘float away’, and the membranes of the nucleus get damaged, possibly buckling under the pressure of the unorganized DNA.

These events damage the genetic information, which can lead to the cell dying or forming tumors. ‘Junk DNA’ therefore appears to participate in fundamental cellular processes across species, a result that opens up several new lines of research.

Introduction

Satellite DNA is AT-rich, non-coding, repetitive DNA that is abundant in centromeric and pericentromeric heterochromatin. Unlike the satellite DNAs that comprise the vast majority of natural centromeres (Willard, 1990; Sun et al., 1997, 2003), the role of pericentromeric satellite DNA remains obscure: although function for a few satellite DNA repeats has been implied in certain cellular processes such as meiotic segregation of achiasmatic chromosomes, X chromosome dosage compensation and formation of lampbrush-like loops on the Y chromosome during male meiosis (Yunis and Yasmineh, 1971; Bonaccorsi et al., 1990; Dernburg et al., 1996; Menon et al., 2014), a unifying theme for pericentromeric satellite DNA function remains elusive. Moreover, highly divergent satellite DNA sequences even among closely related species has led to the idea that satellite DNA does not serve a conserved function and is mostly a selfish element or junk (Doolittle and Sapienza, 1980; Walker, 1971). Pericentromeric satellite DNA repeats are proposed to be sources of genomic instability, as their misexpression is associated with the formation of genotoxic R-loops and DNA damage (Zhu et al., 2011; Zeller et al., 2016; Zeller and Gasser, 2017). Most studies on pericentromeric heterochromatin have focused on the mechanisms to repress satellite DNA transcription, and accordingly, a clear rationale for the existence of most pericentromeric satellite DNA is still lacking.

Cytologically, it is well documented that pericentromeric satellite DNA from multiple chromosomes is clustered into chromocenters in interphase nuclei in diverse eukaryotes including Drosophila, mouse and plants (Figure 1A) (Jones, 1970; Pardue and Gall, 1970; Gall et al., 1971; Fransz et al., 2002). While multiple factors such as epigenetic modifications and transcription of repetitive DNA from pericentromeric DNA sequences are known to be required for chromocenter formation (Peters et al., 2001; Probst et al., 2010; Bulut-Karslioglu et al., 2012; Pinheiro et al., 2012; Hahn et al., 2013), the ultimate consequences of disrupted chromocenter formation has never been addressed, leaving the function of chromocenters unknown.

Figure 1. Multi-AT-hook proteins, D1 and HMGA1, are required for chromocenter formation in Drosophila and mouse cells.

(A) Schematic of pericentromeric heterochromatin being organized into the chromocenter. (B) FISH against {AATAT}n satellite (red) on the Drosophila neuroblast mitotic chromosomes co-stained with DAPI (blue) indicating the location of {AATAT}n in the Drosophila genome. (C) FISH against {AATAT}n satellite (red) in spermatogonial cells immunostained for H3K9me2 (blue) and D1 (green). Dotted lines indicate nucleus. Bars: 5 µm. (D) Drosophila neuroblast mitotic chromosomes stained for D1 (green), phospho-histone H3 Serine 10 (pH3-S10) (blue) and Cid/CENP-A (red). (E–G) FISH against the mouse major satellite (green) on C2C12 mitotic chromosomes co-stained with DAPI (blue) (E), in interphase MOVAS cells co-stained for DAPI (blue) and HMGA1 (red) (F) and in MOVAS cells expressing GFP-D1 (blue) stained for HMGA1 (red) (G). (H, I) FISH against {AATAT}n satellite (red) in control (D1LL03310/+) (H) and D1LL03310/Df (I) spermatogonial cells stained for DAPI (blue) and Vasa (green). (J) Quantification of spermatogonial cells with disrupted chromocenters (+/+ control n = 117, D1LL03310/Df n = 89) from three independent experiments. p-Value from student’s t-test is shown. Error bars: SD. (K, L) FISH against the major satellite (green) in siControl (K) and siHMGA1 (L) transfected MOVAS cells co-stained with DAPI (blue). (M) Quantification of cells with disrupted chromocenters from siControl (n = 304) and siHMGA1 (n = 329) from three independent experiments.

Figure 1.

Figure 1—figure supplement 1. Multi-AT-hook proteins, Drosophila D1 and mouse HMGA1, localize to chromocenters in various mouse cell types.

Figure 1—figure supplement 1.

(A, B) FISH against the mouse major satellite (red) in C2C12 (A) and RAW 264.7 (B) cells stained for HMGA1 (green) and DAPI (blue). (C, D) Colocalization of GFP-D1 (green) with DAPI-dense chromocenters in C2C12 (C) and RAW 264.7(D) cells. DAPI (red). Scale bars: 5 μm.

Figure 1—figure supplement 2. Drosophila D1 and mouse HMGA1 are required for chromocenter formation.

Figure 1—figure supplement 2.

(A–C) Testes from control (+/Df) (A) and two D1 mutant (D1LL03310/Df (B) and D1EY05004/Df (C)) flies were stained for DAPI (blue), Phalloidin (red) and D1 (green). Asterisks indicate the apical tip of the testis. Bars: 5 μm. (D, E) FISH against {AATAT}n (red) in control (D1EY05004/+) (D) and D1EY05004/Df (E) spermatogonial cells stained for DAPI (blue) and Vasa (green). Bars: 2.5 μm. (F, G) FISH against {AATAT}n (red) in control (D1LL03310/+) (F) and D1LL03310/Df (G) spermatocytes stained for DAPI (blue) and Vasa (green). (H, I) FISH against {AATAT}n (red) in control (D1LL03310/+) (H) and D1LL03310/Df (I) accessory gland cells stained for DAPI (blue). Bars: 5 μm. (J, K) FISH against the major satellite (green) in siControl (J) and siHMGA1 transfected (K) C2C12 cells. Dotted lines indicate nucleus. (L) Quantification of cells with disrupted chromocenters in siControl (n = 304) and siHMGA1 (n = 298) transfected C2C12 cells from three independent experiments. p-Value from student’s t-test is shown. Error bars: SD.
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In this study, we explored the role of pericentromeric satellite DNA/chromocenters by studying multi-AT-hook proteins, D1 from Drosophila melanogaster and HMGA1 from mouse. D1 and HMGA1 are known to bind specific pericentromeric satellite DNA, and we show that these proteins are required for chromocenter formation. When chromocenters are disrupted in the absence of these proteins, cells exhibited a high frequency of micronuclei formation, leading to DNA breakage and cell death. We show that micronuclei are formed during interphase by budding from the nucleus. We further show that D1 binding to the target DNA sequence is sufficient to bring it to the chromocenter. High-resolution imaging revealed chromatin threads positive for D1/HMGA proteins and satellite DNA that connect heterologous chromosomes. Taken together, we propose that chromocenter formation via bundling of satellite DNA from heterologous chromosomes functions as a mechanism to encapsulate the full complement of the genome into a single nucleus. We suggest that satellite DNA function as a critical constituent of chromosomes and may serve an evolutionarily conserved role across eukaryotic species.

Results

The multi-AT-hook satellite DNA binding proteins, Drosophila D1 and mouse HMGA1, localize to chromocenters

D1 in Drosophila melanogaster and HMGA1 in mouse are multi-AT-hook proteins; D1 contains 10 AT-hooks, HMGA1 contains three AT-hooks and both proteins contain C-terminal acidic domains (Aulner et al., 2002). D1 and HMGA1 are known to bind the Drosophila {AATAT}n satellite DNA (~8% of the Drosophila male diploid genome) and mouse major satellite DNA (~6% of the mouse genome), respectively (Goodwin et al., 1973; Rodriguez Alfageme et al., 1980; Levinger and Varshavsky, 1982b; Levinger and Varshavsky, 1982a; Lund et al., 1983). The {AATAT}n satellite is distributed across 11 loci on multiple chromosomes as visualized by DNA fluorescence in situ hybridization (FISH) on mitotic chromosome spreads (Figure 1B) (Lohe et al., 1993; Jagannathan et al., 2017). However, it is typically clustered into a few foci in Drosophila interphase nuclei, colocalizing with the D1 protein (Figure 1C). The D1/{AATAT}n foci stained positively for H3K9me2 in interphase nuclei (Figure 1C), a well-established characteristic of constitutive heterochromatin/chromocenters (Guenatri et al., 2004). Consistently, D1 localized near the centromere (marked by Drosophila CENP-A, Cid) on mitotic chromosome spreads (marked by phospho-H3 S10) (Figure 1D). These results suggest that D1 is a chromocenter-localizing protein, via its binding to the {AATAT}n satellite DNA.

The mouse HMGA1 protein was originally identified as an abundant non-histone component of mammalian chromatin (Goodwin et al., 1973; Lund et al., 1983) with subsequent studies demonstrating its binding to satellite DNA (Strauss and Varshavsky, 1984; Radic et al., 1992). Mouse major satellite, which is present in pericentromeric regions of all chromosomes (Figure 1E) (Lyon and Searle, 1989), clustered into DAPI-dense chromocenters positive for HMGA1 protein (Figure 1F, Figure 1—figure supplement 1A,B), revealing an analogous relationship to D1/{AATAT}n satellite in Drosophila. Interestingly, we found that Drosophila D1 protein localizes to major satellite/chromocenters when ectopically expressed in multiple mouse cell lines (Figure 1G, Figure 1—figure supplement 1C,D), suggesting that D1 and HMGA1 may possess an orthologous and conserved function as satellite DNA/chromocenter-binding proteins.

D1 and HMGA1 are required for organizing chromocenters

We next examined the effects of D1 mutation and siRNA-mediated knockdown of HMGA1 on chromocenters. We used two D1 alleles, D1LL03310 and D1EY05004, which we show to be protein null alleles, evidenced by near-complete loss of anti-D1 antibody staining (Figure 1—figure supplement 2A–C). When these alleles were combined with the D1 deficiency allele, Df(3R)BSC666, it led to severe declustering of {AATAT}n satellite DNA (Figure 1H–J, Figure 1—figure supplement 2D–E), suggesting that D1 is required for clustering of pericentromeric satellite DNA into chromocenters. We observed D1’s requirement for chromocenter formation in multiple cell types (Figure 1—figure supplement 2F–I), but we largely focused on spermatogonial cells, where the phenotypes (such as cell death) were most penetrant and severe.

We also examined the requirement for HMGA1 in mouse chromocenter formation. Following siRNA-mediated knockdown of HMGA1, which led to near complete loss of HMGA1 protein (see Figure 2D,E and Figure 2—figure supplement 1A–B,D–E for efficiencies of HMGA1 knockdown), we observed chromocenter disruption in multiple mouse cell lines (Figure 1K–M, Figure 1—figure supplement 2J–L). These results suggest that D1 and HMGA1 have an orthologous function to organize pericentromeric satellite DNA into chromocenters.

Figure 2. D1/HMGA1 loss-of-function results in micronuclei formation, and defective nuclear envelope integrity.

(A, B) Control (D1LL03310/+) (A) and D1LL03310/Df mutant (B) spermatogonial cells stained for DAPI (red), Vasa (blue) and LaminDm0 (green). Arrow indicates micronucleus. Bars: 5 μm. (C) Quantification of micronuclei-containing cells from +/+ control (n = 269) and D1LL03310/Df (n = 334) from three independent experiments. p-Value from student’s t-test is shown. Error bars: SD. (D, E) siControl (D) and siHMGA1 transfected (E) MOVAS cells stained for DAPI (blue), HMGA1 (red) and Lamin (green). Arrow indicates micronucleus. (F) Quantification of micronuclei-containing cells in siControl (n = 518) and siHMGA1 (n = 588) transfected cells from four independent experiments. (G, H) Control (D1LL03310/+) (G) and D1LL03310/Df (H) spermatogonia expressing nls-GFP (green) stained for Vasa (blue) and LaminDm0 (red). nlsGFP was observed in cytoplasm in D1LL03310/Df spermatogonia. (I) Quantification of spermatogonia with cytoplasmic GFP (>1 μm exclusions or pan-cytoplasmic) in D1LL03310/+ (n = 810) and D1LL03310/Df (n = 780) testes from two independent experiments. (J, K) D1LL03310/+ (J) and D1LL03310/Df (K) spermatogonia expressing ER-GFP marker (green) stained for Vasa (blue) and LaminDm0 (red). Arrowhead points to ER marker-positive micronucleus. Arrows point to site of weak nuclear LaminDm0 staining. (L, M) Control (D1LL03310/+) (L) and D1LL03310/Df (M) spermatogonia stained for Vasa (blue) and LaminDm0 (green) and Otefin (red). Arrowhead points to Otefin-containing micronucleus. Arrows point to site of weak nuclear LaminDm0 staining.

Figure 2.

Figure 2—figure supplement 1. Formation of micronuclei upon chromocenter disruption in C3H10T1/2 and C2C12 mouse cells.

Figure 2—figure supplement 1.

(A, B) siControl (A) and siHMGA1 (B) transfected C3H10T1/2 cells stained for DAPI (blue), HMGA1 (red) and LaminB (green). Arrowhead indicates micronuclei. Bars: 5 μm. (C) Quantification of micronuclei containing cells from siControl (n = 291) and siHMGA1 (n = 303) transfected cells from three independent experiments. p-Value from student’s t-test is shown. Error bars are SD. (D, E) siControl (D) and siHMGA1 (E) transfected C2C12 cells stained for DAPI (blue), HMGA1 (red) and LaminB (green). Arrowhead indicates micronuclei. Bars: 5 μm. (F) Quantification of micronuclei containing cells from siControl (n = 953) and siHMGA1 (n = 699) transfected cells from three independent experiments. p-Value from student’s t-test is shown. Error bars are SD.
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Loss of D1/HMGA1 leads to micronuclei formation

To explore the function of chromocenters and satellite DNA, we examined the effects of D1 mutation/HMGA1 knockdown, which showed strikingly similar phenotypes. We found that D1 mutation as well as siRNA-mediated HMGA1 knockdown in multiple mouse cell lines resulted in a significant increase in micronuclei formation (Figure 2A–F, Figure 2—figure supplement 1A–F).

Micronuclei are known to have compromised nuclear envelope integrity, leading to DNA damage and catastrophic chromosomal rearrangement therein (Crasta et al., 2012; Hatch et al., 2013). Therefore, we first examined a possible defect in nuclear envelope integrity in D1 mutant. We found that loss of D1 led to breaching of the nuclear envelope both in major and micronuclei, visualized by the cytoplasmic leakage of nuclear GFP (nlsGFP) (Figure 2G–I), suggesting that nuclear envelope integrity might be generally compromised. Consistently, we observed mislocalization of nuclear envelope proteins in D1 mutant spermatogonia. We frequently observed that lamin surrounded the nucleus incompletely in D1 mutant (1.9% in control (n = 52) and 68.9% in D1 mutant (n = 58)) (Figure 2J,K, arrows indicate lamin-negative regions on the nuclear membrane). We also observed cytoplasmic ‘holes’, which resemble the nucleus in that they exclude cytoplasmic proteins such as Vasa (Figure 2K, arrowhead), but are devoid of nuclear lamin (Figure 2K, arrowhead). These ‘holes’ were often surrounded by an ER marker, which normally surrounds the nuclear envelope (Figure 2J) (Dorn et al., 2011). Similarly, Otefin, an inner nuclear membrane LEM-domain protein (Barton et al., 2014), also showed perturbed localization (2.7% in control (n = 109) and 24.5% in D1 mutant (n = 106)) (Figure 2L,M, arrows indicate lamin/Otefin negative regions on the nuclear envelope while the arrowhead indicates Otefin-positive micronuclei). Taken together, these results show that D1 mutant cells exhibit compromised nuclear envelope integrity, which is associated with micronuclei formation.

Loss of D1/HMGA1 leads to accumulation of DNA damage

It has been shown that defects in nuclear envelope integrity can lead to extensive DNA damage in the major nucleus and micronuclei (Crasta et al., 2012; Hatch et al., 2013; Zhang et al., 2015; Denais et al., 2016; Raab et al., 2016). Nuclear envelope defects and extensive DNA damages therein lead to catastrophic chromosomal breaks/rearrangements termed chromothripsis (Crasta et al., 2012; Hatch et al., 2013). Such catastrophic DNA breaks/rearrangements are speculated to lead to tumorigenesis (Hatch and Hetzer, 2015).

Consistent with defective nuclear envelope integrity, we observed extensive DNA damage (revealed by γ-H2Av) in both major and micronuclei (Figure 3A–F, arrows point to damaged DNA in micronuclei in B and D). Likely as a result of DNA damage and defective nuclear envelope integrity, D1 mutant testes rapidly degenerated (Figure 3—figure supplement 1A,B). When Omi, a gene required to promote germ cell death (Yacobi-Sharon et al., 2013), was knocked down in D1 mutant testes, it restored the cellularity in D1 mutant testis (Figure 3—figure supplement 1C–D), but the surviving cells showed a dramatic increase in DNA damage (Figure 3—figure supplement 1E–F). Under these conditions, we observed that surviving germ cells in D1 mutant testes showed a high frequency of chromosome breaks compared to control, revealed by FISH on metaphase chromosome spreads from spermatocytes (3.7% in control (n = 27) vs. 15.8% in D1 mutant (n = 57)) (Figure 3G,H, arrowheads indicate sites of chromosome breaks). These results show that loss of D1/HMGA1 results in compromised nuclear envelope integrity, leading to extensive DNA damage and chromosomal breaks.

Figure 3. D1 mutation/HMGA1 depletion leads to an increase in DNA damage.

(A, B) Control (D1LL03310/+) (A) and D1LL03310/Df (B) spermatogonia stained for DAPI (blue), Vasa (green) and γ-H2Av (red). Dotted lines indicate nucleus and arrow points to DNA damage in micronuclei. (C, D) siControl (C) and siHMGA1 (D) transfected MOVAS cells stained for DAPI (blue), γ-H2Av (red) and LaminDm0 (green). Arrow points to DNA damage in micronuclei. (E) Quantification of γ-H2Av positive cells in D1LL03310/+ (n = 317) and D1LL03310/Df (n = 242) spermatogonia from three independent experiments. (F) Quantification of cells containing >6 γ-H2Ax foci in siControl (n = 304) and siHMGA1 (n = 309) transfected cells from three independent experiments. (G, H) FISH against the rDNA intergenic spacer (IGS) (green), {AATAT}n (red) and {AAGAG}n (blue) on chromosome spreads from meiotic spermatocytes from control (nos > OmiRNAi, n = 27) and D1 mutant (nos >Omi RNAi; D1LL03310/Df, n = 57) testes co-stained for DAPI (grey). OmiRNAi was used to block DNA damage-induced cell death. Arrowheads point to chromosome breaks.

Figure 3—source data 1. Quantification of g-H2Ax foci in mouse cells.
g-H2Ax foci were scored in siControl and siHMGA1 transfected MOVAs cells. At least 100 cells were scored from three biological replicates. Percentage of cells containing the indicated number of g-H2Ax foci is listed in the table.
elife-34122-fig3-data1.xlsx (9.4KB, xlsx)
DOI: 10.7554/eLife.34122.010

Figure 3.

Figure 3—figure supplement 1. Chromocenter disruption results in germ cell death in Drosophila in an Omi-dependent manner.

Figure 3—figure supplement 1.

(A, B) Representative images of 14-day-old control (D1LL03310/+, n = 18) and D1LL03310/Df (n = 12) testes stained for DAPI (blue) and the germ cell marker, Vasa (green). Asterisk indicates apical tip. Bars: 25 μm (C, D) Representative images of D1 mutant testes (D1LL03310/Df) without (C) and with (D) germ cell death suppression by Omi knockdown (nos > OmiRNAi), stained for DAPI (blue), Vasa (green) and LaminDm0 (red) at 7 days post eclosion. (E, F) Representative images of control (nos > OmiRNAi, D1LL033010/+, n = 10) and D1 mutant (nos > OmiRNAi; D1LL03310/Df, n = 13) testes stained for DAPI (blue), Vasa (green) and γ-H2Av (red).
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Micronuclei formation in D1 mutant/HMGA1 knockdown cells is due to budding from the nucleus during interphase

It has been shown that micronuclei form by lagging chromosomes (Crasta et al., 2012). Thus, we examined whether D1 mutation/HMGA1 knockdown resulted in mitotic chromosome segregation errors, causing micronuclei formation. However, we did not observe an increase in lagging chromosomes in D1 mutant spermatogonia or HMGA1-depleted mouse cells (Figure 4—figure supplement 1A–G), suggesting an alternative route for micronuclei formation. Instead, time-lapse live observation showed that micronuclei formed by budding from the interphase nucleus both in Drosophila spermatogonia and mouse cells (Figure 4A–D). In Drosophila spematogonia, nuclear contents were visualized by a GFP-tagged nuclear protein, Df31, and RFP-tagged histone H2Av. Control cells stably maintained nuclear contents for a prolonged time period (only 1 event of nuclear blebbing without concurrent micronuclei formation (as detected by H2Av-RFP) over 1552 min of live imaging) (Figure 4A). In contrast, D1 mutant cells showed budding off of nuclear contents and micronuclei formation in interphase (15 nuclear breaches with eight micronuclei formed over 3427 min of live imaging with a total budding duration of 172 min) (Figure 4B). Similarly, live imaging in mouse cells using the Hoechst DNA dye revealed that HMGA1 knockdown also resulted in micronuclei formation during interphase (siControl – no micronuclei formation over 253 min of live observation, siHMGA1 – three micronuclei formed by budding over 5962 min of live imaging with a total budding duration of 310 min) (Figure 4C,D). These results show that micronuclei in D1 mutant/HMGA1-knockdown cells are generated during interphase, via budding from the nucleus.

Figure 4. D1/HMGA1 loss of function results in micronuclei formation due to nuclear budding during interphase.

(A, B) Time-lapse live imaging of control (+/+) (A) and D1LL03310/Df (B) spermatogonial cells expressing Df31-GFP as a nuclear marker and H2Av-RFP as a DNA marker. (C, D) Time-lapse live imaging of siControl (C) and siHMGA1 (D) MOVAS cells stained with Hoechst 33342. Arrowheads indicate site of micronucleus budding. Time is indicated in mm:ss. Scale bars: 5 μm.

Figure 4.

Figure 4—figure supplement 1. Micronuclei formation upon chromocenter disruption is not a result of mitotic lagging chromosomes.

Figure 4—figure supplement 1.

(A, B) Examples of normal and lagging mitotic chromosomes in Drosophila spermatogonia stained for Vasa (blue) and pH3-S10 (green). (C) Quantification of spermatogonia with lagging chromosomes from control (D1LL033010/+, n = 43) and D1LL03310/Df (n = 47) from three independent experiments. P value from student’s t test is shown. Error bars are SD. (D, E) Examples of normal and lagging mitotic chromosomes in mouse cells stained for DAPI (red) and α-tubulin (green). Bars: 5 μm. (F) Quantification of mitotic cells with lagging chromosomes from siControl (n = 149) and siHMGA1 (n = 174) transfected MOVAS cells from three independent experiments. (G) Quantification of mitotic cells with lagging chromosomes from siControl (n = 110) and siHMGA1 (n = 129) transfected C2C12 cells from three independent experiments.
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D1 bundles satellite DNA from multiple chromosomes to form chromocenter

Based on the above results, we postulated that chromocenter formation, that is clustering of satellite DNA from multiple chromosomes, might be a mechanism to bundle heterologous chromosomes together to prevent individual chromosomes from floating out of the nucleus. In this manner, the full set of chromosomes may be retained within a single nucleus. In the absence of chromocenter formation, individual chromosomes may bud off the nucleus, leading to micronuclei formation.

Previous in vitro experiments indicated that HMGA1 is capable of crosslinking multiple DNA strands with individual AT-hooks binding AT-rich DNA strands (Vogel et al., 2011). Bundling of DNA in this manner by D1/HMGA1 could explain how pericentromeric satellite DNA from multiple chromosomes may be clustered to form chromocenters. A few lines of evidence support this idea. When Drosophila D1 was expressed in mouse cells, it localized to the chromocenter as described above (Figure 1G), and its overexpression enhanced chromocenter formation in a dose-dependent manner (Figure 5A–C): the higher the amount of D1 that was expressed in mouse cells, the fewer chromocenters per cell was observed (i.e. more clustering). These results suggest that D1 is sufficient to bundle its binding target, tethering it to chromocenter. Consistent with this idea, we found that artificial tethering of D1 protein to euchromatic LacO repeat DNA sequences was sufficient to bring LacO repeats to the chromocenter. D1 protein or D1-LacI fusion protein was expressed in a Drosophila strain in which LacO repeats are inserted in the distal regions of the 2nd chromosome (Figure 5D, arrows). In control spermatogonial cells expressing wild type D1, LacO repeats were observed far away from the {AATAT}n satellite foci/chromocenters (Figure 5E,G, arrow indicates site of LacO repeats in interphase nucleus). However, in cells expressing the LacI-D1 chimeric protein, we observed recruitment of the LacO repeats close to {AATAT}n/chromocenters (Figure 5F,G, arrow indicates site of LacO repeats recruited to the chromocenter), demonstrating that D1’s binding to a DNA sequence is sufficient to incorporate the target sequence into chromocenters.

Figure 5. D1 bundles satellite DNA from heterologous chromosomes to form chromocenter.

Figure 5.

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(A, B) C2C12 cells expressing GFP only (blue) (A) or GFP-D1 (blue) (B) stained for DAPI (red). Dotted lines indicate nucleus. (C) Quantification of chromocenter number relative to expression level of GFP (n = 29) or GFP-D1 (n = 47). P value and R2 value are indicated from linear regression analysis. (D) FISH against LacO (red) and {AATAT}n (green) on mitotic neuroblast chromosomes from the LacO strain stained for DAPI (blue), indicating the sites of LacO insertion (arrows). (E, F) FISH against LacO (red) and {AATAT}n (green) in spermatogonia expressing GFP-D1 (blue) (E) or GFP-LacI-D1 (blue) (F). Arrows indicate location of LacO sequence. (G) AATAT-LacO distance (nm) in GFP-D1 (n = 97) and GFP-LacI-D1 (n = 69) expressing spermatogonia. P value from student’s t-test is shown. Error bars: SD. All scale bars: 5 μm.

Figure 5—source data 1. Quantification of the relative expression of GFP/GFP-D1 and number of chromocenters in mouse cells.
Images of cells expressing GFP (n=27) or GFP-D1 (n=47) were acquired from a single slide/transfection with all image acquisition parameters held constant and care taken to avoid pixel oversaturation. Total GFP fluorescence per cell was quantified using ImageJ software. Relative expression was calculated by dividing each individual fluorescence value over the maximum fluorescence value obtained over the course of the experiment. DAPI staining was used to calculate number of chromocenters per cell.
elife-34122-fig5-data1.xlsx (9.7KB, xlsx)
DOI: 10.7554/eLife.34122.014
Figure 5—source data 2. Quantification of LacO-AATAT distance (nm) in cells expressing GFP-D1 and GFP-LacI-D1.
LacO-AATAT distance (nm) was measured in spermatogonial cells expressing GFP-D1 (n=97) and GFP-LacI-D1 (n=69) using Leica LAS X software.
elife-34122-fig5-data2.xlsx (10.3KB, xlsx)
DOI: 10.7554/eLife.34122.015

Although it cannot be visualized how DNA strands from multiple chromosomes might be bundled in these interphase chromocenters, deconvolution microscopy of D1/HMGA1 proteins on early mitotic chromosomes revealed proteinaceous threads between chromatin in the process of condensation (Figure 6A,B, arrows indicate D1/HMGA1 threads), which we speculate contributed to bundling of chromosomes in the previous interphase. These threads were also detectable by DNA FISH against {AATAT}n and the mouse major satellite (Figure 6C,D, dotted lines are alongside the satellite DNA threads), suggesting that satellite DNA bound by D1/HMGA1 can form threads. These threads likely connect heterologous chromosomes, as we see threads between chromosomes that are clearly distinct in their morphology (e.g. Figure 6C). These D1/HMGA1 threads are reminiscent of ‘DNA fibers’, which were observed among mitotic chromosomes, although their function has never been appreciated (Takayama, 1975; Burdick, 1976; Kuznetsova et al., 2007).

Figure 6. D1/HMGA1 and satellite DNA form chromatin threads that link chromosomes.

Figure 6.

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(A) Deconvolution microscopy performed on Drosophila mitotic neuroblasts stained for D1 (cyan) and pH3-S10 (magenta). Arrows in magnified images indicate D1-positive thread connecting two chromosomes. (B) Deconvolution microscopy performed on CSK-extracted RAW 264.7 macrophages stained for HMGA1 (cyan) and DAPI (magenta). Arrows in magnified images indicate HMGA1-positive thread connecting two chromosomes. (C) Deconvolution microscopy performed on neuroblast mitotic chromosomes stained for DAPI (magenta) and FISH against {AATAT}n (cyan) from a Drosophila strain containing AATAT-rich B chromosomes (Bauerly et al., 2014). Dotted lines in magnified images indicate AATAT-positive threads connecting heterologous chromosomes. (D) Deconvolution microscopy performed on RAW 264.7 macrophages stained for DAPI (magenta) and FISH against major satellite (cyan). Dotted lines in magnified images indicate major satellite-positive threads connecting two chromosomes. (E) The model of chromosome bundling by D1/HMGA1 and satellite DNA.

Taken together, these results support a model, in which D1/HMGA1 bind their target sequences (satellite DNA) on multiple chromosomes and bundle them into chromocenters, likely via their multivalent DNA-binding domains (multiple AT-hooks) (Figure 6E).

Discussion

The function of chromocenters, as well as that of satellite DNA, has remained enigmatic, even though cytological association of pericentromeric satellite DNA into chromocenters was identified almost 50 years ago (Jones, 1970; Pardue and Gall, 1970). Pericentromeric heterochromatin has most often been studied and discussed in the context of how to maintain its heterochromatic, repressed nature (Nishibuchi and Déjardin, 2017), based on the assumption that the underlying sequences are mostly selfish, which have negative phenotypic consequences when derepressed in cells (Zeller and Gasser, 2017).

Although satellite DNA’s function has been speculated and implicated in several examples (Yunis and Yasmineh, 1971; Bonaccorsi et al., 1990; Dernburg et al., 1996; Menon et al., 2014), the non-coding nature and lack of conservation in repeat sequence among closely related species led to the idea that they are mostly junk DNA, serving no essential function (Walker, 1971; Doolittle and Sapienza, 1980). Instead, we propose that satellite DNA is a critical constituent of eukaryotic chromosomes to ensure encapsulation of all chromosomes in interphase nucleus. Our results may also explain why the sequences of pericentromeric satellite DNA are so divergent among closely related species, a contributing factor that led to their dismissal as junk. Based on our model that pericentromeric satellite DNA serves as a platform for generating heterologous chromosome association to form chromocenters, the essential feature of satellite DNA is that they are bound by protein(s) capable of bundling multiple DNA strands. If so, the underlying sequence does not have to be conserved. Instead, the binding of satellite DNA by a chromocenter bundling protein may be a critical feature of pericentromeric satellite DNAs. Based on this idea, chromocenter bundling proteins and pericentromeric satellite DNA may be co-evolving.

We observed perturbation of nuclear envelope integrity upon chromocenter disruption. Understanding the mechanisms underlying perturbation of nuclear envelope integrity in D1 mutant awaits future investigation. Previous studies have documented that cytoskeletal forces are transmitted to chromatin through nuclear envelope and external mechanical forces can cause temporary nuclear envelope breaches (King et al., 2008; Denais et al., 2016; Hatch and Hetzer, 2016; Raab et al., 2016). Therefore, we speculate that chromosome bundling in the form of chromocenter may help prevent cytoskeletal forces from shearing chromosomes and nuclear envelope: when chromosomes are not bundled, cytoskeletal forces may be transmitted to individual chromosomes and associated nuclear envelope, resulting in shearing of nuclear envelope, disrupting its integrity.

In summary, our study provides the first evidence for a conserved function of pericentromeric satellite DNA and chromocenters. Our data suggest that the multi-AT-hook proteins, D1 and HMGA1, play an evolutionarily conserved role in the formation of chromocenters, likely via their ability to bind and bundle satellite DNA from heterologous chromosomes. Heterologous chromosome association, mediated by chromocenter-binding proteins, may represent a third mode of chromosomal ‘gluing’ after meiotic homologous pairing and sister chromatid cohesion. Through heterologous association, the chromocenter plays a fundamental role in maintaining the full complement of the genome, which is divided into multiple chromosomes, into a single nucleus. This function of the chromocenter may be conserved in eukaryotic species that contain pericentromeric satellite DNA, thereby bringing about a signature characteristic of eukaryotic cells.

Materials and methods

Key resources table.

Reagent type (species
) or resource		 Designation Source or reference Identifiers Additional information
Genetic Reagent (D. melanogaster) D1EY05004 Bloomington Drosophila Stock Center ID_BDSC:17340
Genetic Reagent (D. melanogaster) Df(3R)BSC666 Bloomington Drosophila Stock Center ID_BDSC:26518
Genetic Reagent (D. melanogaster) UAS-OmiRNAi Bloomington Drosophila Stock Center ID_BDSC:55165
Genetic Reagent (D. melanogaster) UAS-GFP-nls Bloomington Drosophila Stock Center ID_BDSC:4776
Genetic Reagent (D. melanogaster) UAS-GFP-ER-SR Bloomington Drosophila Stock Center ID_BDSC:59042
Genetic Reagent (D. melanogaster) D1LL03310 Kyoto Stock Center ID_DGRC:140754
Genetic Reagent (D. melanogaster) Df31-GFP Kyoto Stock Center ID_DGRC:110806
Genetic Reagent (D. melanogaster) nos-gal4 PMID: 9501989
Genetic Reagent (D. melanogaster) hs-flp;nos-FRT-stop-FRT-gal4,UAS-GFP PMID: 24465278
Genetic Reagent (D. melanogaster) UAS-H2A-YFP PMID: 11146626
Genetic Reagent (D. melanogaster) B1 LacO PMID: 12225662
Genetic Reagent (D. melanogaster) mtrm126 + B PMID: 24478336 Gift of Dr. Scott Hawley
Recombinant DNA Reagent pUASt-GFP-attB PMID: 24465278
Recombinant DNA Reagent pUASt-GFP-D1-attB This Paper
RecombinantDNA Reagent pUASt-GFP-LacI-D1-attB This Paper
Recombinant DNA Reagent pCDNA3 Gift of Dr. Cheng-Yu Lee
Cell Line MOVAS Gift of Dr. Daniel Eitzman
Cell Line C2C12 Gift of Dr. David Bridges
Cell Line RAW264.7 Gift of Dr. Harry Mobley
Cell Line C3H10T1/2 Gift of Dr. Stephen Weiss
siRNA ON-TARGET plus Mouse HMGA1 siRNA SMARTpool Dharmacon/GE Healthcare ID_Dharmacon: L-049293–01
siRNA ON-TARGET plus Non-targeting pool Dharmacon/GE Healthcare ID_Dharmacon: L-001810–10
Antibody anti-Vasa Santa Cruz Biotechnology ID_SCB: d-26
Antibody anti-H3K9 dimethyl Abcam ID_abcam: ab32521
Antibody anti-Otefin Gift of Dr. Georg Krohne
Antibody anti-D1 This Paper Peptide - CDGENDANDGYVSDNYNDSESVAA
Antibody anti-LaminDm0 Developmental Studies Hybridoma Bank ID_DSHB: ADL84.12
Antibody anti-γ-H2Av Developmental Studies Hybridoma Bank ID_DSHB: UNC93-5.2.1
Antibody Phalloidin-Alexa546 ThermoFisher ID_ThermoFisher: a22283
Antibody anti-HMGA1 Abcam ID_abcam: ab129153
Antibody anti-LaminB (C20) Santa Cruz Biotechnology ID_SCB: 2616
Antibody anti-α-tubulin Developmental Studies Hybridoma Bank ID_DSHB: 4.3
Antibody anti-γ-H2Ax S139 Cell Signaling Technologies ID_CST: 2577
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Fly husbandry and strains

All fly stocks were raised on standard Bloomington medium at 25°C. The following fly stocks were used: D1EY05004(BDSC17340), Df(3R)BSC666 (BDSC26518), UAS-OmiRNAi (BDSC55165), UAS-GFP-nls (BDSC4776) and UAS-GFP-ER-SR (BDSC59042) were obtained from the Bloomington Drosophila stock center. D1LL03310 (DGRC140754) and Df31-GFP (DGRC110806) were obtained from the Kyoto stock center. nos-gal4 (Van Doren et al., 1998), hs-flp;nos-FRT-stop-FRT-gal4,UAS-GFP (Salzmann et al., 2013), UAS-H2A-YFP (Bellaïche et al., 2001) and B1 LacO (Vazquez et al., 2002) have been previously described. A stock containing B chromosomes, mtrm126 +B (Bauerly et al., 2014), was a kind gift from Scott Hawley. Chromocenter disruption was scored in Drosophila testes by assessing {AATAT}n morphology in GFP +cells that were generated as follows in control (hs-flp; nos-FRT-stop-FRT-gal4,UAS-GFP) and D1 mutant (hs-flp;nos-FRT-stop-FRT-gal4, UAS-GFP;D1LL03310/Df) flies. Testes were dissected 24 hr following a 20 min heat shock at 37°C. Chromocenters were considered disrupted in Drosophila and mouse when satellite DNA adopted thread-like morphology in interphase nuclei. Micronuclei were scored in 0-3 d testes where early germ cell chromosomes were labeled with H2A-YFP. The genotypes used were, control – nos > H2 A-YFP and D1 mutant – nos > H2 A-YFP; D1LL03310/Df.

Transgene construction

For construction of UAS-GFP-D1, the D1 ORF was PCR-amplified from cDNA using the following primer pair, 5’-GATCAGATCTATGGAGGAAGTTGCGGTAAAG-3’ and 5’-GATCCTCGAGTTAGGCAGCTACCGATTCGG-3’. The amplified fragment was subcloned into the BglII and XhoI sites of pUASt-EGFP-attB (Salzmann et al., 2013) resulting in UAS-GFP-D1. For UAS-GFP-LacI-D1, the LacI ORF (lacking 11 C-terminal residues) (Straight et al., 1996) was synthesized using GeneArt (Thermofisher) and inserted into the BglII site of UAS-GFP-D1 resulting in UAS-GFP-LacI-D1. Transgenic flies were generated by PhiC31 integrase-mediated transgenesis into the attP40 site (BestGene). For expression of GFP and GFP-D1 in mouse cells, GFP and GFP-D1 was subcloned from pUASt-EGFP-attB into pCDNA3 (gift from Cheng-Yu Lee) using EcoRI and XhoI sites.

Cell lines

Mouse MOVAS cells were obtained from Daniel Eitzman. Mouse C2C12 cells were obtained from David Bridges. Mouse RAW264.7 cells were obtained from Dr. Harry Mobley. Mouse C3H10T1/2 cells were obtained from Stephen Weiss. MOVAS, C2C12 and RAW264.7 cells were maintained in Dulbecco's minimal essential medium (DMEM) (Gibco) supplemented with 10% fetal bovine serum (FBS). C3H10T1/2 cell line was maintained in alpha minimal essential media (Gibco) supplemented with 10% fetal bovine serum. All cell lines used were authenticated as mouse cells by the presence of mouse-specific satellite DNA as is shown throughout the manuscript. Two major cell lines used in this study, C2C12 and MOVAS cells, were treated with Plasmocin (Invivogen) prior to use as a precaution for mycoplasma infection.

siRNA and transfections

RNA interference (RNAi) against HMGA1 was performed using ON-TARGET plus Mouse HMGA1 siRNA SMARTpool (Dharmacon, L-049293–01) consisting of the following target sequences, CCAUUUAGCCGCAGCCCGA, AGGCAAACGGGCACCAACA, GGGCGCAGCAGACUGGUUA, GUUCAUUCUUAGAUACCCA. ON-TARGET plus Non-targeting pool (Dharmacon, D-001810–10) consisting of the following sequences, UGGUUUACAUGUCGACUAA, UGGUUUACAUGUUGUGUGA, UGGUUUACAUGUUUUCUGA, UGGUUUACAUGUUUUCCUA, was used as a negative control. siRNA transfections were performed using DharmaFECT four reagent (Dharmacon, Lafayette, CO) according to the manufacturer's protocol. 25 nM of siControl and siHMGA1 were transfected using DharmaFECT 4 (Dharmacon) according to the manufacturer’s protocol. Cells were fixed for immunostaining/in situ hybridization 6 days post-transfection. Transient transfection of GFP and GFP-D1 was performed using Fugene HD (Roche) reagent according to the manufacturer’s protocol.

Immunofluorescence staining and microscopy

For Drosophila tissues, immunofluorescence staining was performed as described previously (Cheng et al., 2008). Briefly, tissues were dissected in PBS, transferred to 4% formaldehyde in PBS and fixed for 30 min. Testes were then washed in PBS-T (PBS containing 0.1% Triton-X) for at least 60 min, followed by incubation with primary antibody in 3% bovine serum albumin (BSA) in PBS-T at 4°C overnight. Samples were washed for 60 min (three 20 min washes) in PBS-T, incubated with secondary antibody in 3% BSA in PBS-T at 4°C overnight, washed as above, and mounted in VECTASHIELD with DAPI (Vector Labs). The following primary antibodies were used: rabbit anti-vasa (1:200; d-26; Santa Cruz Biotechnology), rabbit anti-H3K9 dimethyl (1:200; Abcam, ab32521), guinea pig anti-Otefin (gift from Georg Krohne, 1:400), chicken anti-Cid (1:500, generated using the synthetic peptide CDGENDANDGYVSDNYNDSESVAA (Covance)), mouse anti-LaminDm0 (ADL84.12, 1:200, Developmental Studies Hybridoma Bank), mouse anti-γ−H2Av (UNC93-5.2.1, 1:400, Developmental Studies Hybridoma Bank), Phalloidin-Alexa546 (ThermoFisher, a22283, 1:200). Adherent mouse cells were fixed in 4% formaldehyde in PBS for 20 min at room temperature on coverslips. Cells were permeabilized in PBS-T for 5 min, rinsed three times with PBS, blocked using 3% BSA in PBS-T for 30 min at room temperature and incubated with primary antibody diluted in 3% BSA in PBS-T overnight at 4°C. Cells were then washed for 30 min (three 10 min washes), incubated with secondary antibody in 3% BSA in PBS-T for 2 hr at room temperature, washed as above and mounted in VECTASHIELD with DAPI (Vector Labs). For nucleoplasmic extraction, cells were incubated with CSK buffer (10 mM PIPES pH7, 100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 0.5% Triton X-100, 1 mM PMSF) for 10 min at room temperature. After CSK extraction, cells were washed with PBS and fixed and immunostained as above. The following antibodies were used: rabbit anti-HMGA1 (1:400, Abcam, ab129153), goat anti-LaminB (C-20) (1:20, Santa Cruz Biotechnology, sc-2616), mouse anti-α-tubulin (4.3, 1:100, Developmental Studies Hybridoma Bank) and γ−H2Ax S139 (2577, 1:200, Cell Signaling Technologies). Images were taken using a Leica TCS SP8 confocal microscope with 63x oil-immersion objectives (NA = 1.4). Deconvolution was performed when indicated using the Hyvolution package from Leica. Images were processed using Adobe Photoshop software.

Time-lapse live imaging

Testes from newly eclosed flies were dissected into Schneider’s Drosophila medium containing 10% fetal bovine serum. The testis tips were placed inside a sterile glass-bottom chamber and were mounted on a three-axis computer-controlled piezoelectric stage. An inverted Leica TCS SP8 confocal microscope with a 63 × oil immersion objective (NA = 1.4) was used for imaging. For mouse live cell imaging, transfected cells were seeded onto a sterile glass-bottom chamber coated with poly-lysine. Cells were incubated with Hoechst 33342 for 10 min, rinsed with PBS and fresh medium was added to the chamber. Cells were imaged using a stage-top Tokai-Hit incubator that was mounted on an inverted TCS SP5 confocal microscope with a 63x oil immersion objective (NA = 1.4). All images were processed using Adobe Photoshop software. Metrics used for quantification of live imaging were total imaging duration (defined as number of cells x imaging duration), total budding duration (defined as number of cells with micronuclei that formed by budding x time with budded micronuclei).

DNA fluorescence in situ hybridization

Whole mount Drosophila testes were prepared as described above, and optional immunofluorescence staining protocol was carried out first. Subsequently, samples were post-fixed with 4% formaldehyde for 10 min and washed in PBS-T for 30 min. Fixed samples were incubated with 2 mg/ml RNase A solution at 37°C for 10 min, then washed with PBS-T +1 mM EDTA. Samples were washed in 2xSSC-T (2xSSC containing 0.1% Tween-20) with increasing formamide concentrations (20%, 40% and 50%) for 15 min each followed by a final 30 min wash in 50% formamide. Hybridization buffer (50% formamide, 10% dextran sulfate, 2x SSC, 1 mM EDTA, 1 μM probe) was added to washed samples. Samples were denatured at 91°C for 2 min, then incubated overnight at 37°C. For mitotic chromosome spreads, testes and larval 3rd instar brains were squashed according to previously described methods (Larracuente and Ferree, 2015). Briefly, tissue was dissected into 0.5% sodium citrate for 5–10 min and fixed in 45% acetic acid/2.2% formaldehyde for 4–5 min. Fixed tissues were firmly squashed with a cover slip and slides were submerged in liquid nitrogen until bubbling ceased. Coverslips were then removed with a razor blade and slides were dehydrated in 100% ethanol for at least 5 min. After drying, hybridization mix (50% formamide, 2x SSC, 10% dextran sulfate, 100 ng of each probe) was applied directly to the slide, samples were heat denatured at 95°C for 5 min and allowed to hybridize overnight at room temperature. Following hybridization, slides were washed thrice for 15 min in 0.2X SSC and mounted with VECTASHIELD with DAPI (Vector Labs). For the in situ experiment described in Figure 4j–m, testes were dissected into PBS and fixed in 4% formaldehyde for 4 min. Tips of fixed testes were firmly squashed with a cover slip and slides were submerged in liquid nitrogen until bubbling ceased. Coverslips were removed with a razor blade and slides were subjected to 5 min washes in 2XSSC and 2XSSC with 0.1% Tween-20. The samples were denatured in freshly made 0.07N NaOH for 5 min, rinsed in 2X SSC. Hybridization mix (50% formamide, 2x SSC, 10% dextran sulfate, 100 ng of each probe) was added directly to the slide and allowed to hybridize overnight at room temperature. Following hybridization, slides were washed three times for 15 min in 0.2X SSC and mounted with VECTASHIELD with DAPI (Vector Labs). The following probes were used for Drosophila in situ hybridization: {AATAT}6, {AAGAG}6, IGS and have been previously described (Jagannathan et al., 2017). LacO probe - 5’-Cy5-CCACAAATTGTTATCCGCTCACAATTCCAC-3’. For interphase mouse cells, optional immunostaining was carried out as above. Subsequently, samples were post-fixed with 4% formaldehyde in PBS for 10 min and rinsed three times in PBS. Post-fixed cells were incubated with 0.1 mg/ml RNase A solution at 37°C for 1 hr, rinsed three times in PBS and denatured in 1.9M HCl for 30 min. After three rinses in ice-cold PBS, hybridization mix (2X SSC, 60% formamide, 5 µg/ml salmon sperm DNA and 500 nM probe) was added to the samples and incubated overnight at room temperature. Following hybridization, coverslips were washed three times for 15 min in 2X SSC and mounted with VECTASHIELD with DAPI (Vector Labs). For mouse mitotic cells, chromosomes were spread on slides as previously described. Subsequently, chromosomes were denatured in 70% formamide in 2XSSC for 1.5 min at 70°C, dehydrated in 100% ethanol and hybridization mix (2X SSC, 60% formamide, 5 µg/ml salmon sperm DNA and 500 nM probe) was added directly to the slide and incubated overnight at room temperature. Following hybridization, slides were washed three times for 15 min in 2X SSC and mounted with VECTASHIELD with DAPI (Vector Labs). The following probe was used: Major satellite - 5’-Cy3-GGAAAATTTAGAAATGTCCACTG-3’.

Acknowledgements

We thank Cheng-Yu Lee, Scott Hawley, Stephen Weiss, Harry Mobley, Dave Bridges, Daniel Eitzman, Georg Krohne, Bloomington Drosophila Stock Center, Kyoto Stock Center, Developmental Studies Hybridoma Bank and Michigan Imaging Laboratory for reagents and resources. We thank the Yamashita lab members, Sue Hammoud, Ajit Joglekar, Puck Ohi, Dan Barbash, and Maurizio Gatti for discussion and comments on the manuscript. This research was supported by the Howard Hughes Medical Institute (YY) and an American Heart Association postdoctoral fellowship (MJ). MJ and YY conceived the project, interpreted the data and wrote the manuscript. All authors contributed to conducting experiments and analyzing data.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Yukiko M Yamashita, Email: yukikomy@umich.edu.

K VijayRaghavan, National Centre for Biological Sciences, Tata Institute of Fundamental Research, India.

Funding Information

This paper was supported by the following grants:

  • Howard Hughes Medical Institute to Yukiko M Yamashita.

  • National Institute of General Medical Sciences to Yukiko M Yamashita.

  • American Heart Association to Madhav Jagannathan.

Additional information

Competing interests

Reviewing editor, eLife.

No competing interests declared.

Author contributions

Conceptualization, Formal analysis, Supervision, Funding acquisition, Investigation, Writing—original draft, Writing—review and editing.

Formal analysis, Validation, Investigation, Writing—review and editing.

Conceptualization, Supervision, Funding acquisition, Writing—original draft, Writing—review and editing.

Additional files

Transparent reporting form
elife-34122-transrepform.docx (249.2KB, docx)
DOI: 10.7554/eLife.34122.017

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Decision letter

Editor: K VijayRaghavan1

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "A conserved function for pericentromeric satellite DNA" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by K VijayRaghavan as the Reviewing Editor and Senior Editor. The following individual involved in the review of your submission has agreed to reveal her identity: Susan Gerbi.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This is an excellent paper with solid and thorough data leading to a conceptually novel model. It includes scholarly citation of papers from the older literature as well as current papers, drawing together many disparate areas. The authors have studied the proteins D1 in Drosophila and its counterpart HMGA1 in mouse which are multi AT-hook satellite DNA binding proteins. First, the authors show that these proteins co-localize with chromocenters in Drosophila and mouse. Next, they demonstrate that deletions or knock-down of these proteins disrupt the chromocenters. Moreover, they find that defective chromocenter formation leads to budding off of micronuclei from the nucleus. In addition, loss of D1/HMGA1 leads to accumulation of DNA damage. An elegant experiment with an ectopic LacO and a D1/LacI fusion shows that the D1/LacI fusion protein can recruit the ectopic LacO to the chromocenter. Microscopy reveals that D1/HMGA1 can bundle threads of satellite DNA, consistent with their hypothesis that this helps to hold heterologous chromosomes together in the interphase nucleus. Most of their experiments are carried out in both Drosophila and mouse, suggesting the universality of the proposed function for the chromocenter.

Although this is primarily a descriptive study, it suggests a mechanism for retention of chromosomes in the nucleus. Several important questions (not articulated by the authors) could be followed up in future studies based on these foundational observations. For example: in S. pombe the homolog of ORC4 is an AT-hook protein, thus raising the question of whether D1/HMGA1 associates with other ORC subunits in satellite DNA. Moreover, since a loss of D1/HMGA1 results in chromosome breakage, it would be interesting to know if this is correlated with a defect in DNA replication. Are the DNA breaks throughout the genome, even though D1/HMGA1 is localized in a few places? If so, how is the genome-wide breakage caused? Could the loss of D1/HMGA1 be causal for chromothripsis (shattering of the genome)? There is no need for the authors to address these and other questions in their paper, but it simply illustrates how the present paper may stimulate future experiments by the field.

Overall, the data are convincing and support the authors' model that satellite DNA at pericentromeres form chromocenters, via satellite DNA-binding proteins, to keep the nucleus intact. This paper makes an important contribution and definitely merits publication if the important suggestions below are speedily addressed.

Essential revisions:

1) Some results are not quantified, such as Figure 2J-M. We would expect some sort of quantification for results that merit inclusion in the main figure set.

2) We did not understand panels G and H in Figure 3. Some additional explanation of how the experiment was designed and how it should be interpreted would be helpful. These panels also lack quantification.

3) The live imaging in Figure 4 should have some quantification, such as how frequently budding events were observed. Also, the experiment could be more informative if it were repeated with markers for chromatin (such as histone H2B, which is commonly used) and nuclear envelope, to reveal more details about how nuclear envelope defects lead to budding and whether chromatin is initially in the bud.

4) In Figure 6 threads between chromosomes are shown. These are said to be between heterologous chromosomes in the Discussion, but we don't see data that clearly distinguish between homologous and heterologous chromosomes. If the authors have such data, they should mention it, and if not they should be clear in the Discussion that heterology is an interpretation. The model schematic (Figure 6E) could be more informative. How do the threads combine parts of different chromosomes? Do they consist of DNA from each chromosome cross-linked by D1/HMGA1 proteins? A little more detail in the schematic would help readers understand the model.

5) What is the connection between chromocenters and nuclear envelope defects? It's not obvious why chromosomes need to be linked to keep the nucleus intact. This idea is crucial for the overall message of the paper, so it merits some discussion even if we don't know the answer yet.

6) The first heading in the Results section states that D1 and HMCA1 bind satellite DNA, but the data do not show direct binding. The heading should probably be limited to what is directly shown by the data. Points supported by data in other papers can be explained elsewhere in the text.

7) Do the authors have any ideas about why the D1 phenotypes are most severe in spermatogonial cells (subsection “D1 and HMGA1 are required for organizing chromocenters”, first paragraph)?

8) Can the authors speculate about why the satellite DNA that forms chromocenters is located near the centromere? For the function of linking different chromosomes together, it's not clear why centromere localization would be important.

9) If chromocenter bundling proteins and pericentromeric satellite DNA are co-evolving (line 237), why does the D1 protein (from fly) also function in mouse cells where the satellite DNA sequences are different? If the fly protein can bind diverse satellite DNAs, then what does the co-evolution mean?

10) What happens in mitosis, when chromocenters on different chromosomes should not be linked to each other? Are the D1/HMGA1 proteins removed in mitosis, or is there some other mechanism to inactivate them?

11) Chromocenters are not always as visible in some cells (e.g., human cell lines) as they are in the mouse and fly cells in this paper. Do the authors think they are less important in these cases, or still important but not as apparent by standard DNA staining for some reason?

12) In Figure 1 and Figure 1—figure supplement 2 the authors present data on% disruption of chromocenters, but it is not clear how disruption is defined. Is there a maximum number of foci that are considered to be not disrupted, or a minimum number of foci that are considered to be disrupted? These numbers would presumably differ between Drosophila and mouse. In most of the photos the difference seems clear, but supplement Figure 1—figure supplement 2J-K is a case where having a definition of chromocenter disruption would seem to be helpful.

13) The authors' intriguing hypothesis about the function of chromocenters is welcome, but we think they should be slightly more cautious in ascribing universality to it. This should start in the title, which we think would be better as "A conserved function for chromocenters", since in Drosophila they have only shown that the AATAT satellite, not other satellites, participates in chromocenter formation. In the Introduction they say "a rationale for the very existence of pericentromeric satellite DNA is still lacking", but selfish DNA is a rationale that we think is widely accepted for some satellites. The use of a more qualified statement like " a clear rationale for the existence of most satellites is lacking" would be more accurate. On the flip side, budding yeast lacks satellites and chromocenters, yet manages to keep 16 chromosomes together, perhaps through attachment of the chromosomes to the spindle pole body. A little nuance in allowing for exceptions would not diminish the appeal of their hypothesis.

14) The authors say that D1 and HMGA1 "may possess an orthologous and conserved function". Are the proteins orthologous? A little more discussion of AT hook proteins and their phylogenetic distribution in eukaryotes would be appreciated, perhaps with a supplemental cartoon of the proteins and their AT hook domains.

15) In the Discussion, do they imagine that all chromocenters are based on AT hook proteins, or might there be other proteins that bundle multiple DNA strands of other satellites? A little more thought about the scope of applicability of their model might improve the Discussion.

eLife. 2018 Mar 26;7:e34122. doi: 10.7554/eLife.34122.020

Author response

1) Some results are not quantified, such as Figure 2J-M. We would expect some sort of quantification for results that merit inclusion in the main figure set.

Quantification for both sets of panels was reported in the text. Quantification for Figure 2J-K and Figure 2L-M can be found in the last paragraph of the subsection “Loss of D1/HMGA1 leads to micronuclei formation”.

2) We did not understand panels G and H in Figure 3. Some additional explanation of how the experiment was designed and how it should be interpreted would be helpful. These panels also lack quantification.

We apologize for the lack of clarity in our experimental design for Figure 3G-H. We sought to determine whether the DNA damage observed in the micronuclei and main nuclei of D1 mutant spermatogonia resulted in chromosomal breaks/shattering and whether these breaks occurred on specific chromosomes or satellite DNAs. To do so, we performed FISH on chromosome spreads from meiotic spermatocytes (the developmental outcome of spermatogonia) in control and D1 mutant testes. In addition, this experiment was performed in testes that were depleted for Omi, which alleviated germ cell death in the D1 mutant (Figure 3—figure supplement 1C-D). When quantified, we observed an increased frequency of chromosome breaks (though not specific to a particular chromosome or satellite DNA) that are indicated in the text (subsection “Loss of D1/HMGA1 leads to accumulation of DNA damage”, last paragraph). In Figure 3H, we show a representative image of chromosome breaks (arrowheads) from D1 mutant testes. We have added a more detailed description to the text (see the aforementioned paragraph) and to the revised figure legend to clarify these points.

3) The live imaging in Figure 4 should have some quantification, such as how frequently budding events were observed. Also, the experiment could be more informative if it were repeated with markers for chromatin (such as histone H2B, which is commonly used) and nuclear envelope, to reveal more details about how nuclear envelope defects lead to budding and whether chromatin is initially in the bud.

The reviewers have brought up a good point and we have addressed these comments by repeating the live imaging experiment in Drosophila using Df31-GFP to mark the nucleus and H2Av-RFP to label chromosomes (revised Figure 4A-B). In D1 mutant spermatogonia, this experiment clearly demonstrates that the nucleus forms a bud, followed by chromatin leaking into it, resulting in micronuclei. In addition, we have quantified all of our live imaging in the form of total imaging time (defined as number of cells x total imaging duration), total budding time (defined as total time with budded micronuclei/total imaging duration) and number of budding events, which are now described in the Materials and methods. These data are reported in the text (subsection “Micronuclei formation in D1 mutant/HMGA1 knockdown cells is due to budding from the nucleus during interphase”) and in the figure legend for Figure 4.

4) In Figure 6 threads between chromosomes are shown. These are said to be between heterologous chromosomes in the Discussion, but we don't see data that clearly distinguish between homologous and heterologous chromosomes. If the authors have such data, they should mention it, and if not they should be clear in the Discussion that heterology is an interpretation. The model schematic (Figure 6E) could be more informative. How do the threads combine parts of different chromosomes? Do they consist of DNA from each chromosome cross-linked by D1/HMGA1 proteins? A little more detail in the schematic would help readers understand the model.

This is a very fair comment and we have edited the language to appropriately reflect the experiment and the conclusions drawn from it (subsection “D1 bundles satellite DNA from multiple chromosomes to form chromocenter”, third paragraph). In Drosophila, we have clearly observed AATAT-containing DNA threads linking heterologous chromosomes using FISH experiments (e.g. Figure 6C). Allied with previous studies showing satellite DNA containing threads connecting heterologous chromosomes (1–3), we are confident that these protein/DNA threads connect heterologous chromosomes. We have also updated our model (Figure 6E), showing that D1/HMGA1 physically link satellite DNA from multiple chromosomes within chromocenters.

5) What is the connection between chromocenters and nuclear envelope defects? It's not obvious why chromosomes need to be linked to keep the nucleus intact. This idea is crucial for the overall message of the paper, so it merits some discussion even if we don't know the answer yet.

We are grateful for this thoughtful comment and we agree with the reviewers that this is an exciting direction for future research. Previous reports have shown a close relationship between heterochromatin and the nuclear envelope (NE); heterochromatin/chromocenters are frequently localized along the nuclear envelope (4). Consistently, the NE is a transcriptionally repressive environment although other possible functions of this stereotypic pattern have not been discussed. Multiple studies have demonstrated that the nuclear envelope is prone to breaches, especially in response to external mechanical forces such as cytoskeletal forces and cellular migration (5–8). Therefore, we imagine that chromosome bundling in the form of chromocenter helps maintain all the chromosomes inside the nucleus in the face of mechanical challenges. In the absence of chromocenter formation (as in D1 mutant), unbundled individual chromosomes may be subjected to mechanical forces, exerting forces in multiple directions, which may tear apart the nucleus. The Discussion section of the paper has been reworked in order to incorporate this comment (third paragraph). Citations have been added where needed.

6) The first heading in the Results section states that D1 and HMCA1 bind satellite DNA, but the data do not show direct binding. The heading should probably be limited to what is directly shown by the data. Points supported by data in other papers can be explained elsewhere in the text.

D1 and HMGA1 have been well-characterized biochemically in their ability to directly bind satellite DNA (9–13). However, this is a fair comment by the reviewers and we have edited the title to “The multi-AT-hook satellite DNA binding proteins, Drosophila D1 and mouse HMGA1, localize to chromocenters”.

7) Do the authors have any ideas about why the D1 phenotypes are most severe in spermatogonial cells (subsection “D1 and HMGA1 are required for organizing chromocenters”, first paragraph)?

We thank the reviewers for pointing out this very interesting feature of D1 mutation in Drosophila. We do not understand the cell type-specificity at this point. In our ongoing study, we have found that another chromocenter protein, exhibits a mild phenotype in gonads but a severe phenotype in imaginal discs and other diploid larval tissues. The only possibility that we can think of at this point is that chromocenter organization may be somewhat different in a cell type-specific manner, influencing the degree of phenotypic severity in different tissues. We aim to address this point in our future studies, as we understand more about chromocenter biology.

8) Can the authors speculate about why the satellite DNA that forms chromocenters is located near the centromere? For the function of linking different chromosomes together, it's not clear why centromere localization would be important.

As the reviewers have observantly pointed out, the overwhelming majority of chromocenter-forming satellite DNAs is present at pericentromeric locations on chromosomes. Although we can only speculate at this point, one possibility is the behavior of the nuclear envelope during the cell cycle. In many higher eukaryotes, the nuclear envelope disassembles during mitosis (Drosophila undergoes a ‘semi-open mitosis’, where the nuclear envelope undergoes extensive perforation, although they may look relatively intact at the level of light microscopy), which creates a challenge for nuclear envelope reassembly at the end of mitosis. Because kinetochores (i.e. centromeres) are pulled to a spindle pole, the centromere and nearby regions naturally come into close proximity at the end of mitosis. Thus, placing chromocenter-forming DNA (i.e. pericentromeric satellites) near the centromere makes sense in terms of re-establishing chromocenters by cross-linking many chromosomes and linking it to nuclear envelope reassembly. However, we feel that this is too speculative to be included in the main text of the manuscript at this point.

9) If chromocenter bundling proteins and pericentromeric satellite DNA are co-evolving (line 237), why does the D1 protein (from fly) also function in mouse cells where the satellite DNA sequences are different? If the fly protein can bind diverse satellite DNAs, then what does the co-evolution mean?

The reviewers have raised an important point regarding our ideas on the co-evolution of satellite DNAs and satellite DNA binding proteins. Please note that our experiments do not show that D1 (from Drosophila) can substitute the function of HMGA1 in mouse cells. Our result indicated that ectopically expressed D1 could bundle satellite DNA in mouse. With that said, the fact that fly D1 can bundle mouse satellite is striking, and naturally leads to a question how it is possible if satellite binding proteins and satellite DNA sequences are co-evolving. Our ongoing research suggests that chromocenter formation involves protein-protein and protein-DNA interactions (also please see response to essential revision #15). Based on this model, we speculate that a satellite DNA binding protein must evolve not only with its cognate satellite DNA but also with other satellite DNA sequences and other satellite DNA binding proteins that are important for chromocenter formation. We suggest that investigating the ability of satellite binding proteins to complement homolog function in closely related species may shed light on this fascinating topic.

10) What happens in mitosis, when chromocenters on different chromosomes should not be linked to each other? Are the D1/HMGA1 proteins removed in mitosis, or is there some other mechanism to inactivate them?

The reviewers highlight a very interesting direction for future research: clearly, chromocenters must be broken down to facilitate mitotic chromosome segregation. Consistently, we observe a significant reduction of chromosome-associated D1/HMGA1 in mitosis. Previous studies have identified that mitotic phosphorylation of HMGA1 dramatically reduces its affinity for DNA (14), which is suggestive of a cell cycle regulated mechanism for chromocenter dissolution in mitosis.

11) Chromocenters are not always as visible in some cells (e.g., human cell lines) as they are in the mouse and fly cells in this paper. Do the authors think they are less important in these cases, or still important but not as apparent by standard DNA staining for some reason?

We appreciate this insightful comment. Although chromocenters are not visible as ‘DAPI-dense foci’ in human cells, pericentromeric satellite DNA in human cells (such as HsatII) is known to be clustered into foci when visualized by FISH (15). Thus, we would suggest that human cells contain chromocenters even though they are not visible by DNA staining. This feature of human cells may reflect different abundance of pericentromeric satellite DNA, relative AT-contents, or even DNA secondary structure that might make satellite DNA refractory to DAPI staining.

12) In Figure 1 and Figure 1—figure supplement 2 the authors present data on% disruption of chromocenters, but it is not clear how disruption is defined. Is there a maximum number of foci that are considered to be not disrupted, or a minimum number of foci that are considered to be disrupted? These numbers would presumably differ between Drosophila and mouse. In most of the photos the difference seems clear, but supplement Figure 1—figure supplement 2J-K is a case where having a definition of chromocenter disruption would seem to be helpful.

We thank the reviewers for pointing this out. We observed that loss of D1/HMGA1 resulted in abnormal shape and thread-like morphology of pericentromeric satellite DNAs. Due to the thread-like appearance of satellite DNA, it is impossible to reliably estimate number of chromocenters per cell, although we stress that satellite DNA clustering is nevertheless defective. We therefore define chromocenter disruption as cells in which satellite DNA FISH adopt thread-like morphology in interphase nuclei, similar to a chromocenter scoring method utilized previously (16). This scoring method accounts for defective chromocenter formation in cells with a higher baseline of chromocenters such as C2C12 cells in Figure 1—figure supplement 2J-K. We have added this clarification to the Materials and methods section.

13) The authors' intriguing hypothesis about the function of chromocenters is welcome, but we think they should be slightly more cautious in ascribing universality to it. This should start in the title, which we think would be better as "A conserved function for chromocenters", since in Drosophila they have only shown that the AATAT satellite, not other satellites, participates in chromocenter formation. In the Introduction they say "a rationale for the very existence of pericentromeric satellite DNA is still lacking", but selfish DNA is a rationale that we think is widely accepted for some satellites. The use of a more qualified statement like " a clear rationale for the existence of most satellites is lacking" would be more accurate. On the flip side, budding yeast lacks satellites and chromocenters, yet manages to keep 16 chromosomes together, perhaps through attachment of the chromosomes to the spindle pole body. A little nuance in allowing for exceptions would not diminish the appeal of their hypothesis.

We appreciate this feedback from the reviewers and we have toned down our language in ascribing universality to satellite DNA and chromocenter function throughout the manuscript.

With regards to the title, the term ‘chromocenter’ in Drosophila is most frequently used to describe the one with highly polytenized chromosomes in the post-mitotic cells in salivary gland. We are afraid that using the term ‘chromocenter’ in the title may mislead the readers that our manuscript is about this specialized chromocenter, whereas our study rather highlights its importance in mitotically dividing cells, and its common characteristics with other cells (mitotically dividing mouse cells). Prior to our initial submission, we have undergone extensive discussion on the title, and we felt that ‘satellite DNA’ is better suited (not to mislead readers to salivary gland chromocenter), particularly because we study distinct satellites from two species. To avoid claiming too much universality, we decided to use ‘A conserved function….’ Instead of ‘The conserved function…’. Because we are not claiming that chromocenter formation is the only function of pericentromeric satellite, we hope that this title is acceptable. However, we very much welcome alternative suggestions that can further tone down implication of universality without misleading readers to salivary gland chromocenter.

14) The authors say that D1 and HMGA1 "may possess an orthologous and conserved function". Are the proteins orthologous? A little more discussion of AT hook proteins and their phylogenetic distribution in eukaryotes would be appreciated, perhaps with a supplemental cartoon of the proteins and their AT hook domains.

HMGA1 and D1 have been previously speculated to be orthologues based on their ability to bind satellite DNA and similar domain organization; they consist of N-terminal AT-hooks and a C-terminal acidic domain (9, 12, 17). However, D1 contains 10 AT-hooks whereas HMGA1 contains 3 AT-hooks, making it difficult to call them orthologous purely based on the primary protein sequences. We have bolstered this hypothesis with our data showing that D1 can localize to mouse chromocenters and enhance the clustering of mouse chromosomes. We have added slightly more detailed description on the protein primary structures in the revised text (subsection “The multi-AT-hook satellite DNA binding proteins, Drosophila D1 and mouse HMGA1, localize to chromocenters”, first paragraph).

15) In the Discussion, do they imagine that all chromocenters are based on AT hook proteins, or might there be other proteins that bundle multiple DNA strands of other satellites? A little more thought about the scope of applicability of their model might improve the Discussion.

Our current research indicates that multiple satellite DNA binding proteins function cooperatively to form chromocenters, especially in organisms with a variety satellite DNA repeats such as Drosophila. They do not seem to be all AT-hook proteins: instead they contain multiple DNA-binding motifs. Our speculation is that multiplicity of DNA binding domains may be the key (such that the protein can bind to multiple chromosome strands for bundling). We speculate that chromocenters are formed by protein-DNA as well as protein-protein interactions.

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Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Supplementary Materials

    Figure 3—source data 1. Quantification of g-H2Ax foci in mouse cells.

    g-H2Ax foci were scored in siControl and siHMGA1 transfected MOVAs cells. At least 100 cells were scored from three biological replicates. Percentage of cells containing the indicated number of g-H2Ax foci is listed in the table.

    elife-34122-fig3-data1.xlsx (9.4KB, xlsx)
    DOI: 10.7554/eLife.34122.010
    Figure 5—source data 1. Quantification of the relative expression of GFP/GFP-D1 and number of chromocenters in mouse cells.

    Images of cells expressing GFP (n=27) or GFP-D1 (n=47) were acquired from a single slide/transfection with all image acquisition parameters held constant and care taken to avoid pixel oversaturation. Total GFP fluorescence per cell was quantified using ImageJ software. Relative expression was calculated by dividing each individual fluorescence value over the maximum fluorescence value obtained over the course of the experiment. DAPI staining was used to calculate number of chromocenters per cell.

    elife-34122-fig5-data1.xlsx (9.7KB, xlsx)
    DOI: 10.7554/eLife.34122.014
    Figure 5—source data 2. Quantification of LacO-AATAT distance (nm) in cells expressing GFP-D1 and GFP-LacI-D1.

    LacO-AATAT distance (nm) was measured in spermatogonial cells expressing GFP-D1 (n=97) and GFP-LacI-D1 (n=69) using Leica LAS X software.

    elife-34122-fig5-data2.xlsx (10.3KB, xlsx)
    DOI: 10.7554/eLife.34122.015
    Transparent reporting form
    elife-34122-transrepform.docx (249.2KB, docx)
    DOI: 10.7554/eLife.34122.017

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