Heritable transcriptional defects from aberrations of nuclear architecture

Abstract

Transcriptional heterogeneity due to plasticity of the epigenetic state of chromatin contributes to tumour evolution, metastasis and drug resistance^1-3. However, the mechanisms that cause this epigenetic variation are incompletely understood. Here we identify micronuclei and chromosome bridges, aberrations in the nucleus common in cancer^4,5, as sources of heritable transcriptional suppression. Using a combination of approaches, including long-term live-cell imaging and same-cell single-cell RNA sequencing (Look-Seq2), we identified reductions in gene expression in chromosomes from micronuclei. With heterogeneous penetrance, these changes in gene expression can be heritable even after the chromosome from the micronucleus has been re-incorporated into a normal daughter cell nucleus. Concomitantly, micronuclear chromosomes acquire aberrant epigenetic chromatin marks. These defects may persist as variably reduced chromatin accessibility and reduced gene expression after clonal expansion from single cells. Persistent transcriptional repression is strongly associated with, and may be explained by, markedly long-lived DNA damage. Epigenetic alterations in transcription may therefore be inherently coupled to chromosomal instability and aberrations in nuclear architecture.

Fig. 1. Transcription defects in newly generated micronuclei.

a, Two patterns of chromosome segregation that generate a micronucleated cell (MN cell) and its sister (MN sister). Filled magenta shapes indicate the mis-segregated chromatid in the micronucleus (MN) and its sister chromatids in the primary nucleus (PN); open magenta shapes indicate sister chromatids of the other homologue. Top, a 1:3 mis-segregation generates monosomy in a MN sister cell and trisomy in a MN cell. Bottom, a 2:2 segregation generates disomy in both cells. In G2 (when cells were isolated), chromosomes in the primary nucleus are replicated but the MN chromatid is poorly replicated. Lollipops represent transcripts (open circles for transcripts from the normal homologue; filled circles for transcripts from the MN homologue; dashed lines for transcripts from the MN chromatid). b, Normalized transcription yield of each chromosome in a MN cell and sister cell pair after 1:3 mis-segregation. Filled and open bars indicate transcription from different homologues assessed by the parental haplotypes; filled magenta bars for the MN homologue (Chr.2B); open magenta bars for the normally segregated homologue (Chr.2A). Monosomic transcription of Chr.2B in the MN cell (bottom) is from the normally segregated chromatid in the primary nucleus and indicates near-complete silencing of the MN chromatid. c, Chromosome-wide silencing of an intact micronucleus generated by a 2:2 segregation, similar to b. The MN homologue is Chr.1B. d, Summary of transcription output in 21 MN cell–MN sister pairs, grouped by the status of MN nuclear envelope (NE) integrity. See also Supplementary Table 2 and Extended Data Fig. 3 for more information related to b–d e, RNAP2-Ser5ph signal (MN:PN ratios of background normalized fluorescent intensities) at the indicated time points after MN formation (left to right, n = 644, 212 and 605 from 2 or 3 experiments). Boxes indicate median ratio with a 95% confidence interval (CI), P values from two-tailed Mann–Whitney test. f, Correlation between micronuclei transcription (RNAP2-Ser5ph intensity) and nuclear pore complex density (POM121, 2 h after mitotic shake-off), (n = 334 from 3 experiments). Two-tailed Spearman’s correlation. g, Left, MN:PN ratios for H3K27ac (left to right, n = 187 and 118 from 2 experiments), analysed as in e. Right, correlation between H3K27ac and RNAP2-Ser5ph signals (2 h after shake-off; n = 187 from 2 experiments). Two-tailed Spearman’s correlation. Source Data

Fig. 2. Variably penetrant memory of MN chromosome transcription compromise after re-incorporation into a normal nucleus.

a, Example of copy number and transcriptional yield after two generations following a 1:3 mis-segregation in generation 1. The MN sister cell generates two monosomic MN nieces (N1 and N2), whereas the MN cell generates MN daughters (D1 and D2). Only one MN daughter is trisomic because the MN chromatid is poorly replicated. See Extended Data Fig. 2a for the outcome of 2:2 mis-segregations. b, Near-complete loss of transcription of a re-incorporated MN chromosome 5 (magenta) after a 1:3 mis-segregation in generation 1. Shown are the normalized transcription yields as in Fig. 1b. Chromosome 13 (green) underwent a 2:2 mis-segregation in generation 1 and displays transcription recovery after re-incorporation. See also Extended Data Fig. 7. c, Transcription output of 44 re-incorporated MN chromosomes from 37 families using Look-Seq2. Defective indicates a significant reduction in the transcriptional yield (Methods, Supplementary Table 2 and Extended Data Fig. 6). d, Transcription status of the U2OS 2-6-3 transcription reporter (n = 70 from 13 experiments). Defective indicates little or no visible MCP–Halo signal. e, Example of defective MN chromosome transcription after re-incorporation. Grey line indicates the mean and s.e.m. of the FI in controls (normal nuclei; n = 23 LacI reporters; Extended Data Fig. 8a). Red horizontal line indicates minimum detectable value in the controls. Black line indicates reporter transcription in a ruptured MN (no detectable generation 1 signal) that does not reach a normal level after re-incorporation (generation 2). f, Example of full transcription recovery, analysed as in e. Red vertical line indicates the time point of MN nuclear envelope rupture. g, Best single focal plane confocal images from a time-lapse series showing defective transcription after re-incorporation. Green, GFP–H2B; blue triangles, reporter locus; magenta triangles, MS2 reporter expression; open arrowheads, cell that enters the field, providing an adventitious MCP–Halo bleaching control. Scale bar, 5 µm. Source Data

Fig. 3. MN bodies exhibit transcription defects and extensive DNA damage.

a, Defective transcription and H3K27ac in MN bodies. Top, scheme of the experiment (time points are approximate). IF, immunofluorescence imaging. Bottom, representative images of a daughter cell with a MN body from a ruptured micronucleus. Magenta dashed lines indicate a MN body with low RNAP2-Ser5ph and low H3K27ac levels. Scale bars, 5 µm. b, Aggregate data of relative MN body fluorescence intensities (FI) for RNAP2-Ser5ph and H3K27ac as in a (left to right, n = 43 and 41 from 8 experiments). Boxes are median with 95% CI; P values from two-tailed Mann–Whitney test comparing the FI ratio between MN and control PN region in the same cell. c, Decrease in RNAP2-Ser5ph in MN bodies verified by fixed imaging. Cells were fixed approximately 45 h after mitotic shake-off. MN bodies were identified on the basis of the endogenous MDC1 signal. Data points represent relative FI of RNAP2-Ser5ph in MN bodies against control regions (n = 1,447 from 12 experiments). Boxes are median with 95% CI; two-tailed Mann–Whitney test. d, Decrease in H3K27ac in MN bodies (n = 341 from 2 experiments). e, DNA damage in MN bodies. FI measurements of γH2AX intensity (94% of MN bodies were positive, >3 s.d. above the mean of the corresponding nuclear background; n = 195 from 2 experiments). f, 53BP1 accumulation within MN bodies as in e (82% of MN bodies were positive for 53BP1; n = 211, from two experiments). Analyses in d–f are similar to c. Source Data

Fig. 4. Damaged MN bodies are more likely to have persistent transcription defects.

a, Transgenerational tracking of MN chromosome fate. Top, cartoon of the gene expressing megaDam. Bottom, scheme of manipulations that restrict DamMN expression to the first interphase when MN form. b, Representative images of re-incorporated MN with high (top) and low (bottom) DNA damage in MN bodies. Top, MN body (^m6A-Tracer, dashed magenta outline) with high γH2AX signal (top quartile, see c on the right) but low RNAP2-Ser5ph labelling. Bottom, MN body with low γH2AX signal (bottom quartile) and normal RNAP2-Ser5ph labelling. c, Aggregate data of relative RNAP2-Ser5ph intensities in MN bodies with DNA damage levels (Extended Data Fig. 10e; left to right, n = 220, 111, 220 and 112 from 4 experiments). Boxes are median with 95% CI; Kruskal–Wallis with Dunn’s multiple comparisons. d, Correspondence between high damage level and low transcriptional activity for re-incorporated MN chromosome 1 using the U2OS 2-6-3 nascent transcription reporter. Top, scheme of the experiment. During live imaging, cells with the reporter in a micronucleus were identified by LacI–SNAP and followed through cell division to identify MN body formation. Transcription activity of the reporter was tracked in live cells (MCP–Halo foci) and, after re-incorporation, was scored for γH2AX-marked DNA damage. Bottom left, MS2 signal used to detect reporter transcription activity as in Fig. 2e (grey line indicates the mean and s.e.m. of FI in controls as shown in Extended Data Fig. 8a). Bottom right, after live imaging, cells were fixed to detect γH2AX and RNAP2-Ser5ph (which correlated with the MS2 signal). Shown are representative images. Yellow arrowheads indicate MN bodies. e, Summary of the transcriptional output and presence of damage for 49 re-incorporated chromosome 1 with the reporter. The correlation between transcription defect (Fig. 2d) and DNA damage is significant (11 experiments, P < 0.0001, two-sided Fisher’s exact test). Scale bars, 5 µm (b,d). Source Data

Fig. 5. Long-term epigenetic and transcriptional consequences of exposure of a chromosome to the cytoplasm.

a, Generation of RPE-1 clones with broken chromosome 4 and control RPE-1 clones. Chromosome 4 bridges were generated by sister chromatid fusion after CRISPR–Cas9 breakage at the subtelomeres of either 4p or 4q (left, bottom to top). ATAC-seq, RNA-seq and DNA-seq were performed on the bridge clones, control clones, and bulk (parental population) controls. b, Average ATAC signal variation in 10 Mb intervals in control (n = 10) and bridge clones (n = 12). White dots indicate the median; black boxes indicate first and third quantiles; red dots indicate a 10 Mb region on chromosome 4 (27–37 Mb) with significantly reduced chromatin accessibility in bridge clones. Regions with significantly reduced ATAC intensities in the control clones (fold change < 0.65) are from pericentric regions of chromosome 1, chromosome 9 and chromosome 15 with few ATAC peaks. c, Copy number normalized ATAC-seq peak intensities in chromosome 4 (26–38 Mb) in the parental clone, control clones and bridge clones. The region with an asterisk in bridge clone III has DNA copy number zero from homozygous deletion and is masked. See Extended Data Fig. 11 for haplotype-specific DNA copy number alterations and rearrangements in bridge clones. There is a significant reduction in the ATAC signal in 6 out of 12 clones (I–V and XII) (P < 0.0001, one-sided permutation test; Methods). Bridge clones are ordered by the level of PCDH7 expression (ascending), the only gene with significant expression within this region. Arrow indicates the PCDH7 promoter. d, Correlation between PCDH7 expression (log-transformed transcripts per million ratio) and chromatin accessibility in the promoter and gene body of PCDH7 (normalized ATAC peak intensity) in control (green dots) and bridge clones (purple dots). Four bridge clones displaying a reduction in both chromatin accessibility and gene expression are on the lower left labelled with sample identifiers.

Extended Data Fig. 1. Overview of experimental and analytical workflows.

(a) Scheme of Look-Seq2. There are two key improvements compared to the original Look-Seq. First, live-cell imaging starts before the first cell division that leads to micronuclei; this enables tracking, isolation, and transcriptome analysis of both the MN cell and its sister cell (“generation 1”). Moreover, we can image cells over two cell divisions (“generation 2”) and analyze both the daughters of the MN cell (“MN daughters”) and the daughters of the MN sister cell (“MN nieces”). The second improvement is that single cells are isolated using a new capture strategy with minimal mechanical perturbation that is illustrated in (b). (b) Second generation experimental strategy for single-cell capture and sequencing. We adapted a previously developed LCM system (Palm Microbeam, Carl Zeiss) and re-designed the imaging and capture setup. The modifications enable the inversion of the membrane rings relative to the microscope objective. This allows medium to be present continuously throughout capture, which provides more time for the capture of family member cells. The setup is also compatible with laser catapulting into 96 well plates, which further increases throughput. See Methods for details. (c) Two measures of transcription yield from single-cell RNA-Seq data: (1) The total transcriptional yield is assessed by the transcripts per million (TPM) calculated from all RNA-Seq fragments overlapping with annotated coding regions. (2) The fraction of transcripts derived from each parental homologue is estimated from the counts of haplotype-specific sequencing reads. The haplotype-specific transcription yield is estimated by multiplying the total transcriptional yield by the haplotype fraction of transcripts. The transcription level of each gene in a single cell is further normalized by its mean in normal RPE-1 cells to obtain the normalized transcription of each gene. Details of the computational analysis are provided in Methods. (d) Normal range of transcriptional variation of each parental homologue derived from single-cell RNA-Seq data of control RPE-1 cells (n = 198; for Chr.12 n = 190 after excluding trisomies). Shown are the range of mean transcription of each chromosome (mean TPM ratio across all genes on a chromosome in each cell; shaded boxes) and the range of haplotype-specific transcription (mean haplotype-specific TPM ratio across all genes on a chromosome in each cell, open boxes) calculated from the total transcription and the haplotype fractions. Box plots indicate the 1st (bottom edge) and 3rd (top edge) quartiles and the median (horizontal line), with whiskers indicating 1.5x the interquartile range. The range of total transcriptional variation is used to estimate the range of normal disomic transcription (i.e., transcription of two copies of a chromosome, either from one copy of both parental homologues or from two copies of one homologue); the range of haplotype-specific transcriptional variation is used to estimate the range of normal transcription from each parental homologue. For the trisomic Chr.10q segment (61Mb-qter), the two haplotype-specific TPM ratios reflect the transcriptional output of the single-copy homologue (A) and the duplicated homologue (B); for Chr.X, the haplotype-specific TPM ratios reflect the transcriptional output of the active X (Xa) and the inactive X (Xi). For the 10q segment and Chr.X, the haplotype-specific TPM ratios are calculated by normalizing the TPM ratio of the intact 10q (A homologue) and Xa to 1. The duplicated 10q segment is appended to the q-terminus of the active X.

Extended Data Fig. 2. General strategy for the inference of haplotype-specific DNA copy number and chromosome mis-segregation events from single-cell RNA-Seq data.

(a) Two segregation patterns of MN chromosomes (left: 2:2 segregation; right 1:3 segregation) generated by nocodazole-block-and-release and the predicted copy-number outcomes over two generations. MN chromatids (filled magenta) and chromatids of the other haplotype (open magenta) are represented in the same fashion as in Figs. 1 and 2. Under 2:2 segregation, the MN sister cell or the MN nieces (dashed boxes) should display bi-allelic disomic transcription but one of the two MN daughter cells (shaded boxes) should display mono-allelic transcription of the intact haplotype (open magenta) due to deficient replication of the MN chromatid; under 1:3 segregation, the MN sister cell or the MN nieces should display mono-allelic transcription from the intact haplotype (open magenta). The predicted monosomic transcription outcomes are used to identify micronuclear chromosomes and their segregation pattern in each experimental family. (b) and (c) Validation of the transcriptional outcomes of MN (“generation 1”) using an experimental strategy (b) of inducing MN with acentric Chr.5q fragments generated by CRISPR-Cas9 as reported in our recent study. The data shown in (c) demonstrate the predicted transcriptional outcome when the micronucleus contains only one copy of Chr.5q fragment arm that most closely resembles the segregation patterns generated by nocodazole block-and-release. Two measures of gene transcription are shown: in the left plot, filled and open magenta circles are the normalized allelic expression of the broken and the intact haplotype in 10 Mb bins; on the right are the cumulative TPM (from low to high expression). The MN sister cell shows normal disomic transcription. In the MN cell, monoallelic transcription of the MN haplotype (filled circles) extending from near 64 Mb to the q-terminus indicates silencing of an acentric Chr.5 fragment partitioned into the micronucleus after Cas9-breaks generated at ~64 Mb. Reduced transcription of Chr.5 in the MN cell is also evident from the cumulative TPM plot on the right that shows a reduction in total transcription relative to normal disomic Chr.5. As the cumulative TPM plot is generated for all genes on Chr.5, it does not distinguish chromosome-wide transcriptional reduction from regional loss of transcription. (d) Identification of chromosomes with non-reference transcriptional states (red dots) in MN families (n = 173 cells) based on reference transcription distributions determined from control RPE-1 cells (Extended Data Fig. 1d). Based on the inferred DNA copy-number states of these chromosomes (assuming proportional transcriptional yield and DNA copy number), we further identify chromosomes with mis-segregation patterns consistent with the predicted outcomes in (a). Error bars represent normal range of transcription estimated based on the 5 % and 95 % values in control cells. Red dots represent chromosomes with significant deviations (Bonferroni corrected P <0.05, two-tailed Z-test; for 48 chromosomes including both homologues).

Extended Data Fig. 3. Additional data on the loss of transcription in newly generated (generation 1) MN.

(a) Summary of the transcriptional yield of MN chromosomes in all generation 1 families. Each bar plot represents the average transcriptional yield of both homologues of the MN chromosome (filled for the micronuclear homologue that is annotated below each plot; open for the intact homologue) in the MN cell (left) and the MN sister (right) in each family. Two families with near identical transcription from all chromosomes (indicating normal transcription yield of the MN chromosome) are not shown. In family F98, the MN haplotype (Chr.4B, green) displays reduced transcription in the MN cell; in all the other families, the MN haplotype in the MN cell displayed near complete silencing as indicated by either near complete loss of transcription (2:2 segregation, left) or close to monosomic transcription (1:3 segregation, right) from the intact sister chromatid of the MN haplotype in the primary nucleus. In family F71, the transcriptional imbalance is restricted to the 1p arm with near complete silencing (see Supplementary Table 2). In family F230 and F203, the transcriptional pattern indicates an extra copy of the MN homologue that is shared between the MN cell and the MN sister reflecting a pre-existing duplication of the MN homologue (i.e., a pre-existing trisomy). The examples shown in panels b (F220) and c (F216) are highlighted. (b) Chromosome-wide transcriptional data of both Chr.2 haplotypes (left) in the MN family shown in Fig. 1b (F220) and plots of cumulative TPM from low to highly expressed genes (right) plots that validate the inference of monosomic and disomic Chr.2 transcription. (c) Chromosome-wide transcriptional data of both Chr.1 haplotypes in the MN family shown in Fig. 1c (F216) and cumulative TPM plots that validate the inference of monosomic and disomic Chr.1 transcription.

Extended Data Fig. 4. Transcription defects and chromatin modifications in newly formed (generation 1) micronuclei.

(a) Transcription is present at a reduced level in intact MN and is nearly absent in ruptured MN. Data points are background normalized MN:PN fluorescence intensity (FI) ratios of RNAP2-Ser5ph at 23 h post mitotic shake-off. RFP-NLS levels were used to assign the micronuclei in the two groups (n = 83 and 82, left to right, from three experiments). Micronuclei with NLS ratios below 0.1 relative to the PN were considered ruptured and above 0.3 were considered intact. Median with 95% confidence interval (CI); Two-tailed Mann–Whitney test. (b) Data from Fig. 1e, but instead of the MN:PN FI ratios what is shown here are the background normalized intensity values at 2, 6 and 23 h post release from nocodazole and mitotic shake-off (n = 644 for 2 h, 212 for 6 h and 605 for 23 h, from two or three experiments). Median with 95% CI; Kruskal-Wallis with Dunn’s multiple comparisons test. (c) In U2OS cells, active transcription (RNAP2-Ser5ph) is also reduced in newly formed MN. Performed and analyzed as in Fig. 1e (n = 88 and 104, left to right, from two experiments). (d) Representative images from the data shown in Fig. 1e. Yellow arrows indicate micronuclei. Scale bars 5 µm. (e) Independent confirmation of the MN transcription defect by 30 min EU pulse labeling. Left, representative images of S/G2 cells with MN (generation 1). Right, correlation between RNAP2-Ser5ph and EU levels. Cells with varying levels of RNAP2-Ser5ph intensity were selected and then EU intensity levels were measured (n = 37, from one experiment). Note the strong EU signal in the nucleoli which lack RNAP2-Ser5ph, because rDNA is transcribed primarily by RNA polymerase I and RNA polymerase III. Two-tailed Spearman’s correlation. Scale bar 5 µm. (f) MN transcription defects verified in MN generated by G2 arrest with CDK1 inhibition, followed by release into an MPS1 inhibitor. This synchronization and MN induction method differs from the nocodazole block and release protocol primarily used in this study because it shortens rather than lengthens mitosis (excluding hypothetical artifacts from prolonged mitotic arrest). Left: scheme of the experiment. RPE-1 cells were analyzed 2 h after release from the G2 block (n = 334, from three experiments). Right: quantification and analysis of the results as in Fig. 1e. (g) Transcription and chromatin defects in spontaneously generated micronuclei. Decreased levels of RNAP2-Ser5ph and H3K27ac in spontaneous micronuclei of untreated RPE-1 (left) and U2OS cells (right). Performed and analyzed as in Fig. 1e (n = 134 and 295, left to right, from two experiments). (h) Representative images from data shown in Fig. 1f. Yellow arrows indicate micronuclei with nucleoporin signal (POM121) and transcription (RNAP2-Ser5ph). In contrast, red arrows indicate a micronucleus with decreased nucleoporin signal and much lower transcription signal. Note that we evaluated the specificity of POM121 staining by confocal microscopy, showing the typical nuclear pore complex dot-like pattern at the nuclear surface and the increased rim signal at the nuclear periphery by imaging a focal plane in the middle of the cell. Scale bar 5 µm. (i) Reduced accumulation of CDK9 and CDK12 in the micronuclei. The levels of CDK9, CDK12 and RNAP2-Ser5ph were analyzed in micronuclei 23 h post release from a nocodazole block followed by a mitotic shake-off. The experiment was performed and analyzed as in Fig. 1e (n = 285 and 291, left to right, from two experiments). An analysis of intact micronuclei also showed the defective accumulation of both CDK9 and CDK12 (MN:PN ratios: 0.35 for CDK9 and 0.09 for RNAP2-Ser5ph;  n = 111, P < 0.0001; 0.28 and 0.08 MN:PN for CDK12 and RNAP2-Ser5ph groups, respectively, n = 102, P < 0.0001; from two experiments). Source Data

Extended Data Fig. 5. Epigenetic alterations in micronuclei.

(a) Modest increase of repressive chromatin marks in a subset of late S/G2 MN. Performed and analyzed as in Extended Data Fig. 4a (n = 129, 179, 114 and 105, left to right, from two experiments for H3K9me2; from one or two experiments for H3K27me3). (b) Left, selected example images from (a) of cells with ruptured MN that show apparent enrichment for H3K9me2 and HK27me3 in the MN. Arrowheads: micronuclei lacking normal RFP-NLS accumulation. Right: related to (a) but comparing intact and ruptured MN for H3K9me2 (n = 48 and 40, left to right, from two experiments) and H3K27me3 (n = 35 and 26, left to right, from two experiments) at 23 h post mitotic shake-off. Performed and analyzed as in Extended Data Fig. 4a. Scale bars 5 µm. (c) Loss of H3K9ac and H3K27ac in MN at the indicated timepoint during interphase. Left: representative images of the indicated histone modifications at 2 h post mitotic shake-off. Right: quantification and analysis of data for H3K9ac as in Fig. 1e (n = 148 and 124, left to right, from two experiments). Scale bars 5 µm. (d) HDAC inhibition is not sufficient to rescue the transcription defect of chromosomes in micronuclei. Cells were analyzed after incubation with a pan-HDAC inhibitor (SAHA) for 23 h post release from a nocodazole block followed by a mitotic shake-off (n = 302 and 335, left to right, from three experiments for both H3K27ac and RNAP2-Ser5ph). For the intact micronuclei, a significant rescue of H3K27ac levels was also observed after HDACi treatment (0.36 and 0.87 MN:PN for H3K27ac in DMSO and HDACi groups, respectively, P < 0.0001) and this was also not detectably accompanied by rescue of the transcription defect (0.14 and 0.07 MN:PN for RNAP2-Ser5ph in DMSO and HDACi groups, respectively) (n = 165 and 131 for both H3K27ac and RNAP2-Ser5ph). Performed and analyzed as in Fig. 1e. All pairwise comparisons between DMSO and HDACi have P < 0.0001. For the intact MN, all pairwise comparisons between MN and PN have P < 0.0001, except for the HDACi of H3K27ac group that has P = 0.502. Source Data

Extended Data Fig. 6. Summary of the transcriptional yield of reincorporated MN chromosomes in all generation 2 families.

Each bar plot shows the transcriptional yield of both homologues of the MN chromosome (filled for the MN homologue that is annotated below each plot; open for the intact homologue) in the MN daughters (two on the left) and one or both MN nieces (on the right) in a family. Families are grouped based on the status of MN nuclear envelope integrity (left: NE disruption during generation 1 interphase; right: intact NE) and the segregation pattern of MN chromosomes (top, 2:2; bottom 1:3). Nine families (NE disruption: F12, F24, F34, F37, F154, F155, F281, F34; intact NE: F25) without MN nieces are shown separately from the remaining families with MN nieces. MN chromosomes with near normal transcriptional yield are shown in green: Under a 2:2 segregation, the MN haplotype displays a transcriptional ratio of 1:0 between the MN daughters and normal (monosomic) transcription in the MN nieces; under a 1:3 segregation, the MN haplotype displays a transcriptional ratio of 2:1 between the MN daughters and complete transcriptional loss in the MN nieces. (See Extended Data Fig. 2a for the segregation patterns.) MN chromosomes with significantly reduced transcription are shown in magenta. For these chromosomes, the MN haplotype shows a statistically significant lower transcription than normal transcription (monosomic transcription under 2:2 segregation and disomic transcription under 1:3 segregation, two-sided z-test). In families F24, F205, F259, and F37, we identified transcription of the MN haplotype in both daughter cells that is consistent with chromosome fragmentation; we combined the transcriptional yield in both MN daughters in these samples to assess the normality of transcription of reincorporated MN chromosomes. All MN chromosomes with deficient transcription are associated with NE disruption. The summary bar charts of normal and deficient transcription in 2:2 segregation samples and 1:3 segregation samples only include MNs with the predicted patterns of transcriptional imbalance. Deviations from the predicted patterns are explained below. In family F233 and F261, the presence of an extra copy of the Chr.12A homologue in all family members indicates a pre-existing duplication of Chr.12A that is a frequent alteration in RPE-1 cells. In family F12, we inferred the MN chromosome to be the active X (the transcription yield of the inactive X is not shown) that also contains a duplicated 10q segment. In seven families (F236, F231, F25, F189, F238, F281, F34), we inferred that the MNs contain only a chromosome arm; in family F238, we inferred the 1p arm was reincorporated into one MN daughter and the 1q arm was persistent in a MN niece cell based on live-cell imaging. We note that for MN chromosomes that underwent 1:3 segregations, the normality of transcription is assessed by comparing the level of MN haplotype-specific transcription to the level of total transcription of normal disomies (2). The proportional gain of transcription of duplicated homologue (2) is verified using observations from 18 spontaneous trisomies.

Extended Data Fig. 7. Complete data of the F258 family.

Data are for the family shown in Fig. 2b. (a) Haplotype-specific chromosomal transcriptional ratios showing non-disomic transcription of Chr.5 (magenta) and Chr.13 (green). The first two cells are the MN daughters; the second two are MN nieces. Regional transcription data of Chr.5 and Chr.13 are shown in (b) and (c). (b) The segregation pattern, expected transcriptional yield, and observed transcriptional levels of Chr.5 in all four cells. The presence of monosomic expression in both nieces and disomic/biallelic expression in both MN daughters indicate a 1:3 segregation of Chr.5. As the two MN daughters both display close to disomic transcription but one or both of them have reincorporated the Chr.5 copy from the micronucleus, we conclude that the reincorporated Chr.5 is not actively transcribed. (c) The segregation pattern, expected transcriptional yield, and observed transcriptional data of Chr.13 in all four cells. In contrast to the pattern of Chr.5, the two nieces both display disomic/biallelic expression, and one MN daughter displays monosomic expression; this pattern establishes a 2:2 segregation of Chr.13. The presence of transcripts phased to the MN haplotype (filled green circles in the bottom cell) indicates transcription of the reincorporated Chr.13 in the bottom cell. We note that there is more regional transcription variation in Chr.13 than in Chr.5 that is partially due to the lower gene density on Chr.13.

Extended Data Fig. 8. Analysis of nascent transcription from reincorporated micronuclei.

(a) Control U2OS 2-6-3 reporters to assess nascent transcription of normally expressing reporters in the main nucleus. Normalized FI of MS2 signal (MCP-Halo) were measured from reporters that were in the main nucleus during both generation 1 and 2 (n = 23 LacI reporters). Grey bar: mitosis. Error bars: mean +/− SEM). Red line: minimum detectable normalized MS2 value of the controls (see Methods). (b) Example of a MN with late G2 rupture in generation 1 that recovered transcription after reincorporation into a daughter nucleus in generation 2. Performed and analyzed as in (a), above. Grey line: mean intensity of the control reporters in main nucleus. (c) Aggregated data of nascent transcription from reincorporated MN assessed by the U2OS 2-6-3 reporter, similar to (a) and Fig. 2e and f. Normalized FI of the MS2 signal (MCP-Halo) were measured from reporters that were in a MN in generation 1 and then incorporated into a daughter nucleus in generation 2. Top, a subset of cells with MN that ruptured during generation 1 interphase and then recovered transcription after reincorporation into a daughter nucleus in generation 2 (n = 7 analyzed out of 19 similar cases). Note that prior to mitosis there is variable MS2 signal because of variable MCP-Halo accumulation in intact micronuclei and because of variability in the timing of MN NE rupture. Bottom, aggregate data for a similar subset of samples where the MN ruptured during generation 1 interphase and then displayed a generation 2 transcription defect after reincorporation (black line, n = 9 analyzed out of 20 similar cases). Note: (1) for ease of visualization, error bars (mean +/− SEM) are shown only for the experimental samples, but not the controls; (2) when there was no detectable MS2 signal in an experimental sample, we assigned the minimal detectable normalized value in control cells (1.7, see Methods) to this sample (This explains the complete overlap between the black and red lines after the 10-hour timepoint). (d) Images from a timelapse series for the experiment in (b), above. Green arrowhead: MN rupture. Red arrowheads: MS2 expression from the reporter after reincorporation into a daughter nucleus in generation 2. Time: hours post release from the G2 block. Scale bars 5 µm. (e) Validation that MN-bodies originate from MN chromosomes, using same-cell live/fixed imaging. Left, images from a time-lapse series (U2OS 2-6-3 system, see Figs. 2e–g, 4d). A cell with a MN harboring Chr.1 (with the reporter integrated in Chr.1p, yellow arrowheads) was identified. The MN ruptured in the interphase that it was formed. After mitosis, the MN chromosome was reincorporated into a daughter cell PN (blue arrowheads, LacI-SNAP) but was not expressed (magenta arrowheads, MCP-Halo), even though it was in a normal nuclear environment. Right, at the end of the time-lapse imaging (t = 42.5 h) cells were fixed and the same cells were analyzed by immunofluorescence microscopy, revealing a large γH2AX-positive MN-body is at the location of the reincorporated MN chromosome identified by LacI-SNAP (open magenta arrowheads). Scale bars 5 µm. (f) Validation of the method to assess MN rupture with the U2OS 2-6-3 reporter system. For all experiments with this reporter, loss of the general nuclear MCP-Halo signal (MCP-Halo contains an NLS) was used to determine the time of MN NE rupture. We verified that MCP-Halo signal loss from MN corresponds to RFP-NLS by two-color live-cell imaging in U2OS 2-6-3 cells expressing both MCP-Halo and RFP-NLS (n = 40 from four experiments; two-tailed Spearman’s correlation). Source Data

Extended Data Fig. 9. Characterization of MN-bodies.

(a) Same-cell live-fixed experiment supporting the fixed imaging shown in Fig. 3a, b. Top, scheme of the experiment. MN were induced in RPE-1 cells and the MN fate was tracked with GFP-H2B and RFP-NLS (to visualize MN NE rupture in generation 1). After most cells progressed into generation 2, they were fixed and labeled to detect γH2AX. Bottom left: representative images of a daughter cell pair, one with and one without an MN-body. Bottom right: summary of 13 cell pairs tracked and analyzed by the same-cell live-fixed experiments (from two experiments). Scale bars 5 µm. (b) MDC1 accumulation on a mitotic chromosome in an RPE-1 cell that had a micronucleus in the prior interphase (generated by nocodazole block and release), shown by immunofluorescence staining of endogenous MDC1 (representative images from two experiments). Note that the micronuclear chromosome can be identified because it is decondensed, a known feature of mitotic micronuclear chromosomes. Scale bar 5 µm. (c) Images from a timelapse series tracking damaged MN chromosomes through cell division and MN-body formation. GFP-H2B: chromosomes; green arrowheads: MN chromosome; RFP-NLS: NE integrity; blue arrowheads: MN NE rupture; red arrowheads: SNAP-MDC1-marked MN DNA damage. Time: hours post release from the nocodazole block for MN induction. Scale bars 5 µm. (d) Durations of MN-bodies assessed by live-cell imaging of SNAP-MDC1 indicate MN-bodies persist throughout most of the generation 2 interphase. Each row shows the lifetime of a MN-body (black bar) and the duration of imaging (light grey bar). In all but four cases, the MN-bodies persisted until the end of the imaging (see Extended Data Fig. 9c for an example of a time lapse series). Note that analysis of the live-cell imaging experiments showed that 68% of cells with MN-bodies were derived from mother cells with a micronucleus that ruptured, 22% were derived from mother cells with intact micronuclei and 10% from non-micronucleated mother cells. (e) Distribution of signal intensities for MN-body by immunofluorescence staining for the endogenous MDC1. Performed and analyzed as in Fig. 3c (n = 341, from two experiments). Median with 95% CI. Two-tailed Mann–Whitney. (f) Determination of the background nuclear RNAP2-Ser5ph signal in nucleoli. We measured the background RNAP2-Ser5ph signal in nucleoli (fibrillarin positive), which should lack active RNA polymerase II, and in nuclear regions lacking nucleoli. These values were then normalized to the density of fluorescence intensity from a nuclear mask excluding the nucleoli. The detection of measurable RNAP2-Ser5ph signal in the nucleoli means that we likely underestimate the extent of RNAP2-Ser5ph signal loss in MN-bodies (see Methods; n = 650, from two experiments). Median with 95% CI. Kruskal-Wallis with Dunn’s multiple comparisons test. (g) Verification of low transcription and H3K27ac loss in MN-bodies in U2OS cells. Performed and analyzed as in Fig. 3c (n = 138, from two experiments). (h) Reduced H3K9ac (left) but not H3K9me2 (middle) or H3K27me3 (right) in MN-bodies. Performed and analyzed as in Fig. 3c (n = 222, 234 and 244, left to right, from two experiments). (i) H3S10ph and H3T3ph levels show no increase but a minor decrease in MN-bodies compared to the control. Performed and analyzed as in Fig. 3c (n = 130 left; n = 124 right, from two experiments). Source Data

Extended Data Fig. 10. DamMN system characterization.

(a) Validation of the DamMN system. Shown are representative single-focal plane confocal images of RPE-1 megaDam cells ~45 h post release from the CDK1-induced G2 block at the start of the experiment (see Fig. 4a and Methods). There is no ^m6A DNA methylation if megaDam transcript is not induced (left, no Dox); if megaDam is not degraded prior to mitotic entry, all primary nuclei show ^m6A DNA methylation because of labeling during mitosis (middle, Dox, no ASV no IAA); if megaDam is degraded prior to mitosis because of size exclusion through the NE by passive import, primary nuclei are mostly not ^m6A methylated (right, Dox, +ASV +IAA). ^m6A methylation is visualized with the ^m6A-Tracer (see Fig. 4a and Methods; four experiments). Scale bars 20 µm. Note that even in the condition of degrading megaDam before mitosis (Dox, +ASV +IAA), many cells still show whole nucleus labeling with the m6A-Tracer. This could either result from cells that were in mitosis at the time of megaDam induction or from cells where nuclear exclusion of megaDam was not complete. (b) Efficient induction and degradation of megaDam. FACS analysis to detect mCherry-tagged megaDam. All samples are unsynchronized RPE-1cells with or without megaDam, with or without megaDam transcriptional induction or megaDam degradation for the indicated periods of time. The controls are RPE-1 cells lacking the megaDam construct showing no background autofluorescence without or with Dox treatment. Shown is the percentage of cells expressing mCherry (PE channel, from two experiments). (c) Western blot to detect megaDam for the indicated samples corresponding to the experiment shown in (b), above. Shown is a cropped image of a gel from the region at the megaDam molecular weight (~130 kDa). Note: the a-mCherry Ab detects non-specific background bands, but megaDam is readily distinguished from these background bands (two experiments). * indicates a background band. For gel source data, see Supplementary Fig. 1. (d) Specific labeling of MN chromosomes in mitotic cells (two examples) using the DamMN system. Top shows an MN chromosome in a prometaphase cell lacking DNA damage. Bottom shows an MN chromosome in a metaphase cell with DNA damage. Note that the MN chromosome is less condensed during mitosis, as has been previously described. Performed as described in Fig. 4a. Yellow dashed line: an MN chromosome positive for γH2AX and m6Tracer (n = 4 experiments). Scale bars 5 µm. (e) Control for Fig. 4c showing the distribution of MN-body γH2AX FI units relative to the general nuclear background (lacking nucleoli). The MN-body region of interest corresponds to the ^m6A-Tracer signal (see Methods). The γH2AX low MN-bodies were designated if the total area of γH2AX positive pixels (>3SD above background, see Methods) occupy less than 21% of the MN-body area (corresponding to the bottom quartile of γH2AX positive MN-bodies). The designation of γH2AX intermediate MN-bodies was between 21% and 65.7% of the MN-body area (the middle two quartiles), and γH2AX high was >65.7% of the MN-body area (the top quartile of MN-bodies) (left to right: n = 220, 111, 220 and 112, four experiments). Error bars: median with 95% CI. Source Data

Extended Data Fig. 11. Haplotype-specific DNA copy number, rearrangements, and transcriptional levels of chromosome 4 in 12 bridge clones.

Top: Haplotype-specific Chr.4 DNA copy number (250 kb bins) determined from newly generated DNA-Seq data (~5X mean sequencing depth) of the bridge clones. Black arcs represent intra-chromosomal rearrangements determined from previous whole-genome sequencing of the same bridge clones and subclones (10-60X mean depth). Rearrangements are phased to each homologue based on haplotype-specific copy-number changes at rearrangement breakpoints. The shaded box denotes the region of 27-37 Mb on Chr.4p with reduced ATAC-Seq signal as shown in Fig. 5c. We detected no clonal or subclonal breakpoints in this region (based on both DNA copy-number changepoints and rearrangement junctions) in bridge clones I, II, IV, VI, VII, IX, X. Bridge clone VIII and XI contain the most breakpoints within this region but display no significant change of ATAC density after normalization for copy-number variation. In bridge clone III, ATAC reduction is most prominent between 27 and 30.5 Mb and this region is far away from rearrangements that affect two segments between 32.19-32.23 Mb. Bridge clones V and XII both contain multiple copy-number and rearrangement breakpoints in this region and display modest but significant ATAC reduction (based on the permutation test). Bottom: Haplotype-specific gene transcription (TPM) ratio on Chr.4. Each dot represents the average TPM ratio of a single gene calculated from all 12 bridge clones, excluding samples with complete DNA deletion. Arrows point to the PCDH7 gene residing in the region with reduced ATAC signal.

Extended Data Fig. 12. Long-term effects on chromatin and expression after chromosome bridge formation.

(a) MN-body-like structures in the daughter cells after chromosome bridge formation. Top, schematic presentation of the experiment. Bottom left, representative images of immunofluorescence analysis for MDC1 and RNAP2-Ser5ph, showing MDC1-positive nuclear bodies (dashed magenta line) after chromosome bridge formation and cell division of RPE-1 cells expressing TRF2-DN (see Methods). We observed a high frequency of cells with MDC1-positive nuclear structures of varying size (~10%) that likely represent reincorporated chromosome bridges. This number is expected, since the frequency of chromosome bridge formation is ~30% per cell division under the conditions described in Methods. Scale bar, 5 µm. Right, quantification of the RNAP2-Ser5ph levels in the MDC1-positive nuclear structures after bridge chromosome reincorporation compared to the PN control. Performed and analyzed as in Fig. 3c (n = 309, from two experiments). (b) Genome-wide ATAC signal variation in control (i-x, left) and bridge (I-XII, right) clones in 1 Mb (top), 5 Mb (middle), and 10 Mb (bottom) intervals. The ATAC change in each interval (1 Mb increment) is assessed by normalizing the observed total ATAC signal (only from peaks) in each interval by the mean ATAC density of the null distribution generated by random permutations of individual peaks (see Methods, n = 2637 of 1 Mb genomic intervals). Bins with less than 10 ATAC peaks/Mb are excluded. Box plots indicate the first (bottom edge) and third (top edge) quartiles and the median (horizontal line), with whiskers indicating 1.5x the interquartile range. In each plot, red dots represent bins overlapping with the region of 27-38 Mb of Chr.4 that displays the most significant ATAC reduction across all bridge clones (see below). (c) Average ATAC signal variation in 10 Mb intervals across all 12 bridge clones. We only consider 10 Mb regions with 100 or more peaks. As the calculation is performed on all 10 Mb intervals with 1 Mb increment, a single region with a significant reduction in ATAC signal may result in multiple 10 Mb intervals with significant ATAC reduction; these consecutive 10 Mb bins are merged. Bins with the most significant ATAC reduction (fold change < 0.8) mostly come from two regions: Red dots are from the 4p region (26-38 Mb) shown in Fig. 5; purple dots are from a region from Chr.13q (54-76 Mb). Among 10 Mb regions with ATAC signal < 0.85, two are from Chr.4 and Chr.13: Chr.4:129-139 Mb (red circles) and Chr.13:78-94 Mb (purple circles). The other regions with ATAC signal < 0.85 are likely to have a non-epigenetic origin: Two regions (Chr.3:88-99 Mb, green dots; Chr.6:58-70 Mb, blue dots) span centromeres and have low confidence; another region on Chr.12p (12-30 Mb, light green dots) shows a similar reduction in the control clones and the variation is likely related to 12p gain or uniparental disomy that are frequent subclonal alterations in RPE-1 cells. The significant reduction in ATAC signal in Chr.4 and Chr.13 is unlikely to reflect random technical variation as they are specific to the bridge clones. For the Chr.13 region, we do not exclude a biological source for this variation, for example, an unidentified trans signaling effect that is related to bridge formation, breakage, or downstream evolution. It is known that certain genomic regions display more intrinsic variability of ATAC-Seq signals and such regions may be more prone to effects from chromosome bridge formation or breakage. Box plots indicate the first (bottom edge) and third (top edge) quartiles and the median (horizontal line), with whiskers indicating 1.5x the interquartile range. (d) Scatter plot of the fold change of ATAC signal (log2 transformed) and the P-value of ATAC signal variation estimated from permutations in bridge Clone I (two-sided permutation test, up to 5 million permutations without additional adjustment; see Methods). The two red dots are both from Chr.4:27-38 Mb. The cap of p-value at 2 x 10⁻⁷ reflects 5 million permutations performed for each interval. Source Data

Extended Data Fig. 13. Transcriptional and epigenetic consequences of micronucleation (Model summarizing the results).

Top, transcriptional outcomes of chromosomes transiently in micronuclei or chromosome bridges. Green line shows the transcriptional yield of chromosomes in MN without persistent DNA damage (generation 2). Red line shows the transcription yield of MN chromosomes with DNA damage that persists into generation 2. Bottom, cellular events leading to heritable transcription defects. Mitosis I: A cell with a lagging chromosome divides, generating the MN cell and its sister (shaded). Deficient nuclear import of MN prevents the establishment of H3K27 acetylation and causes significantly reduced or complete loss of transcription. If the MN nuclear envelope remains intact and the MN chromosome does not acquire DNA damage during mitosis (top), the MN chromosome can recover transcription after Mitosis II. If the MN chromosome acquires DNA damage either due to MN nuclear envelope rupture during interphase (bottom, red) or subsequently during mitosis (dashed arrow), the damaged chromosome may form MN-bodies with varying degrees of transcriptional silencing (bottom, with red MN-body in the primary nucleus) after Mitosis II. With partial penetrance, transcriptional silencing may persist for multiple generations, generating transcriptional heterogeneity that can be subject to selection.