the molecular mechanism underlying fibril growth
and potentially will suggest ways to interfere
with fibril formation and growth.
REFERENCES AND NOTES
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The authors gratefully acknowledge the continued support of
D. Riesner and G. Büldt. We thank P. J. Peters for advice and helpful
discussions, H. Duimel for help with sample preparation, and the
M4I Division of Nanoscopy of Maastricht University for microscope
access and support. The authors gratefully acknowledge the
computing time granted by the Jülich Aachen Research Alliance
High-Peformance Computing (JARA-HPC) Vergabegremium
and VSR commission on the supercomputer JURECA at
Forschungszentrum Jülich. Computational support and
infrastructure was provided by the Center for Information
and Media Technology (ZIM) at the University of Düsseldorf
(Germany). The authors acknowledge access to the Jülich-Düsseldorf
Biomolecular NMR Center. D. W. was supported by grants from
the Portfolio Technology and Medicine, the Portfolio Drug
Design, and the Helmholtz-Validierungsfonds of the Impuls und
Vernetzungs-Fonds der Helmholtzgemeinschaft. D. W. is a
paid scientific advisor for the Institut de Biologie Structurale (IBS),
CEA, DSV, IBS, 38027 Grenoble (France). H.H. was supported
by the Entrepreneur Foundation at the Heinrich-Heine-University
of Düsseldorf and by the Deutsche Forschungsgemeinschaft (DFG)
(HE 3243/4-1). Support from an European Research Council (ERC)
Consolidator Grant (grant agreement no. 726368) to W.H. is
acknowledged. The 4.0-Å EM density map of the Ab(1– 42) fibril
has been deposited in the Electron Microscopy Data Bank with
accession code EMD-3851; the coordinates of the atomic model
have been deposited in the Protein Data Bank under accession
code 5OQV. The NMR data have been deposited in the Biological
Magnetic Resonance Data Bank (BMRB) under accession number
27212. The authors declare no competing financial interests.
Materials and Methods
Figs. S1 to S13
Tables S1 to S7
Movies S1 to S3
References ( 22–47)
3 July 2017; accepted 24 August 2017
Published online 7 September 2017
Mitotic transcription and waves of
gene reactivation during mitotic exit
Katherine C. Palozola,1,2 Greg Donahue,2 Hong Liu, 3 Gregory R. Grant, 4
Justin S. Becker,1,2 Allison Cote, 5 Hongtao Yu, 6 Arjun Raj, 5 Kenneth S. Zaret1,2*
Although the genome is generally thought to be transcriptionally silent during mitosis,
technical limitations have prevented sensitive mapping of transcription during mitosis and
mitotic exit. Thus, the means by which the interphase expression pattern is transduced to
daughter cells have been unclear. We used 5-ethynyluridine to pulse-label transcripts
during mitosis and mitotic exit and found that many genes exhibit transcription during
mitosis, as confirmed with fluorescein isothiocyanate–uridine 5′-triphosphate labeling,
RNA fluorescence in situ hybridization, and quantitative reverse transcription polymerase
chain reaction. The first round of transcription immediately after mitosis primarily
activates genes involved in the growth and rebuilding of daughter cells, rather than cell
type–specific functions. We propose that the cell’s transcription pattern is largely retained
at a low level through mitosis, whereas the amplitude of transcription observed in
interphase is reestablished during mitotic exit.
During mitosis, chromatin condenses (1), gene regulatory machinery is largely evicted from chromatin (2– 4), and transcription is thought to be silenced ( 5–7 ). Yet reac- tivation of a specific gene expression program is needed to maintain cell identity during
exit from mitosis. Long-distance interactions
across the genome are lost during mitosis ( 8), as
is hypersensitivity at distal enhancers, but not
at promoters ( 9). “Bookmarking” transcription
factors remain bound in mitosis to a subset of
their interphase sites ( 10–15). Knockdown of
these factors during mitosis delays reactivation
of target genes ( 10, 11, 13), although the proper
transcriptome is eventually regenerated. Thus,
the basis for identity maintenance during mitosis
remains unclear, and the hierarchy by which
genes are reactivated during mitotic exit is not
Because of nuclear envelope breakdown in mi-
tosis and hence the inability to isolate nuclei for
direct labeling of transcripts ( 16), genome-wide
studies during mitotic exit used RNA polymerase
II (RNAP2) cross-linking to assess active transcrip-
tion ( 4, 17 ) and found a burst in RNAP2 binding
to promoters 60 to 90 min after release from
mitotic arrest ( 17). However, the dynamic range
of antibody-based methods is much less than
from direct measurements of nascent transcrip-
tion, and cross-linking artifactually causes pro-
tein exclusion from mitotic chromatin ( 14, 18).
Transcription elongation inhibition of prometa-
phase HeLa cells elicits paused RNAP2 at pro-
moters, suggesting the presence of elongating
enzyme, even though elongating RNAP2 was not
detected directly ( 19). The study also mapped non-
polyadenylated, chromatin-associated RNAs from
prometaphase cells, but it was unclear whether
these RNAs were transcribed during mitosis or, as
suggested by the authors, at the G2/M transition.
A study of pulse-labeled transcripts in arrested
MCF- 7 human breast cancer cells used nuclear
isolation for bromouridine- 5'-triphosphate labeling
and hence did not appear to be assessing mitotic
cells ( 20).
To define the timing of transcription events
during mitotic exit, we used the cell-permeable
5-ethynyluridine (EU) to pulse-label nascent transcripts ( 21) in intact HUH7 human hepatoma
cells during nocodazole-induced mitotic arrest,
mitotic exit, and in asynchronous cells. Arrested
cells, enriched by mitotic shake-off, were highly
pure (fig. S1, A to D) and reenter G1 (fig. S1, E
to K). Previously, we labeled transcripts with EU
during mitotic exit in HUH7 cells and attached
azide-fluorophore, discovering that bulk global
transcription initiates approximately 80 min after
nocodazole wash-out ( 11). On the basis of this assessment of global reactivation, we pulse-labeled
transcripts at 0, 40, 80, 105, 165, and 300 min after
nocodazole wash-out in HUH7 cells, but instead
conjugated azide-biotin to the EU-RNA in order
to measure the relative changes over time (Fig.
1A). The addition of biotin allowed us to use
streptavidin beads to isolate EU-labeled transcripts from total RNA and generate cDNA
libraries on the beads for sequencing (figs. S2,
A to E, and S3A and table S1). For direct comparison of transcription in asynchronous versus
mitotic cells, we designed and generated bio-tinylated RNAs to add as spike-in controls (fig.
S2, C and F to H, and tables S2 to S4).
1Institute for Regenerative Medicine, Perelman School of
Medicine, University of Pennsylvania, Philadelphia, PA
19104-5157, USA. 2Department of Cell and Developmental
Biology, Perelman School of Medicine, University of
Pennsylvania, Philadelphia, PA 19104-5157, USA. 3Department
of Biochemistry and Molecular Biology and Tulane Center for
Aging, Tulane University Health Sciences Center, New
Orleans, LA 70112, USA. 4The Institute for Translational
Medicine and Therapeutics, Department of Genetics,
Perelman School of Medicine, University of Pennsylvania,
Philadelphia, PA 19104, USA. 5Department of Bioengineering,
University of Pennsylvania, Philadelphia, PA 19104, USA.
6Howard Hughes Medical Institute, Department of
Pharmacology, University of Texas Southwestern Medical
Center, Dallas, TX 75390, USA.
*Corresponding author. Email: firstname.lastname@example.org