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RESEARCH ARTICLE
Chromatin dynamics and the transcriptional
competence of HSV-1 genomes during lytic
infections
MiYao HuID
1,2☯
, Daniel P. DepledgeID
3☯¤
, Esteban Flores CortesID
2
, Judith BreuerID
3
, Luis
M. SchangID
1,2
*
1University of Alberta, Department of Biochemistry, Edmonton, AB, Canada, 2Cornell University, College of
Veterinary Medicine, Baker institute for animal health, Ithaca, NY, United States of America, 36401Division
of Infection and Immunity, University College London, London, United Kingdom
☯These authors contributed equally to this work.
¤Current address: New York University School of Medicine, Department of Medicine, New York, NY, United
States of America.
*lms428@cornell.edu
Abstract
During latent infections with herpes simplex virus 1 (HSV-1), viral transcription is restricted
and the genomes are mostly maintained in silenced chromatin, whereas in lytically infected
cells all viral genes are transcribed and the genomes are dynamically chromatinized. Histones
in the viral chromatin bear markers of silenced chromatin at early times in lytic infection or of
active transcription at later times. The virion protein VP16 activates transcription of the imme-
diate-early (IE) genes by recruiting transcription activators and chromatin remodelers to their
promoters. Two IE proteins, ICP0 and ICP4 which modulate chromatin epigenetics, then acti-
vate transcription of early and late genes. Although chromatin is involved in the mechanism of
activation of HSV- transcription, its precise role is not entirely understood. In the cellular
genome, chromatin dynamics often modulate transcription competence whereas promoter-
specific transcription factors determine transcription activity. Here, biophysical fractionation of
serially digested HSV-1 chromatin followed by short-read deep sequencing indicates that
nuclear HSV-1 DNA has different biophysical properties than protein-free or encapsidated
HSV-1 DNA. The entire HSV-1 genomes in infected cells were equally accessible. The acces-
sibility of transcribed or non-transcribed genes under any given condition did not differ, and
each gene was entirely sampled in both the most and least accessible chromatin. However,
HSV-1 genomes fractionated differently under conditions of generalized or restricted tran-
scription. Approximately 1/3 of the HSV-1 DNA including fully sampled genes resolved to the
most accessible chromatin when HSV-1 transcription was active, but such enrichment was
reduced to only 3% under conditions of restricted HSV-1 transcription. Short sequences of
restricted accessibility separated genes with different transcription levels. Chromatin dynam-
ics thus provide a first level of regulation on HSV-1 transcription, dictating the transcriptional
competency of the genomes during lytic infections, whereas the transcription of individual
genes is then most likely activated by specific transcription factors. Moreover, genes tran-
scribed to different levels are separated by short sequences with limited accessibility.
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 1 / 37
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OPEN ACCESS
Citation: Hu M, Depledge DP, Flores Cortes E,
Breuer J, Schang LM (2019) Chromatin dynamics
and the transcriptional competence of HSV-1
genomes during lytic infections. PLoS Pathog 15
(11): e1008076. https://doi.org/10.1371/journal.
ppat.1008076
Editor: Donna Neumann, LSU Eye Center
Department of Ophthalmology, UNITED STATES
Received: April 29, 2019
Accepted: September 11, 2019
Published: November 14, 2019
Copyright: ©2019 Hu et al. This is an open access
article distributed under the terms of the Creative
Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in
any medium, provided the original author and
source are credited.
Data Availability Statement: All primary data is
uploaded in the NCBI BioProject repository and
publicly available under accession number
PRJNA550980
Funding: The research performed in the Schang
lab was funded by CIHR Operating Grants
MOP106452 and 301689, start up funds by The
Baker Institute and Cornell University College of
Veterinary Medicine and a CONCyT studentship to
EFC. The research performed in the Breuer lab was
funded by UCl/UCLH NIHR Biomedical Research
Author summary
Although chromatin epigenetics modulate transcription of the nuclear replicating DNA
viruses, and play major roles in the process of establishment of, and reactivation from,
latency, the specific mechanisms of this modulation are not totally clear. Chromatin often
regulates the transcriptional competency of cellular genes, rather than the actual level of
transcription of individual genes. Here, we show that chromatin dynamics regulate the
transcription competency of entire herpes simplex virus 1 (HSV-1) genomes, rather than
the actual transcription level of individual genes. Moreover, CTCF/ insulator containing
sequences flanking the immediate-early gene loci are more inaccessible when these genes
are highly transcribed in a context of little transcription from the rest of the genome than
when no gene was highly transcribed or all genes were. We postulate that chromatin
dynamics modulate the transcriptional competency of the HSV-1 genome. Genes in
genomes rendered transcriptionally inactive by chromatin dynamics cannot be tran-
scribed, whereas transcription of individual genes, or of group of genes, is regulated sepa-
rately in the transcriptionally competent genomes.
Introduction
Herpes simplex virus 1 (HSV-1) is a double stranded nuclear replicating DNA virus. It estab-
lishes latent infections, in which viral transcription is restricted, and lytic infections, in which
all viral genes are transcribed. The HSV-1 DNA is maintained in silenced chromatin during
latency [1–4] but is in most dynamic chromatin during lytic infections [5–7]. Consequently,
micrococcal nuclease (MCN) cleaves HSV-1 DNA into mono- to poly-nucleosome sized frag-
ments in latent infection [1], or to heterogeneously sized fragments in lytic infections [5,8–
11]. A subset of the latter fragments resolve together with cellular mono- or di-nucleosomes
during differential centrifugations [5,6], and associates with all core histones [12]. Complete
MCN digestion has suggested that several regions of the HSV-1 genome could be nucleosome-
free [13], whereas “serial” MCN protection assays indicate that HSV-1 DNA in lytic infections
is chromatinized in highly dynamic nucleosomes [5,6]. HSV-1 DNA may also associate pref-
erentially with ICP4 rather than with histones [14,15].
Transcription of the five HSV-1 immediate early (IE) genes is activated by the tegument
protein VP16 immediately after the HSV-1 genomes enter the nucleus. VP16 interacts with
the host cell factor 1 (HCF-1) and octamer-binding transcription factor 1 (Oct-1) to be
recruited to the TAATGARAT motifs on the promoters of HSV-1 IE genes[16–18]. This com-
plex then recruits chromatin remodelers to allow IE gene transcription [16,19–26]. Transcrip-
tion of the early genes (E) is activated by the IE proteins ICP4, ICP0 and ICP22. ICP4, which is
essential for the activation of transcription of E and L genes [27], recruits chromatin remode-
lers to the HSV-1 genomes [28,29] and induces histone dynamics [7]. ICP0, which enhances
IE, E, and L gene expression, induces the degradation of silencing histone variants and affects
apparent histone occupancy [30–34]. DNA replication occurs after E proteins are expressed,
and late genes (L) are transcribed after the onset of HSV-1 DNA replication. The mechanisms
preventing E and L gene transcription before IE proteins are expressed or DNA replication
starts, respectively, remain only partially understood [35,36].
Via many mechanisms such as recruitment of transcription factors or heterochromatin
proteins and nucleosome stability, chromatin normally regulates nuclear DNA accessibility
and, consequently, DNA-dependent processes such as transcription, replication or DNA
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 2 / 37
Centre. The funders had no role in study design,
data collection and analysis, decision to publish, or
preparation of the manuscript.
Competing interests: The authors have declared
that no competing interests exist.
repair [37–40]. HSV-1 DNA associates with histones bearing markers of silencing early in
infection or with markers of transcription at later times [11,19,32,41–43]. Inhibition of cer-
tain histone demethylases modulates HSV-1 transcription [44–47]. Histone acetyltransferases
(HATs) such as GCN5, PCAF, and CBP/p300 are recruited to HSV-1 chromatin by VP16 [16,
19,48], although knockdown of these HATs has limited effects on HSV-1 transcription [49].
Inhibition of histone deacetylases (HDACs), such as CLOCK, SWI/SNF and REST/Co-REST
complexes, increases HSV-1 transcription or replication [29,50–53]. Chromatin thus performs
important roles in the regulation of HSV-1 transcription, and epigenetic modifications affect
HSV-1 transcription and replication. However, the precise mechanisms of HSV-1 transcrip-
tion regulation by chromatin remain incompletely understood.
Chromatin often regulates cellular transcription competence. DNA chromatinized to
silenced states, often via the recruitment of silencing proteins to specific histone modifications
and exclusion of transcription proteins, is not accessible to transcription complexes. With the
notable exception of binding by the pioneering factors, the promoter sequences in this chro-
matin often become accessible to transcription factors only after the chromatin is remodelled
[54,55]. Binding of the promoter-specific transcription factors to the now accessible promot-
ers then determines the transcription activity of each specific gene.
Here, we explored the dynamics of HSV-1 chromatin in lytically infected cells by biophysi-
cal fractionation of serially digested HSV-1 chromatin and short-read deep sequencing analy-
ses of each fraction. Within the constraints of genome-wide population analyses, which
preclude the observation of changes in minority subsets of genomes, HSV-1 genomes were
dynamically chromatinized as units and the global dynamics of the viral chromatin were
related to transcription activity. The subpopulation of the viral genomes in the most dynamic
chromatin including fully sampled genes was enriched under conditions of active transcription
but depleted when transcription was restricted. Short sequences with limited accessibility sepa-
rated loci with different transcription levels.
Materials and methods
Cells and virus
Vero African green monkey kidney fibroblast cells (originally purchased from ATCC and main-
tained in the Schang lab since 2000) were maintained in Dulbecco’s modified minimum Eagle’s
medium (DMEM) supplemented with 5% fetal bovine serum (FBS). Low-passage (p11 to 15)
HSV-1 strain KOS was used throughout this study. Viral stocks were propagated and titrated on
monolayers of Vero cells as previously described [5,6,56]. Vero cells were infected with 10 plaque
forming unit (pfu) per cell in serum-free medium as previously described [5,6,56], with modifi-
cations. Briefly, Vero cells were inoculated for 1 h at 37˚C, inocula were then removed, and the
cells were washed twice with 4˚C phosphate-buffered saline (PBS– 1 mM KH
2
PO
4
, 154 mM
NaCl, 3 mM Na
2
HPO
4
[pH 7.4]) or SFM. Fresh pre-warmed DMEM-5% FBS was added to the
infected cells, which were then harvested at 7 hours post-infection (hpi) unless otherwise stated.
Drugs
Cycloheximide (CHX) 5 mg/ml stock solution was prepared in serum free medium (SFM) and
diluted to 50 μg/ml in DMEM-5% FBS, SFM or PBS before use. Cells were pre-treated with
50 μg/ml CHX in DMEM-5% FBS for 1 h prior to infection. CHX was maintained in the inoc-
ula, PBS or SFM washes and DMEM-5% FBS until harvest.
Phosphonoacetic acid (PAA) 100 mM stock solution was prepared in SFM and adjusted to
pH 7.0 with 1 N NaOH. The stock was diluted to 400 μM in DMEM-5% FBS and added to the
cells after removing inocula and washing with PBS.
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 3 / 37
Roscovitine (Rosco) 100 mM stock solution was prepared in dimethyl sulfoxide (DMSO).
The stock was diluted to 100 μM in pre-warmed DMEM-5% FBS and added to the cells after
removing the inocula and washing with PBS or SFM.
Chromatin extraction
Chromatin of Vero cells was extracted as described [5,6], with minor modifications. Briefly,
each 1 x 10
7
cells were washed with 4˚C PBS, trypsinized, and resuspended in 15 ml of
DMEM-5% FBS. Cells were then pelleted and resuspended in 15 ml 4˚C PBS. Cells were pel-
leted again, and resuspended in 15 ml of 4˚C reticulocyte standard buffer (RSB, 10 mM Tris
[pH 7.5], 10 mM NaCl, 5 mM MgCl
2
). Cells were pelleted again, resuspended in 10 ml 4˚C
RSB and incubated on ice for 10 min. The cell membrane was lysed by adding 0.5 ml 10% (vol/
vol) triton X-100 to every 10 ml of cells in RSB and incubating on ice for 10 min, flipping the
tubes upside down thrice. Nuclei were then pelleted by centrifugation at 3,250 xg for 25 min at
4˚C.
Each 1 x 10
7
nuclei were resuspended and lysed in 1.5 ml chromatin extraction buffer (1
mM ethylene glycol-bis(beta-aminoethyl ether)-N, N, N’, N’,- tetra-acetic acid (EGTA), 2%
Triton X-100, 3 mM MgCl
2
, 2 mM Tris [pH 8]) on ice for 30 min. Every 5 min, nuclei were
mixed by pipetting up and down 10 times using a P1000 at a setting of 750 μl. Chromatin was
spun down at 2,000 xg for 10 min at 4˚C, and resuspended in 200 μl per 1 x 10
7
cells MCN
digestion buffer (10 mM Tris [pH 8], 1 mM CaCl
2
).
Nuclease protection assay
Serial MCN digestion was performed basically as described [5,6]. Briefly, 5% of the chromatin
suspension was taken as unfractionated and undigested chromatin, directly subjected to pro-
teinase K digestion (1mg/ml proteinase K with 0.5% SDS) overnight at 37˚C and purified by
phenol: chloroform extraction and ethanol precipitation. The DNA purified from this no-
MCN digested chromatin was then sonicated into smaller fragments prior to resolution in aga-
rose gels or sequencing. The remaining chromatin was pelleted at 2,000 xg for 5 min. The
chromatin pellet was disrupted by racking on a rack, resuspended in 40 μl of MCN digestion
buffer with 0.05 U MCN per each 1 x 10
7
cells, and immediately spun down at 800 xg for 5
min. The supernatant (soluble chromatin) was removed and the MCN was immediately
quenched with 10 μl of 0.05 M EGTA to prevent further digestion of the DNA released in the
soluble chromatin. The pellet (insoluble chromatin) was disrupted as before and resuspended
with fresh 40 μl of MCN digestion buffer with 0.05 U MCN per each 1 x 10
7
nuclei. The entire
procedure was repeated six times. The supernatants from the six rounds of digestion were
pooled and resolved together in sucrose gradients.
Encapsidated or protein-free HSV-1 DNA
Vero cells were infected with HSV-1 at a multiplicity of infection (moi) of 5. After inoculation
for 1 h at 37˚C, the inoculum was removed, the cells were washed and fresh DMEM-5% FBS
was added to the infected cells. The supernatant was collected after 24 h, to minimize contami-
nation with cellular DNA from lysed cells. Virions were spun down at 10,000 xg for 2 h, resus-
pended in serum free media (SFM), and titrated. HSV-1 membranes were lysed with 0.5%
Triton X-100 on ice for 10 min by pipetting up and down using P1000 at the setting of 500 μl.
The capsids were then spun down at 15,000 xg for 1 h.
For deproteinized HSV-1 DNA, an aliquot of the purified HSV-1 capsids were subject to
proteinase K digestion (10 mg/ml proteinase K, 1% SDS) overnight at 37˚C. Deproteinized
HSV-1 DNA was purified by phenol-chloroform extraction and ethanol precipitation, and
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 4 / 37
then resuspended in TE (10 mM Tris (pH8.0), 1 mM ethylenediaminetetraacetic acid (EDTA)
buffer.
Sucrose gradient ultracentrifugation
Continuous 0 to 10% sucrose gradients were prepared using a Gradient Master instrument
with sucrose gradient buffer (SGB) (10 mM Tris [pH 8.0], 1.5 mM MgCl
2
, and 0.5 M EGTA)
containing 200 mM NaCl. Soluble chromatin was loaded on top of the gradients and centri-
fuged at 200,000 xgin a SW-40 Ti rotor for 180 min at 4˚C. Approximately 1 ml fractions were
then collected from the bottom of the tube. The pellet was recovered in 1 ml of STE (10 mM
Tris [pH 7.5], 100 mM NaCl, 1 mM EDTA) after the removal of the final fraction.
To analyze the DNA, each fraction was subjected to proteinase K digestion (1mg/ml, 0.5%
SDS) at 37˚C overnight. The DNA was extracted by phenol-chloroform, precipitated with iso-
propanol, washed twice with 70% ethanol, and resuspended in TE (10mM Tris, 1mM EDTA,
pH8.0) buffer. Ten percent of the DNA in each fraction was resolved in 2% agarose gels and
visualized by incubating in 0.1mg/ml EtBr for 1h. The remaining DNA was sealed and shipped
on ice packs for sequencing.
To analyze proteins, each chromatin fraction was subjected to trichloroacetic acid (TCA)
precipitation, to a final concentration of 25% (V / V), incubated on ice for 30 min. Fractions
were spun down at 20,000 xg for 30 min at 4˚C, and the pellet was washed twice with -20˚C
acetone. The final protein pellet was resuspended in loading buffer (2% SDS, 10% glycerol, 60
mM Tris (pH 7.4), 0.01% bromophenol blue, 10 mM DTT), and denatured at 100˚C for 10
min. Proteins were resolved in 14% SDS gels for resolution of histones, or 10% SDS gels for
resolution of ICP4 or ICP8, at 15 V for 1 h and then 30 V until the front dye ran out of the gel.
Hybridization
After visualization, agarose gels were soaked in 1 M HCl for 30 min on a shaker at room tem-
perature, rinsed with double distilled water, and soaked in alkaline transfer buffer (0.4 N
NaOH, 1 M NaCl) for 15 min at room temperature on a shaker. The alkaline transfer buffer
was replaced for another 15 min at room temperature. Gels were rinsed with double distilled
water, and soaked in neutralizing buffer (1 M Tris (pH7.4), 1.5 M NaCl) for 15 min at room
temperature on a shaker. BioDyne B0.45 nylon membranes were cut to size and soaked in 10x
SSC buffer (1.5 M NaCl, 150 mM trisodium citrate, pH 7.0). DNA in the agarose gels was capil-
lary transferred according to standard protocols. Membranes were prehybridized with 10 ml
rapid hybrid buffer (Amersham Biosciences, Piscataway, NJ) for one hour at 75˚C in a rotating
oven. The probe (HSV-1 EcoRI restriction fragment JK, [5,6]) was labeled by random priming
according to the manufacturer’s instructions (Amersham Biosciences). The denatured probe
was added to 5 ml of rapid hybrid buffer pre-warmed at 75˚C, and hybridized in the rotating
oven for 4 h at 75˚C. Membranes were washed twice for 15 min with hybridization wash buffer
1 (75 mM NaCl, 7.5 mM sodium citrate, 0.5% SDS) at room temperature, rinsed with distilled
water, and exposed to Kodak PhosphorImager screens. Signal was quantitated in a Bio-Rad FX
molecular imager.
Western blot
Gels were soaked in transfer buffer (25 mM Tris, 190 mM glycine) for 30 min. PVDF mem-
branes were cut to size, activated in 100% methanol for 5 min, and soaked in transfer buffer
for 20 min. Proteins were transferred to the PVDF membranes in a TransBlot Turbo transfer
system (Bio-Rad) at 1 A for 30 min. PVDF membranes were air dried and re-activated with
100% methanol for 2 min prior to blocking. Membranes were blocked in 10 ml Odyssey
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 5 / 37
blocking buffer (Cat. 427–40000) for 1 h at room temperature with shaking. Antibodies against
histone H2A (Abcam, rabbit, Cat. ab88770), H2B (Abcam, mouse, Cat. ab52484), H3 (Abcam,
rabbit, Cat. ab1791), H4 (Abcam, mouse, Cat. ab31830) were diluted 1:1000 in 10 ml PBS with
0.05% tween-20. Antibodies against ICP4 (Virusys, mouse, Cat. P1101), VP5 (Virusys, mouse,
Cat. HA018) and ICP8 (Abcam, mouse, Cat. ab20194) were diluted 1:500, 1:250, or 1:50,
respectively, in the same buffer. Membranes were probed for 2 h at room temperature for his-
tones, or for 18 h at 4˚C for ICP4, VP5 and ICP8, and then washed twice for 15 min each with
10 ml PBS at room temperature. Membranes were incubated with goat anti-mouse (680LT),
goat anti-mouse (Alexa 488), or anti-rabbit (800LT) secondary antibodies diluted 1:1000 in 10
ml PBS, 0.05% tween-20 for 1 h at room temperature. Membranes were washed once with 10
ml PBS, 0.05% tween-20 for 15 min, and once with 10 ml PBS for 10 min. Secondary antibod-
ies were visualized in a Li-Cor Odyssey system.
Chromatin Immunoprecipitation
The samples containing soluble, insoluble, and undigested chromatin where adjusted to 540 μl
with MCN-digestion buffer (10 mM Tris [pH 8], 1 mM CaCl
2
) and cross-linked by adding
135 μl of buffered 5% formaldehyde solution (5% CH
2
O, 100 mM Na
2
HPO
4
pH 8.67), to a
final concentration of 1%, for 10 min at 4˚C with constant mixing. Cross-linking was
quenched with 45 μl of 2 M glycine, to a final concentration of 125 mM, for 5 min at room
temperature. The soluble chromatin was subjected to sucrose gradient fractionation, whereas
the insoluble and undigested chromatin was centrifuged at 4,000 xg for 10 min at 4˚C, resus-
pended with 1 ml of chromatin immunoprecipitation binding buffer and kept on ice. Fraction-
ation was analysed by gel electrophoresis (5% of each fraction).
The insoluble chromatin, the undigested chromatin, and the chromatin fractions were dia-
lyzed against ChIP binding buffer (1% Triton X-100, 10 mM Tris pH 8.0, 150 mM NaCl, 2
mM EDTA pH 8.0) overnight at 4˚C. The insoluble chromatin, the undigested chromatin and
fractions 1–6, which contained chromatin fragments longer than 1,500 bp, were sheared by
sonication. Histone H3 antibody (ab1791; 0.375 μg) or nonspecific IgG (ab171870; 0. 375 μg)
per ChIP reaction were conjugated to 56 μg of Dyna Beads (Invitrogen 10001D) for ~20 hrs at
4˚C with constant rotation. After setting aside 2% to evaluate the input DNA, equal amounts
of each fraction were incubated with H3 or IgG bead-ab complexes for ~25 hrs at 4˚C with
constant rotation. Immunopurified complexes were rinsed consecutively with 1 ml of washing
solution 1 (1% Triton, 2 mM EDTA, 20 mM Tris pH 8.0, 150 mM NaCl), 2 (2 mM EDTA, 20
mM Tris pH 8.0, 150 mM NaCl), 3 (1% Triton, 2 mM EDTA, 20 mM Tris pH 8.0, 500 mM
NaCl), 4 (2 mM EDTA, 20 mM Tris pH 8.0, 500 mM NaCl), 5 (1% NP-40, 1% NaDOC, 1 mM
EDTA, 10 mM Tris pH 8.0, 250 mM LiCl), and 6 (1 mM EDTA, 10 mM Tris pH 8.0, 250 mM
LiCl). Immunopurified complexes were eluted with 600 μl of ChIP elution buffer (10 mM
Tris-HCl pH 8.0, 150 mM NaCl, 1 mM EDTA, 0.5% SDS) and 6 μl of proteinase K (20 mg/
ml), and then incubated at 65˚C for ~19 hrs to decrosslink and deproteinize. DNA was ex-
tracted using phenol-chloroform, isopropanol precipitated, ethanol washed, and subjected to
saturating PCR (Fast SYBR Green master mix) with 300 nM each the of UL25 (5’-AGCCGTC
CCTTCGGAGGCCA-3’, 5’-TCGGCCCGGTACAGCGGAAGCAC-3’), gE (5’-CCACCCCCG
ACCCCAGCCGA-3’, 5’-GGGGTCCCGCTGGCGGGAGT-3’), ICP4 (5’-CTGTCGCGACGA
GACGGCGT-3’, 5’-GCCCCCGCGGCGGGCACCGA-3’) or GAPDH (5’- CCTCCTGCACC
ACCAACTGCTT -3’, 5’- GTCCCATTCCCCAGCTCTCATACC -3’) primers, or 75 nM each
of the UL46 (5’-TCCTCGTAGACACGCCCCCCGT-3’, 5’-ACGCCCCCTACGAGGACGAC
GAGT -3’) primers. Purified DNA from the insoluble chromatin and the undigested chroma-
tin were diluted 1:10 before PCR. PCR conditions: 20 sec denaturation at 95˚C followed by 28
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 6 / 37
cycles of 3 sec denaturation at 95˚C and 30 sec annealing/elongation at 60˚C, followed by a
final 5 min elongation at 65˚C (all viral primers), or 20 sec denaturation at 95˚C, followed by
35 cycles of 3 sec denaturation at 95˚C and 30 sec annealing/elongation at 60˚C, followed by a
final 5 min elongation at 65˚C (GAPDH). PCR products were analysed by gel electrophoresis.
Library preparation and Illumina sequencing
Total DNA was quantitated using a Qubit HS DNA assay (Thermo Fisher). Sequencing librar-
ies were constructed using 200 ng of input DNA. End repair, A-tailing and adapter ligation
were performed using reagents from the SureSelect XT Reagent Kit (Agilent), each step fol-
lowed by a cleanup procedure using AMPure XP beads (Beckman Coulter) at 1.8 x concentra-
tion. Following adapter ligation, four rounds of PCR (Herculase II, Agilent) were performed
to enrich for correctly adapter-ligated fragments. Initial denaturation was at 98˚C for 2 min,
followed by 4 cycles of denaturation (98˚C, 30 s), annealing (65˚C, 30 s) and extension (72˚C,
1 min), and finally a 10 min extension step at 72˚C. Libraries were purified as described above
and diluted 1:6 with nuclease free water. A second round of PCR (6 cycles) using 15 μl of input
was performed using indexing primers available in the SureSelect XT Reagent Kit and PCR
conditions as outlined above except for an adjustment to the annealing temperature (to 57˚C).
Final libraries were analysed using a Qubit Fluorimeter and Tapestation prior to sequencing
across 8 runs on an Illumina MiSeq (2x150 bp PE reads, 300 cycle kit).
RNASeq libraries were prepared using the SureSelect Strand-Specific RNA Library Prep for
Illumina Multiplexed Sequencing (Protocol version D0, 2015), multiplexed and sequenced on
a single Illumina NextSeq run (2x34 bp PE reads, 75 cycle v2 high output).
Sequence data analyses
Sequence run data were de-multiplexed using bcl2fastq2 v2.17 under stringent conditions
(—barcode mismatches 0). Demultiplexed datasets were trimmed using TrimGalore software
(http://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) to remove low quality 3’
ends and aligned against the HSV-1 strain 17 genome (NC_001806) using BWA [57].
Genome-wide coverage and read depth data were calculated using custom scripts following
the generation of pileup files using SAMTools [58]. RNASeq data were processed as above,
except that alignment of reads against the HSV-1 strain 17 genome was performed using the
splice-aware BBMap (http://sourceforge.net/projects/bbmap/) with default parameters.
DNA per fraction. Percentage of HSV-1 DNA to total DNA was calculated by the ratio of
HSV-1 DNA reads to total DNA reads in each fraction. HSV-1 DNA genome copy equivalents
in each fraction were then calculated by multiplying the genome copy equivalents in the total
DNA by the percentage of HSV-1 DNA in each fraction. The number of HSV-1 genome copy
equivalents in each fraction was divided by the number of genome copy equivalents in the
total undigested and unfractionated HSV-1 DNA to give the percentage of HSV-1 genome
copy equivalents recovered in each fraction, or to the addition of the number of HSV-1
genome copy equivalents in all fractions and pellet to give the percentage of HSV-1 genome
copy equivalents in each fraction.
Normalized DNA reads. DNA reads (250 bp window) were generated for the entire
HSV-1 genome for each fraction. Total DNA reads of soluble and insoluble chromatin at any
given genomic position were divided to the DNA reads in the unfractionated chromatin at the
same position, to give the normalized DNA reads for that position in each chromatin fraction.
HSV-1 DNA locus in each fraction. HSV-1 DNA reads in each fraction for each position
were corrected by the number of HSV-1 genome copy equivalents in that fraction. The cor-
rected HSV-1 DNA reads in each position were then normalized to the total HSV-1 DNA
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 7 / 37
reads in that position in the same fraction, to give the number of HSV-1 genome copy equiva-
lents in each genome position in every fraction. The sum of the number of HSV-1 genome
copy equivalents in each position in all fractions was then normalized to the number of HSV-1
genome copy equivalents in the same position of the undigested and unfractionated chroma-
tin, to give the recovery of each HSV-1 locus after digestion.
Gene sampling. The number of DNA reads equal or larger to one mapping to every 250
bp window was manually counted as 1. Then, the manually corrected reads were assigned to
each gene according to their positions. Gene sampling was calculated as the sum of the manu-
ally corrected DNA reads of that gene, normalized to the sampling of the same gene in the
undigested and unfractionated chromatin. Sampling of all genes in each kinetic class was aver-
aged to give the average sampling of that kinetic class. Gene sampling of all genes in each frac-
tion was averaged to give the mean sampling. Mean sampling in each fraction was divided by
the overall mean sampling to analyze sampling enrichment in each fraction.
CTCF chromatin immunoprecipitation
Vero cells (6 x 10
6
on a 100 mm dish) were cross-linked with 5 ml of DMEM supplemented
with 100 μl of 1M Na
2
HPO
4
pH 8.96 and 142 μl of 37% formaldehyde, to a final concentration
of 1%, for 10 min at 37˚C. Crosslinking was quenched with 5 ml of DMEM supplemented
with 714 μl of 2 M glycine, to a final concentration of 250 mM. After vacuuming, monolayers
were further incubated with 5 ml of DMEM supplemented with 333 μl of 2 M glycine, to a
final concentration of 125 mM, for 5 min at room temperature. Cells were then washed with
10 ml 4˚C PBS supplemented with protease inhibitor (Pierce Protease Inhibitor Tablets
EDTA-free [A32965]). Monolayers were scraped in 15 ml of 4˚C PBS supplemented with pro-
tease inhibitor and transferred to a 50 ml conical tube; the dishes were then washed with other
15 ml of 4˚C PBS, for a final volume of 30 ml. Cells were pelleted and resuspended in 6 ml of
4˚C reticulocyte standard buffer (RSB, 10 mM Tris [pH 7.5], 10 mM NaCl, 5 mM MgCl
2
) sup-
plemented with protease inhibitor. Cells were re-pelleted, resuspended in 6 ml 4˚C RSB sup-
plemented with protease inhibitor and incubated on ice for 20 min. The plasma membrane
was lysed by adding 6 ml of RSB 1% triton X-100 supplemented with protease inhibitor and
incubating on ice for 15 min, flipping the tubes upside down thrice. Nuclei were pelleted by
centrifugation at 3,250 xg for 25 min at 4˚C, resuspended in 1 ml of RSB and re-pelleted at
21,000 xg for 1 min at 4˚C. Nuclei were resuspended and lysed in 1 ml chromatin extraction
buffer (1 mM ethylene glycol-bis(beta-aminoethyl ether)-N, N, N’, N’,- tetra-acetic acid
(EGTA), 2% Triton X-100, 3 mM MgCl2, 2 mM Tris [pH 8]) supplemented with protease
inhibitor on ice for 60 min, pipetting up and down 10 times every 10 min. Chromatin was pel-
leted at 4,000 xg for 10 min at 4˚C, resuspended in 1 ml of ChIP binding buffer (1% Triton, 10
mM Tris pH 8.0, 150 mM NaCl, 2 mM EDTA), sheared by sonication, flash frozen, and stored
at -80˚C.
Sheared chromatin was thawed on ice and diluted with ChIP binding buffer, to a final con-
centration of 16 ng/μl. Ten micrograms per ChIP reaction of CTCF antibody (07–729; EMD
Millipore) or nonspecific IgG (12–370; EMD Millipore) were conjugated to 1.5 mg of Dyna
Beads (Invitrogen 10001D) for ~20 hrs at 4˚C, with constant rotation. Chromatin (8 μg) was
incubated with bead-Ab complex for ~25 hrs at 4˚C, with constant rotation. Immunopurified
complexes were rinsed consecutively with 1 ml of each washing solution 1 (1% Triton, 2 mM
EDTA, 20 mM Tris pH 8.0, 150 mM NaCl), 2 (2 mM EDTA, 20 mM Tris pH 8.0, 150 mM
NaCl), 3 (1% Triton, 2 mM EDTA, 20 mM Tris pH 8.0, 500 mM NaCl), 4 (2 mM EDTA, 20
mM Tris pH 8.0, 500 mM NaCl), 5 (1% NP-40, 1% NaDOC, 1 mM EDTA, 10 mM Tris pH 8.0,
250 mM LiCl), and 6 (1 mM EDTA, 10 mM Tris pH 8.0, 250 mM LiCl). Immunopurified
HSV-1 transcriptional competence
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complexes were eluted with 600 μl of ChIP elution buffer (10 mM Tris-HCl pH 8.0, 150 mM
NaCl, 1 mM EDTA, 0.5% SDS) and 6 μl of proteinase K (20 mg/ml), and then incubated at
65˚C for approximately 19 hrs for decrosslinking and deproteinization. Immunoprecipitated
or input (0.8 μg of chromatin) DNA was phenol-chloroform extracted, isopropanol precipi-
tated, ethanol washed, and subjected to qPCR (Fast SYBR Green master mix) with 300 nM each
the B9# (5’- TCCTGTTGATCGATACGGGG -3’, 5’- ATCACAAACACTGGCAGCAG -3’)
B1# (5’- TCTAACGTTACACCCGAGGC -3’, 5’- GTATATATGCGCGGCTCCTG-3’), ICP4
(5’- GAGGATCCCCGCGGCGCCGTA -3’, 5’- TGGCCATGAGCCGCCGATACGAC -3’),
CTRS3 (5’- CACGAACGACGGGAGCG -3’, 5’- CACCCAAGGTGCTTACC -3’), CTRS1/2
(5’- GCCCCTCGAATAAACAACGCTA -3’, 5’- GTTGTGGACTGGGAAGGCGC -3’), or
CTa’m (5’- GGCTGCCACAGGTAAAACA -3’, 5’- TGTAGCAGGAGCGGTGTG -3’). DNA
from the untreated infections was diluted 1:1,000. qPCR conditions (except for the ICP4 peak)
were 20 sec denaturation at 95˚C, followed by 40 cycles of 3 sec denaturation at 95˚C and 30 sec
annealing/elongation at 60˚C, followed by 15 sec denaturation at 95˚C and 60 sec annealing/
elongation at 60˚C, followed by a temperature ramp from 60 to 95˚C in 1% increments every 15
sec and a final 15 sec incubation at 60˚C. The ICP4 peak was amplified by 20 sec denaturation
at 95˚C, followed by 40 cycles of 3 sec denaturation at 95˚C and 30 sec annealing/elongation at
70˚C, followed by 15 sec denaturation at 95˚C and 60 sec annealing/elongation at 60˚C, fol-
lowed by a temperature ramp from 60 to 95 in 1% increments every 15 sec and a final 15 sec
incubation at 60˚C. Viral copy number was calculated from a standard curve using 10 fold dilu-
tions of 300,000 genome copies of cell free HSV-1 DNA in 2.8 ng of purified Vero cell DNA
and sheared by sonication. Immunoprecipitation efficiencies were calculated by subtracting the
genome copy equivalents precipitated with the non-specific IgG from those precipitated with α-
CTCF antibody and then dividing by the total genome copy equivalent in the input.
Results
Most intracellular HSV-1 DNA is neither protein-free nor encapsidated at
7 hours after infection
Prior studies had suggested that HSV-1 DNA is organized in highly dynamic nucleoprotein
complexes that otherwise behave like cellular chromatin, as evidenced by hybridization of
chromatin resolved after serial micrococcal nuclease digestion [5,6]. However, it is also possi-
ble that protein-free HSV-1 DNA simply co-fractionated with cellular chromatin. To test for
this possibility, the DNA in each fraction was co-immunoprecipitated with histone H3. The
fractionation pattern of the soluble chromatin was not obviously affected by cross-linking (Fig
1A). At 7 hours after infection, HSV-1 DNA from the immediate-early (ICP4), early (gE) or
late (UL25 and UL46) genes co-immunoprecipitated with histone H3 in all fractions, as tested
by saturating PCR (Fig 1B). The co-immunoprecipitation of both cellular and viral DNA in
the most inaccessible fractions was less efficient than in the more accessible ones. Intracellular
HSV-1 DNA thus fractionated in complexes with the hydrodynamic ratios of mono- to poly-
nucleosomes in which it interacts with histone H3.
Knowing that the HSV-1 DNA in all fractions was protected to nucleosome sizes in com-
plexes with the hydrodynamic ratios (and therefore shape and size) of cellular mono- to poly-
nucleosomes and which contain histone H3, we next evaluated the distribution of protein-free,
encapsidated and intracellular HSV-1 DNA after serial MCN digestion by differential centrifu-
gation and deep sequencing (Fig 2A). Any protein-free HSV-1 DNA would be digested to small
fragments, which would fractionate mostly to the very top of the gradients, or to short polynu-
cleotides, which would not be detected by sequencing (Fig 2A). Sporadic nucleosomes would
partially protect HSV-1 DNA and result in digestion to mono- to short poly-nucleosomes
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which would fractionate to only the top fractions of the gradient (Fig 2A). Fully (stably or unsta-
bly) chromatinized HSV-1 DNA would be sufficiently protected to limited digestion to result in
mono- to long poly-nucleosomes, which would fractionate throughout the gradients (Fig 2A).
Cellular chromatin resolved as mono- to poly-nucleosomes throughout the entire gradient, as
expected, with DNA and all core histones present in each fraction (Fig 2B and 2C).
DNA in each chromatin fraction and in the unfractionated chromatin was then subjected
to deep sequencing. The fraction of viral to total (cellular plus viral) reads and the total HSV-1
genome copy equivalent were used to calculate the number of HSV-1 genome copy equivalents
in each fraction (Fig 2D). The fractionation of protein-free and encapsidated DNA marked the
migration of naked or fully protected DNA, respectively. Protein-free HSV-1 DNA (approxi-
mately 800 genome copy equivalents per cell) added to chromatin of mock infected cells was
poorly protected and resolved almost exclusively to the lowest density fractions (10 to 12—Fig
2D). Encapsidated HSV-1 DNA (approximately 800 genome copy equivalents per cell) added
to the chromatin of mock infected cells resolved to the highest density fractions (Fig 2D),
clearly peaking with the insoluble chromatin as expected.
Fig 1. At after infection, HSV-1 DNA was protected from serial MCN digestion to sizes of mono to poly-nucleosomes that fractionated in
complexes with the hydrodynamic ratios of mono to poly-nucleosomes and contain histone H3. Nuclei of infected cells were subjected to
serial MCN digestion and then cross-linked. (A) Cross-linked soluble chromatin separated by hydrodynamic ratios in a 0–10% sucrose gradient,
and resolved in 2% agarose gel electrophoresis, stained with EtBr. The left pointing arrowheads at the right indicate the migration of mono- to
octa-nucleosome-sized DNA. (B) Chromatin immunoprecipitation of HSV-1 DNA with histone H3. EtBr stained agarose gels of saturating
PCR-amplified cellular (GAPDH) or HSV-1 (UL25, UL46, ICP4, or gE) DNA co-immunoprecipitated with histone H3. (C) Cartoon presenting
the positions of the PCR amplicons in the HSV-1 genome. In, input (2%); H3, anti-histone H3 antibody; IgG, isotype antibody control (labeled
only in the No Dig sample for simplicity); No Dig, undigested unfractionated chromatin; Ins, insoluble chromatin.
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There were approximately 845 HSV-1 genome copy equivalents per cell at 7 hours after
infection, constituting 6% of the nuclear DNA. Sixty percent of all the intranuclear HSV-1
DNA was recovered after MCN digestion and fractionation, which is consistent with previous
results using hybridization [5,6]. In contrast to protein free or encapsidated HSV-1 DNA,
intracellular HSV-1 DNA migrated throughout the gradient. Nonetheless, it peaked in frac-
tions 9 to 13, with the major peak in fractions 10 to 12, containing the smallest nucleosome
complexes (i.e. the most accessible chromatin) and fraction 1 and the insoluble chromatin,
containing the largest poly-nucleosome complexes (i.e. the least accessible chromatin) (Fig
2D).
We considered the possibility that a large fraction of the protected HSV-1 DNA was still
encapsidated. However, VP5 was hardly detectable (~10%) when all soluble chromatin was
analyzed together (Fig 3), and undetectable in any individual fraction (S7 Fig). ICP4 and ICP8
are two major HSV-1 DNA binding proteins. Approximately 30% ICP4, and 20% ICP8
resolved in the soluble chromatin, while approximately 70% and 80%, respectively, was present
in the insoluble chromatin. By contrast, only ~25% HSV-1 DNA was in the insoluble chroma-
tin at this time. The HSV-1 DNA present in the insoluble chromatin may thus have been pro-
tected in part by ICP4 or ICP8 (or other non-chromatin proteins). Most ICP8 in the soluble
chromatin resolved to fractions 10 to 12 (Fig 3), whereas ICP4 more evenly fractionated
through fraction 5 to 13.
Transcribed and non-transcribed HSV-1 genes are in similarly accessible
chromatin
Besides other effects such as recruitment of transcription factors, differential chromatinization
of individual HSV-1 genes could regulate the accessibility of each gene and therefore its tran-
scription. To modulate the transcription level of specific groups of genes, we used cyclohexi-
mide (CHX) and roscovitine (Rosco). Cycloheximide inhibits protein synthesis, thereby
indirectly inhibiting transcription of most HSV-1 genes while allowing high levels of transcrip-
tion of the IE ones. Rosco inhibits transcription of all HSV-1 genes. When E and L gene tran-
scription was inhibited with CHX, there were approximately 35 HSV-1 genome copy
equivalents per cell (infected at an moi of 10) at 7 hpi. When HSV-1 transcription was inhib-
ited with Rosco, there were approximately 42 HSV-1 genome copy equivalents per cell. As dis-
cussed, there were approximately 845 HSV-1 genomes per cell at 7 hpi in the untreated
infections.
To evaluate the relative accessibility of the IE, E or L genes, the number of DNA reads for
each IE, E or L gene in each fraction were subjected to cluster analysis (Fig 4A). If transcription
of a specific gene was related to its accessibility, then the transcribed IE genes in infections
treated with CHX would cluster together with all transcribed genes in untreated infections,
whereas the non-transcribed E and L genes in CHX-treated infections would cluster with all
non-transcribed genes in the infections treated with Rosco. However, all IE, E and L genes in
the untreated infections were equally overrepresented in the least and most accessible
Fig 2. Intracellular HSV-1 DNA is differentially protected from MCN digestion than deproteinized or encapsidated
DNA. (A) Cartoon presenting the different models proposed for the HSV-1 chromatin and the expected fractionation
patterns after serial digestion and sucrose gradients of: (i) mostly non-chromatinized HSV-1 DNA, (ii) HSV-1 DNA
associated with nucleosomes only sporadically, (iii) fully (stably or unstably) chromatinized HSV-1 DNA. (B) DNA from each
fraction was resolved in 2% agarose gel electrophoresis and stained with EtBr. (C) Western blot for core histones (H2A, H2B,
H3 and H4) in all fractions, each resolved by SDS-PAGE and probed with corresponding antibodies. (D) Bar graph showing
the number of HSV-1 genome copy equivalents in each fraction. Chromatin of HSV-1 infected cells was digested and
fractionated. DNA in each fraction was subjected to deep sequencing. Ins, insoluble chromatin; GCE, genome copy
equivalents. Results from one experiment representative of three.
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chromatins (with the caveat that the top fractions may also contain some naked DNA that was
somewhat not digested by MCN) and thus clustered together, and away from IE, E and L
genes in the infections treated with Rosco or CHX. The IE, E and L genes in the infections
treated with CHX also clustered together, as they were more evenly distributed in all fractions
than in the untreated infections (Fig 4A and 4B). They were less depleted in the intermediate
fractions and less overrepresented in the most accessible fractions. The DNA of some IE and E
genes had a somewhat unique distribution among all genes in the infections treated with
Rosco, clustering closer to genes in the CHX-treated infections. Therefore, the accessibility of
a given gene, at the population level, does not directly correlate with its transcription levels.
To further test whether the accessibility of any individual gene was related to its transcrip-
tion, we evaluated individually the fractionation of each known HSV-1 gene. For the accessi-
bility of a gene to be related to its transcription level, the entire gene (but not necessarily its
promoter) would be expected to be similarly accessible. Consequently, the entire gene would
fractionate as a unit in the relevant fractions and deep sequencing would sample DNA
sequences covering most of the gene in those fractions. As HSV-1 genes range from 170bp to
9,490bp, we analyzed the relative sampling of each gene in each chromatin fraction or in the
Fig 3. Fractionation of the major HSV-1 DNA binding proteins. (A) Western blots of VP5, ICP4 and ICP8 in the insoluble
and soluble chromatin fractions. Results representative of two independent repeats. (B) Fractionation of ICP4, ICP8, and HSV-1
DNA after serial MCN digestion and sucrose gradient centrifugation. Western blots of ICP4 and ICP8, and line graph
presenting the percentage of ICP4, ICP8 and HSV-1 DNA in each fraction. VP5 was below sensitivity levels in all fractions. Ins,
insoluble chromatin fraction; sol, soluble chromatin fraction. Average of three (ICP8) or two (ICP4) independent experiments.
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Fig 4. All HSV-1 genes resolved together to the least and most accessible chromatin fractions. (A) Cluster analyses of the number
of DNA reads for IE, E or L genes in each fraction. Cluster analysis was performed with Cluster 3.0 from Stanford University,
visualized with Java Treeview. Color key on the left indicates ratio of reads in fraction over reads in the total DNA (ranges are in log
2
).
7, 7 hpi, untreated infections; C, CHX- treated infections (7 hpi); R, roscovitine-treated infections (7 hpi). (B) Gene sampling of each
HSV-1 gene in each fraction. Sampling of each HSV-1 gene was calculated and normalized to the sampling of the same gene in the
undigested and unfractionated chromatin. Each gene is color coded according to kinetic class. Blue, IE genes; black, E genes; dark
purple, unclassified L genes; light purple, early L genes; brown, true L genes; grey, unclassified genes. Ins, insoluble chromatin fraction.
Results from one experiment representative of three.
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unfractionated chromatin. The fraction of a given gene sampled in each fraction was then nor-
malized to the fraction of the same gene sampled in the unfractionated chromatin and
expressed as percentage (gene sampling).
The sampling of each gene was calculated in each fraction and plotted according to their
kinetic groups (Fig 4B). At 7 h after infection, the least (insoluble chromatin and fractions 1
and 2) and most (fractions 11 to 13) accessible chromatin were enriched in fully sampled
HSV-1 genes, whereas the intermediately accessible chromatin contained only random sam-
plings of each gene. Fractions 3 and 10 sampled approximately half of each HSV-1 gene,
whereas the rest of the intermediate fractions had sampled only less than 20% of each gene. No
individual HSV-1 gene was differentially sampled in any fraction.
When transcription of E and L genes was inhibited (in infections treated with CHX), the
transcriptionally active IE genes still fractionated as the transcriptionally inactive E and L
genes (Fig 4B). The least (insoluble chromatin and fraction 1) and most (fractions 12 to 13)
accessible chromatin were enriched in completely sampled HSV-1 genes. In the intermediately
accessible chromatin each gene was sampled less frequently—albeit not nearly to the same
extent as in the untreated infections. Fractions 2 to 5 and fraction 11 sampled 40–60% of each
HSV-1 gene, and fractions 6 to 10, 25% - 40%.
No HSV-1 genes fractionated differently when HSV-1 transcription was inhibited with
Rosco either, and the least (insoluble chromatin and fraction 1) and most (fractions 12 to 13)
accessible chromatin were enriched in mostly fully sampled HSV-1 genes (Fig 4B). As in the
infections treated with CHX, the depletion in fully sampled genes in the intermediately accessi-
ble chromatin was only partial. Fractions 2 to 4 and fraction 11 sampled 40–60% of each HSV-
1 gene, and factions 5 to 10, 25–40%. The depletion in fully sampled genes in the intermediate
accessible chromatin could have been an artifact resulting from too little HSV-1 DNA. How-
ever, fully sampled HSV-1 genes resolved to the insoluble chromatin, and to fractions 1, 12, 13
and the insoluble chromatin in infections treated with CHX or Rosco. These fractions con-
tained as little HSV-1 DNA as the intermediate fractions in the untreated infection, which con-
tained only partially sampled genes. From the comparison with the CHX-treated infections,
the accessibility of all genes at the population level related to the overall levels of transcription
of the genomes, whereas the accessibility of any single gene, at the population level, did not.
To evaluate the global accessibility of IE, E and L genes, the average gene sampling for each
group was calculated in each fraction. At 7 h after infection, the least and most accessible chro-
matins were enriched in fully sampled HSV-1 genes of all three kinetic classes. The other frac-
tions were in contrast partially depleted of fully sampled genes (Fig 5A). The enrichment in
fully sampled IE, E, or L genes in only the least and most accessible chromatins was not altered
when only IE genes were transcribed to high levels (CHX), no gene was transcribed to high
levels (Rosco), or all IE, E and L genes were transcribed to high levels (Fig 5A). In contrast, the
depletion in fully sampled genes in the intermediately accessible chromatin was less marked in
CHX or Rosco-treated infections. We concluded that transcription and accessibility of genes
in any kinetic class are not directly related (Figs 4and 5).
Sampling of all genes in all fractions was averaged. Sampling of all genes in each fraction
was then expressed as enrichment over the average sampling and plotted together with the per-
centage of HSV-1 DNA in each fraction (Fig 5B). At 7 h after infection, the least (insoluble
chromatin to fraction 2) and most (fractions 11 to 13) accessible chromatins that were
enriched in fully sampled HSV-1 genes had most of the HSV-1 DNA. The most accessible
chromatin that was enriched in fully sampled genes had 31% of the HSV-1 DNA, and the least
accessible chromatin that was equally enriched in fully sampled genes, 35%. Although the frac-
tions containing the most accessible chromatin could also contain some naked DNA, the pres-
ence of fully sampled genes indicates that the DNA in these fractions had not been digested
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Fig 5. The most and least accessible chromatin enriched in fully sampled HSV-1 genes contained more HSV-1 DNA
when transcription was active. (A) Bar graphs showing average sampling of all genes in each HSV-1 gene kinetic groups
in each fraction. Sampling of each HSV-1 gene was calculated and normalized to the sampling of the same gene in the
undigested and unfractionated chromatin. The normalized sampling of all genes in each group was then averaged. Blue,
IE genes; black, E genes; dark purple, unclassified L genes; light purple, early L genes; brown, true L genes; grey,
unclassified genes. (B) Area graphs and pie charts presenting relative enrichment in gene sampling and percentage of
HSV-1 DNA in each fraction. Gene sampling in each fraction is expressed as ratio to the average sampling to present the
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extensively, contrary to what would had been expected from naked DNA. The intermediately
accessible chromatin (fractions 3 to 10), which contained 34% HSV-1 DNA, contained only
partially sampled genes (Fig 5B). This depletion in fully sampled genes was not a result of a
mix of chromatinized and protein-free HSV-1 DNA. Protein-free HSV-1 DNA did not resolve
to these intermediate fractions (Fig 2D) and DNA from an E and a L gene in these fractions
co-immunoprecipitated with histone H3 (Fig 1B). The distribution of HSV-1 DNA was
markedly distinct from that of the cellular DNA (dashed black line), indicating biophysical dif-
ferences between the viral and cellular chromatin under these conditions.
When most HSV-1 transcription was inhibited with CHX, the least accessible chromatin
that was enriched in fully sampled HSV-1 genes (insoluble chromatin and fraction 1) had 22%
of the HSV-1 DNA. In contrast, the most accessible chromatin that was enriched in fully sam-
pled genes (fractions 12 and 13) had only 2% of the HSV-1 DNA (Fig 5B). Similarly, when
HSV-1 transcription was inhibited with Rosco, the least accessible chromatin that was
enriched in fully sampled HSV-1 genes (insoluble chromatin and fraction 1) had 15% of the
HSV-1 DNA whereas the most accessible chromatin that was also enriched in fully sampled
genes (fractions 12 and 13), had only 4% of the HSV-1 DNA (Fig 5B). The intermediately
accessible chromatin, containing only partially sampled genes, contained 75–80% of the HSV-
1 DNA. In both conditions, the depletion in fully sampled genes in the intermediately accessi-
ble chromatin was only partial and progressive, with less depletion in the least accessible inter-
mediate chromatin (fractions 1–6) and more in the most accessible ones (fractions 7–9). In
both conditions, the distribution of the viral DNA very much replicated that of the cellular
DNA, indicating similar biophysical properties between the viral and cellular chromatin when
most of the viral genome is not transcribed.
In summary, the HSV-1 chromatin had similar biophysical properties to the cellular chro-
matin when there was limited HSV-1 transcription, but it was far more accessible when the
genomes were transcribed. Moreover, the levels of HSV-1 DNA in the most accessible chroma-
tin that was enriched in fully sampled genes related to the overall transcription level of the
genomes.
Changes in the accessibility of HSV-1 chromatin along infection
We next evaluated the HSV-1 chromatin as infection progressed. At 2h after infection with 10
pfu/cell, there were approximately 20 HSV-1 genome copy equivalents per cell, constituting
0.2% of the DNA in the cell. The HSV-1 DNA was enriched in the least accessible chromatin
(insoluble chromatin and fractions 1–2), with a very minor relative enrichment in the most
accessible chromatin (fractions 9–11), and depleted in the intermediately accessible chromatin
(fractions 3–8) (Fig 6). The distribution of the HSV-1 DNA changed at 4h, when there were
approximately 90 HSV-1 genome copy equivalents per cell, constituting 0.8% of the nuclear
DNA. HSV-1 genomes were then enriched in the least (mostly in the insoluble chromatin) and
in the most (fractions 9–11) accessible chromatin, while they were relatively depleted in the
intermediately accessible chromatin (fractions 2–8). As already discussed, there were approxi-
mately 850 HSV-1 genome copy equivalents per cell, constituting 6% of the nuclear DNA at
after infection. The HSV-1 genomes were also enriched in the most and, less so, in the least
relative enrichment in fully sampled genes in each fraction. The distribution of the cellular chromatin is indicated by the
dotted black line. Pie charts indicating the percentages of the HSV-1 DNA in the most accessible chromatin fractions
containing completely sampled HSV-1 genes (brown), in the intermediate accessible chromatin fractions containing
random samplings of each gene (grey), or in the least accessible chromatin fractions containing completely sampled
HSV-1 genes (purple). Ins, insoluble chromatin fraction. Results from one experiment representative of three.
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accessible chromatin and depleted in the intermediately accessible chromatin (Fig 2D–pre-
sented again in Fig 6 for comparison). At 15h after infection, there were approximately 2,650
HSV-1 genome copy equivalents per cell, constituting 20% of the nuclear DNA. At this time,
the HSV-1 DNA was also enriched in the least (insoluble chromatin and fraction 1) and most
(fractions 9–11) accessible chromatin (Fig 6).
The accessibility of all HSV-1 genes increases in synchrony as the infection
progresses
No particular HSV-1 gene was differentially accessible at 7 hpi, regardless of whether tran-
scription was inhibited or not (Fig 4), and the most accessible chromatin was enriched in fully
sampled genes only when there was abundant transcription (Fig 5B). Both the most and least
accessible chromatins contained fully sampled HSV-1 genes (Fig 4). One possibility was that
the most accessible chromatin contained transcriptionally competent genomes and the least
accessible one, transcriptionally silenced ones (and potentially encapsidated DNA although
VP5 was barely detectable at this time), whereas the intermediately accessible chromatin con-
tained genomes transitioning between states. Then, the fractionation of the HSV-1 genomes
would be expected to change from the least accessible chromatin toward the most accessible
one as infection progresses. To test this model, we analyzed the accessibility of the viral chro-
matin as the infection progressed.
When evaluated by their fractionation, IE, E and L genes clustered together according to
time after infection, not to kinetic class (Fig 7A). All IE, E, and L genes were relatively under-
represented in the most accessible chromatin and thus clustered the furthest away at 2 h, when
there is limited HSV-1 transcription. IE, E and L genes were all enriched in the most and least
accessible chromatin at 4 h, when there is already widespread HSV-1 transcription. At 12 h,
the depletion in the intermediately accessible chromatin was even more uniform and marked.
Genes of all kinetic classes resolved similarly and were similarly sampled in each fraction at all
infection times (Fig 7B).
To analyze potential differences among IE, E or L genes, we averaged the sampling of all
genes in each kinetic class. At 2 h, the least accessible chromatin fractions (insoluble, fractions
Fig 6. Differential fractionation of HSV-1 DNA throughout infection. Bar graphs showing HSV-1 genome copy equivalents in each fraction. (�)Data
from Fig 2 presented again for comparison. Ins, insoluble chromatin fraction; GCE, genome copy equivalent. Results from one experiment representative
of two.
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Fig 7. All HSV-1 genes resolved together to the least and most accessible chromatin fractions as infection progressed. (A) Cluster analyses
of the number of DNA reads from IE, E and L genes in each fraction. Cluster analysis was performed using Cluster 3.0 from Stanford
University, visualized with Java Treeview. (B) Bar graphs showing sampling of each HSV-1 genes at 2h, 4h, and 15h after infections. Sampling
of each HSV-1 gene was calculated and normalized to the sampling of the gene in the undigested and unfractionated chromatin. The sampling
of each gene was plotted according to kinetic class. Blue, IE genes; black, E genes; dark purple, unclassified L genes; light purple, early L genes;
brown, true L genes; grey, unclassified genes. Ins, insoluble chromatin fraction. Results from one experiment representative of two.
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1 and 2) were enriched in fully sampled HSV-1 genes of all kinetic classes (Fig 8A). The fully
sampled genes were depleted in the intermediately accessible fractions, although not to the
extent as in 7 h infections. The most accessible chromatin fractions (fractions 11 to 13) were
only partially enriched in fully sampled genes (Fig 8A). As infection progressed, the least acces-
sible chromatin fractions (insoluble and fractions 1 and 2 at 2 h, or insoluble and fractions 1 at
4 or 15 h) were enriched in fully sampled HSV-1 genes of all kinetic classes, as were the most
accessible fractions (fractions 11 to 13 at 2 h, or fractions 10 to 13 at 4 or 15 h) (Fig 8A).
Sampling of all genes in all fractions was again averaged and the sampling in each fraction
was expressed as enrichment over the average sampling and plotted together with the percent-
age of HSV-1 DNA in each fraction as in Fig 5B. At 2 h after infection, the least accessible
(insoluble chromatin to fraction 2) chromatin that was enriched in fully sampled HSV-1 genes
had 72% of the HSV-1 DNA (Fig 8B). The intermediately accessible chromatin that was only
partially depleted of fully sampled genes contained 21% HSV-1 DNA (fractions 2 to 10). The
most accessible (fractions 11 to 13) chromatin that was only partially enriched in fully sampled
HSV-1 genes (20–40% of each gene) had only 7% of the HSV-1 DNA (Fig 8B). The distribu-
tion of the viral DNA was somewhat similar to that of the cellular DNA, albeit more enriched
in the least accessible chromatin. At 4h, the least (insoluble chromatin to fraction 1) and most
(fractions 11 to 13) accessible chromatin that were enriched in fully sampled HSV-1 genes had
57% or 21% of the recovered HSV-1 DNA, respectively. At this time, there was a more marked
depletion in gene sampling in the intermediately accessible chromatin, and the distribution of
the viral DNA started to more markedly differ from that of the cellular DNA, with a clear
enrichment of viral DNA in the most accessible chromatin. At 7 h, as already discussed (Fig
5B–presented again in Fig 8B for comparison), the least (insoluble chromatin to fraction 2)
and most (fractions 11 to 13) accessible chromatin that was enriched in fully sampled HSV-1
genes had 35% or 31% HSV-1 DNA, respectively. There was a marked depletion of HSV-1
DNA in the intermediately accessible chromatin and the distribution of cellular and viral
DNA differed the most. At 15 h, the least (insoluble chromatin to fraction 1) and most (frac-
tions 10–13) accessible chromatins that were enriched in fully sampled HSV-1 genes had 50%
or 30% of HSV-1 DNA, respectively (Fig 8). At this time, HSV-1 DNA was also enriched,
together with fully sampled HSV-1 genes, in the intermediately accessible chromatin in frac-
tion 4 (4%).
Selected short HSV-1 DNA sequences are less underrepresented in the
digested and fractionated HSV-1 chromatin than in the undigested and
unfractionated one
To analyze if there were large scale differences in the accessibility of different HSV-1 loci, the
number of HSV-1 DNA reads for each genome position in each fraction was corrected by the
HSV-1 genome copy equivalents in the same fraction. The addition of the genome copy equiv-
alents in each position in all fractions were normalized to the HSV-1 DNA genome copy
equivalents in the same position in the unfractionated and undigested chromatin to give the
relative genome copy equivalents of that position after digestion and fractionation. The nor-
malized values were plotted against genome position using a 250bp sliding window. The cover-
age of HSV-1 RNA were plotted likewise to evaluate the transcription levels at each locus
(Fig 9).
Consistently with the cluster analyses, the entire HSV-1 genomes were similarly accessible
at 7 h, regardless of the transcription level at any specific locus. No genomic region was obvi-
ously enriched or depleted in any fractions (S1 Fig), and the total recovery of the HSV-1 DNA
in each locus in all fractions was equally proportional to the total HSV-1 DNA in that locus
HSV-1 transcriptional competence
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Fig 8. The most and least accessible chromatin enriched in completely sampled HSV-1 genes contained more HSV-1
DNA as the infection progressed. (A) Bar graphs showing average sampling of the genes in each HSV-1 gene kinetic class
in each fraction at 2, 4 and 15 hours after infection. Sampling of each HSV-1 gene was calculated and normalized to the
sampling of the gene in the undigested and unfractionated chromatin. The normalized sampling of all genes in each group
was then averaged. Blue, IE genes; black, E genes; dark purple, unclassified L genes; light purple, early L genes; brown, true L
genes; grey, unclassified genes. (B) Area graphs showing relative enrichment in completely sampled genes and percentage of
HSV-1 DNA in each fraction. Gene sampling in each fraction is expressed as ratio to the average sampling (relative
HSV-1 transcriptional competence
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before digestion and fractionation (Fig 9). One sequence in the repeat region appeared to have
been relatively enriched to some extent in the digested and fractionated over the undigested
and unfractionated chromatin; it mapped to a CTCF binding site [59] (empty downward
arrow). As the HSV-1 genome is widely transcribed at 7 h after infection, we analyzed the
infections treated with CHX or Rosco to test transcribed and non-transcribed loci.
When E and L gene transcription was inhibited with CHX, only the IE genes were tran-
scribed to a high level (Fig 9), as expected. However, the transcribed IE loci did not fractionate
differently from the non-transcribed E or L loci. No obvious HSV-1 genome region, and par-
ticularly no IE locus, was enriched in any fraction and compensatory depleted from other (S2
Fig). A few sequencing artifacts map to the same positions in several fractions (fractions 8
through 10, for example) and the undigested and unfractionated chromatin fraction. All HSV
loci, including the IE, were also proportionally recovered in the digested and fractionated
chromatin over the undigested and unfractionated one (Fig 9). However, 16 short sequences
of less than 250bp were relatively overrepresented in the digested and fractionated chromatin
over the undigested and unfractionated one (Fig 9). Seven of them, including five of the six
most overrepresented, map to previously mapped CTCF binding sites (empty arrows) or chro-
matin insulator-like elements (black solid circles) in strain 17 [60].
When HSV-1 transcription was inhibited with Rosco, the levels of transcription were low
through the entire genome, including the IE genes, as expected, with some low levels of tran-
scription through the genome and lower levels of IE transcription than in CHX treated infec-
tions. As in all other conditions, no specific HSV-1 locus was enriched in any fraction and
compensatory depleted in others (S3 Fig). The same sequencing artifacts detected in the CHX
samples were evident in the same positions (for example, fractions 7 through 10). As in the
previous cases, however, 15 short sequences of less than 250bp were slightly and relatively
overrepresented in the digested and fractionated chromatin over the undigested and unfractio-
nated one, albeit not nearly as overrepresented as in the CHX-treated infections (Fig 9). One
of these sequences map to chromatin insulator-like elements (black solid circle) [59].
To evaluate a condition in which only some loci are not highly transcribed in a context of
globally active transcription, we used PAA. No gene loci fractionated differently in infections
treated with PAA either, in which HSV-1 IE and E genes were transcribed to high levels
whereas the L genes were not. Nine short sequences were again relatively overrepresented in
the digested and fractionated chromatin over the undigested and unfractionated one, albeit
again not nearly as much as in the CHX-treated infections. Two map to a previously mapped
chromatin insulator like elements (black solid circle) [59], and another to a previously mapped
CTCF binding site (empty downward arrow) [60].
The short sequences overrepresented in the chromatin fractions flank
highly transcribed HSV-1 genes
As the relatively most overrepresented short sequences in the digested and fractionated chro-
matin mapped to the internal and terminal repeats, which encode most of the IE genes, the
LAT, and some other genes, the HSV-1 repeats were evaluated in more detail (Fig 10). In infec-
tions treated with CHX, the six dominant peaks flanked the highly transcribed IE loci,
enrichment). The distribution of the cellular chromatin is indicated by the dotted black line. (�)Data from Fig 5 presented
again for comparison. Pie charts showing percentage of HSV-1 DNA in the most accessible chromatin fractions containing
completely sampled HSV-1 genes (brown), in the intermediate accessible chromatin fractions containing random sampling
of each gene (grey), or in the least accessible chromatin fractions containing completely sampled HSV-1 genes (purple). Ins,
insoluble chromatin fraction. Results from one experiment representative of two.
https://doi.org/10.1371/journal.ppat.1008076.g008
HSV-1 transcriptional competence
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separating them from the rest of the genome (which is non-transcribed, orange line on top).
No similar dominant peaks flanking the IE genes were observed in the untreated infections, in
which there is abundant transcription across the entire genome, or in the infections treated
with Rosco, in which there was little transcription across the genome, or with PAA, in which
there is abundant transcription of IE and E but not L genes (Fig 10). No peaks thus separated
the IE genes from the rest of the genome when the overall transcription level was high
(untreated or PAA treated), or low (infections treated Rosco) across the entire genomes.
Fig 9. No HSV-1 loci fractionated differently regardless of transcription levels, but short overrepresented sequences flanked transcribed genes when
transcription was restricted. Line graphs showing HSV-1 number of genome copy equivalents (GCE) in each genome position in untreated infections, or in
infections treated with CHX, Rosco, or PAA at 7 hpi, normalized to the number of HSV-1 genome copy equivalents in the respective position of undigested and
unfractionated chromatin (blue line). Orange line graphs, HSV-1 RNA reads. X-axes, genome position (cartoon on top). Black downward empty arrows, CTCF
binding sites in strain 17; black solid circles, chromatin insulator-like elements in strain 17; black dots, other overrepresented sequences;purple bars underneath
genome plots, IE genes; dark green bars underneath genome plots, LAT; light green bar, stable LAT; GCE, genome copy equivalent. Results from one experiment
representative of three.
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HSV-1 transcriptional competence
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The relatively overrepresented sequences were often underrepresented in both the undi-
gested and unfractionated and some also in the digested and fractionated chromatin, which is
more obvious when the data are plotted in a logarithmic scale (S4 and S5 Figs). When so, how-
ever, they were less underrepresented in the digested and fractionated chromatin than in the
undigested and unfractionated chromatin.
There was only one just visible overrepresented short sequence in the digested and fraction-
ated chromatin in the untreated infections, when the overall transcription level was high, nine
Fig 10. Enlargement of the repeats region, from position 100kbp to 152kbp, of the plots presented in Fig 9.Orange lines graphs, HSV-1 RNA reads. X-
axes, genome position (cartoon on top). Blue lines, HSV-1 number of genome copy equivalents (GCE) in each genome position. Black downward empty
arrows, CTCF binding sites in strain 17; black solid circles, chromatin insulator-like elements; black dots, other overrepresented short sequences; purple bars
underneath genome plots, IE genes; dark green bars underneath genome plots, LAT; light green bar, stable LAT; GCE, genome copy equivalent. Results from
one experiment representative of three. ++ sequences co-immunoprecipitated with CTCF by ChIP in two independent experiments; + sequences co-
immunoprecipitated with CTCF by ChIP in one of two independent experiments;—sequences not co-immunoprecipitated with CTCF by ChIP in either of
the two independent experiments.
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HSV-1 transcriptional competence
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in PAA treated infections, when L genes are not transcribed, sixteen, including six very pre-
dominant ones, in the chromatin of CHX treated infections, when only the IE genes were tran-
scribed, and fifteen in the chromatin of Rosco treated infections, when the overall
transcription level was low.
The sequences underrepresented in the undigested and unfractionated chromatin of CHX-
treated infections that had not been experimentally mapped as CTCF binding sites in HSV-1
KOS included sequences recognized by CTCF in strain 17 or predicted in silico (S8 Fig) to be
potential CTCF binding sites in strain KOS. We thus explored whether these peaks comprised
sequences recognized by CTCF. Sequences on the previously described CTRS3, CTRS1/2 and
CTa’m [59], as well as the CTCF binding sites recognized by primers B1# and B9# [60] in
strain 17, and the proposed new CTCF binding site in the ICP4 gene, all co-immunoprecipi-
tated with CTCF above background in two independent experiments in infections treated with
cycloheximide (Fig 10). Consistently with the much reduced magnitude of the overrepresenta-
tion of all peaks in the roscovitine-treated infections in the MCN-seq, only the CTRS1/2
sequences and those amplified with primes B1# and B9# co-immunoprecipitated with CTCF
above background and each in only one of two independent experiment under these condi-
tions (Fig 10). Consistently with the lack of underrepresented sequences in the undigested and
unfractionated chromatin in the MCN-seq experiments, none of these sequences co-immuno-
precipitated with CTCF above background in the untreated infections in either of the two
independent experiments (Fig 10).
The number of short sequences that are less depleted in the digested and
fractionated chromatin decreases as the infection progresses
We next evaluated the number and location of the relatively overrepresented short sequences
as the infection progressed. At 2h after infection, 41 short sequences were clearly overrepre-
sented, including 30 predominant ones (Fig 11). Although the total HSV-1 genome copy
equivalents were similar to those in the infections treated with CHX, the distribution of these
relatively overrepresented sequences was markedly different. The relatively overrepresented
short sequences were evenly distributed through the genome at 2 h, rather than clustered
flanking the IE genes as in the infections treated with CHX. Nonetheless, most IE genes were
still flanked by sequences with limited accessibility (Fig 11B). There were 23, 1 or no short
sequences relatively overrepresented at 4, 7 or 15 h, respectively. The overrepresented peaks
included, again, previously mapped chromatin-insulator like sequences (solid black circles)
[59] and CTCF binding sites (empty downward arrows) [60], or both (black downward
arrows) (Fig 11).
Discussion
Chromatin remodelers are often recruited, via previous histone post-translational modifica-
tions, to cellular genes to increase chromatin dynamics and activate transcription [54,55,61–
64]. One could have expected the histone dynamics on transcribed and non-transcribed indi-
vidual HSV-1 genes to differ, too. The genes in the more dynamic chromatin would be more
accessible to transcription factors and MCN than those in less dynamic chromatin. Under this
model, the DNA of IE genes would be more accessible early in infection than that of E and L
genes. The HSV-1 genes would therefore fractionate according to their kinetic class after serial
MCN digestion and sucrose gradient centrifugation. When analyzing their fractionation pat-
terns, however, all HSV-1 genes clustered together according to the global level of transcrip-
tion of the genome, regardless of the level of transcription of the individual gene (or kinetic
class). Neither the clustering of HSV-1 genes nor the sampling of each gene support the
HSV-1 transcriptional competence
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Fig 11. The number of overrepresented short sequences decreases as infection progresses. Line graphs showing number of HSV-1 genome copy
equivalents in each genome position at 2, 4, 7 (�), or 15hpi, normalized to the number of HSV-1 genome copy equivalentsin the same position of the
HSV-1 transcriptional competence
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simplest model that chromatin dynamics are the major regulator of the transcription of indi-
vidual HSV-1 genes. Instead, they rather support an alternate model in which chromatin
dynamics regulate the transcriptional competence of entire HSV-1 genomes. As a caveat, pop-
ulational level approaches preclude the evaluation of differential dynamics of specific genes in
small subsets of the viral genomes.
The global accessibility of the HSV-1 DNA differed as infection progressed. At 2h after
infection, most of the HSV-1 DNA was in the least accessible chromatin. At this time after
infection, the DNA in the promoters of IE (ICP0), E (TK) and L (VP16) genes co-immunopre-
cipitates with histones bearing silencing markers, and the efficiency of immunoprecipitation
of HSV-1 chromatin is comparable to that of cellular chromatin [11,12,23,32,42,65,66],
always constrained by the intrinsic limitations of populational analyses and infection synchro-
nization. Recently, Dr. O’Hare’s group used EdC-labeled HSV-1 virions to analyze the nuclear
topology of the HSV-1 DNA, enables discriminative analyses of different genomes within an
infected cell [67]. As infection progress from 0.5h to 2h, the EdC signal, representing the HSV-
1 genomes, became less condensed and more dispersed. These changes are consistent with
HSV-1 genomes becoming less compacted, and therefore more accessible, as the infection pro-
gresses [67]. Depletion of a chromatin remodeler ATRX also increases mRNA levels of all
genes in three HSV-1 kinetic groups at any time of infection up to 8 hours [43]. In our experi-
ments, genes of all three kinetic classes were enriched in the least accessible chromatin at the
earlier times (Figs 7and 8).
As the infection progresses, the efficiency of co-immunoprecipitation of HSV-1 DNA with
chromatin decreases, and silencing markers become depleted from the viral chromatin [11,
32]. Overall histone dynamics in the nucleus of infected cells also increase at 4h after infection
[7,68–70]. ChIP of HSV-1 DNA in ATRX depleted cells indicated that histone H3K27me3
occupancy on the ICP27 and ICP8 gene promoters is not significantly affected up to 4 hours of
infection, but reduced to about half at 8 hours after infection [43]. In our experiments, HSV-1
DNA started to be enriched in the most accessible chromatin (and perhaps some naked DNA)
at this time, too (Figs 7and 8). Furthermore, DNA from IE, E and L genes mapping to the Ul,
Us, and Rs regions of the genome co-immunoprecipitated with histone H3 (Fig 1). These
results are consistent with more dynamic interactions of histone H3 with HSV-1 DNA than
with cellular DNA.
The increase in histone dynamics and the enrichment in the most accessible HSV-1 chro-
matin (which may also contain perhaps some naked DNA) could both have resulted from
HSV-1 DNA replication. However, PAA has no obvious effects on HSV-1 chromatin. The
decrease in efficiency of co-immunoprecipitation of HSV-1 DNA with histones as infection
progresses is independent of HSV-1 DNA replication [32], as are the changes in dynamics of
histone H1, H2B, H2A, H3.3, and H4 [68–70]. Neither is HSV-1 DNA accessibility to MCN
digestion altered by inhibition of HSV-1 DNA replication [6]. Consistently, Dr. O’Hare’s
work, using approaches that allow to resolve different genomes in a single cell, showed that
inhibition of HSV-1 DNA replication by PAA or acyclovir did not affect HSV-1 genome
decompaction as infection progressed [67], and using populational approaches Dr. Knipe’s
group showed that PAA does not affect the level of H3K9me3 on HSV-1 chromatin [43].
respective undigested and unfractionated chromatin. X-axes, genome position (cartoon on top). �: data from Fig 9 presented again for comparison.
Black downward empty arrows, CTCF binding sites; black solid circles, chromatin insulator-like elements; black downward solid arrows, CTCF
binding- and chromatin insulator-like elements; black dots, peaks of the DNA plots; black diamonds underneath the plots, the seven overrepresented
sequences that do not contain the AT-rich motifs; purple bars underneath genome plots, IE genes; dark green bars underneath genome plots, LAT;
light green bar, stable LAT; GCE, genome copy equivalent. (A) Presentation of relative HSV-1 DNA plots through the entire genome; (B)
Enlargement of the relative HSV-1 DNA plots of the repeat region, from position 100kbp to 152kbp. Results from one experiment representative of
two.
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Several models have been proposed for the potential structures of the HSV-1 chromatin in
lytic infections. Only specific regions of HSV-1 genes were protected to nucleosome-sized frag-
ments after complete MCN digestion [13]. Most of the promoters of IE genes, and 60% of the
promoters of E genes and L genes, were not detected in the MCN-resistant fraction. The HSV-
1 DNA fragments of nucleosome size tended to cluster in the center of the longer genes,
whereas short genes and the 5‘and 3‘ends of most genes were mostly not detected in the DNA
subpopulation protected as nucleosome sized fragments [13]. Had these patterns of protection
reflected different static levels of nucleosome occupancy, then the serial MCN digestion would
have yielded similar protection patterns. The poly-nucleosome fragments of regular HSV-1
chromatin would have resolved to the higher density fractions in the sucrose gradient, whereas
the short nucleosome fragments of the partially chromatinized HSV-1 DNA would have
resolved to the lighter ones, and the naked DNA would have mostly been digested, or would
have resolved to the very top fractions. In contrast, complete HSV-1 genomes partitioned to
every fraction, with clear enrichment of HSV-1 DNA in the most and least accessible chroma-
tin (Figs 2D and 5). The patterns of protection from complete MCN digestion [13] are there-
fore more likely to reflect the dynamics of the HSV-1 chromatin than differential static
nucleosome occupancy.
HSV-1 DNA is protected from serial MCN digestion to sizes consistent with nucleosome
DNA in complexes that have the same hydrodynamic ratios as mono- to poly-nucleosomes [5,
6] in which it interacts with histone H3 (Fig 1). The most inaccessible chromatin contained 1/
3 of the intracellular HSV-1 DNA at 7 hours after infection (Fig 5B). VP5 was barely detectable
in the insoluble (or soluble) chromatin at this time. Therefore, most of the DNA in the insolu-
ble chromatin fraction at 7 hours is not likely to be encapsidated. However, ICP4 and ICP8
fractionated to the most inaccessible fractions at this time (Fig 3) and could thus have pro-
tected some of the HSV-1 DNA in them. We would not expect to detect much PML at 7 hours
after infection. We have not evaluated the fractionation of PML at earlier times or in the pres-
ence of cycloheximide or roscovitine and the approaches we have used do not evaluate whether
other non-chromatin proteins could have possibly also protected some of the HSV-1 DNA.
Although it is thus possible that PML, ICP4, ICP8 or other non-chromatin proteins could have
also participated in the protection of the HSV-1 DNA under some conditions, the protected
HSV-1 DNA fragments have the same hydrodynamic ratios than mono- to poly-nucleosomes
in all tested conditions. Any protecting nucleoprotein complex would thus have to have struc-
tures with similar hydrodynamic ratios to actual nucleosomes.
Intracellular HSV-1 DNA pulled down using EdU-labelling and click chemistry was appar-
ently associated mostly with ICP4, VP16 and ICP22 [14]. In our analyses, ICP8 was enriched
only in the most accessible chromatin (Fig 3), whereas ICP4 fractionated similarly to HSV-1
DNA, with a shift towards the more inaccessible fractions (Fig 3). ICP4 co-localized with the
HSV-1 genomes immediately after infection, but such co-localization was disrupted as the infec-
tion progressed. At 3hpi, ICP4 was enriched at the periphery of the replication compartments,
where the HSV-1 DNA signal was relatively low [67]. It is possible that EdU-mediated purifica-
tion enriches for the most dynamic HSV-1 chromatin in which the EdU label would be more eas-
ily accessible. Such an enrichment would result in the apparent lack of enrichment in histones,
particularly under a stringent enrichment criterion [14,15] (four-fold increase in the number of
the reads over the number in cells infected with non-labelled HSV-1, a very high threshold for
most abundant nuclear proteins such as histones). The HSV-1 DNA pulled down by EdU-medi-
ated purification was associated with a number of chromatin remodelers [14], which is consistent
with the recovery of mostly HSV-1 DNA chromatinized to a highly dynamic state.
Chromatin epigenetics affect HSV-1 transcription. Inhibition of proteins involved in chro-
matin epigenetic regulation (Set1, LSD1, Co-REST, JMJD2, or CLOCK complex), by small-
HSV-1 transcriptional competence
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molecule inhibitors, knockdown, or knockout inhibits HSV-1 transcription or replication [42,
46,47,71]. The HSV-1 transcription activators ICP0, ICP4 and VP16 all interact with chroma-
tin remodelers [19,20,29,47,50,51,72]. ICP0, an E3-ubiquitin ligase, blocks histone deacety-
lation by interacting with histone deacetylases and the REST/Co-REST complexes [32,50,51,
73,74]. It also induces degradation of silencing histone variants such as Cenp-A and Cenp-B
[30,31]. VP16 recruits histone acetylases, methyltransferases and demethylases to activate
HSV-1 transcription [19,20,47]. ICP4 increases the dynamics of all core histones [7,68–70],
and interacts with a variety of chromatin remodelers, including the CLOCK complex [29]. All
the IE epigenetic regulators that functionally interact with the HSV-1 transcription regulators
could very well modulate the dynamics of the HSV-1 chromatin through lytic infections.
Dr. Enquist’s group created a pseudorabies virus (PRV) strain containing three fluorescent
proteins, each flanked by different recombination sites [75]. Infection of a cre-expressing cell
with a single genome of this PRV strain can result in expression of only one randomly recom-
bined fluorescent protein [75]. Infection of a single cell with multiple genomes would be antic-
ipated to result in expression of a mixture of all three fluorescent proteins, each from different
randomly recombined genomes. However, it resulted in expression of only one, randomly
recombined, fluorescent protein [75]. Moreover, infections with an equal mixture of PRV
expressing dTomato, EYFP, or cyan at high moi resulted in most of the infected cells express-
ing only one of them, or at most two [75,76]. The simplest mechanism leading to such effects
is that only a few viral genomes are biologically active in each infected cell [75,76]. Recent
studies found similar results for HSV-1 [77]. Infection with an equal mixture of 14 HSV-1 dif-
ferent recombinants resulted in an average of only 3.4 or 7.6 of the co-infecting genomes repli-
cated in a given cell at moi of 10 or 100, respectively [77]. The number of replicating genomes
correlated with the levels of HSV-1 transcription in each cell, suggesting that HSV-1 transcrip-
tion and replication competence are regulated together at the genome level. Inhibitors of his-
tone deacetylases (TSA, SBX or VPA) reduced the numbers of transcription competent
genomes in each infected cell [78], suggesting that HDACs modulate the transcription compe-
tence of the infecting HSV-1 genomes [78]. Histone H3.3 was also enriched in the chromatin
of entire HSV-1 genome at 2 or 6h during quiescent infections, without a significant difference
between the promoter or the gene body [79]. We show here that some HSV-1 genomes are in
most accessible chromatin whereas others are in least accessible chromatin. The accessible
genomes were enriched under conditions of ongoing HSV-1 transcription and depleted when
HSV-1 transcription was inhibited, which is consistent with the recent results in which the
decondensation of the HSV-1 genome was inhibited by restricting HSV-1 transcription using
actinomycin D or CHX [67], and the enrichment of silencing histone maker H3K9me3 on
HSV-1 chromatin during quiescent infections [79]. It is thus tempting to speculate that the
biologically active genomes are in the most accessible chromatin, and the biologically inactive
ones, in the least accessible one.
Activation of transcription HSV-1 IE, E or L genes is differentially regulated. The promot-
ers of the IE genes contain multiple transcription factors binding sites, including multiple
TAATGARAT, Oct1 and Sp1 binding sites and a TATA box. The promoters of the E genes are
simpler. They typically contain only an NF1, a Sp1 binding site, and a TATA box. Those of the
L genes are even simpler; they contain only a TATA box. HSV-1 E and L gene promoters are
thus likely unable to recruit as many transcription activators or epigenetic modifiers as the
promoters of the IE genes do. Therefore, E and L gene promoters may only be able to drive
transcription once the IE proteins have disrupted the global silencing of the viral genomes.
Unexpectedly, we found several short sequences that were relatively overrepresented in the
MCN-digested and fractionated HSV-1 chromatin, including an overrepresentation of previ-
ously described chromatin insulator-like elements and CTCF binding sites. CTCF binding
HSV-1 transcriptional competence
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sites and chromatin insulator-like elements function as boundaries between transcriptionally
active and inactive genes [59,80]. CTCF binding was enriched on latent HSV-1 DNA, and
CTCF enrichment at CTRL2, CTa’m and CTRS3 was depleted at early times of reactivation
[81]. Co-immunoprecipitation of HSV-1 genes with CTCF during lytic infections showed no
enrichment of CTCF in CTRL2, CTa’m or CTRS3 clusters [60]. CTCF was instead enriched in
the promoters of UL8, UL26 and UL36 genes, on the gene bodies of ICP0 and ICP34.5, and at
the 3‘of UL25, UL33, UL36 and UL37 genes [60]. Knockdown of CTCF increased H3K9 and
H3K27 trimethylation on the promoters and gene bodies of IE (ICP0 and ICP4), E (TK, ICP8)
and L (VP16) genes [60], which is consistent with global regulation of HSV-1 genomes by
chromatin dynamics. Consistently, studies from Han et al., showed that EGFP expression in
absence of any viral immediate early proteins increased when the CMV promoter and the
EGFP gene were flanked together by CTRL1 and CTRL2. CTRL1 and CTRL2 decreased the
levels of the repressive markers H3K9 and K27 trimethylation in the CMV promoter, but did
not alter the levels of repressive markers nor the transcription levels of the neighborhood
genes UL51 and UL52 [82]. These studies support role for CTCF in delimiting transcription
activity.
The transcribed IE genes (in CHX treated infections) were, interestingly, flanked by the rel-
atively most overrepresented short sequences. Five of these six short sequences mapped to pre-
viously experimentally mapped CTCF binding sites or chromatin insulator-like elements in
either KOS or 17 (Figs 9and 10) [59,60]. As expected, in silico analyses identify at least one
potential CTCF binding site in each of these four experimentally mapped binding sites (S8
Fig), and the ChIP assays provided experimental evidence that all six of them are indeed recog-
nized by CTCF in CHX-treated infections (Fig 10). We propose that these sequences with lim-
ited accessibility are insulator elements, separating transcribed from non-transcribed genes or
genes with vastly different transcription levels. Whether the observed CTCF binding limits the
transcription to the IE loci or is a consequence of transcription being limited to these loci
remains to be determined.
The short sequences with limited accessibility may also anchor the HSV-1 genome to the
nuclear matrix, lamin, or other nuclear structures. In silico analysis of the peaks that did not
map to previously mapped (or predicted) CTCF or insulator-like elements (http://meme-suite.
org/tools/meme) identified a TA-rich motif (TG/TTTTT/ANA/TA/G in about half of them
(25 of 49; S6C Fig). Only fourteen of the peaks in the genomic region encoding most IE genes
had neither a CTCF/Insulator or an A/T rich motif. A/T rich motifs are often present in at
least a subset of scaffold/matrix attachment regions (S/MARs) [83]. These sequences of limited
accessibility are progressively lost as the infection proceeds, perhaps reflecting the lack of
genome boundaries when the entire genomes are transcribed or fewer and fewer anchoring
points as the infection progresses. Consistently with the latter model, HSV-1 genomes are
more compacted when transcription is restricted and genomes decondense as infection pro-
gresses [67].
Several of the short sequences relatively enriched in the digested and fractionated chroma-
tin were actually underrepresented in both the undigested and unfractionated and the digested
and fractionated chromatins. The relative enrichment of these sequences in the chromatin
fractions resulted from their larger underrepresentation in the undigested and unfractionated
than in the digested and fractionated chromatin. These sequences are different from the
sequencing artifacts, which map to different loci, are conserved in CHX or Rosco-treated
infections, and are overrepresented in both in the undigested and unfractionated and digested
and fractionated chromatin (S2 and S3 Figs). Chromatin insulators bind to many chromatin
insulator binding factors, including CTCF, and interact with multiple nuclear structures to
form chromatin loops [84–86]. Like cellular DNA, intracellular HSV-1 DNA also interacts
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 30 / 37
with nuclear structures such as lamin [87]. DNA involved in these anchoring interactions is
usually difficult to extract, most likely resulting in its depletion in the unfractionated and undi-
gested DNA. Although nucleases like MCN cannot digest DNA in direct interaction with such
structures, they do digest adjacent DNA, thus reducing the size of the DNA trapped in these
interactions and therefore decreasing the underrepresentation. The 250bp slide window does
not allow for the detection of short missing DNA sequences. The DNA likely trapped in these
complex nuclear structures was more depleted in the undigested (and unfractionated) samples
than in the MCN digested (and fractionated) ones.
We propose a model whereby chromatin dynamics provides a first level of regulation of
HSV-1 gene expression. Cells attempt to silence the infecting HSV-1 genomes by chromatiniz-
ing the protein-free infecting HSV-1 DNA to a silenced compacted chromatin. If not all HSV-
1 genomes are successfully silenced, then a lytic infection is established. The promoters of the
E and L genes are unable to recruit multiple transcription factors to disrupt silencing, whereas
those of the IE genes do so via the presence of multiple transcription factors and epigenetic
modifiers binding sites. The insulator-like elements anchor the genomes to nuclear structures
and prevent transcription from extending to neighboring regions. The viral transcriptional
activators, such as ICP0 and ICP4, either as virion proteins or expressed de novo in the infected
cell, then increase the dynamics of the chromatin in entire HSV-1 genomes, resulting a more
relaxed HSV-1 chromatin state and also upregulating E and L gene transcription. Only a few
genomes need not be silenced early in infection and become biologically active, while most of
the infecting genomes would remain constrained and compacted in silenced chromatin and
remain biologically inactive.
Supporting information
S1 Fig. No HSV-1 genomic region was overrepresented in some fractions and compensa-
tively underrepresented in others. Line graphs showing the number of HSV-1 genome copy
equivalents (GCE) at each locus in each soluble fraction, the insoluble fraction, the overlap of
all insoluble and soluble fractions, and undigested and unfractionated chromatin fraction at 7
hours after infection. Y-axis in logarithmic scale. X-axes, genome position (cartoon on top).
Ins, insoluble chromatin fraction.
(TIFF)
S2 Fig. No HSV-1 genomic region was overrepresented in some fractions and compensa-
tively underrepresented in others when only the IE genes were transcribed. Line graphs
showing the number of HSV-1 genome copy equivalents (GCE) at each locus lots for each sol-
uble fraction, the insoluble fraction, the overlap of all insoluble and soluble fractions, and undi-
gested and unfractionated chromatin fraction at 7 hours after infection treated with CHX. Y-
axis in logarithmic scale. X-axes, genome position (cartoon on top). Ins, insoluble chromatin
fraction.
(TIFF)
S3 Fig. No HSV-1 genomic region was overrepresented in some fractions and compensa-
tively underrepresented in others when HSV-1 transcription was restricted. Line graphs
showing the number of HSV-1 genome copy equivalents (GCE) at each locus in each soluble
fraction, the insoluble fraction, the overlap of all insoluble and soluble fractions, and undi-
gested and unfractionated chromatin fraction at 7 hours after infection treated with Rosco. Y-
axis in logarithmic scale. X-axes, genome position (cartoon on top). Ins, insoluble chromatin
fraction.
(TIFF)
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 31 / 37
S4 Fig. The overrepresented peaks result from fewer DNA reads in the unfractionated,
undigested HSV-1 DNA in untreated infections or infections treated with Rosco or CHX.
Line graphs showing the number of HSV-1 genome copy equivalents (GCE) in each genome
position in all fractions (blue) and in the undigested and unfractionated chromatin (black), in
untreated infections, or infections treated with CHX or Rosco. Y-axis in logarithmic scale. X-
axes, genome position (cartoon on top); upward arrows, the peaks overrepresented in Fig 9;
purple bars underneath genome plots, IE genes; dark green bars underneath genome plots,
LAT; light green bar, stable LAT.
(TIFF)
S5 Fig. The overrepresented peaks result from fewer DNA reads in the unfractionated,
undigested HSV-1 DNA as infection progresses. Line graphs showing the number of HSV-1
genome copy equivalents (GCE) in each genome position in all fractions (blue) and in the
undigested and unfractionated chromatin (black), in 2, 4, or 15hpi. Y-axis in logarithmic scale.
X-axes, genome position (cartoon on top); upward arrows, the peaks overrepresented in Fig
11; purple bars underneath genome plots, IE genes; dark green bars underneath genome plots,
LAT; light green bar, stable LAT.
(TIFF)
S6 Fig. Many of the short DNA sequences with limited accessibility not previously mapped
to CTCF or insulator binding sites contain predicted CTCF binding sites or T/A-rich
motifs. (A) Line graphs showing HSV-1 number of genome copy equivalents (GCE) in each
genome position in infections treated with CHX, showing two potential CTCF binding sites
(CGCCCCCTTGGGGC; GAACTGCC) as predicted by CTCFBSDB 2.0 (http://insulatordb.
uthsc.edu/home_new.php). �:these data are from Fig 11 presented again for clarity, as this in
silico analysis has no experimental support at this time. (B) Line graphs showing number of
HSV-1 genome copy equivalents in each genome position at 2 and 4 hpi showing the 25 poten-
tial T/A rich motifs as predicted by MEME (http://meme-suite.org/tools/meme). �: these data
are from Fig 10 presented again for clarity, as this in silico analysis has no experimental sup-
port at this time. Red bar, a copy of the consensus sequence. (C) The predicted consensus
sequence.
(TIFF)
S7 Fig. VP5 was below detection levels in any fraction. Western blots of VP5, in the insoluble
chromatin and all soluble chromatin fractions after serial MCN digestion and sucrose centrifu-
gation. Results of four independent experiments. The three top blots are over-exposed to better
show the lack of signal.
(TIFF)
S8 Fig. In silico predicted CTCF binding sites in the six sequences overrepresented in the
digested and fractionated chromatin over the undigested and unfractionated one. The line
graph from Fig 10 showing the HSV-1 number of genome copy equivalents (GCE) in each
genome position in infections treated with CHX, are presented again to show the predicted
CTCF binding sites in each of the overrepresented sequences. In silico prediction was performed
with CTCFBSDB 2.0 (http://insulatordb.uthsc.edu/home_new.php). �:these data are from Fig
10 and are presented again for clarity, to separate experimental results from in silico predictions.
(TIFF)
Author Contributions
Conceptualization: Luis M. Schang.
HSV-1 transcriptional competence
PLOS Pathogens | https://doi.org/10.1371/journal.ppat.1008076 November 14, 2019 32 / 37
Data curation: MiYao Hu, Daniel P. Depledge, Esteban Flores Cortes, Luis M. Schang.
Formal analysis: MiYao Hu, Daniel P. Depledge, Esteban Flores Cortes, Luis M. Schang.
Funding acquisition: Judith Breuer, Luis M. Schang.
Investigation: MiYao Hu, Daniel P. Depledge, Esteban Flores Cortes, Luis M. Schang.
Methodology: Daniel P. Depledge, Esteban Flores Cortes, Luis M. Schang.
Project administration: Judith Breuer, Luis M. Schang.
Resources: Judith Breuer.
Software: Daniel P. Depledge.
Supervision: Luis M. Schang.
Validation: MiYao Hu, Daniel P. Depledge, Luis M. Schang.
Visualization: MiYao Hu, Daniel P. Depledge, Luis M. Schang.
Writing – original draft: MiYao Hu, Daniel P. Depledge, Esteban Flores Cortes.
Writing – review & editing: MiYao Hu, Daniel P. Depledge, Esteban Flores Cortes, Judith
Breuer, Luis M. Schang.
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