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Blood First Edition Paper, prepublished online March 7, 2017; DOI 10.1182/blood-2016-06-725093
Ssb1 and Ssb2 cooperate to regulate mouse hematopoietic stem and progenitor cells by
resolving replicative stress
Wei Shi1,15,*, Therese Vu1,2,12, Didier Boucher1,13, Anna Biernacka3, Jules Nde4, Raj K Pandita5,
Jasmin Straube1, Glen M Boyle1, Fares Al-Ejeh1, Purba Nag1,6, Jessie Jeffery1, Janelle L Harris1,
Amanda L Bain1, Marta Grzelak3, Magdalena Skrzypczak3, Abhishek Mitra4, Norbert Dojer4,14,
Nicola Crosetto7, Nicole Cloonan1, Olivier J Becherel2,8, John Finnie9, Jeffrey R Skaar10, Carl R
Walkley11, Tej K Pandita5, Maga Rowicka4, Krzysztof Ginalski3, Steven W Lane1,2,12*, Kum Kum
Khanna1,*
1.
QIMR
Berghofer
Medical
Research
Institute,
300
Herston
Road,
Herston
QLD
4006,
Australia
2.
University of Queensland, Brisbane, Australia
3.
Laboratory
of
Bioinformatics
and
Systems
Biology,
Centre
of
New
Technologies,
University of Warsaw, Zwirki i Wigury 93, 02-089 Warsaw, Poland
4.
Dept.
of
Biochemistry
&
Molecular
Biology,
Institute
for
Translational
Sciences,
University of Texas Medical Branch, Galveston, TX 77555-1071, USA
5.
Department of Radiation Oncology, Houston Methodist Research Institute, Houston, TX,
77030, USA.
6.
School of Natural Sciences, Griffith University, 170 Kessels Road, Nathan Brisbane QLD
4111, Australia
7.
Science for Life Laboratory, Division of Translational Medicine and Chemical Biology,
Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm,
Sweden
8.
UQ Centre for Clinical Research (UQCCR), Cancer and Neuroscience, The University of
Queensland, Brisbane, QLD 4029, Australia
9.
SA
Pathology
and
School
of
Veterinary
Science,
University
of
Adelaide,
Adelaide,
Australia
10. Department of Pathology, NYU Cancer Institute, New York University School of Medicine,
522 First Avenue, New York, NY 10016, USA
11. St Vincent's Institute of Medical Research and Department of Medicine, St Vincent’s
Hospital, University of Melbourne, 9 Princes St, Fitzroy 3065, Australia
12. Department of Haematology, Royal Brisbane and Women’s Hospital. Brisbane, Australia
13. Current affiliation: Cancer & Ageing Research Program, Institute of Health and
Biomedical Innovation, Translational Research Institute, Queensland University of
Technology, Brisbane, Australia.
14. Current affiliation: Institute of Informatics, University of Warsaw, Banacha 2, 02-097
Warszawa, Poland
15. . Co-first author
1
Copyright © 2017 American Society of Hematology
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*Correspondence:
Wei Shi, Ph.D., [email protected]
Steven Lane, MBBS, Ph.D., [email protected]
Kum Kum Khanna, Ph.D., [email protected]
Short Title: Ssb1 and Ssb2 regulate mouse HSPCs
Text Word Count: 4457
Abstract Word Count: 217
Figures/Tables: 7+7/7
Reference Count: 67
Scientific category chosen during submission: Hematopoiesis and Stem Cells
2
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Key Points
Combined loss of
Ssb1/Ssb2
induces rapid lethality due to replication stress associated loss of
hematopoietic stem and progenitor cells.
Functionally, loss of
Ssb1/Ssb2
activates p53 and IFN pathways causing enforced cell cycling in
quiescent HSPCs and apoptotic cell loss.
Abstract
Hematopoietic stem and progenitor cells (HSPCs) are vulnerable to endogenous damage and
defects in DNA repair can limit their function. The two single-stranded DNA binding proteins
SSB1 and SSB2 are crucial regulators of the DNA damage response; however their overlapping
roles during normal physiology are incompletely understood. We generated mice where both
Ssb1
and
Ssb2
were constitutively or conditionally deleted. Constitutive
Ssb1/Ssb2
double
knockout (DKO) caused early embryonic lethality, while conditional Ssb1/Ssb2 double knockout
(cDKO) in adult mice resulted in acute lethality due to bone marrow failure and intestinal
atrophy featuring stem and progenitor cell depletion, a phenotype unexpected from the
previously reported single knockout models of
Ssb1
or
. Mechanistically, cDKO HSPCs
Ssb2
showed altered replication fork dynamics, massive accumulation of DNA damage, genome-wide
double strand breaks (DSBs) enriched at Ssb binding regions and CpG islands, together with the
accumulation of R-loops and cytosolic ssDNA. Transcriptional profiling of cDKO HSPCs revealed
the activation of p53 and interferon pathways which enforced cell cycling in quiescent HSPCs
resulting in their apoptotic death. The rapid cell death phenotype was reproducible in
in-vitro
cultured cDKO-HSCs, which was significantly rescued by nucleotide supplementation or after
depletion of p53. Collectively, Ssb1 and Ssb2 control crucial aspects of HSPCs function including
proliferation and survival in vivo by resolving replicative stress to maintain genomic stability.
3
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Introduction
The ability to maintain genome integrity upon endogenous DNA damage is critical for cell
survival, self-renewal, proliferation and differentiation. Cells employ a tightly-coordinated DNA
damage response (DDR) to either remove or repair the damage or activate an apoptotic cell
death program. A defective DDR underlies a number of human diseases and developmental
disorders.
A key class of proteins involved in the DDR are the single-stranded DNA (ssDNA) binding
proteins (SSBs), which are recruited to DNA damage sites to protect ssDNA1. Replication Protein
A (RPA) was previously believed to be the sole SSB protein complex in eukaryotes, essential for
DNA replication, repair and recombination, and modulation of gene expression2. Our group
identified two additional human SSB proteins, designated as SSB1 and SSB2 (also known as
NABP2/OBFC2B/SOSS-B1 and NABP1/OBFC2A/SOSS-B2) conserved from archaea to mammals3.
SSB1 and SSB2 share 73% sequence identity, with highly conserved N-terminal OB-fold domains
but divergent C-terminal regions. Depletion of SSB1 in cells results in increased radiosensitivity,
defective repair of DNA double strand breaks (DSBs), oxidative DNA damage and failure to
restart stalled replication forks3-8. Moreover, Ssb1 has been shown to mediate telomere
homeostasis by protecting newly replicated G-overhangs of leading- and lagging-strand
telomeres9,10.
SSB1
is recurrently mutated in various cancers while an
SSB2/RARA
fusion gene
has been described in variant acute promyelocytic leukemia11.
SSB1 and SSB2 independently form complexes with C9Orf80/INIP and INTS3, a component of
Integrator Complex4-6. The Integrator Complex is a 14 subunit, RNA Polymerase II binding
complex that controls the 3’ end processing of small-nuclear RNAs12. Recent studies indicate
that the Integrator Complex is required in many steps of the transcription cycle: 3’-end
processing and termination of non-polyadenylated snRNA and replicative histone genes, pause
release at immediate early genes, and biogenesis of transcripts required from distal regulatory
elements (enhancers)13-17. The association of SSB1/2 with the INTS3 complex indicates the
potential for SSBs to influence transcription and RNA processing15. Furthermore, the target sites
of INTS3-SSB complexes are favorable to the formation of DNA:RNA hybrids (R-loops),
structures in which nascent RNA transcripts fall back on the template DNA, leaving the non4
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template ssDNA exposed18. R-loops are formed normally during transcription and if not
resolved properly, can become a source of genomic instability19.
In mice, Ssb1 is ubiquitously expressed whereas Ssb2 is mainly expressed in the thymus and
testis. Deletion of
Ssb1
dorsal rib cage9,20-22.
leads to perinatal lethality due to highly abnormal patterning of the
Ssb1
conditional knockout
20
or
Ssb1
hypomorphic mice9 are viable long
term and show increased tumor incidence after late latency and are radiosensitive. However,
Ssb2
knockout mice develop to term and have no overt pathological phenotype23. Strikingly,
Ssb2 shows pronounced upregulation in
and hypomorphic
Ssb1
F/F
thymus and spleen from
−/−
Ssb1
tissues, mouse embryonic fibroblasts (MEFs),
tissues9,20,21, while a modest upregulation of Ssb1 is observed in
Ssb2-/-
mice and
Ssb2-/-
MEFs23. This compensatory upregulation
suggests Ssb1 and Ssb2 may have overlapping functions in vivo.
Here, we report that constitutive
Ssb1/Ssb2
double knockout (DKO) mice are early embryonic
lethal, whilst conditional Ssb1/Ssb2 double knockout (cDKO) in adult mice results in unexpected
acute bone marrow failure and intestinal atrophy due to loss of rapidly proliferating progenitor
cell populations, phenotypes which are reminiscent of acute radiation toxicity. We observed
replication stress, DSBs and R-loop accumulation accompanied by transcriptional activation of
p53 and interferon (IFN) pathways in cDKO HSPCs. This resulted in enforced cell cycle entry of
quiescent HSPCs followed by apoptotic cell death. In conclusion, Ssb1 and Ssb2 coordinately
restrict HSPC proliferation and promote HSPC survival by resolving replication/transcription
associated DNA damage and R-loop accumulation.
5
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Materials and Methods
Experimental mice and phenotypic analysis
All experimental animals were maintained on a C57BL/6J strain in a pathogen-free animal
facility. All procedures were approved by the QIMR Berghofer animal ethics committee
(A11605M and A0707-606M). Peripheral blood was collected by retro-orbital venous blood
sampling and analyzed on a Hemavet analyzer (Drew Scientific). Tissues were collected and
fixed in 10% buffered formalin fixative, embedded in paraffin blocks and stained with
hematoxylin and eosin (H&E) for histological examination. Immunostaining methods and
antibodies used are described in supplemental experimental procedures. All Western analyses
were performed on the LICOR platform (Biosciences). Bone marrow (BM) cells were harvested
by flushing femur and tibia bones. Various BM stem and progenitor populations were purified
as described24. For cell cycle analysis, cells were fixed and permeabilized (FIX & PERM kit,
Invitrogen) and stained with Ki-67 (B56) and Hoechst 33342 (20ug/ml, Invitrogen). All flow
cytometric analysis was performed on a FACS LSR Fortessa (BD Biosciences).
Competitive BM transplantation
BM cells derived from 6- to 8-week-old control or cDKO mice (expressing CD45.2) were
combined with equal numbers of CD45.1 congenic competitor bone marrow cells, and injected
into the lateral tail vein of lethally irradiated (11 Gy in 2 separate fractions at least 3 h apart)
CD45.1/CD45.2 congenic recipient mice (Animal Resource Centre, Western Australia).
In vitro
apoptosis rescue assay
T2
BM cells were harvested under sterile conditions from naïve Rosa26-CreER
(n=5) and
T2
Rosa26-CreER
flfl
;Ssb1
fl/fl
Ssb2
+/+
;Ssb1
+/+
Ssb2
mice
mice (n=5). LKS cells were purified as previously
described24. Retroviral Hoxb8-producing fibroblasts were seeded in a 10cm plate at 1x105 in
low glucose DMEM supplemented with 10% FCS. After 24 h, 5x105 sorted LKS cells were
cultured atop a layer of Hoxb8-transformed fibroblasts in the presence of 0.25ng/ml IL-325.
After 4 days in culture, non-adherent cells were passaged into 12-well plates and used in
subsequent apoptosis assays by staining with Annexin V (BD Biosciences) and Sytox blue
(Invitrogen). EmbryoMax® nucleoside supplement (Merck Millipore) was added to individual
wells where indicated at 1:10026,27. To knockdown p53, cells were plated on Retronectin-coated
6
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plates (Takara) and spinoculated with lentiviral p53-shRNA28 or luciferase-shRNA (control) at a
MOI of 10, in the presence of 4ug/ml polybrene at 2,500 RPM at 30ºC for 90 mins.
DNA damage and genomic instability analysis
For immunostaining, DNA combing and comet assay on HSPCs, whole BM were harvested from
Ctrl and cDKO littermate mice at 48 h post 4 mg tamoxifen (TAM), and sorted for LKS+
(lineagelowc-Kit+Sca-1+) cells. Cells were cultured for 16 h prior to processing as described29.
Preparation of metaphases and chromosome aberration analysis was done as described20,30.
Telomere fluorescence in situ hybridization (FISH) at metaphases was performed as described
previously31.
Direct
in situ
single-nucleotide resolution labeling and capture of genome-wide DSBs in nuclei
were performed using BLESS technique as described previously and in Supplementary
Experimental Procedures32.
7
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Results
Ssb1 and Ssb2 are essential for early embryogenesis
+/-
Ssb1
Ssb2
-/-
mice were intercrossed to generate embryos at specific stages. Timed matings
revealed that, whilst Ssb1+/+Ssb2-/-,
+/-
Ssb1
Ssb2
-/-
and Ssb1-/-Ssb2-/- (DKO) were recovered at the
expected Mendelian ratios at E7.5, no DKO embryos could be recovered at E10.5 (Figure 1A and
quantified in Table S1). Instead, resorbed embryos with apoptotic bodies were observed in the
expected proportion (Figure 1B).
Somatic deletion of Ssb1 and Ssb2 in adult mice triggers rapid lethality
We employed a conditional approach to delete Ssb1 and Ssb2 across a broad range of tissues in
adult mice using the TAM-inducible
Ssb2
were generated as follows:
(1ko,2het),
fl/+
Ssb1
fl/fl
Ssb2
-CreERT2 strain33. Individual genotypes of
Ssb1
Rosa26
+/+
Ssb1
+/+
Ssb2
(1het,2ko) and
(WT),
fl/fl
Ssb1
fl/fl
Ssb2
fl/+
Ssb1
Ssb2
fl/+
(Dbl het),
fl/fl
Ssb1
and
fl/+
Ssb2
(cDKO). Cre-mediated recombination
was induced with TAM (1mg/d by intraperitoneal injection for five consecutive days). cDKO
mice displayed 15% body weight loss within 7 days post TAM induction and became moribund,
whereas mice of all the other genotypes maintained normal body weight in the same period
(Figures 1C, S1A and S1B). The knockout efficacy in BM, spleen and thymus was confirmed
(Figures S1C).
cDKO causes bone marrow failure and small intestine atrophy
cDKO spleens and thymuses were smaller and paler than controls (Figures 1D and S1D). cDKO
small intestines showed profound shortening of villi and marked thinning of the mucosa,
resembling villous atrophy due to damage to crypt resident proliferative progenitors (Figure
34,35
1E)
while other tissues remained grossly intact (data not shown). BM hypocellularity was
observed in cDKO sections, featuring trilineage reduction in hematopoiesis with fatty
replacement (Figures 1F and 1G). Analysis of the cDKO peripheral blood showed leukocytopenia
and anemia (Figures 1H, 1I, S1E and S1F), with preservation of platelets (Figure S1G). The acute
BM and intestinal damage of both highly proliferative tissues is comparable to that found in
acute ionizing radiation toxicity36. These findings indicate that
Ssb1
and
Ssb2
are collectively
essential for maintaining tissue homeostasis in vivo.
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cDKO causes loss of hematopoietic stem and progenitor cells (HSPC) by apoptosis and impairs
long term hematopoietic stem cell (LTHSC) function
All blood cell lineages are thought to be derived from LTHSCs, which have the ability to selfrenew and differentiate into multipotent and lineage-committed blood cells37. Recent data
suggests homeostatic hematopoiesis is supported primarily by progenitor populations38,39. The
longevity and proliferative capacity of myeloid progenitor cells renders them particularly
susceptible to DNA damage40. Analysis of BM populations by flow cytometry revealed a
dramatic reduction in committed myeloid progenitor cells (Lineagelowc-Kit+Sca-1-; LKS-) in cDKO
BM (Figure 2A and S2A), while lineagelowc-Kit+Sca-1+ (LKS+, enriched for HSPC) cDKO cells were
expanded in frequency, but not absolute number due to BM hypocellularity (Figure 2A), and
exhibited a marked induction of Sca-1 expression and slight decrease in c-Kit intensity (Figure
S2A). Phenotypic common myeloid progenitors (CMP; lineagelowc-Kit+Sca-1-CD34+CD16/32-) and
granulocyte-macrophage
progenitors
(GMP;
lineagelowc-Kit+Sca-1-CD34+CD16/32+)
were
proportionally expanded in cDKO BM (Figures S2A and S2B) but reduced in absolute numbers
compared to WT (Figure 2A). Megakaryocyte-erythroid progenitors (MEP; lineagelowc-Kit+Sca-1CD34-CD16/32-) were drastically diminished in both frequency and absolute number in cDKO
BM (Figures 2B and S2A). Long-term hematopoietic stem cells (LTHSCs; lineagelowc-Kit+Sca1+CD150+CD48-) were markedly reduced in cDKO BM (Figures 2C and S2A). To exclude the
impact of Sca-1 induction in cDKO BM, we quantified lineagelowc-Kit+CD150+CD48- cells. Indeed,
we still observed a profound reduction of lineage low c-Kit+ cells and lineage low cKit+CD150+CD48- cells in cDKO BM (Figure S2C and S2D). Furthermore, both cDKO LKS+ and
CD150+ cells showed a marked increase in Annexin V+ cells (Figures 2D, 2E and S2E), suggesting
that cDKO BM cells were depleted due to apoptotic cell death. Furthermore, we found that
expression of a single allele of either Ssb1 or Ssb2 is sufficient to rescue the cell loss phenotype
and restore peripheral blood leukocytes/hematocrit and BM LKS+/LTHSCs (Figures S2F-a-d).
To investigate the functional potential of cDKO LTHSCs in greater depth, we performed
in vivo
competitive bone marrow transplantation assays (Figure 2F). Once equivalent engraftment of
donor cells of all genotypes was confirmed at 4 weeks, the recipient mice were treated with
1mg of TAM once daily for 5 days. The induced genetic deletion of
Ssb1
and
Ssb2
led to the
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gradual depletion of cDKO donor cell chimerism, as evidenced by the progressive loss of CD45.2
cells from the peripheral blood, while all other genotype cohorts sustained stable engraftment
for 24 weeks post-transplant (Figure 2G). When the BM was assessed in the recipients at 24
weeks post-transplant, cDKO CD45.2, LKS- and LKS+ populations were dramatically reduced
compared to other groups, indicating that cDKO cells were unable to sustain long-term
hematopoiesis (Figures S2G, S2H and S2I). These findings demonstrate that both Ssb1 and Ssb2
are required for in vivo HSPC maintenance in a cell-autonomous manner.
cDKO causes replication stress and DNA damage in HSPCs
To shed light on the mechanism of HSPCs loss, we analyzed DNA replication fork dynamics. BM
LKS+ cells were isolated from Ctrl or cDKO mice at 48 h after one injection of 4 mg TAM,
cultured for 16 h and pulse labeled with the thymidine analogue chlorodeoxyuridine (CldU),
followed by iododeoxyuridine (IdU). DNA was combed onto glass slides and CldU and IdU
incorporation in nascent DNA fibers was detected by fluorescent staining (Figure 3A)29. Ctrl
LKS+ cells displayed longer elongating fiber lengths representing a fork speed distribution
centered around a mean fork velocity of 1.61 kb/min, whilst cDKO cells had shorter fiber
lengths overall with a mean velocity of 0.99 kb/min, highlighting a significantly slower DNA
replication fork rate in cDKO cells (Figure 3B). We did not observe asymmetric replication in
cDKO cells as found previously in old HSCs (Figure S3A)29. We did however observe a
significantly increased percentage of stalled replication forks (Figure 3C). These findings
correlated with a strong enrichment of RPA foci and phosphorylated RPA (S4/8) in cDKO HSPCs
(Figures 3D and 3E), indicating the presence of extensive ssDNA at replication forks, a hallmark
of replication stress.
Altered DNA replication, in particular increases in DNA replication stalling events, can generate
DSBs. Consistent with this, we observed focal accumulation of the DSB marker phosphorylated
H2AX (γH2AX) in cDKO HSPCs (Figures S3B-G), suggesting defective DSB repair. Furthermore,
using alkaline comet assays, we showed that cDKO LKS+ cells displayed higher baseline and
radiation-induced DNA damage compared to control cells (Figures 3F and 3G). cDKO bone
marrow metaphases demonstrated a significant increase in spontaneous chromatid and
chromosomal breakage (Figures 3H-J), as well as telomere signal loss (Figures 3K and 3L), which
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was only evident in the conditional
Ssb1-/-
genotoxic insult20. These results provide
bone marrow metaphases after exposure to
in vivo
evidence to suggest that Ssb1 and Ssb2 may
have overlapping roles in regulating HSPC function in genome maintenance by resolving
replication stress.
To investigate if replication stress is causative of apoptotic death in cDKO LKS+ cells, we first
established Hoxb8-immortalized LKS+ cells
in vitro
to directly monitor their growth and cell
death after 4OHT induction, independent of what is likely a strongly pro-inflammatory
environment in the cDKO animal. Overexpression of Hoxb8 has been shown to immortalize
interleukin-3 (IL-3)-dependent myeloid progenitor cells by blocking differentiation of these cells
25,41
to arrest them in a self-renewing state
induce apoptosis
in
. 4-OHT-mediated cDKO in LKS+ cells was sufficient to
by day 6 (Figure S3H). We next treated cells with EmbryoMax®
vitro
nucleosides (Merck Millipore) to relieve cDKO cells of replicative stress26,27 and observed
decreased apoptosis in 4OHT-induced cDKO cells supplemented with nucleosides compared to
those treated with vehicle (Figure 3M and S3I). These findings demonstrate that replication
stress is a major effector of cell death in cDKO.
cDKO activates the interferon and p53-mediated apoptosis pathways in HSPCs
Next, we performed microarray analysis on purified MEP and GMPs from Ctrl and cDKO mice
two days after TAM induction to gain insight into the molecular determinants and cellular
pathways involved. The key upregulated genes in cDKO MEP and GMP were interferon-α and -β
(IFNα/β) and p53 target genes (Figure 4A), which belong to an integrated network (Figure 4B).
Notably,
Ssb1
(Obfc2b) was the most downregulated gene (Figure 4A). Ingenuity Pathway
Analysis (IPA) identified p53, antigen presentation, aryl hydrocarbon receptor (AhR), and ATM
signalling as the top canonical pathways affected by cDKO in MEP or GMP (Figures 4C). Gene
Set Enrichment Analysis (GSEA) also revealed enrichment of apoptosis and immune response
functions in cDKO regulated genes (data not shown). This finding was validated by quantitative
real time PCR (qPCR) on independent samples where cDKO HSPC exhibited significant
downregulation of
and
Ssb1
Ssb2
(data not shown) and upregulation of IFN regulated
transcripts (Myd88, Oas1g, Oas2, Ifitm3, and Irgm1, Figures S4A-B) and p53 DNA damage genes
(JunB,
,
Gadd45b
,
Gadd45g
, and
Mdm2
Cdkn1a,
Figures S4C-D). These analyses suggest that
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depletion of both Ssb1 and SSb2 activates transcriptional IFNα/β and p53 responses which may
converge to induce apoptosis.
To validate if IFN and p53 were also activated in cDKO HSCs, we performed RNAseq analysis on
sorted CD48-LineagelowcKit+ESAM+ (LKE+; ESAM used in place of Sca142) HSCs from Rosa26CreERT2Ssb1+/+/Ssb2+/+ (Ctrl) and Rosa26-CreERT2Ssb1 fl/fl/Ssb2fl/fl (cDKO) mice at 48 h post TAM
induction. Gene expression was distinct for each genotype (Figure S4E). GSEA analysis revealed
the loss of a LKS self-renewal signature43 in cDKO HSCs (Figure 4D). p53 pathway was
upregulated in cDKO HSCs (Figure 4E) and the IFN genes were dramatically enriched in cDKO
HSCs (Figures 4F-G, S4E-F) suggesting transcriptional activation of IFN and p53 pathways
accompany this replicative and pro-apoptotic phenotype. Similarly, this IFN gene signature is
present in in vitro cultures of Hoxb8-immortalized LKS+ cDKO cells after 4OHT induction (Figure
S4G), indicating the HSC-autonomous nature of IFN production.
Cytosolic ssDNA in cDKO cells prime HSCs to exit quiescence and enter the cell cycle
Accumulation of unrepaired DNA lesions has recently been shown to activate the type I IFN
pathway and inhibit stem cell function through the release of ssDNA into the cytoplasm44-46.
Consistent with this, cDKO HSPCs and BM sections showed the presence of ssDNA in the
cytoplasm (Figures 5A and S5A), like that of positive control cells (Ctrl HSPCs treated with
Aphidicolin or LPS) (Figure S5B). Functionally, IFN signaling is reported to force HSCs to exit
quiescence and enter the cell cycle, leaving them vulnerable to DNA damage47,48. We analyzed
the cell cycle profile of BM LKE+
42
from day 0 to day 4 after a single dose of 4 mg TAM IP
injection. Notably, a significantly increased percentage of cDKO LKE+ cells lost quiescence (G0)
and became more proliferative (S/G2/M) on D3 and D4 (Figure 5B). This finding was validated in
highly enriched HSC fractions LineagelowcKit+CD150+CD48- on D3 post cDKO (Figure S5C).
Similarly, immunostaining on BM tissues from the corresponding time course showed a
transient burst of proliferation followed by p53 stabilization and apoptotic cell death (Figures
5C-D).
In an effort to rescue the phenotype by knocking down p53, LKS+ cells were isolated from a
non-TAM treated Rosa26CreERT2Ssb1fl/flSsb2fl/fl donor mouse and transduced with control
(shLuc-GFP) or p53 shRNA (shP53-GFP)28, sorted for GFP+ cells, combined with competitor
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CD45.1 BM cells, and then transplanted into irradiated syngeneic WT recipient mice (Figure
S5D). TAM was injected 4 weeks post-transplant. A trend of increased reconstitution capacity
was observed after knocking down p53 (Figure S5E) although this was inadequate to rescue
function long-term (Figure 5E), suggesting that either additional pathways limit HSPC function
in cDKO HPSC or complete p53 knockout may be required for full rescue. To demonstrate p53mediated apoptosis is functional short-term in cDKO, we transduced Hoxb8-immortalized LKS+
cells with control (shLuc-GFP) or p53 (shP53-GFP) shRNAs. We observed a significant increase in
apoptosis in 4OHT-induced shLuc-transfected cDKO cells, but no increase in 4OHT-induced cells
transfected with shP53, suggesting that p53 knockdown
in vitro
is able to temporarily prevent
cDKO-induced cell death (Figure 5F and S5F).
cDKO induces specific genome-wide double strand breaks enriched at CpG islands and tRNAs
To map the distribution of genome-wide DSBs at nucleotide (nt) resolution, direct in situ Breaks
Labeling, Enrichment on Streptavidin and next-generation Sequencing (BLESS)32,49 was applied
on whole BM from control and induced cDKO mice at day 4 after a single dose of 4 mg TAM IP
injection (Figure 6A).
At the resolution of 1250 nt and at a p-value threshold of p=0.001 (hypergeometric test) we
detected 43757 genomic regions enriched in spontaneous DSBs (in control BM cells) and 2053
cDKO-induced fragile regions (Supplementary Webpage, Table S3). Strikingly, cDKO-induced
breaks occur in different locations than spontaneous DSBs in control BM cells (hypergeometric
test, p-value < 10-323) (Figure 6B), and did not correlate with previously described common
fragile sites50,51. Fragility of whole genes was independent of gene length (Supplementary
webpage, Table S4). Strikingly, we observed very significant enrichment of CpG islands at
different resolutions in DSB enriched regions (Figures 6C). We also detected borderline
significant enrichment of cDKO-induced DSBs in 2kb promoter-proximal region (Figure S6A).
Moreover, we observed significant enrichment of DSBs in the vicinity of sentinel highly
expressed genes (tRNAs) and all transcripts, versus lowly expressed genes (retrogenes),
suggesting that cDKO associated DSBs are related to highly expressed genes (Figures S6B and
S6C). We also analyzed co-localization of DSBs and SNPs, to gain evolutionary perspective.
Common SNPs were 1.2-fold enriched and SNPs in coding regions showed up to 2.7-fold
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enrichment at cDKO associated breaks (Figure S6D). Again, enrichment was highest within a 1
to 3kb vicinity of a coding SNP.
Furthermore, we reanalyzed HIT-Seq data, where Skaar et al. have mapped genome-wide
binding sites of Ssb1 and Ssb215, using the same method applied for the BLESS data and
analyzed the enrichment of Ssb1 and Ssb2 binding sites (from HIT-Seq data) in the regions
enriched with cDKO-induced DSBs (from BLESS data) at different resolutions. We observed
significant enrichment of Ssb1 and Ssb2 binding sites in intervals enriched with cDKO-induced
DSBs (Figure 6D and Table S6). Moreover, we detected enrichment of CpG islands in intervals
enriched with Ssb1 and Ssb2 binding sites (Figure 6E and Table S7) as well as in those enriched
with the cDKO-induced DSBs (Figure 6C). When we specifically considered the aforementioned
LKS (stemness) signature (Figure 4D), there were nine genes allocated within the regions
enriched with cDKO specific DSBs identified by BLESS. Amongst these genes, seven were
differentially expressed with six showing a decrease in expression in cDKO compared to Ctrl
(FDR =<0.05). The exception was Gbp10, an interferon inducible gene, which was predictably
increased (Figure S6E).
Altogether, assuming that induction of DSBs is rare and the ones we observe occur in a small
fraction of cells and are detectable only due to very sensitive BLESS detection technique, we
hypothesize that the deleterious effects of cDKO-induced DSBs occur initially in the context of
transcriptionally active genes, but also through dominant effects of downstream transcriptional
pathways such as the IFN and p53 pathways.
cDKO leads to R-loop accumulation
INTS3-SSB-complexes have previously been shown to be enriched at the GC-rich regions of
transcription start sites (TSS) and transcription termination sites (TTS) as well as open
18,52,53
chromatin states favourable to R-loop formation
. Interestingly, R-loops are also
commonly observed at CpG islands, where significant DSB accumulation was observed in cDKO
BM cells by BLESS. We assessed the formation of R-loops and γH2AX on BM tissue sections. Rloop accumulation was observed between 1 and 2 days following 4 mg TAM injection and
increased with time post cDKO. γH2AX staining overlapped with R-loop staining and showed a
similar increase during the time course in BM (Figures 7A-C), small intestine (Figures S7A-C),
14
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spleen, and thymus (data not shown). Furthermore, RNase H treatment markedly reduced the
intensity of the R-loop signal, validating the specificity of antibody for R-loop detection (Figure
S7D). Altogether, these results suggest that R-loop accumulation is one of the early events
when both Ssb1 and Ssb2 are disrupted, which accompanies DSB generated most likely due to
conflict with replication.
15
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DISCUSSION
SSB1 and SSB2 have crucial roles in the repair of extrinsic DNA damage in human cells3-8. Our
generation and phenotypic analysis of cDKO mice has unmasked the compensatory and
essential functions of Ssb1/Ssb2 in maintaining tissue homeostasis, which was unexpected from
analysis of single knockout mouse models20,23. Here, we studied hematopoiesis to demonstrate
the requirement for Ssb1 and Ssb2 in protecting stem and progenitor cells from endogenous
DNA damage. Our study is the first to report acute BM failure and severe intestinal atrophy due
to stem and progenitor cell death in
Ssb1/Ssb2
cDKO mice. The observed phenotypes likely
manifest due to the rapidly dividing nature of BM and small intestine, however, the effects of
cDKO are likely to be relevant to other rapidly dividing cells in culture. Consistent with this, we
observed that
in vivo
in vitro
cultured Hoxb8-immortalized cDKO HSPCs corroborate and validate our
findings in BM.
Functionally, cDKO triggers quiescent HSPCs to proliferate followed by apoptotic cell loss, which
ultimately results in hematopoietic failure. We observed that HSPCs have an intrinsic
requirement for Ssb1/Ssb2 as their deletion resulted in a loss of hematopoiesis in BM
transplant recipients. Mechanistically, we observed replication stress, DSBs and cytosolic ssDNA
accumulation, intrinsic transcriptional activation of interferon, p53 and DNA damage pathways
in cDKO HSPCs, followed by induction of significant cell death and BM failure. Moreover, in vitro
cultured Hoxb8-immortalized HSC undergo significant cell death within 6 days of induction of
cDKO. This phenotype can be rescued by supplementation with nucleosides or by knockdown of
p53, suggesting that replication stress associated DNA damage and p53 induction are major
effectors of cell death in cDKO cells. We propose that ssDNA generated from unrepaired DNA
44-46
damage induces cell-intrinsic activation of the interferon pathway
, which perturbs HSC
quiescence, primes HSPCs for p53-mediated apoptotic cell death and ultimately contributes to
HSPC depletion (Figure 7D)29,48,54. These findings indicate that Ssb1 and Ssb2 coordinately
protect organs from endogenous replication stress during normal physiology and are essential
genome guardians for homeostasis of BM stem and progenitor cells.
The acute BM failure seen in adult cDKO mice was completely penetrant, as compared to Ssb1
or Ssb2 single knockouts or knockouts of other DNA repair genes including genes in the Fanconi
16
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anemia (FA) pathway, which show much milder phenotypes and longer latencies that in some
cases are only evident during aging or stress55-58. FA is a polygenic human syndrome
characterized by aberrant DNA repair and with stem cell defects, leading to BM failure56,59,60. In
murine models of FA, BM failure can only be induced by physiological activation of HSCs out of
a quiescent state (e.g. through the induction of type 1 interferon)48. Notably, in cDKO mice BM
failure occurred rapidly and with full penetrance in the absence of extrinsic genotoxic stressors.
Ssb2 has been proposed as a HSC marker61,62; its expression is higher in more immature HSCs
and downregulated with lineage maturation63. This trend is distinct from other repair genes
that show increased expression during lineage commitment and differentiation63. However, the
function of Ssb2 in BM can be well compensated by Ssb1 when Ssb2 is abolished since Ssb2-null
mice show no defect23.
The human SSB proteins are components of the Integrator Complex, which has recently been
shown to play a broader role in transcription including in promoter proximal pause release and
elongation, and 3’-end processing and termination13-17. Most of the target sites of the
components of integrator complex including SSB1 and SSB2 maintain a constitutively open
chromatin state that is favorable to R-loop formation18. Defects in transcription initiation and
termination can lead to accumulation of R-loops and consequent genomic instability as the
exposed non-templated ssDNA becomes more susceptible to DNA damage, which may restrict
transcription and slow down or block replication forks. Blocked replication forks could further
lead to fork stalling, collapse, and DSBs64,65.
In situ
mapping of DSBs revealed that cDKO-
induced specific DSBs are enriched in Ssb1 and Ssb2 binding sites15, CpG islands and near TSSs
of highly expressed genes; all of these regions are favorable to R-loop formation. Consistently,
rapid R-loop accumulation was observed after induction of cDKO and progressively increased
over time concomitant with γ-H2AX staining. This evidence supports potential roles of Ssb1 and
Ssb2 in preventing accumulation of R-loop and associated DNA damage in concert with the
Integrator Complex during transcription. However it remains to be determined whether Rloops play a causative role in genomic instability and cell death observed in cDKO cells, as our
attempts to rescue the phenotype by RNAseH overexpression led to non-specific toxicity in WT
and cDKO HSPCs.
17
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In conclusion, this study has elucidated the novel roles of Ssb1 and Ssb2 which function as
guardians of genome stability by resolving endogenous replication stress in HSPCs and later
components of the blood hierarchy. The cDKO model exhibits rapid cell death of stem and
progenitors and functional loss of BM and intestine, which resembles the changes seen in acute
radiation toxicity. This warrants further studies into these two proteins, for their roles in
mediating chemotherapy and radiation resistance by protecting the genome against the DNA
damaging effects of cytotoxic cancer treatments. Alternatively, inducing DNA damage (for
example, through interferon therapy) together with inhibition of SSB1/2 may synergize as an
anti-neoplastic therapy. Importantly, accumulating evidence suggests that networks that
coordinate normal stem cell self-renewal may lead to tumorigenesis upon overactivation, or to
premature aging once their functionality declines66. As genomic instability is a major hallmark
of actively self-renewing and proliferating cancer cells67, further study on the function of SSB1
and SSB2 in cancer formation and progression will be of great interest to the field.
Acknowledgements
We thank Michael Mcguckin (University of Queensland), Robert Ramsay and Jordane Malaterre
(Peter MacCallum Cancer Centre) for thoughtful discussion on the intestinal phenotype. We
thank David Curtis and Ross Dickins (Monash, Melbourne) for provision of plasmid constructs.
We thank Axia Song, Emma Dishington, Stephen Miles (QIMR Berghofer Medical Research
Institute) and Jian Gong (Central South University) for technical assistance. This work is
supported by a National Health and Medical Research Council (NHMRC) grant NHMRC1085367
to KKK and SWL. KKK was supported by NHMRC Senior Principal Research Fellowship, SWL is a
NHMRC Career Development Fellow and TV is a Leukaemia Foundation PhD Scholar. TKP is
supported by NIH grants CA129537, CA154320, and GM109768. MR, JN and ND are supported
by NIH grant GM112131. KG, MS, AB and MG are supported by NCN grants
2011/02/A/NZ2/00014, 2014/15/B/NZ1/03357 and 2015/17/D/NZ2/03711 and by FNP grant
TEAM.
18
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Author Contributions and Conflict-of-interest Statements
WS, TV, SWL and KKK designed the study, analyzed data and wrote the manuscript. WS, TV, DB,
AB, RKP, GMB, JJ, JLH, ALB, MG, MS and PN performed the experiments. JN, JS, FA, AM, ND, NC,
NC, OJB, JF, JRS, CRW, TKP, MR and KG analyzed data. All authors read and edited the
manuscript. All authors claimed no potential conflicts of interest.
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Figure Legends:
Figure 1. Early embryonic lethality in constitutive
acute mortality in conditional
Ssb1/Ssb2
Ssb1/Ssb2
double knockout (DKO) mice and
double knockout (cDKO) mice due to BM failure and
small intestine atrophy.
(A) Embryos at E7.5 (upper panel) and E10.5 (lower panel) from
Ssb1+/-; Ssb2-/-
intercrossed,
timed matings. (B) Histologic analyses of resorbed embryos from E10.5. Apoptotic bodies are
indicated by black arrows. (C) Body weight change on day 7 of TAM-induced adult Ctrl and
cDKO mice. (D) Representative images of spleen and thymus recovered from Ctrl and cDKO
mice on day 7 post induction with TAM (1mg/day by intraperitoneal injection for 5 consecutive
days). (E) H&E staining of small intestine sections. (F) H&E staining of BM sections. (G) BM cell
count. (H-I) Leukocyte and hematocrit counts in peripheral blood from Ctrl and cDKO mice on
day 7 post induction with TAM (1mg/day by intraperitoneal injection for 5 consecutive days).
Statistical analysis represents t-test (* p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001). Each
point represents an individual mouse/biological replicate. See also Figure S1.
Figure 2. cDKO causes loss of hematopoietic stem and progenitor cells (HSPCs) through
apoptotic cell death and loss of repopulating potential in hematopoietic stem cells (HSCs).
(A-C) Cell numbers of BM progenitors (A), myeloid progenitors (B), and LTHSCs (C) from 2 hind
limbs. (D-E) Frequency of apoptotic cells in LKS+ (D) and CD150+ cells (E). (F) Experimental
scheme. Whole BM cells from either non-TAM treated WT or cDKO animals (expressing the
CD45.2 allele) were mixed with an equal number of congenic whole BM (CD45.1), and injected
into lethally irradiated recipients (CD45.1/CD45.2). (G) Peripheral blood was monitored to
measure bone marrow chimera establishment in recipient mice on week 4. cDKO was induced
in 4 weeks post transplantation. TAM or vehicle control was administrated, and the percentage
of donor chimerism was measured at indicated time points. Statistical analysis represents t-test
or two-way ANOVA, * p<0.05, ** p<0.01, *** p<0.001, **** p<0.0001. Each point represents an
individual mouse/ biological replicate. See also Figure S2.
Figure 3. Replication stress and genomic instability in cDKO HSPCs and BM.
23
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(A-C) DNA fibre assay on HSPCs. (A) LKS+ cells were isolated on day 2 post 4 mg of TAM,
cultured for 16 hours and sequentially pulsed with two different thymidine analogues.
Replication fork movement was measured by incorporation of CldU (green) and IdU (red). (B)
Distribution of red fibre length of ongoing forks. At least 300 structures were measured per
sample per experiment and quantification on 4 independent experiments was presented. (C)
The frequency of terminated fibres that did not incorporate the second label in at least 300
structures per sample per experiment from 4 independent experiments. (D) Representative
images of immunofluorescent staining of RPA (red) and phosphorylated RPA (S4/8) (green). (E)
Quantification of relative Mean Fluorescence Intensity (MFI) from 4 independent experiments.
Each data point represents the relative MFI in cDKO HSPCs normalized to that in Ctrl HSPCs. n
represents total number of cells. (F) Alkaline comet assays on LKS+ cells and (G) olive tail
moment from indicated groups before or at 1h post 2 Gy of ionizing radiation. Each point
represents an individual cell from pooled biological replicates from 4 independent experiments.
(H-L) Bone marrow metaphase spreads on day 3 post 4 mg of TAM. Representative images for
Giemsa (H) and telomere FISH staining (K), chromosome aberration (I and J) and telomere signal
loss (L) analysis are shown. Radial chromosomes, chromosomal breakages and telomere signal
loss are indicated by arrows. (M) EmbryoMax® nucleoside supplement or vehicle (dH2O) was
added to Hoxb8-immortalized cells with indicated treatment. Annexin V was analyzed for
apoptotic cell death 5 days after
in vitro
4OHT induction of cDKO. Each point represents a
biological replicate. Statistical analysis represents t-test or one-way ANOVA for two or multiple
groups respectively, *** p<0.001, **** p<0.0001. See also Figure S3.
Figure 4. cDKO mediates p53 pathway and IFN system activation in HSPCs.
(A) Heat map of commonly up- and down-regulated transcripts in cDKO MEP and GMP from
microarray analysis. (B) Networks enriched in cDKO MEP and GMP by Ingenuity Pathway
Analysis of protein-protein interaction databases only; network name: Cell Cycle, Antimicrobial
Response, and Inflammatory Response. (C) Top 4 overlapping canonical signaling pathways and
-values for enrichment in cDKO MEP and GMP. (D-G) Gene Set Enrichment Analysis (GSEA) of
p
RNA-seq analysis of LKE+CD48- cells showing loss of stemness signature (D), p53 pathway
activation (E) and interferon activation (F and G) in cDKO HSCs. See also Figure S4.
24
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Figure 5. Cytosolic ssDNA primes cDKO HSPC exit from quiescence, and p53 activation leads to
apoptotic cell death in cDKO BM.
(A) Immunofluorescent staining shows ssDNA (green) presence in the cytoplasm of LKS+ cells
(left panel) and immunohistochemistry staining of cytosolic ssDNA (brown) on BM sections of
cDKO mice on day 3 post 4 mg of TAM. (B) Cell cycle analysis of BM LKE+ cells in a time course
of 4 days post cDKO. (C) Histological analysis of H&E and IHC staining of Ki67, p53, and ApopTag.
(D) Western blot showing Ssb1, Ssb2 and p53 levels in cDKO BM samples on D0 and D4 post 4
mg of TAM. (E) LKS+ cells (CD45.2) were isolated from donors, transduced with shLuc-GFP or
shP53-GFP, mixed in a 1:1 ratio with competitor BM (CD45.1), and transplanted into irradiated
recipient mice. cDKO was induced in 4 weeks post-transplantation. Peripheral blood was
monitored to measure bone marrow chimera percentage in recipient mice every 4 weeks post
TAM (cDKO) or vehicle control administration until week 16. (F) Hoxb8-immortalized LKS+ cells
were transduced with control (shLuc-GFP) or p53 (shP53-GFP) shRNAs.
measured 5 days after
in vitro
Apoptosis was
4OHT induction of cDKO. Statistical analysis represents two-way
ANOVA, * p<0.05, ** p<0.01. Each point represents an individual mouse/ biological replicate.
See also Figure S5.
Figure 6. Enrichment of CpG islands and tRNAs in cDKO-induced breaks and all DSBs in cDKO BM.
(A) Experimental scheme of BLESS analysis to map genome-wide DSBs. From pooled BM
samples of Ctrl and cDKO mice on day 4 post 4 mg of TAM, intact nuclei were purified, and DSBs
were ligated to a biotinylated linker (proximal). Genomic DNA (gDNA) was extracted and
fragmented, and labeled fragments were captured by streptavidin and ligated to a secondary
linker (distal), PCR amplified, and sequenced. (B) Venn diagram showing regions with cDKOinduced DSBs (BLESS cDKO vs. BLESS Ctrl) and spontaneous DSBs (BLESS Ctrl vs. gDNA Ctrl) with
a p-value threshold p<0.001 and 1250nt resolution. (C) Enrichment of CpG islands in cDKOinduced breaks. (D) Enrichment of tRNAs in all DSBs in DKO BM. (E) Enrichment of Ssb1 and
Ssb2 binding sites from HIT-Seq data (Skaar et al., Cell Res 2015) in intervals enriched with DSBs
from BLESS data. (F) Enrichment of CpG islands in intervals enriched with Ssb1 and Ssb2 binding
sites using the same method as that in BLESS data analysis (described in Supplemental
25
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Materials and Methods, # borderline significant, 0.05<p<0.1, * p<0.05, ** p<0.01). Significance
of enrichment is calculated by permutation test, ** p<0.01. See also Figure S6.
Figure 7. R-loop and DNA damage accumulation lead to BM failure in cDKO mice.
(A) Representative image of immunostaining of R-loop (green), γH2AX (red), and nuclei (blue,
DAPI staining) on BM sections on day 4 after 4 mg of TAM. (B-C) Ratio of BM cells with positive
R-loop (B) and γH2AX (C) during the 4-day time course post cDKO. For each experiment 4
sections from 4 individual mice were analysed at 5 time points. Statistical analysis represents ttest, *** p<0.001. (D) Proposed model of genomic instability in cDKO cells. cDKO induces R-loop,
replication stress and DSB accumulation, cytosolic ssDNA with consequent activation of IFN,
p53 and DDR pathways, and apoptotic cell death, hence disruption of stem cell homeostasis.
See also Figure S7.
26
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Prepublished online March 7, 2017;
doi:10.1182/blood-2016-06-725093
Ssb1 and Ssb2 cooperate to regulate mouse hematopoietic stem and
progenitor cells by resolving replicative stress
Wei Shi, Therese Vu, Didier Boucher, Anna Biernacka, Jules Nde, Raj K. Pandita, Jasmin Straube, Glen
M. Boyle, Fares Al-Ejeh, Purba Nag, Jessie Jeffery, Janelle L. Harris, Amanda L. Bain, Marta Grzelak,
Magdalena Skrzypczak, Abhishek Mitra, Norbert Dojer, Nicola Crosetto, Nicole Cloonan, Olivier J.
Becherel, John Finnie, Jeffrey R. Skaar, Carl R. Walkley, Tej K. Pandita, Maga Rowicka, Krzysztof
Ginalski, Steven W. Lane and Kum Kum Khanna
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