2735
Research Article
Bacterial genotoxin triggers FEN1-dependent RhoA
activation, cytoskeleton remodeling and cell survival
Lina Guerra1,*, Riccardo Guidi1,*, Ilse Slot1, Simone Callegari1, Ramakrishna Sompallae1, Carol L. Pickett2,
Stefan Åström3, Frederik Eisele4, Dieter Wolf4, Camilla Sjögren1, Maria G. Masucci1 and Teresa Frisan1,‡
1
Department of Cell and Molecular Biology, Karolinska Institutet, Box 285, S-171 77 Stockholm, Sweden
Department of Microbiology and Immunology, Chandler Medical Center, University of Kentucky, Lexington, KY 40536-0298, USA
Department of Developmental Biology, Wenner-Grens Institutet, Stockholm University, 106 91 Stockholm, Sweden
4
Institute of Biochemistry, University of Stuttgart, 70569 Stuttgart, Germany
2
3
*These authors contributed equally to this work
‡
Author for correspondence (Teresa.Frisan@ki.se)
Journal of Cell Science
Accepted 15 April 2011
Journal of Cell Science 124, 2735-2742
© 2011. Published by The Company of Biologists Ltd
doi:10.1242/jcs.085845
Summary
The DNA damage response triggered by bacterial cytolethal distending toxins (CDTs) is associated with activation of the actinregulating protein RhoA and phosphorylation of the downstream-regulated mitogen-activated protein kinase (MAPK) p38, which
promotes the survival of intoxicated (i.e. cells exposed to a bacterial toxin) cells. To identify the effectors of this CDT-induced survival
response, we screened a library of 4492 Saccharomyces cerevisiae mutants that carry deletions in nonessential genes for reduced
growth following inducible expression of CdtB. We identified 78 genes whose deletion confers hypersensitivity to toxin. Bioinformatics
analysis revealed that DNA repair and endocytosis were the two most overrepresented signaling pathways. Among the human orthologs
present in our data set, FEN1 and TSG101 regulate DNA repair and endocytosis, respectively, and also share common interacting
partners with RhoA. We further demonstrate that FEN1, but not TSG101, regulates cell survival, MAPK p38 phosphorylation, RhoA
activation and actin cytoskeleton reorganization in response to DNA damage. Our data reveal a previously unrecognized crosstalk between
DNA damage and cytoskeleton dynamics in the regulation of cell survival, and might provide new insights on the role of chronic bacteria
infection in carcinogenesis.
Key words: Cytolethal distending toxin, DNA damage, FEN1, Cell survival, TSG101, RhoA, Actin cytoskeleton
Introduction
Cytolethal distending toxins (CDTs) are produced by a variety of
Gram-negative bacteria, such as Escherichia coli, Aggregatibacter
actinomycetemcomitans, Haemophilus ducreyi, Shigella
dysenteriae, Campylobacter sp. and Helicobacter sp., and
Salmonella enterica (reviewed in Smith and Bayles, 2006). CDT
activity requires the function of three genes (cdtA, cdtB and cdtC)
and the active holotoxin is a tripartite complex (Lara-Tejero and
Galan, 2001; Scott and Kaper, 1994), formed by the three subunits
CdtA, CdtB and CdtC. The CdtB component is the active subunit,
that shares structural and functional homology with the mammalian
deoxyribonuclease I (DNase I) (Elwell et al., 2001; Lara-Tejero
and Galan, 2000; Nesic et al., 2004). The CdtA and CdtC subunits
are ricin-like lectin domains (Nesic et al., 2004), important for the
cellular internalization of the toxin (Lee et al., 2003; McSweeney
and Dreyfus, 2004; McSweeney and Dreyfus, 2005).
The capacity of CDT to induce DNA double-strand breaks
(DSBs) was confirmed in intoxicated (i.e. cells exposed to a
bacterial toxin) mammalian cells by pulsed-field gel electrophoresis
(PFGE) (Frisan et al., 2003). Similar results were obtained by
Hassane and co-workers who observed DSBs in Saccharomyces
cerevisiae cells transfected with the active CdtB subunit (Hassane
et al., 2001). In both cases, mutations within the DNase-conserved
motives prevented induction of DNA damage.
The cellular effects of CDTs are similar to those induced by
exposure to ionizing radiation (g-irradiation), a well-characterized
DNA-damaging agent. Both CDT and ionizing radiation stimulate
the phosphorylation of histone H2AX and the relocalization of the
DNA repair protein complex Mre11–Rad50–Nbs1 (MRN) (Hassane
et al., 2003; Li et al., 2002). Both agents also activate cell cycle
checkpoints in a cell-type-dependent manner (human primary
fibroblasts are arrested in G1 and G2 phases, whereas HeLa cells are
arrested in G2) (Cortes-Bratti et al., 2001; Hassane et al., 2003).
In adherent cells, g-irradiation or CDT intoxication are associated
with the formation of actin stress fibers (Cortes-Bratti et al., 1999;
Gelfanova et al., 1999). This effect is regulated by the activation
of the small GTPase RhoA, and promotes cell survival in
intoxicated cells (Frisan et al., 2003). Activation of RhoA and actin
stress fiber formation in response to CDT is dependent on the
RhoA-specific guanine nucleotide exchange factor (GEF) Net1
(Guerra et al., 2008a).
The DNA-damage-dependent Net1–RhoA signaling diverges
into two different effector cascades: one dependent on the RhoA
kinases ROCKI and ROCKII that controls the formation of actin
stress fibers, and one regulated by the mitogen-activated protein
kinase (MAPK) p38 and its downstream target MAPK-activated
protein kinase 2 (MAPKAPK2) that promotes cell survival (Guerra
et al., 2008a).
The survival of cells with damaged DNA may promote genomic
instability and favor tumor initiation and/or progression (reviewed
in Kastan and Bartek, 2004; Shiloh, 2003). Characterization of
the survival signals in response to CDTs is, therefore, relevant to
understand how bacterial infections contribute to carcinogenesis.
To identify new CDT-induced survival signals, we have screened
a yeast deletion library in cells that express the active CdtB
subunit under the control of a galactose-inducible promoter. This
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Journal of Cell Science 124 (16)
screen identified 78 deletion mutants with a reduced growth rate
following the inducible expression of CdtB. Bioinformatics
analysis revealed that 12 human orthologs of these genes interacts
with the RhoA signaling pathway. Functional studies in
mammalian cells showed that flap structure-specific endonuclease
1) FEN1 (yeast ortholog RAD27) promotes RhoA activation,
MAPK p38 phosphorylation and, ultimately, cell survival in
response to CDTs.
Results
Journal of Cell Science
Screening of the S. cerevisiae deletion library
CDT-induced DNA damage is accompanied by activation of the
small GTPase RhoA, actin cytoskeleton rearrangements and
phosphorylation of the downstream-regulated MAPK p38, which
leads to delayed cell death (Guerra et al., 2008a). To identify the
effectors of this survival response, we expressed the active subunit
CdtB of the Campylobacter jejuni CDT under the control of a
galactose-inducible promoter in the S. cerevisiae strain BY4743.
Induction of CdtB expression by addition of galactose to the culture
medium (CdtB ON) induced G2 arrest (supplementary material
Fig. S1A) and inhibited cell growth (supplementary material Fig.
S1B) in cells transformed with the pDCH-CdtB plasmid, whereas
no effect was observed in cell grown in presence of glucose (CdtB
OFF) or in cells transformed with the vector control.
Having confirmed the capacity of CdtB to induce cell cycle
arrest, which was demonstrated to be dependent on the DNase
activity of the toxin (Hassane et al., 2001), we performed a genomewide screen using the EUROpean Saccharomyces cerevisiae
ARchive for Functional analysis (EUROSCARF) library, derived
from the BY4743 strain (Cherry et al., 1998), where nonessential
genes are individually deleted. Deletion strains that showed growth
aberrations, such as respiration deficiency (petite strains), were
excluded. Of the 4793 deletion mutants 4085 were recovered on
YPGluc plates and were successfully transformed with the CdtBexpressing pDCH–CdtB plasmid using the high-throughput yeast
transformation method (Schafer and Wolf, 2005). To identify
mutants with increased sensitivity to CdtB, serial dilutions of the
transfected yeast were performed on plates containing selective
medium supplemented with either glucose (CdtB OFF) or
galactose/raffinose (CdtB ON) to regulate CdtB expression.
As expected, the growth of wild type (wt) cells was reduced in
galactose/raffinose, compared with those grown in glucose. However,
several deletion mutants showed a more pronounced inhibition of
growth upon induction of CdtB expression (Fig. 1A, left panel). We
defined as CdtB hypersensitive, deletion strains that did not grow
after the third dilution step upon induction of CdtB expression. A
total of 121 hypersensitive transformants were identified in three
independent screens (Fig. 1A, right panel). To ensure that the
reduced growth was due to the expression of CdtB rather than an
intrinsic property of the deletion, each of the 121 mutants was
transformed with a control empty plasmid, and a serial dilution
screen in the presence or absence of galactose/raffinose was
performed (data not shown). This analysis restricted the number of
CdtB-hypersensitive mutants to 78 (Table 1). PCR analysis,
performed for 30 of the 78 mutants, demonstrated that all the clones
tested carried the correct deletion and validated the quality of the
library used in this study (supplementary material Tables S1 and S2).
The identity of the yeast genes whose deletion confers
hypersensitivity to CdtB is summarized in Table 1. On the basis of
the annotated function in the Saccharomyces Genome Database
(http://yeastgenome.org), we classified these genes into twelve
Fig. 1. Screening of the EUROSCARF deletion library. (A) Replica plating
of fivefold serial dilutions of the wild-type (wt) cells or deletion mutants,
transformed with the pDCH-CdtB plasmid, were incubated on plates that
contained selective medium supplemented with 2% glucose (CdtB OFF) or
2% galactose/raffinose (CdtB ON) at 30°C for 48 hours. The left panel shows
a selection of the tranformants. The right panel summarizes the growth pattern
of all 4085 transformants. Each diamond represents the last dilution at which
cell growth was detected. Strains that failed to grow after the third dilution
(1/125) were scored as CdtB hypersensitive. (B) Pathway enrichment analysis
was performed as described in Materials and Methods using as data set the 59
human orthologs of the 78 yeast genes found in the EUROSCARF library.
groups. Five genes could not be grouped because they encode
proteins with unknown function. As expected, the majority of the
genes conferring hypersensitivity to CdtB encode for proteins
involved in the regulation of the DNA damage checkpoint
responses, genome integrity and DNA repair (39%, groups 1-4).
The next largest functional group comprised genes encoding for
effectors involved in the control of vesicular transport and
endocytosis (13%, group 5).
Using the Inparanoid database (O’Brien et al., 2005) and BLAST
sequence alignment we identified 59 human orthologs of the 78
yeast genes found in the EUROSCARF library screening (Table
1). In agreement with the functional classification in the yeast data
set, pathway enrichment analysis performed on the human orthologs
demonstrated that DNA repair and endocytosis were the two most
over-represented signaling pathways, followed by glucose
metabolism and ribosome biogenesis (Fig. 1B and supplementary
material Fig. S2).
Survival signals induced by bacterial genotoxin
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Table 1. Genes identified in the EUROSCARF screen
Journal of Cell Science
Group
Function
Number of genes
Yeast genes
1
Replication recombination repair
13
2
3
Checkpoint
Chromatin silencing telomeres
5
8
4
5
Mitotic chromosome transmission
Vacuolar/Golgi/ endocytosis
5
10
6
Mitochondrial
8
7
6
8
Cytokinesis, spindle,
cytoskeleton rearrangements
Translation/Ribosomal
9
10
12
Transcription/ RNA metabolism
Ubiquitin proteasome system
Others
3
2
6
DCC1; DDC1; DIA2; ELG1; MMS4;
MUS81; RAD27; RAD50; RAD51;
RAD57; SAE2; SHU1; SRS2
CHK1; MEC3; MRC1; RAD9; TOF1
CST6; ESC1; NAM7; NAT3; NMD2;
NPT1; UPF3; YPL205C
CHL1; CHL4; CTF4; MCM21; MCM22
DRS2; PEP7; PEP12; PMR1; SNF7;
VPS3; VPS4; VPS8; VPS21; VPS23;
VPS24
ADK1; ALT1; FMP37; HFA1; MRPL10;
OAR1; PDB1; SOD2
CTF3; NIP100; PTC1; RVS161; SLT2;
SRV2
KAP123; RPL27A; RPL36A; RPS0B;
RPS17A; ZUO1
BUR2; CKA2; NPL3
DOC1; SHP1
LAT1; LIP5; MKC7; RTS1; TPD3; TPS1
13
Unknown
5
6
YBR099C; YDR109C; YEL045C;
YMR252C; YOR376W
78 genes
Identification of candidate proteins regulating the RhoAdependent survival signals
The aim of the genome-wide screen was to characterize new genes
that regulate cell survival through RhoA activation in response to
CDT-induced DNA damage in mammalian cells. To identify
possible candidates, a RhoA protein–protein interaction network
was generated using the Protein Interaction Network Analysis
(PINA) (Wu et al., 2009). We next assessed whether any of the
genes present in our data set encoded for proteins that share
common interacting partners with RhoA. As shown in Fig. 2, the
products of 12 human orthologs directly bind with RhoA-interacting
proteins (Fig. 2). Three of these proteins belong to the two most
over-represented pathways identified in Fig. 1B: FEN1 (DNA
repair), TSG101 and RAB5A (endocytosis).
On the basis of this analysis, we selected two proteins, FEN1
and TSG101, one from each of the enriched signaling pathways,
to assess whether these effectors regulate cell survival, RhoA
activation and actin cytoskeleton remodeling in response to DNA
damage in higher eukaryotic cells.
Human orthologs
DSCC1, DIAPH3, MUS81, FEN1,
RAD50, RAD51, XRCC3, UBA2,
UBE2B
CHEK1, PIP3-E, MRC1L.1, RAD9A
CST6, UPF1, NAT5, UPF2, SLC17A1,
UPF3A
DDX11, CENPN, REPS2, NFIC
ATP8A2, LAP3, STX12, ATP2C1,
CHMP4B, VPS4B, VPS8, RAB5A,
TSG101, VPS24
AK2, GPT, MRPL10, PDHB, SOD2
PTCH2, BIN3, MAPK7, CAP1
IPO4, RPL27A, RPL36AL, LOC387867,
RPS17, DNAJC2
CST4, CSNK2A1, TAF15,
TBX6, NSFL1C
DLAT, LIAS, PPP2R5C, PPP2R1A
TPSAB1/TPSB2
59 genes
the activated form of the pro-apoptotic protein Bax by using the
conformation-dependent antibody 6A7 in immunofluorescence
assays, and through assessment of the number of cells with nuclear
fragmentation – a late marker of death (Fig. 3). CDT-intoxicationinduced Bax activation in ~20% of the untransfected cells (data not
shown) or in cells transfected with the control siRNA (scRNA)
(Fig. 3C, left panel). A similar ratio of cell death was observed
when expression of TSG101 or CSNK2B was downregulated by
RNA interference (Fig. 3C, right panel). By contrast, knockdown
of FEN1 resulted in a significant increase of Bax-positive cells
upon toxin treatment, corresponding to an approximately fourfold
FEN1 prolongs cell survival in response to DNA damage
Expression of the endogenous TSG101 and FEN1 proteins was
downregulated through RNA interference (RNAi) by transfection
of specific small interfering RNAs (siRNAs) in HeLa cells 48
hours prior to toxin treatment or g-irradiation. As negative control
we included a scramble siRNA (scRNA) and siRNA that
specifically targets CSNK2B, which was not identified in the yeast
screen as a relevant gene to promote cell survival in response to
DNA damage. Two independent siRNAs were used for each protein.
Transfection with the two specific siRNAs induced ~80–90%
reduction in the levels of expression of TSG101 and FEN1 (Fig.
3A). Knockdown of TSG101, FEN1 or CSNK2B did not prevent
CDT-induced cell cycle arrest (data not shown).
The first set of experiments aimed to investigate the role of
TSG101 and FEN1 in inhibition of cell death upon treatment with
CDT. Forty-eight hours after siRNA transfection, cells were left
untreated or exposed to CDT. The number of cells undergoing cell
death was assessed 48 hours after treatment through detection of
Fig. 2. RhoA protein–protein interaction network. The RhoA protein–
protein interaction network was generated as described in Materials and
Methods. The figure shows only interacting partners that are shared between
RhoA and the products of the human orthologs present in our data set (green
boxes). Proteins that belong to the two most-enriched pathways according to
the analysis presented in Fig. 1B, are labelled in red.
Journal of Cell Science
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fragmentation in ~5% of intoxicated untransfected cells (data not
shown) and in ~5% cells transfected with the control siRNA
(scRNA) (Fig. 3D,E). The percentage of dying cells was not
significantly increased upon downregulation of the endogenous
levels of TSG101 and CSNK2B by siRNA. By contrast, 15–20%
of intoxicated cells showed chromatin fragmentation upon FEN1
knockdown (Fig. 3E, left panel), corresponding to a three- to
fourfold increase compared with intoxicated untransfected cells
(Fig. 3E, right panel).
The late occurrence of CDT-induced cell death observed upon
FEN1 knockdown was similar to that observed in cells that express
the dominant-negative form of RhoA (RhoAN19) and in cells
where the expression of the RhoA-specific GEF Net1 was knocked
down by siRNA (Frisan et al., 2003; Guerra et al., 2008a).
Induction of DNA DSBs stimulates the activation of the MAPK
p38 in a RhoA-dependent manner; and this is required to delay cell
death in response to toxin treatment or g-irradiation (Guerra et al.,
2008a). Since FEN1 appears to be required to promote cell survival,
we tested whether it is also important for the activation of MAPK
p38. A four- to fivefold increase in MAPK p38 activity was observed
4 hours after g-irradiation in control HeLa cells or cells transfected
with non-silencing siRNA, as assessed by western blot analysis
using an antibody specifically recognizing phosphorylated MAPK
p38 (Fig. 4). Similar levels of MAPK p38 phosphorylation were
detected in irradiated HeLa cells, where expression of the endogenous
levels of TSG101 or CSNK2B where knocked down with specific
siRNA. Importantly, downregulation of FEN1 expression
significantly decreased activation of MAPK p38 (Fig. 4).
RhoA activation and actin remodeling
Fig. 3. FEN1 knockdown increases the rate of cell death upon induction of
DNA damage. (A) HeLa cells were transfected with non-silencing siRNA
(scRNA) or two specific siRNAs (indicated as _1 and _2) targeting TSG101,
FEN1 or CSNK2B. The individual siRNAs have been labeled as TSG101_1
and TSG101_2; FEN1_1 and FEN1-2; CSNK2B_1 and CSNK2B_2.
Expression levels of the endogenous proteins were analyzed by western blotting
48 hours after transfection. Actin expression was assayed as loading control. (B)
Cells transfected with non-silencing siRNA (scRNA), TSG101_1, FEN1_1, or
CSNK2B_1 specific siRNA were left untreated (images not shown), or were
exposed to CDT (2 g/ml) for 48 hours. Apoptosis was assessed by
immunofluorescence, using the conformation-dependent anti-Bax 6A7 antibody
(magnification 63⫻). (C) Quantification of Bax-positive cells as percentage of
Bax-positive cells (left panel) and as fold increase of Bax-positive cells (right
panel). Fold increase is the ratio between the number of Bax-positive cells in
intoxicated cells transfected with each specific siRNA and the number of Bax
positive cells in intoxicated untransfected cells. Mean ± s.d. of three independent
experiments. (D) HeLa cells transfected with non-silencing siRNA (scRNA),
TSG101_1, FEN1_1, or CSNK2B_1 specific siRNA were left untreated (picture
not shown) or exposed to CDT (2 g/ml) for 48 hours. Nuclei were
counterstained with Hoechst 33258 dye. (E) Quantification of cell death is
shown as percentage of cells with fragmented nuclei (left panel) and as fold
increase of cells with fragmented nuclei (right panel). Fold increase is the ratio
between the number of cells with fragmented nuclei in intoxicated cells
transfected with a specific siRNA versus the number of cells with fragmented
nuclei in intoxicated untransfected cells. Mean ± s.d. of three independent
experiments. For each experiment 100 cells were counted.
increase compared with the intoxicated untransfected cells (Fig.
3B,C). Similar data were obtained by monitoring the number of
cells with fragmented nuclei. Intoxication-induced chromatin
We then assessed whether TSG101 and FEN1 are involved in the
activation of RhoA and reorganization of the actin cytoskeleton in
intoxicated or irradiated cells. As expected, intoxication or girradiation induced an approximately threefold increase of RhoA
activation in control cells or cells transfected with non-silencing
siRNA 4h after treatment (Fig. 5A). Similar levels of GTP-bound
RhoA were observed upon downregulation of the endogenous
levels of TSG101, whereas knockdown of FEN1 completely
prevented activation of this protein (Fig. 5A).
We also investigated whether FEN1-dependent inhibition of
RhoA correlated with the reduction of stress fiber formation upon
induction of DNA damage. Intoxication or g-irradiation induced
actin stress fibers in 50% of control cells or cells transfected with
non-silencing siRNA 48 hours after treatment (Fig. 5B,C).
Knockdown of the endogenous levels of TSG101 using siRNA did
not significantly change stress fiber formation in treated cells. By
contrast, FEN1 knockdown strongly reduced the number of cells
carrying stress fibers upon exposure to CDT or g-irradiation (Fig.
5B,C).
As a complementary evaluation of the level of actin
polymerization upon induction of DNA damage, we have assessed
the mean fluorescence intensity of phalloidin staining per cell
area, measured using the ImageJ software (Fig. 5C, right panel).
Intoxication or g-irradiation induced a small increase of the mean
fluorescence intensity in control cells, or in cells transfected with
non-silencing siRNA or with the TSG101-specific siRNA
compared with control untreated cells. By contrast, we observed
a 50% reduction of the mean fluorescence intensity upon induction
of DNA damage in cells whose levels of endogenous FEN1 were
knocked down by siRNA. This effect was similar to that observed
in cells expressing the dominant-negative form of RhoA
Journal of Cell Science
Survival signals induced by bacterial genotoxin
2739
Fig. 4. FEN1-knockdown prevents phosphorylation of MAPK p38.
Untransfected HeLa cells or cells transfected with non-silencing siRNA
(scRNA), TSG101_1, FEN_1 or CSNK2B_1 specific siRNA were left
untreated (–DSB), or exposed to ionizing radiation (20Gy, +DSB), and further
incubated for 4 hours in complete medium. Samples were analyzed by western
blot, using specific antibodies against phosphorylated MAPK p38 (p-p38) and
MAPK p38 (p38) antibody. The bar graph represents the quantification of
MAPK p38 activation (mean ± s.d. of three independent experiments). Data
are the ratio between the optical density of the p-p38-specific band in
irradiated cells and the optical density of the p-p38-specific band in untreated
cells.
(RhoAN19) and in cells where the expression of the RhoAspecific GEF Net1 was knocked down by siRNA (Frisan et al.,
2003; Guerra et al., 2008a).
In conclusion, our study demonstrates that FEN1 is a new
effector in the RhoA-dependent signaling pathway that is activated
in response to genotoxic stress (Fig. 6).
Discussion
The exact mechanism by which bacteria contribute to
carcinogenesis is still poorly characterized. Several Gram-negative
pathogenic bacteria have been shown to produce CDTs that induce
DNA damage in infected cells. Since incorrect repair of DNA
damage may lead to genomic instability and tumour development,
it is conceivable that chronic infection with CDT-producing bacteria
can be a risk factor for cancer development. One key event in the
tumorigenic process is activation of survival signals, which can
prevent cell death induced by DNA damage and/or oncogene
activation (reviewed in Halazonetis et al., 2008). We used S.
cerevisiae transformed with a CdtB-expressing plasmid as a model
for intoxicated cells and screened a yeast deletion library to identify
novel genes required for the CdtB-induced survival response (Fig.
1A and Table 1).
A genome-wide screen for the cellular responses to CdtB in S.
cerevisiae has been previously described by Kitagawa et al.
(Kitagawa et al., 2007). The focus of their study was to characterize
the DNA damage repair mechanisms induced by CDTs, whereas
we were interested to identify RhoA-dependent effectors that
regulate cell survival in intoxicated cells. The significant overlap
between the genes identified in this study and those reported by
Kitagawa and co-workers or Bennett et al. (Bennett et al., 2001),
who identified effectors that confer resistance to ionizing radiation,
validates our screen and support the reliability of the data presented.
An expected result of the screen was the demonstration that
proteins involved in DNA repair were important for cell survival
Fig. 5. RhoA activation and actin stress fiber formation upon knockdown of
FEN1 and TSG101. (A) Untransfected HeLa cells or cells transfected with nonsilencing siRNA (scRNA), TSG101_1, or FEN1_1 specific siRNA were left
untreated (–DSB) or exposed to CDT (2 g/ml; +DSB) and were then further
incubated for 4 hours. Activation of RhoA was assessed by using the RhoAspecific G-LISATM kit (mean ± s.d. of three independent experiments). Pos,
internal positive control of the RhoA-specific G-LISATM assay.
(B) Untransfected HeLa cells or cells transfected with non-silencing siRNA
(scRNA), TSG101_1 or FEN1_1 specific siRNA were left untreated (–DSB) or
exposed to CDT (2 g/ml) for 48 hours (+DSB). The actin cytoskeleton was
visualized by TRITC-phalloidin staining (magnification 63⫻).
(C) Quantification of cells carrying stress fibers. The left panel shows the
percentage of positive cells. Cells exhibiting more than five stress fibers were
scored as positive. The right panel shows the ratio between the mean
fluorescence intensity of the phalloidin staining and the cell area, quantified
using the ImageJ software, as described in Materials and Methods. Mean ±
s.e.m. of three independent experiments; 100 cells were counted for each
experiment.
in cells expressing CdtB. More surprising was the enrichment of
genes encoding for proteins that regulate endocytosis, specifically
those controlling sorting of ubiquitylated cargos at multivesicular
bodies (Fig. 1B and Table 1).
The role of FEN1 in the activation of RhoA, phosphorylation of
MAPK p38 and formation of actin stress fibers in response to DNA
DSBs has not been reported previously. FEN1 is a multifunction
nuclease that possesses endonuclease activity required for the
maturation of the Okazaki fragments during DNA replication and
long-patch DNA-base-excision repair (Liu et al., 2004). FEN1 also
has a 5⬘-exonuclease activity (EXO) required for correct homologous
DNA recombination and a gap-dependent endonuclease (GEN)
activity, which cleaves the single-stranded DNA region of gapped
DNA duplex or DNA forks that generate DNA DSBs (Parrish et al.,
2003; Zheng et al., 2005). It has been shown that the EXO and GEN
activities can be abrogated by a Glu-to-Asp mutation in position 60
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Journal of Cell Science 124 (16)
Journal of Cell Science
Fig. 6. Summary of the RhoA-dependent survival pathway. DNA damage
leads to FEN1-dependent activation of RhoA, actin cytoskeleton remodeling,
phosphorylation of MAPK p38 and prolonged cell survival.
(E60D), leading to frequent spontaneous mutations both in yeast and
mammalian cells, and promotion of cancer progression in a knockin mouse model (Zheng et al., 2007). These effects of FEN1 E60D
are well in line with its role in maintenance of genomic stability.
Interestingly, mouse embryo fibroblasts that expressing FEN1 E60D
also show an increased susceptibility to cell death in response to UV
irradiation compared with the cells that express wild-type FEN1,
indicating that the EXO and/or GEN activities are required for cell
survival in response to certain genotoxic stress. Our data demonstrate
that the role of FEN1 in cell survival is also relevant in the response
to DNA damage caused by the bacterial genotoxin CDT – and this
is associated with activation of RhoA, formation of actin stress
fibers and phosphorylation of MAPK p38 (Figs 3, 4 and 5). However,
how FEN1 may induce activation of the small GTPase RhoA is
presently not known.
We can envisage several possibilities. FEN1 is a protein that has
been shown to regulate multiple pathways, such as genome integrity
and fragmentation of apoptotic DNA (reviewed in Zheng et al.,
2011). Its multitasking role is dependent on the sets of proteins that
directly interact with FEN1, and can be further determined by its
subcellular localization: nuclear and cytosolic (Qiu et al., 2001).
To date 35 FEN1-interacting partners have been identified, which
can be grouped into five functional categories: partners that regulate
(i) DNA replication, (ii) DNA repair, (iii) DNA fragmentation
during apoptosis, (iv) telomere stability and (v) post-translational
modification of FEN1 (phosphorylation, acetylation and
methylation) (reviewed in Zheng et al., 2011). In addition, a high
throughput yeast two-hybrid screen has indicated that FEN1 directly
interacts with the Rho GDP dissociation inhibitor (GDI) a
(ARHGDIA) (Stelzl et al., 2005), suggesting that FEN1 performs
additional activities not related to its endonuclease function.
ARHGDIA maintains the small GTPase RhoA in an inactive form
(DerMardirossian and Bokoch, 2005). It is conceivable that the
fraction of FEN1 that is present in the cytosol sequesters
ARHGDIA, allowing activation of RhoA upon of DNA damage.
An independent DNA repair function has been previously
demonstrated for other components of the DNA repair machinery.
The DNA-dependent protein kinase DNA-PK, which forms a
complex with the Ku heterodimer (Ku70–Ku80), is an essential
component of the non-homologous end-joining recombination
(NHEJ) (reviewed in Weterings and Chen, 2007). However, a
fraction of DNA-PK is localized in the lipid raft microdomains of
the plasma membrane (Lucero et al., 2003), and can phosphorylate
Akt at Ser473 (Feng et al., 2004). Furthermore, the Ku70–Ku80
heterodimer has been shown to regulate cell adhesion to the
extracellular matrix and to protect cells from apoptosis through
suppression of the translocation of the pro-apoptotic protein Bax
at the mitochondrial membrane (reviewed in Muller et al., 2005).
Another example of a membrane-associated protein that regulates
NHEJ recombination is the cell polarity protein Par-3. This has long
been considered to be a component of a complex that functions in
the assembly of tight junctions, cell polarity and regulation of actin
dynamics through interaction with the small GTPase Rac, which
belongs to the Rho subfamily (Chen and Macara, 2005; Hurd and
Margolis, 2005; Nishimura et al., 2005). Recently, Fang and
colleagues have shown that Par-3 is also present in the nucleus and
directly binds to the Ku heterodimer. This interaction is enhanced
when cells are exposed to genotoxic stress (g-irradiation or the anticancer agent etoposide phosphate), and knockdown of the protein by
RNAi delays the DNA repair response and significantly decreases
cell survival in response to ionizing radiation (Fang et al., 2007;
Lees-Miller, 2007). These data demonstrate that there is an active
crosstalk between the DNA repair machinery and proteins that are
traditionally considered to be cytosolic and membrane related, such
as Par-3 and small GTPases of the Rho family.
It is also possible that the FEN1-dependent activation of RhoA
is a secondary effect of the DNA repair process. We have previously
shown that DNA damage induced by ionizing radiation or CDT
promotes dephosphorylation of the RhoA-specific GEF Net1 at the
inhibitory site Ser152 (Alberts et al., 2005; Guerra et al., 2008b).
Therefore, we cannot exclude that the FEN1-dependent DNA repair
mechanisms, evoked by CDT or g-irradiation, indirectly trigger
Net1 dephosphorylation through the activation of a specific
phosphatase or the inhibition of a kinase.
On the basis of the pathway enrichment analysis shown in Fig.
1B and the reported interaction with the RhoA effector ROCKI
(Morita et al., 2007), we also tested the role of the ESCRT protein
TSG101 in the regulation of RhoA activation and cell survival in
HeLa cells exposed to CDT. Our results indicate that knockdown
of TSG101 does not influence the rate of cell death and RhoA
activation (Figs 3, 4 and 5). Our data suggest that the role of
ESCRT proteins in cell survival upon induction of DNA damage
is organism dependent. It is also possible that TSG101 regulates
signaling that controls the extent of the cell cycle arrest upon
induction of DNA damage rather than cell survival in S. cerevisae
and, therefore, its deletion is associated with a prolonged arrest in
G2 phase, which results in the marked delay in cell growth observed
in the library screening (Fig. 1 and Table 1). Our data contribute
to the understanding of poorly characterized aspects of cellular
responses to genotoxic stress, such as the crosstalk between DNA
repair and the cytosolic small GTPase RhoA (Fig. 6).
The identification of survival signals that are triggered by chronic
infections of CDT-producing bacteria is crucial to understand
whether these infections promote genomic instability and favor
malignant transformation. The only bacterium classified as a human
carcinogen is Helicobacter pylori (Crowe, 2005). However, a
possible involvement in oncogenesis has been suggested for other
bacteria, such as the Gram-negative bacterium Salmonella typhi
(Lax, 2005). Establishing an association between infection with
Survival signals induced by bacterial genotoxin
CDT-producing bacteria and cancer promotion and/or progression
might help the development of specific therapeutic protocols aimed
at early and rapid eradication of the bacterial infection, thereby
preventing an initial step of tumor development.
Materials and Methods
Cells
HeLa cells were obtained from ATCC and cultivated in RPMI 1640 medium
supplemented with 10% fetal calf serum (FCS), 5 mM L-glutamine, penicillin (100
units/ml) and streptomycin (100 g/ml) (complete medium) at 37°C in a humid
atmosphere containing 5% CO2.
Yeast strains and plasmids
Journal of Cell Science
The complete EUROpean Saccharomyces cerevisiae ARchive for Functional analysis
(EUROSCARF) library, purchased from Open Biosystems, Thermo Scientific,
consists of mutants where nonessential genes are individually deleted (Cherry et al.,
1998). The pDCH-CdtB and the control plasmids have been previously described
(Hassane et al., 2001).
The library and the control BY4743 strain (MATa/MATa, his31/his31,
leu2/leu2met150/MET15, LYS2/lys20, ura30/ura30) were transformed with
the pDCH-CdtB (Hassane et al., 2001) or a vector control plasmid. Dropout synthetic
medium was prepared according to the manufacturer’s instructions (Clontech).
Transformants were selected and maintained in synthetic dropout medium without
leucine (selective medium) supplemented with 2% glucose. For induction of CdtB
expression, cells were grown in selective medium supplemented with 2% galactose
and 2% raffinose.
Yeast transformation
Systematic yeast transformation experiments were performed, with minor
modifications, using the S. cerevisiae direct transformation in a 96-well format, as
previously described by (Schafer and Wolf, 2005). Yeast deletion strains were grown
on yeast extract/peptone/glucose (YPGluc) square plates (12cm⫻12cm, Greiner
Bio-one) for 48 hours at 30°C. Twenty-five microliters of the transformation solution
(27% PEG600, 200 mM Lithium acetate, 50 mM DTT, carrier DNA 5 g/l, and 15
g of the pDCH-CdtB or control plasmids) were added in 96-well plates and the
yeast deletion strains were transferred using a 48-pin replicator, mixed well and
incubated for 2 hours at 42°C. After mixing, 10 l aliquots of the mixture were
transferred on plates of synthetic dropout selective medium containing 2% glucose
using a multi-channel pipette. Plates were let to dry at 22°C and incubated for 4 days
at 30°C.
Confirmation PCR for the yeast deletion mutants
PCR was performed as previously described in the Single Tube Confirmation PCR
Protocol (http://www-sequence.stanford.edu/group/yeast_deletion_project/single_
tube_protocol.html). Briefly, one colony was suspended in 50 l of 60U/ml zymolase
20T solution (Seikagaku Corporation) and incubated at 37°C for 1 hour, followed
by 10 minutes at 95°C. PCR was performed as followed: 100 pmol of each primer
was added in the PCR tubes containing 5 l of yeast suspension. The PCR master
mix was added and the PCR was carried out accordingly using the following
conditions: Step1, 3 minutes at 95°C; Step 2, 15 seconds at 94°C, 15 seconds at
57°C, 1 minutes at 72°C, repeated for 35 cycles; Step3, 3 minutes at 72°C. PCR
products were analyzed in 1.5% agarose gels.
2741
Bioinformatic analysis
The 59 human orthologs of yeast genes were identified using the Inparanoid database
(O’Brien et al., 2005), BLAST sequence alignment against all human protein
sequences and literature search. These genes were used as input into the DAVID
bioinformatic resource available at http://david.abcc.ncifcrf.gov (Dennis et al., 2003).
Pathway enrichment analysis was conducted using the functional annotation clustering
tool for testing the over-representation of genes in particular pathways from the
Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway database (Kanehisa
et al., 2010).
This test uses a standard Fisher exact test. A Bonferroni multiple testing and a Pvalue (Ease score) of 0.1 and fold enrichment of at least 1.5 were applied as standard
cut-off level.
Identification of the proteins interacting with the RhoA signaling pathway
The RhoA protein–protein interaction network was generated using the protein
interaction network analysis (PINA) (Wu et al., 2009) integrated platform. PINA
integrates protein–protein interaction data from six curated databases (MINT, IntAct,
BioGRID, HPRD amd MIPS/MPact) including experimentally validated and
computationally predicted interaction data (only experimentally reported interactions
were used for our analysis). We then examined whether any of the proteins encoded
by the human orthologs present in our data set share common interacting partners
with RhoA.
CDT and treatments
Production of the H. ducreyi CdtB was as previously described (Frisan et al., 2003).
His-tagged CdtA and CdtC were constructed by PCR amplification of the H. ducreyi
cdtA and cdtC genes from the pAF-tac1cdtA and pAF-tac1cdtC plasmids, respectively
(Frisk et al., 2001) using the following primers: CdtA_F 5¢-ATTCGGATCCATGTTCATCAAATCAACGAATGA-3⬘, CdtA_R 5⬘-TACCGAATTCTTAATTAACCGCTGTTGCTTCTAAT-3⬘, CdtC_F 5⬘- ATTCGGATCCAAGTCATGCAGAATCAAATCCTGA-3, CdtC_R 5⬘-5⬘-TACCGAATTCTTAGCTACCCTGATTTCTT3⬘. Each PCR fragment was cloned into the BamHI and EcoRI restriction sites of the
pACYC-Duet-1 expression vector (Novagen). Purification of the His-tagged CdtA and
CdtC subunits from inclusion bodies was performed as previously described (Moberg
et al., 2004). Reconstitution of the active holotoxin (named as CDT) was as previously
described (Frisan et al., 2003).
Toxin treatment
Cells were incubated for the indicated time periods with CDT (2 g/ml) in complete
medium.
Ionizing radiation
Cells were irradiated (20 Gy), washed once with PBS and incubated for indicated
time periods in complete medium.
RNA interference and transfections
A total of 105 HeLa cells were plated in 12-well plates in 1 ml of medium. Transfection
was performed using the forward protocol with the INTERFERinTM reagent (Polyplus
TransfectionTM), according to the manufacturer’s instructions. Gene silencing was
assessed by western blot analysis 48 hours after transfection. The following duplex
small interfering RNAs (siRNAs) were used: Hs_FEN1_6 #SI02663451 (FEN1_1);
Hs_TSG101_6 #SI02655184 (TSG101_1); Hs_TSG101_7 #SI02664522 (TSG101_2);
Hs_CSNK2B_5 #SI00605185 (CSNK2B _1); Hs_CSNK2B_6 #SI00605192
(CSNK2B _1); Allstars Negative Control siRNA #102780 (scRNA), all from Qiagen;
and FEN1 s5103 (FEN1_2) from Applied Biosystems, Ambion.
Immunofluorescence
Characterization of CdtB hypersensitive strains
To identify the CdtB hypersensitive mutants, transformants were picked, transferred
to 100 l of sterile H2O in 96-well plates, and diluted to an optical density at 600
nm (OD600) of 1. Five microliters of fivefold dilution series were transferred onto
10-cm diameter round plates containing selective medium supplemented with 2%
glucose (CdtB OFF) or 2% galactose/raffinose (CdtB ON) using a 48-pin replicator
and incubated at 30°C for 48 hours. Yeast strains that did not grow after the third
dilution step in raffinose/galactose in three independent screens were considered as
CdtB hypersensitive.
Flow cytometry analysis
Yeast cells were centrifuged at 3000 g for 5 minutes and the cell pellet was
resuspended and fixed with 1 ml 70% ethanol at 4°C for at least 24 hours. After an
additional centrifugation at 3000 g for 5 minutes, cells were resuspended in 0.8 ml
RNase solution (50 mM Tris-HCl pH 7.8, 20 g/ml RNase) and incubated overnight
at 37°C. Samples were centrifuged and resuspended in 0.5 ml PI solution (200 mM
Tris-HCl pH 7.5, 211 mM NaCl, 78 mM MgCl2, 25 g/ml propidium iodide), and
sonicated for 5 seconds at medium voltage. Thirty microliters of cell suspension was
mixed in a tube together with 0.6 ml 50 mM Tris-HCl pH 7.5. Samples were
analyzed with a FACS Calibure flow cytometer. Data from ~30,000 cells were
collected and analyzed using the CellQuest software.
Immunofluorescence analysis was performed as previously described (Frisan et al.,
2003; Guerra et al., 2008a), using the conformation-dependent monoclonal antibody
(6A7, BD Pharmingen), which recognizes the active form of Bax. The actin
cytoskeleton was visualized by staining with TRITC-phalloidin, as previously
described (Frisan et al., 2003). Nuclei were counterstained with DAPI (Vector
Laboratories Inc). Slides were mounted and viewed using a Leica DMRXA
fluorescence microscope with a CCD camera (Hammamatsu), and images were
captured using Improvision Openlab v.2 software. The TRITC-phalloidin fluorescence
intensity and the cell area were measured using ImageJ software
(http://rsbweb.nih.gov/ij/).
RhoA activation
RhoA activation was assessed using the G-LISATM RhoA Activation Assay Biochem
KitTM (Cytoskeleton), according to the instructions of the manufacturer.
Western blot analysis
Proteins were fractionated by SDS-polyacrylamide gels, transferred to PVDF
membranes (Millipore) and probed with the antibodies against phosphorylated
MAPK p38, MAPK p38, FEN1 (Cell Signaling), TSG101 and actin (Sigma). Blots
were developed with enhanced chemoluminescence, using the appropriate horseradish
peroxidase-labelled secondary antibody, according to the instructions of the
manufacturer (GE Healthcare).
2742
Journal of Cell Science 124 (16)
Statistical analysis
To evaluate the significance of the results, the independent two-sample t-test was
performed using the SPSS® software from IBM. Plotting histograms and boxwhisker graph was used to check the assumption of Normality of continuous data.
This work has been supported by the Swedish Research Council, the
Swedish Cancer Society, the Åke-Wiberg Foundation, the Magnus
Bergvall Fundation and the Karolinska Institutet to T.F., the Robert
Lundberg Memorial Fundation to L.G., the European Community
Integrated Project on Infection and Cancer (INCA), Project no. LSHCCT-2005-018704 to M.G.M. The Swedish Cancer Society supports T.F.
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/124/16/2735/DC1
Journal of Cell Science
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