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Molecular and Cellular Biology, December 2006, p. 9244-9255, Vol. 26, No. 24
0270-7306/06/$08.00+0 doi:10.1128/MCB.01538-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Southern Alberta Cancer Research Institute, Departments of Biochemistry & Molecular Biology and Oncology, Faculty of Medicine, University of Calgary, Calgary, Alberta, Canada T2N 4N1
Received 17 August 2006/ Accepted 26 September 2006
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), and p33ING1b protein showed
synergy with TNF-
in inducing apoptosis, which correlated with
reduced NF-
B-dependent transcription. These findings are
consistent with previous reports that HSP70 promotes
TNF-
-mediated apoptosis by binding I-
B kinase
gamma and impairing NF-
B survival signaling. Induction of
HSP70 required the amino terminus of ING1b but not the plant
homeodomain region that was recently identified as a histone binding
domain. Regulation of HSP70 gene expression by the ING tumor
suppressors provides a novel link between the INGs and the
stress-regulated NF-
B survival pathway important in hypoxia
and
angiogenesis. |
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Heat shock
proteins (HSPs) are a large family of evolutionarily conserved proteins
which function as molecular chaperones
(5). These specialized
proteins play important roles in cellular defense mechanisms against
protein aggregation and misfolding by binding nonnative states of other
proteins and assisting them in reaching a correctly folded and
functional conformation. They are also involved in protein
translocation across membranes to different organelles for final
packaging, degradation, or repair through their unfoldase activities
(20,
22). More than a dozen
HSPs, such as HSP27, -40, -60, -70, -90, and -110, etc., have been
identified so far, and they are named according to their molecular mass
(5,
63). Among them, HSP70 is
one of the most conserved, and it is also the best
characterized. In most mammalian cells there are two
prominent isoforms of HSP70, an abundant constitutive member called
HSP73 and the highly stress-inducible HSP72. HSP70 isoform expression
is enhanced by a number of different stress factors, including heat,
cold, glucose, alcohol, heavy metals, and ischemia
(38). Expression of HSP70
protects cells from stress
(46) and is generally
thought to play cytoprotective roles through antagonizing components of
apoptotic pathways, such as apoptosome formation and caspase complex
activation (53). Thus,
HSPs are believed to be involved in the proteasome-mediated degradation
of apoptosis-regulatory proteins in addition to having previously
identified chaperoning functions(45,
52). However, some
studies show that elevated HSP70 inhibits cellular proliferation
(34) and promotes T-cell
receptor/CD3- and Fas/APO-1/CD95-mediated Jurkat T-cell apoptosis
(32). The heat
shock response also sensitizes primary endothelial cells to
apoptosis through the inhibition of NF-
B
activity (11), and the
expression of HSP70 correlates with susceptibility to apoptosis in
acute myeloid leukemia cells
(6). As seen in
studies of endothelial cells, elevation of HSP70 to a significant level
favored tumor necrosis factor alpha (TNF-
)-mediated apoptosis
via inhibiting the NF-
B survival pathway
(45). This finding is
consistent with several studies reporting that HSP70 blocks
NF-
B activation and NF-
B-dependent gene expression
through inhibition of both I-
B kinase (IKK) activation
and subsequent degradation of I-
B
(35,
36,
46,
49,
62).
To examine the
mechanism by which ING proteins affect apoptosis in a genetically
normal model, we tested the effect of altering ING levels on global
gene expression by cDNA microarray analysis in the Hs68 strain of
primary human diploid fibroblasts. A subset of genes was specifically
and reproducibly affected by overexpression of the related
p33ING1b and p32ING2 proteins, including the
70-kDa heat shock protein HSP70. ING proteins also sensitized cells to
TNF-
-mediated apoptosis by impairing the NF-
B
signaling pathway via upregulation of HSP70 expression, and this effect
was dependent upon the amino terminus, but not the PHD, of
ING1.
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Generation of adenoviral constructs and infection of cells. Adenoviral constructs were generated using a modified "pAdEasy" system (24). ING1a, ING1b, ING2, and p53 were subcloned into pAdTrack-CMV, which contains a separate enhanced green fluorescent protein (GFP) expression cassette, and were recombined with pAdEasy-1 in Escherichia coli BJ5183. Recombinant clones were screened and subsequently verified by a series of enzymatic digestions. Recombinants were reamplified in XL1-Blue (Clontech), linearized by PacI (NEB), and transfected into 293 cells for packaging. Viral clones were plaque purified, selected for expression, amplified, and purified by CsCl2 gradient centrifugation. Plaque assays were routinely performed to ensure accurate viral titers. Optimized adenoviral infections were done at multiplicities of infection of 100 for young Hs68 cells and WI38 cells and of 10 for HeLa and HCT116 cells, giving >95% infectivity as monitored by GFP expression. No toxicity was observed when adenoviruses were used at these multiplicities of infection.
Analysis of cDNA microarrays. Hs68 cells were infected either with GFP alone or with GFP-ING adenoviruses. At 24 h after infection, cells were harvested and total RNA was extracted using TRIzol (Invitrogen). DNase treatment preceded the confirmation of RNA quality by examining the 260/280 absorbance ratio on a UV spectrometer (Pharmacia), by visual inspection after 1% agarose gel electrophoresis, and by PCR with GAPDH (glyceraldehyde-3-phosphate dehydrogenase) primers by use of RNA as the template and genomic DNA as a control. Reverse transcription generated cDNA from 10 µg of total RNA by use of a FairPlay microarray labeling kit (Stratagene). An indirect labeling method provided with the above-mentioned kit (Stratagene) was used to generate Cy3 and Cy5 (Amersham Biosciences) fluorescence-labeled cDNA. The labeled cDNA samples were then purified to remove uncoupled fluorescent dye and subsequently combined together with yeast tDNA (Stratagene) and hybridized to 14,000 human oligonucleotide chips (Southern Alberta Microarray Facility) by incubating at 37°C under a humidified condition for 18 h. After hybridization, arrays were washed for 3 to 4 min with 2x SSC (1x SSC is 0.15 M NaCl plus 0.015 M sodium citrate)-0.2% sodium dodecyl sulfate at room temperature (RT), for 5 min with 0.2x SSC at RT, and with 0.05x SSC for 5 min at RT. Slides were quickly dried and scanned using a fluorescence laser microarray scanning device (Virtex). Three independent replicates, including one dye reversal experiment, were performed for each ING gene assay. Replicate data were first quantitated by Array-Pro software (Media Cybernetics) and then transferred to GeneTraffic bioinformatic software (Iobion Informatics Company) for further data normalization, annotation, and management.
Primers and reverse transcription-PCR (RT-PCR). Hs68, WI38, or HCT116 cells were harvested 24 h after infection, total RNA was extracted using TRIzol (Invitrogen), and RNA quality was confirmed as described above. cDNA was made from 1 to 5 µg total RNA immediately after DNase treatment by use of a two-step reverse transcription method as described below. First, RNA treated with DNase was denatured with 1 µl of 500-ng/ml random primer (Amersham) and 1 µl of 10 mM deoxynucleoside triphosphate mixture (Amersham) at 70°C for 10 min and quickly chilled on ice. Then, 2 µl of 5x Superscript II buffer, 2 µl of 0.1 M dithiothreitol, and 1 µl of 32-U/µl RNase inhibitor (Invitrogen) were added, and the samples were incubated at RT for 10 min before the addition of 400 U of Superscript II (Invitrogen) to each tube. After incubation at 42°C for 1 h, cDNA samples were heated to inactivate enzymes at 70°C for 15 min and were subsequently used for PCR to estimate relative levels of HSP70, 25-kDa brain-specific protein (p25), mitotic kinesin-related protein/M-phase phosphoprotein (MPP), DNA polymerase theta (POLQ), clathrin heavy chain 2 (clathrin), L-type, calcium channel alpha-2/delta subunits(CANCLB), and ING1b and ING2 by comparison to internal controls (GAPDH or ß-actin) by use of the primer-dropping method (59). PCR primers used were as follows: HSP70, 5'-CGACCTGAACAAGAGCATCA-3'(forward) and 5'-TCGTCCTCCGCTTTGTACTT-3'(reverse); p25, 5'-ATCGGAGGGTGCTGGTGAGG-3'(forward) and 5'-GGTGCCTGCGTGCTTGTAGC-3'; MPP, 5'-TACGGGCTAACCAATTCAGG-3'(forward) and 5'-AGGCAACACTCTGGTGCTTT-3'(reverse); POLQ, 5'-TCAAAAGGCATAGCTCTCCT-3'(forward) and 5'-CCTCAAACATAGGTGTAACCAG-3'; CANCLB, 5'-GCATTGGAACTGGGACTTGT-3'(forward) and 5'-TTCCGAATCCTTCATTTTGC-3'(reverse); Clathrin, 5'-TGTGAATTATGCAACCAACG-3'(forward) and 5'-CCCCTCAGCAGAAAGATCC-3'; ING1b, 5'-GAAGATCCAGATCGTGAGCC-3'(forward) and 5'-GAGACCTGGTTGCACAGACA-3'(reverse); ING2, 5'-AAAATCGGGCAAGACAAATG-3'(forward) and 5'-GAAGCTTCCCTTTCCTGCTT-3'(reverse); GAPDH, 5'-GTCAGTGGTGGACCTGACCT-3'(forward) and 5'-AGGGGTCTACATGGCAACTG-3'(reverse); and ß-actin, 5'-GAACCCTAAGGCCAACCGTGA-3'(forward) and 5'-AGGAAGAGGATGCGGCAGTGG-3'(reverse). Aliquots of PCR products equalized to give equivalent signals from the internal control mRNAs (GAPDH/ß-actin) were electrophoresed through 2% agarose gels (Ultrapure; Pharmacia), stained with 0.2 µg/ml ethidium bromide, and analyzed by computerized densitometric scanning of the images by use of Kodak imaging software normalized using internal controls.
Transfection and dual luciferase reporter assays.
HeLa or Hs68 cells were seeded into
24-well tissue culture plates (8 x 104 cells/well)
16 to 18 h prior to transfection. Cells were first
cotransfected with either pHSE-luc reporter plasmid (a generous gift
from K. Yoshihara) or NF-
B-dependent luciferase reporter
construct, and with the pRL-TK (Promega) control vector to control for
transfection efficiency, by use of Lipofectamine 2000 (Invitrogen).
Sixteen to 18 h after transfection, cells were infected with
GFP, GFP-ING, or GFP-p53 adenoviruses. Positive control experiments
using heat shock were performed by incubating cells at 42°C for
30 min and then transferring back to 37°C for recovery.
Twenty-four h after infection or heat shock, luciferase activities were
measured by use of a luminometer (Berthold Technology Junior LB 9509)
with the dual luciferase reporter assay system (Promega) according to
the manufacturer's instructions. All experiments were performed in
triplicate, and statistical data analyses were performed using
Microsoft Excel software. TNF-
(Sigma) treatments began
12 h after infection and continued for another 24
h.
Western blotting.
Hs68, WI38, HCT116, or HeLa cells
were infected with the adenoviral constructs indicated. In some
experiments, cells were treated with TNF-
(Sigma) either at 25
ng/ml for 15 min or at 50 ng/ml for 24 h (unless otherwise
indicated) 12 or 24 h after infection. At the end of the
TNF-
incubations, cells were lysed in sodium dodecyl sulfate
loading buffer, and samples were electrophoresed and
blotted with anti-HSP72/73 monoclonal (Stressgen),
anti-I-
B polyclonal, anti-I-
B phospho-32/36
monoclonal (Cell Signaling), anti-poly(ADP-ribose) polymerase,
anti-cIAP2, anti-actin polyclonal, anti-FLIP, anti-GFP monoclonal
(Santa Cruz), anti-ING1 monoclonal, or anti-ING2 polyclonal (SACRI
Antibody Services) antibodies.
Apoptosis assays and cell viability. For microscopic visualization of chromatin condensation and fragmentation, Hs68 cells on coverslips were fixed in 3.7% formaldehyde in phosphate-buffered saline (PBS) for 5 min, permeabilized for 5 min in 0.5% Triton X-100 in PBS, and stained with 1 µg/ml DAPI (4',6'-diamidino-2-phenylindole) in PBS. Coverslips were then mounted in 1 mg/ml paraphenylenediamine in PBS-90% glycerol. Digital imaging was performed using a 14-bit cooled charge-coupled-device camera (Princeton Instruments) mounted on a Leica DMRE immunofluorescence microscope. Annexin V kits (Roche) were used according to the manufacturer's instructions to identify apoptotic cells by use of a FACScan flow cytometer in combination with BD Lysis II software (Becton-Dickinson). The viability of cells was assessed by trypan exclusion assay. The floating dead cells in the medium and the cells that remained attached to the plates were collected by trypsinization and counted using a hemacytometer in the presence of 0.4% trypan blue reagent (Sigma). All the experiments were done in triplicate, and statistical data analysis was performed using Microsoft Excel software.
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View this table: [in a new window] |
TABLE 1. Genes
with increased expression in response to
p33ING1ba
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View this table: [in a new window] |
TABLE 2. Genes
decreased in expression by
p33ING1ba
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View this table: [in a new window] |
TABLE 3. Genes
increased in expression by
p32ING2a
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FIG. 1. Verification
of array hits by primer-dropping RT-PCR. (A) Confirmation of
microarray targets. The genes that were reproducibly and significantly
regulated in microarray analyses were further tested by primer-dropping
RT-PCR (59). Hs68 cells
were infected with GFP, GFP-p33ING1b, or
GFP-p32ING2 adenovirus and harvested 24 h later,
and total RNA was extracted and treated with DNase. Duplex RT-PCR was
then carried out to detect HSP70, 25-kDa brain-specific protein (p25),
MPP, POLQ, L-type, CANCLB, and clathrin heavy chain 2 (Clathrin) by use
of sequence-specific primers for each gene and either ß-actin
primer or GAPDH primer as an internal control. (B) HSP70
analyses. Primary WI38 and Hs68 fibroblasts and immortalized HCT116
cells were infected with the indicated adenoviral constructs, and
24 h later total RNA was extracted. Primer-dropping RT-PCRs
were used to detect mRNA expression levels of HSP70 in the linear range
of the reaction using sequence-specific priming with ß-actin
primer as an internal control. Non, mock infected.(C) Summary of HSP70 induction data. The graph shows the
average level of HSP70 gene expression (normalized to ß-actin)
from three independent primer-dropping RT-PCR experiments and is
plotted ± standard
deviation.
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To determine whether higher levels of HSP70 mRNA resulted in increased protein levels, lysates of control and infected WI38, Hs68, and HCT116 cells were analyzed. As shown in Fig. 2A, p33ING1b and p32ING2 strongly induced levels of HSP72 and HSP73 in primary fibroblasts compared to the control, whereas p47ING1a was less effective. In contrast, although modest effects were seen at the mRNA level (Fig. 1C), none of the ING proteins tested markedly affected HSP70 protein levels in HCT116 (Fig. 2B), which is most likely due to the constitutively high levels of HSP70 expression previously noted for these cells (38). In addition, only a single HSP70 protein band was detected in this highly mutated line. ß-Actin was used as a loading control (Fig. 2A and B), and GFP blots were used to confirm that adenovirus infection gave equal levels of GFP expression in different samples (data not shown).
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FIG. 2. HSP70
protein levels are elevated by ING proteins in normal cells but not in
cancer cells. (A) Normal human skin primary diploid
fibroblasts (Hs68 cells), normal human embryo lung fibroblasts (WI38
cells), and (B) the HCT116 human colon carcinoma cell line
that contains wild-type p53 were either mock infected (non) or infected
with GFP, GFP-p47ING1a, GFP-p33ING1b,
GFP-p32ING2, or p53 adenovirus as indicated. Twenty-four h
later, cells were harvested, and lysates were used in Western blots
with HSP70 antibody (Stressgen) (top panels), ING1 and ING2 antibodies
(SACRI Antibody Services) (middle two panels), and actin antibody
(Santa Cruz) as a loading control (bottom panel). The ING2 panel for
HCT116 was generated from the same membrane as for ING1 and shows
simultaneous staining for p47ING1a, p33ING1b, and
p32ING2. (C to F) HSP70 gene promoter assays. Normal Hs68
cells and HeLa cells were cotransfected with the heat shock-responsive
pHSE-luc reporter construct and PRL-TK (Renilla luciferase
construct as an internal control for transfection efficiency). Eighteen
h later, cells were mock infected or infected with adenovirus encoding
GFP, GFP-p47ING1a, GFP-p33ING1b,
GFP-p32ING2, or p53 and incubated for a further 24
h before being harvested for luciferase assays. To induce heat shock,
cells were incubated at 42°C for 30 min and then incubated at
37°C for 24 h before harvesting. Increasing amounts
of plasmid encoding ING1b were cotransfected with reporter, as shown in
panel E, while the indicated plasmid constructs in a pCI backbone were
cotransfected with reporter, as shown in panel F. Amino acid residues
demarcating exon 1 of ING1b (Ex1), the novel conserved region (NCR) of
the ING proteins, the nuclear localization sequence (NLS), and PHD are
indicated at the bottom of panel F. Infected or transfected cells were
lysed in passive lysis buffer and subjected to a dual luciferase assay
as described previously (Promega). All panels are shown as a
representative of at least two replicate experiments. Values have been
equalized relative to the value obtained with non or control vector
(set at 1) and are the averages of three independent determinations
with standard deviations shown by the error bars. The differences seen
between non/vector and all of the other groups (indicated as
*)
are significant at the level of P
< 0.01 as estimated by analysis of variance. V, vector; WT,
wild
type.
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To further test for possible effects of p53 on HSP70 expression and to compare the effects of the ING proteins to those of heat shock treatment by use of an alternative assay and different cell types, an HSP70 promoter-driven luciferase reporter construct was cotransfected with control (GFP), ING, or p53 expression construct into HeLa and Hs68 cells. This reporter construct contains the HSP70 promoter, including the heat shock element responsible for inducing the expression of HSP70 in response to stress, upstream of the luciferase gene. As seen in Fig. 2C, the ING proteins induced HSP70 expression as effectively as heat shock in HeLa cells and nearly as effectively in Hs68 cells. Consistent with previous reports, we found that overexpression of the p53 protein inhibited HSP70 transcription in both cell types (1). Together, our results suggest that the ING proteins induce HSP70 by a mechanism independent of p53.
Dose-dependent expression of HSP70 depends upon the amino terminus of ING1 and is independent of the PHD. Next, we investigated the effect of increasing concentrations of the ING1b expression plasmid on the induction of the HSP70 promoter-driven reporter in primary fibroblasts. As shown in Fig. 2E, ING1b produced a dose-dependent increase in HSP70 promoter activity. Next, we carried out experiments to map out the determinants in ING1b that specify the induction of the HSP70 promoter. We tested the abilities of several deletion constructs of ING1b that are stably expressed to induce the HSP70 promoter. Interestingly, these data showed that the PHD region, which binds methylated histones (amino acids 211 to 256), and the adjacent polybasic region, which binds phospholipids (amino acids 256 to 279), were not required for induction (Fig. 2F). In contrast, deletion of the first 70 amino acids completely blocked the ability of ING1b to induce HSP70 (Fig. 2F). This region of the ING1b protein contains a PCNA-interacting protein domain as well as a partial bromodomain(14). ING2, which shares partial homology with ING1b in the amino terminus (23), was somewhat effective in inducing the HSP70 promoter, whereas ING4, which is divergent in this region, did not. Together these data suggest that the conserved N-terminal regions of ING1b and ING2 are required for these ING proteins to induce HSP70 expression.
Induction of apoptosis by TNF-
is enhanced by p33ING1b.
Since HSP70 was recently reported to
promote TNF-
-mediated apoptosis via interference with
NF-
B signaling
(45), we asked whether
the ING proteins combined with TNF-
treatment would more
efficiently induce apoptosis as a result of the concerted upregulation
of HSP70 expression and the subsequent disruption of NF-
B
signaling. In these studies, we used the ING1b protein, since it
produced a consistent induction of HSP70 in all of the different assays
used. Consistent with previous reports
(57), p33ING1b
alone induced a certain amount of apoptosis/cell death in Hs68 cells,
as evidenced by characteristic DAPI staining (chromatin condensation
and/or fragmentation in Fig.
3A), trypan blue exclusion (Fig.
3B), annexin V membrane
exposure (Fig. 3C), and
PARP cleavage (Fig. 3D).
When cells overexpressing p33ING1b were treated with
TNF-
(Fig. 3A to
D), much higher levels of apoptosis were noted with all
assays. This effect appears to be synergistic, since when combined,
p33ING1b and TNF-
result in levels of apoptosis
higher than the sum resulting from the use of both agents individually.
For example, the annexin V data indicate that 18% of cells are
apoptotic in response to p33ING1b and 10% are apoptotic
after TNF-
treatment but that when combined, approximately 60%
of Hs68 primary human diploid fibroblasts, which are generally
resistant to apoptosis, initiate programmed cell death (Fig.
3C). In contrast to these
effects, TNF-
treatment led to a much lower degree of
apoptosis in the ING1a-expressing cells (Fig.
3B and C). Together, these
data indicate that enhanced apoptotic response to TNF-
stimulation by ING1b correlates with its ability to induce
HSP70.
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FIG. 3. p33ING1b
and TNF- act synergistically to induce apoptosis.
(A) Hs68 cells infected with the indicated adenoviral
expression constructs were incubated for 12 h and
subsequently treated either with vehicle (upper panels) or with
TNF- (50 ng/ml) (lower panels) for another 24 h.
Cells were then fixed, stained by DAPI, and observed under the
fluorescence microscope. (B to D) Hs68 cells were infected with the
indicated expression constructs and 12 h later were treated
with vehicle or with TNF- (50 ng/ml) for another 24
h. Cells were harvested and subjected to trypan exclusion assay to
assess viability (B), annexin V assay (Roche) (C), or Western blotting
with PARP antibody (D). Lower panels show control Western blots to
verify that ING expression levels and protein loads were
similar.
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. As shown in Fig.
3A and B by use of
independent assays, p35 reduced apoptosis in TNF-
-treated
ING1b-expressing cells by approximately half. These data are consistent
with p33ING1b having effects upon the receptor-mediated
pathway via the TNF-
pathway and through effects upon the
mitochondrial apoptosis pathway, perhaps by inducing the p53 target
gene Bax.
The p33ING1b protein sensitizes cells to TNF-
-mediated apoptosis by inhibiting NF-
B signaling.
As
noted previously, HSP70 promotes TNF-mediated apoptosis by binding
IKK-
and impairing NF-
B survival signaling. To test
whether p33ING1b-induced HSP70 accumulation affected
apoptosis by inhibiting this pathway, we transfected cells with an
NF-
B-responsive promoter driving luciferase expression and
asked whether the expression of p33ING1b would interfere
with the ability of TNF-
to induce NF-
B activity. As
shown in Fig.
4A, reporter activity was increased in a TNF-
dose-dependent
manner as expected. Overexpression of p33ING1b in Hs68 cells
reproducibly inhibited NF-
B-dependent gene activation in both
the absence and the presence of TNF-
, consistent with
p33ING1b inhibiting NF-
B survival signaling in
response to TNF-
. In addition, when overexpressed,
p33ING1b reduced the TNF-
-dependent phosphorylation
and subsequent degradation of I-
B (Fig.
4B). Moreover, induction
of the antiapoptotic NF-
B target genes IAP2 and FLIP by
TNF-
was inhibited more by the expression of GFP plus
p33ING1b than by GFP alone (Fig.
4C). Blotting for GFP
confirmed that cells were uniformly infected with adeno-GFP and
adeno-GFP-ING1b, and ß-actin served as a protein loading
control (Fig. 4B and C).
These data suggest that the expression of p33ING1b
interferes with the ability of TNF-
to induce NF-
B
activity, thus switching from a pathway of survival to an apoptotic
pathway, as diagrammed in the model shown in Fig.
5.
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FIG. 4. p33ING1b
expression blocks TNF- -induced NF- B activity.
(A) Hs68 cells were cotransfected with NF- B-Luc
reporter construct and PRL-TK (Renilla luciferase construct as
an internal control) and 12 h later were infected with GFP or
with GFP-p33ING1b adenovirus as indicated. Twelve hours
postinfection, cells were treated with vehicle or with TNF-
(50 or 80 ng/ml) as indicated for another 24 h. Cells were
then lysed and subjected to a dual luciferase assay (Promega).
(B) HeLa cells were infected with GFP or with
GFP-p33ING1b adenovirus as indicated for 24 h and
then treated with vehicle or with TNF- (50 ng/ml) for 15 min.
Cells were quickly lysed and subjected to Western blotting with
I- B, phosphorylated I- B, ING1, GFP, and
ß-actin antibodies. (C) Hs68 cells were infected with
GFP or with GFP-p33ING1b adenovirus as indicated for
18 h and then treated with vehicle or with TNF- (50
ng/ml) for another 24 h. Cells were lysed and subjected to
Western blotting with IAP2, FLIP, ING1, GFP, and ß-actin
antibodies.
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FIG. 5. Model
linking HSP70 induction to p33ING1b-induced apoptosis.
Previous studies have linked HSP70 to TNF- -mediated apoptosis
by impairing NF- B survival signaling through binding
IKK- . p33ING1b has also been implicated in the
p53-inducible expression of the proapoptotic Bax member of the Bcl2
family. In this study, we show that p33ING1b induces
significant levels of HSP70, that it inhibits NF- B activity,
and that it acts synergistically with TNF- to induce
apoptosis. The p33ING1b protein, therefore, is proposed to
induce apoptosis by at least two mechanisms: by activating p53 and
inducing signaling through the mitochondrial pathway and by inhibiting
the NF- B survival pathway through the induction of
HSP70.
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to induce apoptosis, as assayed by chromatin
condensation, trypan blue dye exclusion, annexin V staining, and
cleavage of PARP. Induction is not a result of adenoviral effects,
since the same virus overexpressing GFP, GFP plus p53, GFP plus an
amino-truncated ING1b, or GFP plus ING4 did not induce HSP70. Finally,
p33ING1b inhibited the ability of TNF-
to induce
NF-
B transcriptional activity, supporting the idea that
p33ING1b-induced expression of HSP70 promotes apoptosis by
inhibiting the NF-
B survival signaling pathway. The ING1 family of type II tumor suppressors regulates gene expression through modulating chromatin structure via interacting with histone acetyltransferase and histone deacetylase complexes (14) and may direct them to specific chromatin locales by binding methylated histones via their PHD regions (37, 44, 50). Moreover, p53 target genes such as p21WAF1 and Bax, which regulate cell cycle progression and apoptosis, have previously been identified as downstream targets of p33ING1 and p32ING2. However, in our study these genes were not identified either by microarray analysis or by RT-PCR. This may be due to the fact that the studies noted above were carried out with different transformed cell lines in which multiple signaling pathways are altered to various degrees, while in this study we used normal Hs68 human primary diploid fibroblasts that have no known mutations in genes encoding tumor suppressors or cell cycle regulators or in proto-oncogenes. A recent microarray study done with the transformed epithelial mouse cell line NMuMG identified 25 genes (1.1% of 2,304 genes examined) whose expression levels were significantly altered upon a decrease in p33ING1b expression, but the set did not include p21WAF1, Bax, or HSP70 (54). In our microarray analysis, the expression levels of 22 (0.15% of 14,000 genes) and 30 (0.21% of 14,000 genes) genes were significantly up- and downregulated, respectively, in response to p33ING1b. The percentages of significant hits were comparable in spite of the different cell types and methodologies used in the two studies. However, the gene targets identified in these two microarray analyses did not overlap, which likely reflects profound cell type differences (immortalized mouse epithelial cells versus normal human fibroblasts) and/or microarray assay differences (cDNAs versus oligonucleotides and the noncomprehensive natures of both chip sets used).
In our analysis, the upregulation of HSP70 was both robust and reproducible in response to p33ING1b in at least two normal fibroblast strains and in two immortalized cell lines. The expression of several other heat shock genes, including those encoding HSP10, -27, -60, -75, -90, -105, and -110, which were included on our array chips, were not affected by the overexpression of ING proteins, suggesting that induction is not a result of a general stress reaction. Indeed, genes encoding HSP40 and HSP47 were downregulated by p33ING1b. HSP40 is a cochaperone protein that regulates complex formation between HSP70 and client proteins, thus facilitating protein folding and enhancing cell survival in response to stresses such as heat shock (13). HSP47 is a collagen-specific chaperone residing in the endoplasmic reticulum (42) that is essential for collagen biosynthesis. The downregulation of HSP40 by p33ING1b and the lack of induction of other HSPs suggest that the upregulation of HSP70 by ING proteins may have downstream effects differing from the cellular protective roles of HSP70 (46). This is consistent with our results showing that the overexpression of ING proteins did not have any effects in protecting cells from the adverse effects of heat shock (data not shown).
It has been known for some time that
cancer cells exposed simultaneously to heat and chemotherapeutic agents
or radiation die at a higher rate than do cells treated with
chemotherapeutic agents or radiation alone, which is the rationale for
the use of hyperthemia in combination with chemotherapy and
radiotherapy to treat cancer in the clinic
(9,
10,
11). Some ING proteins
have also been implicated in chemosensitivity and radiosensitivity
(4,
7,
19,
47,
48,
55,
64). It is possible,
therefore, that the upregulation of HSP70 by ING proteins is one
mechanism by which ING proteins confer chemosensitivity and
radiosensitivity, consistent with a link to the arm of the
TNF-
pathway that impinges on NF-
B-regulated gene
expression.
Several groups have now reported that
p33ING1b and p32ING2 induce apoptosis in
p53-dependent or p53-independent manners in different cell types
(25,
41), but the underlying
mechanisms remain unclear. Here we find that ING proteins sensitize
cells to TNF-
-mediated apoptosis through upregulation of
HSP70, which inhibits NF-
B survival signaling as outlined in
the model shown in Fig. 5.
Thus, in addition to a previous report linking p33ING1b to a
p53-associated mitochondrial apoptotic pathway
(7), this study supports
the contention that ING proteins also have a significant impact upon
p53-independent receptor-mediated apoptosis through NF-
B
signaling pathways.
HSP70 mRNA expression is regulated, at least
in part, by the direct effects of ING1 on the HSP70 promoter. This idea
is supported by previous reports showing ING binding to promoter
regions of genes (29) and
by the results shown in Fig.
2C, in which the
cotransfection of ING is shown to activate HSP70 promoter-driven
reporter constructs in a dose-dependent manner. However, the Hs68
primary fibroblasts show approximately three- to fourfold-higher
reporter activity in response to the ING proteins (Fig.
2C), but HSP70 mRNA and
protein levels increase to a much greater degree (
10- to
15-fold [Fig. 1B and C]).
Furthermore, small interfering RNA against ING1 did not markedly reduce
HSP70 expression (data not shown) or affect the ability of
TNF-
to induce apoptosis, indicating that the upregulation of
HSP70 promoter activity by ING1, while contributing to induction to
some degree, may not be the dominant mechanism responsible for the
ability of ING proteins to induce HSP70. Another mechanism that might
contribute to the ING-specific induction of HSP70, but not that of
other stress-inducible proteins, might be related to ING protein
activation of a pathway that impinges upon HSP70. Since the ING
proteins have been linked to the transduction of stress signals via
binding rare phosphatidylinositol monophosphates
(21,
28), it is possible that
the overexpression of ING in the absence of additional stimuli may
produce activated forms of ING protein, initiating a stress response.
The mechanism by which this response is generated could be through the
formation of aggresomes by ING proteins, which is known to induce HSP70
expression (30), and in
many cases is linked directly to the induction of apoptosis
(2), as is seen upon ING1
overexpression (Fig. 3).
This idea is supported by the observation that the amino terminus of
ING1b that tends to form aggregates (data not shown), but not the more
hydrophilic downstream domains containing the nuclear localization
sequence, PHD, and polybasic regions of ING1b, is effective in inducing
HSP70 expression (Fig.
2F).
Traditionally,
HSP70 has been thought to protect cells by interfering with the
mitochondrial apoptotic pathway
(20,
53); however, a growing
body of evidence shows that heat shock or elevated HSP70 can also
promote receptor-mediated apoptosis
(32,
60). In particular, HSP70
can enhance cell death when it is overexpressed in cells that are also
exposed to TNF-
via inhibiting the NF-
B signaling
cascade (45). This is
reasonable considering the fact that it is well established that
NF-
B signaling is negatively regulated by HSP70
(22,
36,
49,
62). In this study, we
show that p33ING1b inhibits TNF-
-induced
NF-
B-dependent gene transcription (Fig.
4), suggesting that
p33ING1b disrupts NF-
B signaling through the
upregulation of HSP70.
Another member of the ING family of tumor
suppressors, named ING4, has also recently been shown to inhibit
NF-
B activity, resulting in reduced expression of
NF-
B target genes
(18). In that study,
evidence consistent with a physical interaction between ING4 and the
p65 subunit of NF-
B, obtained primarily by use of
overexpression studies, was presented, although links to HSP70 and
TNF-
signaling
(58) were not examined.
Given these data, it is possible that ING1b and ING4 negatively
regulate NF-
B activity through mechanisms involving direct
binding to p65 and via HSP70 induction. Alternatively, ING1 and ING4
may inhibit the NF-
B signaling pathway by distinct mechanisms.
Our finding that ING2 overexpression also leads to HSP70 induction but
that ING4 does not (Fig. 2
and 3), together with the
conserved features of the ING proteins
(23), suggests that ING1b
and ING2 downregulate the NF-
B pathway by use of a common
mechanism related to the amino termini of these proteins. Testing if
p33ING1b directly interacts with components of the
NF-
B pathway will further refine the model presented in Fig.
5 and provide a better
understanding of how the ING family of type II tumor suppressors exert
their effects upon apoptosis and cell growth
control.
This study was supported by grants from the Canadian Institutes of Health Research and the Alberta Cancer Board to K.R. X.F. holds Alberta Heritage Foundation for Medical Research (AHFMR) and ACB/CIHR TRTC doctoral studentships, S.B. is an AHFMR Scholar, and K.R. is a Scientist of the AHFMR.
Published ahead of print on 9 October 2006. ![]()
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