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Molecular and Cellular Biology, October 1999, p. 6963-6971, Vol. 19, No. 10
0270-7306/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Altered Telomere Nuclear Matrix Interactions and
Nucleosomal Periodicity in Ataxia Telangiectasia Cells before and after
Ionizing Radiation Treatment
Lubomir B.
Smilenov,
Sonu
Dhar, and
Tej K.
Pandita*
Center for Radiological Research, College of
Physicians and Surgeons, Columbia University, New York, New York
10032
Received 4 April 1999/Returned for modification 24 May
1999/Accepted 14 July 1999
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ABSTRACT |
Cells derived from ataxia telangiectasia (A-T) patients show a
prominent defect at chromosome ends in the form of chromosome end-to-end associations, also known as telomeric associations, seen at
G1, G2, and metaphase. Recently, we have shown
that the ATM gene product, which is defective in the
cancer-prone disorder A-T, influences chromosome end associations and
telomere length. A possible hypothesis explaining these results is that
the defective telomere metabolism in A-T cells are due to altered
interactions between the telomeres and the nuclear matrix. We examined
these interactions in nuclear matrix halos before and after radiation treatment. A difference was observed in the ratio of soluble versus matrix-associated telomeric DNA between cells derived from A-T and
normal individuals. Ionizing radiation treatment affected the ratio of
soluble versus matrix-associated telomeric DNA only in the A-T cells.
To test the hypothesis that the ATM gene product is
involved in interactions between telomeres and the nuclear matrix, we
examined such interactions in human cells expressing either a
dominant-negative effect or complementation of the ATM gene. The phenotype of RKO colorectal tumor cells expressing ATM fragments containing a leucine zipper motif mimics the altered interactions of telomere and nuclear matrix similar to that of A-T
cells. A-T fibroblasts transfected with wild-type ATM gene had corrected telomere-nuclear matrix interactions. Further, we found
that A-T cells had different micrococcal nuclease digestion patterns
compared to normal cells before and after irradiation, indicating
differences in nucleosomal periodicity in telomeres. These results
suggest that the ATM gene influences the interactions between telomeres and the nuclear matrix, and alterations in telomere chromatin could be at least partly responsible for the pleiotropic phenotypes of the ATM gene.
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INTRODUCTION |
Ataxia telangiectasia (A-T) is a
rare autosomal human recessive disorder characterized by progressive
neurological degeneration, growth retardation, premature aging,
telangiectasia, specific immunodeficiencies, high sensitivity to
ionizing radiation, gonadal atrophy, genomic instability, and
predisposition to cancer (9, 26, 65). Cells derived from A-T
individuals exhibit a variety of abnormalities in culture such as a
higher requirement for serum factors, hypersensitivity to ionizing
radiation, and cytoskeletal defects (50, 72). Primary
fibroblasts from humans and mice with a defective ATM gene
(see below for description) grow slowly in culture and appear to
undergo premature senescence in culture (4, 26, 50, 72).
They also show a prominent chromatin defect at chromosome ends in the
form of chromosome end-to-end associations seen at different phases of
the cell cycle (63, 64, 74), and these associations are
enhanced by stress such as ionizing radiation treatment
(74). Chromosome end associations involve telomeres composed
of repetitive DNA sequences of TTAGGG arrays concealed by a
complex of specialized proteins that protect ends from exonucleolytic
attack, fusion, and incomplete replication. Telomeric associations
correlate with genomic instability and carcinogenesis (15, 63,
64).
Telomeres shorten as a function of age in cells derived from normal
human blood, skin, and colonic mucosa (1, 17, 27, 43, 80).
As a result of this shortening, it is thought that critical genes at
the ends of the chromosomes either become deleted or are activated,
leading to cell growth arrest (41, 57, 58, 86). Recovery of
proper telomere length by the activation of telomerase prolongs the
life span of a cell (8, 81). Shortening or loss of telomeres
in a variety of cancers and immortalized cell lines is correlated with
chromosome end associations that could be the cause of genomic
instability and gene amplification (15, 46, 55, 61, 63, 75,
77).
There is growing evidence suggesting that both the shielding of
telomeric ends and their elongation by telomerase are dependent on
telomere binding proteins. Mammalian telomeres are packaged in
telomere-specific chromatin (76). Human and mouse cell lines have their telomeric tracts attached to the nuclear matrix, which is a
proteinaceous subnuclear fraction (16, 44). There is a
difference in nucleosomal organization of telomeres compared to bulk
DNA, and telomeric histone H4 is hypoacetylated (40, 47, 59,
78). Telomere length homeostasis in yeast requires the binding of
a RAP1p molecule along the telomeric tract (36, 45, 48), and
change in the telomeric matrix binding site occurs at least once in
every kilobase of the telomeric tract in tumor-derived cell lines
(44). It has been suggested that mammalian telomeres have
frequent multiple interactions with the nuclear matrix over a large
domain that encompasses the entire telomeres of most of the chromosome
ends (44). Whether the ATM gene influences the interaction of telomeres with the nuclear matrix is not yet known.
The gene that mediates the disease A-T has been designated
ATM (A-T, mutated), and its product shares the
phosphatidylinositol 3'-kinase signature of a growing family of
proteins involved in the control of cell cycle progression, processing
of DNA damage, and maintenance of genomic stability (28, 31, 68,
69). The protein shows similarity to several yeast and mammalian
proteins involved in meiosis of fission yeast and to the TOR proteins
of yeast and mammals (54, 69). In mitotic cells, ATM is
required for a DNA damage-dependent signal transduction cascade that
activates multiple cell cycle checkpoints (50, 52, 72). The
presence of a leucine zipper (LZ) in the ATM protein suggests possible dimerization of the protein or interaction with additional proteins (68, 69). The only known proteins that interact with ATM are p53, c-Abl, and 
-adaptin (5, 32, 42, 71). The ATM protein has been shown to contribute to the induction of c-Abl activity (5, 71), a tyrosine kinase activated by ionizing radiation, and certain other DNA-damaging agents (33). Hawley and
Friend (28) have suggested that an ATM-like protein has a
critical role in maintaining chromosome condensation in the vicinity of recombination intermediates. In support of the role of the
ATM gene in chromosome condensation, it has been reported
that mei-41 (homologue of the ATM gene)-bearing
oocytes exhibit diffused chromatin (25). Because of the
homology of ATM to TEL1 mutants of yeast (23), it has further been suggested that mutations in
ATM could lead to defective telomere maintenance. Mammalian
telomeres have gained importance because of their possible link with
carcinogenesis (15, 17, 27, 63). Since patients with A-T are
prone to develop cancer and the ATM gene influences telomere
metabolism (74), we have studied the differences among A-T
and normal cells in telomere-nuclear matrix interactions, telomere DNA
structure, and nucleosomal periodicity before and after ionizing
radiation treatment.
To directly test whether the ATM gene product influences
telomere-nuclear matrix interactions, we examined cells with
dominant-negative as well as complementing activity with respect to ATM
function. Expression of the dominant-negative ATM fragments in RKO
cells leads to decreased clonogenic survival, increased chromosomal aberrations, radioresistant DNA synthesis after treatment with ionizing
radiation, and defective telomere metabolism (53, 74). The
ATM protein or fragment containing the kinase domain complemented radiosensitivity, the S-phase checkpoint, irradiation-induced activation of c-Abl, reduced chromosome aberrations after treatment with gamma rays, and reduced frequency of cells with telomere fusions
in simian virus 40-transformed fibroblasts derived from A-T individuals
(5, 53, 71, 87).
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MATERIALS AND METHODS |
Cell culture.
Derivations of the cell strains used are given
in Table 1. All fibroblast strains were
maintained according to procedures described earlier (62, 63,
74). Cell viability was monitored by the trypan blue exclusion
test, and cell population densities were determined by hemacytometer
and electronic counting (Coulter Electronics Inc., Hialeah, Fla.).
The colorectal carcinoma (RKO) cells, with and without the
ENA/FB2F-expressing ATM fragment which contains the LZ motif, were
grown as described previously (
53,
74). A-T cells with and
without full-length cDNA (AT22IJE-T, AT22IJE-TpEBS7, and
AT22IJE-TpEBS7-YZ5)
were obtained from Yossi Shiloh, Tel Aviv
University, Israel,
and the conditions for maintaining the clones were
the same as
described previously (
87). Expression of the ATM
LZ fragments
in the RKO cell line and the ATM full-length cDNA was
determined
as described earlier (
53,
87). Relevant
characteristics of
the isogenic cells used are summarized in Table
2.
Determination of terminal restriction fragment length.
DNA
was isolated from plateau-phase cells by a procedure described earlier
(64, 74). For measuring terminal restriction fragment
lengths, DNA was digested with restriction enzyme RsaI or
HinfI, which do not cut TTAGGG sequences,
processed for fractionation, and hybridized with a
32P-labeled (TTAGGG)5 probe.
Detection and measurement for terminal restriction fragment length were
performed as described earlier (63, 64). The mean length of
the telomere terminal restriction fragment was measured by using
ImageQuant (version 1.2, build 039; Molecular Dynamics.
Determination of telomere-nuclear matrix association.
Plateau-phase cells were used to prepare the nuclear matrix halos,
which were isolated by removing histones and other loosely bound
proteins. Nuclear halos are morphologically defined as nuclear structures that remain after the selective removal of perinuclear components with ionic detergent. The halos are thought to represent relaxed lengths of loops of DNA with periodical attachment to the
nuclear matrix, which is a residual framework of nucleoskeletal proteins. The procedure used for the isolation of lithium
diiodosalicylate (LIS)-generated halo structures is a modification of
the LIS technique described by Mirkovitch et al. (51),
Dijkwel and Hamlin (18), Luderus et al. (44),
Berezney and Coffey (6), and Berezney et al. (7).
Cells were trypsinized, washed twice with cold phosphate-buffered
saline (PBS) and twice with 25 ml of cold cell wash buffer (CWB; 50 mM
KCl, 0.5 mM EDTA, 0.05 mM spermidine, 0.05 mM spermine, 0.25 mM
phenylmethylsulfonyl fluoride [PMSF], 0.5% thiodiglycol, 5 mM
Tris-HCl [pH 7.4]), pelleted at 1,000 × g for 5 min,
and then suspended in 12 ml of CWB containing 0.1% digitonin
(Boehringer Mannheim). The cells were passed through a 20-gauge needle,
and lysis was monitored by phase-contrast microscopy. The 2-ml
suspension was loaded on 3 ml of 10% glycerol cushion in CWB and spun
for 10 min at 800 × g; the nuclei were washed with CWB
containing 0.1% digitonin, suspended in CWB with 0.1% digitonin and
0.5 mM CuSO4 but without EDTA, and incubated for 20 min at
37°C. About 19 volumes of LIS solution (10 mM LIS, 100 mM lithium
acetate, 0.1% digitonin, 0.05 mM spermine, 0.125 mM spermidine, 0.25 mM PMSF, 20 mM HEPES-KOH [pH 7.4]) was added, and the mixture was
incubated for 10 min at room temperature. Halos were collected by
centrifugation for 10 min at 2,800 rpm in a benchtop Eppendorf
centrifuge (model 5403) and washed three times with matrix wash buffer
(MWB; 20 mM KCl, 70 mM NaCl, 10 mM MgCl2, 10 mM Tris HCl
[pH 7.4]) with 0.1% digitonin. The resulting halo structures contain
naked chromosomal DNA and the nuclear matrix. The nuclear halos were
then washed with a restriction enzyme buffer, 6 × 106
halos were cleaved in a volume of 0.5 ml containing 1,000 U of restriction enzyme StyI for 3 h at 37°C, and the
nuclear matrices were pelleted by centrifugation. To purify released
and attached DNA fragments to the nuclear matrix, both fractions were
treated with proteinase K in a solution containing 10 mM EDTA, 0.5%
sodium dodecyl sulfate, and 10 mM Tris-HCl (pH 7.4) and incubated
overnight at 37°C. DNA was purified as described previously (64,
74). Agarose gel electrophoresis was performed for the
fractionation of DNA (64). For Southern blot analysis, equal
volumes from about 106 halos were fractionated on 0.8%
agarose gels. Prior to DNA loading, RNase was added to a final
concentration of 10 µg/ml. Fractionation of DNA, transfer to a
Hybond-N membrane, slot blotting of DNA, hybridization with a
32P-labeled (TTAGGG)5 probe, and
detection were done as described previously (64).
Quantitation and comparison of the telomeric DNA among total, released,
and telomeric DNA fragments attached to the nuclear matrix were
achieved by phosphorimaging.
Preparation of chromatin, digestion with MNase, and detection of
telomeric nucleosomes.
Cells were grown to plateau phase,
trypsinized, and washed twice with growth medium. Cell viability was
monitored by trypan blue exclusion, and cell counts were determined by
hemacytometer and electronic counting (Coulter). Cells were kept on ice
and were used immediately for chromatin preparation. All manipulations were done at 4°C. Cells were suspended in a buffer consisting of 10 mM Tris-HCl (pH 7.5), 50 mM KCl, 3 mM MgCl2, 1 mM
CaCl2, and 0.5 mM PMSF at 2 × 106 cells
per ml and washed three times with the same buffer. Cells were lysed by
addition NP-40 to a final concentration of 0.5% and incubation on ice
for 15 min. Lysis was monitored by visualizing the nuclei under a
microscope. Nuclei were washed twice with buffer to remove the
detergent and aliquoted in 100-µl volumes containing 5 × 106 nuclei. Micrococcal nuclease (MNase) (Nuclease S7;
Boehringer Mannheim) was added to aliquots of nuclei at 30°C for 5 min to give final concentrations ranging from 0 to 8,000 U/ml, and
digestion was terminated by the addition of an equal volume of solution containing 10 mM Tris-HCl (pH 7.5), 10 mM EDTA, 0.1% sodium dodecyl sulfate, and proteinase K at a final concentration of 50 µg/ml. The
solutions were incubated for 2 h at 37°C, and DNA was purified by phenol-chloroform extraction, precipitated with isopropanol in the
presence of 0.3 M sodium acetate (pH 5.5), and dissolved in 200 µl of
10 mM Tris-HCl. DNA (5 µg) was loaded into a 1.5% agarose gel, and
blotting and hybridization for TTAGGG were performed as
described previously (64).
Determination of TRF1 and TRF2 expression.
Total RNA was
isolated from fibroblasts by using an RNeasy kit (Qiagen, Santa
Clarita, Calif.). cDNA was prepared from equal amounts of RNA as
described previously (73). The primers used for
amplification of the human telomere binding factor (hTRF1 and hTRF2) genes were designed from cDNA sequences (10,
14). Primer information will be provided on request. Equal
amounts of cDNA from different cell types were used for PCR
amplification and quantification of the TRF1 and TRF2 products.
SSCP.
To determine the mutations in the hTRF1 and
hTRF2 genes, we analyzed cDNA by the cold single-strand
conformational polymorphism (SSCP) protocol (24, 29). RNA
isolation and cDNA preparation were done as described above. Five sets
of primers covering the hTRF1 cDNA and six sets of primers
covering the hTRF2 cDNA were made. Primer information will
be provided on request. PCR conditions were the same as described
previously (19, 24). Cold SSCP was performed according to
the instructions of Novex (San Diego, Calif.). Electrophoresis was
carried in ready-made precast polyacrylamide gels (Novex), using a
ThermoFlow cold SSCP Novex unit. A positive control that has one allele
mutated and produces four bands was also run.
Immunostaining.
Antibodies against TRF1 and TRF2 proteins
obtained as a kind gift from Titia de Lange (Rockefeller University,
New York, N.Y.) were used. Cells were grown on coverslips, washed with
PBS, fixed in 2% paraformaldehyde in PBS for 10 min, permeabilized for
20 min in 0.5% NP-40, and then stained with the primary antibodies and
anti-rabbit Cy-3-labeled donkey antibody (Jackson Research Laboratories).
Gene expression profile obtained by using total RNA.
We used
human Atlas cDNA expression arrays (Clontech Laboratories, Palo Alto,
Calif.) to determine the differences in gene expression among A-T and
normal fibroblasts, using the procedure described in the Clontech
manual. The Atlas cDNA membrane contains 588 known human genes. In
brief, poly(A)+ RNA was isolated from the fibroblasts by
using an Oligotex Direct mRNA mini kit (Qiagen). The
poly(A)+ RNA was treated with RNase-free DNase I
(Boehringer Mannheim) and then used for cDNA synthesis and labeled with
-32P. The labeled cDNA was hybridized to the Atlas
arrays, followed by washing and exposure of the membrane in a
PhosphorImager (Molecular Dynamics). Image analysis and quantification
were done individually for each dot by using the Scion Image program
(Scion Corp., Frederick, Md.).
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RESULTS |
Studies of telomere-nuclear matrix interactions and telomere
nucleosomal periodicity in A-T and normal individuals were carried out
on primary fibroblasts. The telomeres of these cells shorten during
proliferation in culture. It is possible that the interactions of
telomeres with the nuclear matrix depend on the length of the terminal
restriction fragment (telomere length) of the chromosomes. Therefore,
we determined the mean telomere length of each cell type at the time
their nuclear matrix interactions and nucleosomal periodicity were
examined. To determine the size of the terminal restriction fragment
length, we digested the DNA with restriction enzyme RsaI or
HinfI, which do not cut TTAGGG sequences. First, we examined several cell strains of A-T and normal individuals to
select those with comparable telomere lengths; five strains were
selected for study. The mean telomere lengths of A-T primary cells
(GM5823 and GM2052; 7.0 ± 1.0 and 10.3 ± 1.7, respectively) were comparable to those of the controls (AG1522, AG6234, and C21F;
8.6 ± 1.3, 9.8 ± 0.9, and 7.2 ± 0.6, respectively).
The mean telomere length observed in cells at different passages (Fig. 1) used for further experiments did not
show any significant changes.
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FIG. 1.
Genomic blotting analysis of telomere length. DNA was
digested with RsaI or HinfI and analyzed by
Southern hybridization using a TTAGGG repeat probe.
Molecular sizes are indicated at the right. Note that one of the A-T
fibroblasts has a greater telomere length than normal fibroblasts. The
results are representative of three experiments done separately. Cell
strain AG1522 represents the telomere size at passage 18 in panel a and
passage 20 in panel b.
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Telomere-nuclear matrix association.
To characterize the
nature of telomere anchorage to the nuclear matrix of different cell
types, plateau-phase cells were processed by the LIS procedure (6,
18, 44) and the resulting nuclear matrix halos were cleaved with
StyI. The nuclear matrix halos are the insoluble
nonchromatin scaffolding of the interphase nuclei. The nuclear remnant
and associated DNA were isolated by centrifugation and suspended in
MWB. For genomic blotting analysis, equal volumes representing DNA from
the identical numbers of halos were fractionated side by side on 1.5%
agarose gels. The amount of telomeric sequence in each sample was
determined by storage phosphorimage analysis. The normal fibroblasts
have about 52 to 60% of the telomeric DNA associated with the nuclear
matrix (attached) (P) fraction and 40% to 48% in the soluble (free)
(S) fraction (Fig. 2a; Table 3). Summation of the P and S values is
equal to total telomeric DNA (T), suggesting that no telomeric DNA was
lost during the extraction procedure. Figures 2b and c show comparison
among A-T and control cells for the P and S fractions of telomeric DNA. In A-T cells (GM2052), more than 92% of the telomeric DNA is attached to the nuclear matrix, whereas in control cells (AG6234) about 52% is
attached (Fig. 2b; Table 3). In another A-T cell strain (GM5823) more
than 95% of the telomeric DNA is attached to the nuclear matrix,
whereas in control cells (AG1522) about 60% is attached (Fig. 2c;
Table 3). The ratio between the S and P fractions of telomeric DNA is
about 1:19 to 1:11.5 in A-T cells, compared to 1:1.5 to 1:1.1 in normal
cells (Table 3). These results suggest that the major portion of
telomeres in A-T cells is associated with the nuclear matrix.
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FIG. 2.
Autoradiograph of telomeric DNA in untransformed
fibroblasts of A-T and normal individuals. Halo preparations were
digested with StyI and centrifuged to separate S and P
telomeric sequences. (a) T, P, and S fractions of telomeric DNA in two
normal fibroblast cell line (C21F and AG6234). Note that the summation
of P plus S values is equal to T. (b) Comparison of P and S fractions
of telomeric DNA between AG6234 and GM2052 cells. Lanes T, S, and P for
AG6234 (normal fibroblast) and lanes T, S, and P for GM2052 (A-T
fibroblast) represent telomeric DNA fibroblasts from similar numbers of
halos. (c) Comparison of P and S fractions of telomeric DNA between
AG1522 and GM5823 cells. Lanes P and S of AG1522 (normal fibroblasts)
and lanes S and P of GM5823 (A-T fibroblasts) represent telomeric DNA
from similar numbers of halos. Note the difference in the ratio of P
versus S fractions of telomerase DNA between A-T and normal cells. The
results are representative of four experiments done separately.
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To determine whether altered interactions of telomeres with the nuclear
matrix are due to ATM function, we examined isogenic
cells with and
without normal ATM function. Two different approaches
were used to
determine the influence of ATM function on telomere
associations with
the nuclear matrix. In the first approach, we
determined P and S values
for RKO cells with and without expression
of the dominant-negative ATM
fragment. We found RKO cells expressing
dominant-negative fragments
have 87% of telomeric DNA in the P
fraction and 13% in the S
fraction, whereas parental RKO cells
have 71% in the P fraction and
29% in the S fraction (Fig.
3a;
Table
4). When ratios between means of P
and S were determined
(Table
4) and compared between RKO and RKOFB2F7
(with expression
of dominant-negative fragment) cells, it was found
that RKOFB2F7
cells have 3.1-fold higher ratio than RKO cells,
suggesting that
inactivation of ATM influences the telomere
associations with
the nuclear matrix. However, the difference in P
values of RKO
cells with and without expression of dominant negative
ATM fragment
was lower than the differences in P values between primary
cells
derived from A-T and normal individuals. The possible reason for
this could be that RKO cells are derived from tumors and thus
may have
other factors that could partly rescue the ATM phenotype.
In the second
approach, we determined whether wild-type ATM could
correct the altered
interactions between telomeres and the nuclear
matrix by examining A-T
(AT22IJE-T) cells with and without expression
of the wild-type ATM
protein. Two different techniques were used
to determine the values of
P and S. By slot blot analysis, differences
in the values of P and S
were distinct between the parental cell
line (ATT221JE-T) and the
derivative cells with the wild-type
ATM gene
(AT221JE-TpEBS7-YZ5) (Fig.
3b; Table
4). Similar results
were obtained
by using Southern analysis (Fig.
3c; Table
4).
AT221JE-TpEBS7-YZ5 cells
with a wild-type
ATM gene have lower
amounts of P fraction
compared to AT221JE-TpEBS7 cells that contain
an empty vector. These
results reveal that the expression of the
wild-type
ATM gene
in A-T cells restored the normal telomere-nuclear
matrix interactions,
as is evident by the decrease in the amount
of the P fraction (Fig.
3;
Table
4).
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FIG. 3.
Autoradiograph of telomeric DNA in RKO cells with and
without expression of the dominant-negative fragment of ATM and A-T
(AT221JE-T) cells with and without expression of the wild-type ATM
protein. (a) Comparison of T, P, and S fractions among RKO,
RKOpBABEpuro, and RKOFB2F7 cells. RKO cells are parental cells,
RKOpBABEpuro cells are RKO cells with empty vector, and RKOFB2F7 cells
are RKO cells expressing dominant-negative fragments of ATM. Note the
difference in the ratio of P versus S fractions of telomeric DNA
between RKO and RKOFB2F7 cells. Two different approaches were used to
quantitate the telomeric DNA in T, P, and S fractions. Panel b compares
T, P, and S fractions of A-T cells with (AT22IJE-TpEBS7-YZ5) and
without (AT221JE-T) expression of the wild-type ATM protein by the slot
blot procedure. Note the difference in P and S values between the A-T
cells with and without the wild-type ATM protein. Panel c represents
use of the Southern procedure to quantitate P and S fractions between
the A-T cells with empty vector (AT221JE-TpEBS7) and A-T cells with
vector containing cDNA of wild-type ATM (AT221JE-TpEBS7-YZ5). Note the
difference in the ratio of P versus S fractions of telomeric DNA
between A-T cells with and without the wild-type ATM protein.
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Influence of ionizing radiation on telomere-nuclear matrix
associations.
The results presented above suggested that
untransformed A-T cells have altered telomere-nuclear matrix
associations. By using the isogenic cells, we further demonstrated that
ATM function influences telomere-nuclear matrix associations. Since
gamma irradiation triggers telomere associations in A-T cells
(74), it was important to determine the effects of ionizing
radiation on the interactions of telomeres with the nuclear matrix.
Plateau-phase cells were treated with a dose of 5 Gy of ionizing
radiation, and proportions of S and P fractions of telomeric DNA were
determined. As shown in Fig. 4, no change
in the ratio of S versus P fractions of telomeric DNA was seen
immediately after treatment with ionizing radiation in either the
control or the A-T cells. However, an increase in telomeric DNA in the
S fraction was seen in A-T cells 1 h after treatment, whereas no
such change was found in normal cells. The ratio of S versus P
fractions of telomeric DNA changed from 1:19 at 0 min to 1:5 at 60 min
postirradiation. Since we did not see any change in this ratio in
normal cells 1 h postirradiation, we wished to determine whether
there are any changes immediately after irradiation. Therefore, we
examined the S and P fractions of telomeric DNA in normal cells at 0, 15, 30, and 60 min postirradiation and found no differences (data not
shown). These observations suggest that the interactions of telomeric
DNA in normal cells are not influenced by exposure to 5 Gy of gamma
rays.
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FIG. 4.
Autoradiograph of P and S fractions of telomeric DNA at
0 and 60 min after cells were irradiated with 5 Gy of gamma rays. Note
there is no change in the S fraction of telomeric DNA after 60 min of
irradiation in normal cells (AG1522), whereas there is a significant
increase in the S fraction of telomeric DNA in the A-T cell line
(GM5823). The results are representative of three experiments done
separately.
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Nucleosomal organization in telomeric chromatin.
To determine
the nucleosomal organization of the telomeric arrays of TTAGGG,
we digested the nuclei with different concentrations of MNase and
detected telomere repeats by Southern analysis using the TTAGGG,
probe. Cells were grown to plateau phase and trypsinized, and the
nuclei were prepared as described recently (73). Equal numbers of nuclei from each cell line were digested with MNase, and the
isolated DNA fragments were fractionated on an agarose gel. We compared
the organization of bulk and telomeric chromatin by determining the
nucleosome periodicity in both samples. This was achieved by
simultaneously digesting nuclei from all cell strains with different
concentrations of MNase and fractionating the isolated DNA fragments on
agarose (Fig. 5). Bulk nucleosome arrays
were detected by ethidium bromide staining, and telomeric arrays were
detected by filter hybridization with the TTAGGG, probe. All
cell lines showed a ladder pattern upon ethidium bromide staining (Fig.
5). Interestingly, we found higher MNase digestion of chromatin in A-T
cells than of normal cells (compare Fig. 5a with Fig. 5c and e). This
suggests that the chromatin in A-T cells might be more loosely
condensed than that in normal cells. As shown in Fig. 5b, normal
(AG1522) cells have a telomere ladder containing partial digestions of
up to seven subunits. In contrast, the telomeric pattern in A-T (GM5823
and GM2052) cells revealed a less extensive MNase-dependent nucleosomal
periodicity (Fig. 5d and f) and telomeric nucleosomal arrays of up to
three subunits. Further, when MNase digestion products were run in
parallel for a longer time, the differences in nucleosomal band
positions between A-T and normal cells became apparent (Fig. 5g).
Normal cells have a telomere ladder containing digestions of up to
seven bands, whereas A-T cells have only three. These results suggest
that the telomeric nucleosome arrays in A-T cells might be less
uniformly spaced and extend over a smaller region than the arrays of
normal cells.
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FIG. 5.
(a to f) Comparison of nucleosomal periodicity of normal
(a and b) and A-T (c to f) fibroblasts. The MNase concentrations used
in each panel were 0 (lane 1), 125 (lane 2), 250 (lane 3), 500 (lane
4), 1,000 (lane 5), 2,000 (lane 6), 4,000 (lane 7), and 8,000 (lane 8)
U/ml. The position of the mononucleosomal band is indicated at the
right of each ethidium bromide-stained set of lanes. A-T cell
fibroblasts show more susceptibility to MNase digestion than normal
cells, as is evident by comparison of lanes 3 to 6 of each panel.
Panels b, d, and f are autoradiographs visualizing telomeric DNA as
detected by the TTAGGG probe. Two kinds of chromatin are
seen. Lane 8 in panel b represents normal fibroblasts, which have
telomeres with about seven nucleosomal subunits, and lanes 8 in panels
d and f represent A-T fibroblasts, which have telomeres with about
three nucleosomal subunits. (g) Comparison among two normal and one A-T
cell strain for the telomeric nucleosomal band positions after
treatment with MNase at 3,000 U/ml. The samples were run for a longer
time. Note the difference in telomeric nucleosomal subunits among
normal and A-T cells. The position of the mononucleosomal bands is
indicated at the right.
|
|
To determine how the altered nucleosomal periodicity in telomeres seen
in A-T cells responded to ionizing radiation, we examined
the influence
of radiation treatment on nucleosomal compaction
in telomeres. Normal
(C21F) fibroblasts in plateau phase were
irradiated with 5 Gy of gamma
rays, collected at 0 and 60 min
after treatment, and subjected to MNase
digestion. There was no
change in the appearance of nucleosomal bands
of bulk chromatin,
as revealed by ethidium bromide staining in normal
cells (data
not shown). Similar to bulk nucleosomes, there was no
change in
the nucleosome banding pattern of telomere in normal cells
(Fig.
6a). However we observed the disappearance of the nucleosomal
bands in the telomere region of A-T cells 1 h after gamma ray
treatment (Fig.
6b), whereas no such
change was seen in normal
cells (Fig.
6a). The disappearance of
nucleosomal bands in A-T
cells suggests that ATM influences the
response of telomere chromatin
to radiation.
View larger version (61K):
[in this window]
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|
FIG. 6.
Comparison of telomere nucleosomes of normal and A-T
fibroblasts after gamma irradiation. The autoradiographs visualize
telomeric DNA as detected by the TTAGGG probe. Cells were
irradiated with 5 Gy and collected after various time points. Chromatin
was digested with MNase (2,000 U/ml) for 5 min, and DNA was resolved on
2% agarose gel. The results are representative of three experiments
done separately. (a) Nucleosomal band in telomeric chromatin in normal
(AG1522) fibroblasts at different time points after irradiation. Note
there is no change in the nucleosomal band. (b) Nucleosomal band in
telomeric chromatin of A-T fibroblast (AG5823) at different time points
after irradiation. Note disappearance of the nucleosomal bands at 60 min postirradiation. The position of the mononucleosomal bands is
indicated at the right.
|
|
Telomere repeat binding factors.
To determine whether the
abnormalities in telomere-nuclear matrix interactions and nucleosomal
periodicity seen in A-T cells are correlated with alterations in
telomere binding factors, we first analyzed the expression of TRF1 and
TRF2 in A-T fibroblasts. Using the reverse transcription-PCR approach,
we found comparable levels of expression of TRF1 and TRF2 in A-T and
normal control cells (data not shown). Despite this finding, mutations
in these genes could lead to altered interactions of telomeres with the nuclear matrix. Therefore, we carried mutational analysis of TRF1 and
TRF2 genes in A-T and control cells. Analysis of TRF1 and TRF2 cDNA in
A-T cells by the cold SSCP protocol detected no mutations (data not shown).
To test whether TRF1 and TRF2 were localized correctly in the A-T
cells, we performed immunostaining of the cells and found
that both
proteins were localized in the nuclei of both A-T and
control cells
(data not shown). These observations suggest that
alterations in the
structure or expression of TRF1 and TRF2 are
not the cause of altered
interactions of the telomeres with the
nuclear matrix and nucleosomal
periodicity changes in A-T
cells.
In an attempt to identify gene products that might be involved with the
altered interactions of telomeres with the nuclear
matrix in A-T cells,
we used the Atlas cDNA microarray to analyze
the expression of genes.
The expression profiles of primary fibroblasts
of A-T and normal
control were compared by using poly(A)
+ RNA for
synthesizing
32P-labeled cDNA, subsequently hybridized
separately to array membranes
(data not shown). No significant
differences in the expression
of the 588 genes on the array were found
between A-T and normal
control
cells.
| |
DISCUSSION |
Cells derived from A-T individuals show a prominent chromatin
defect at chromosome ends in the form of chromosome end-to-end associations seen at G1, G2, and metaphase
(34, 63, 64). A-T cells also show an accelerated loss of
telomeres (49, 63, 64, 74, 82). Whether the chromosome end
associations are the cause or the effect of the accelerated loss of
telomeres is unclear. With the cloning of the gene for A-T, it has been
suggested that defective telomere maintenance could be due to the
ATM gene because of its homology to TEL1 mutants
of yeast and mei-41 mutants of Drosophila
(23, 25, 54, 68, 69). Since these genes have a
phosphatidylinositol 3'-kinase domain, the chromosome end association
defect could be due to a defective kinase activity. Recently, we
reported that the ATM gene influences chromosome end
associations as well as telomere length (74); however, it is
not clear how chromosome end associations are formed in cells derived
from A-T patients.
What other factors influence chromosome end association? One possible
factor is loss or shortening of telomeres, as suggested by Counter et
al. (15); another is altered chromatin structure. In our
previous studies, we reported that the frequency of cells with
chromosome end associations is higher in G1 phase than in G2 phase followed by metaphase, and for each phase of the
cell cycle, the frequency of cells with end associations was
significantly higher in A-T than in normal cells (63, 64,
74). It is probable that the end associations seen at mitosis
reflect a continuation of interphase chromosome behavior, perhaps
indicating interactions or linkages between chromosome ends and the
nuclear matrix. Since the telomeric signals are seen at the chromosome
end association sites (64), it is possible that in the
absence of ATM function, the chromosome end associations are the
consequences of the failure of the nuclear matrix with holding the
telomeres together. The telomeric signals at the chromosome end
association sites in A-T cells suggest that chromosome end associations
could be the primary event that subsequently lead to the shortening of
telomeres. This interpretation is consistent with the recent findings
of van Steensel et al. (79), who also reported that the
telomeric signals were present at sites of chromosome end associations
and that shortening of telomeres is not a prerequisite for chromosome
end associations. Telomeric signals at the chromosome end association
sites and changes in the frequency of cells with chromosome end
associations through the cell cycle raise the possibility that A-T
cells have an altered nuclear matrix, leading to defective interactions
between telomeric DNA and the nuclear matrix.
Telomeres are important components of chromosomes, as they have been
implicated in several cellular functions involved in aging and cancer
development. Telomeres have been shown cytologically as well as
biochemically to be tethered to the nuclear matrix. The nuclear matrix
is a proteinaceous scaffold in the interphase nucleus isolated by
removing most of the nuclear DNA and RNA, along with histones and
loosely bound proteins (6, 7). Our present study shows that
telomeres of primary fibroblasts are associated with the nuclear
matrix, and such observations are consistent with the previous
observations of de Lange (16) and Luderus et al.
(44). However, we found a significant difference in the
ratio of the P versus S fractions of telomeric DNA between A-T and
normal control cells. This difference could be attributed to
alterations in the interactions between telomeric DNA and the nuclear
matrix. Our studies demonstrate that changes in the lengths of the
telomeric DNA were not involved in these differences. The present
results are consistent with the hypothesis that the telomere nucleoprotein structure or nuclear matrix structure is different in A-T
cells. The fact that telomere binding to the matrix is greater in A-T
cells and is specifically influenced by irradiation shows that changes
in telomere-matrix association could be involved in the
chromosome-destabilizing function of the ATM gene. The role
of ATM function in telomere nuclear matrix interactions is further
strengthened by the fact that cells expressing dominant-negative fragment of the ATM gene have altered telomere nuclear
matrix interactions. The altered telomere nuclear matrix interactions seen in A-T cells were reversed by expression of the wild-type ATM gene. An influence of the ATM gene product on
the interactions of telomeres with the nuclear matrix might be an
important modulator of cellular processes influencing cellular
senescence and cellular transformation.
Genomic DNA is compacted within the nucleus as chromatin (nucleoprotein
complex), and the basic unit of chromatin is the nucleosome that
consists of 146 bp of DNA wrapped around an octamer of histones (35). The nucleosomal sizes in bulk as well as telomeric DNA are similar between A-T and normal cells. The differences lie in the
periodicity of the nucleosomes in the telomeric region in A-T versus
normal cells. This is further indicative of altered nuclear matrix
composition. Nucleosomes in telomeres of A-T cells are loosely spaced,
and this state of nucleosomal periodicity in telomeres could not be
attributed to the length of telomeres, as the telomeres of the
untransformed A-T and normal fibroblasts examined were similar in size.
Chromatin structure is an important factor in determining protein-DNA
interactions, with consequences for DNA metabolism and transcription
control (60, 67, 83). Since the nucleosomal model emerged,
there has been considerable progress in elucidating how chromatin
structure at the level of nucleosome organization can either repress or
potentiate transcription (2, 39, 84, 85). It has been
demonstrated that nuclear structure is very important for the
site-specific initiation of DNA replication (22). ATM is a
nuclear protein (11, 13, 20, 37) that also colocalizes with
chromatin associated proteins on meiotic chromosomes (30,
66). Recently, Gately et al. (20) have provided
biochemical evidence that ATM is associated with chromatin in somatic
cells. Our studies demonstrate that the altered telomere-nuclear matrix
interactions seen in A-T cells could be the reason for aberrant
radioresistent DNA synthesis in A-T cells. The genes involved in signal
transduction could influence chromatin structure, and that may explain
the basis of the cell cycle checkpoint defect in A-T cells (38,
56, 70) and the prevalence of chromosome damage. Since a
chromatin defect in A-T cells is pronounced at telomeres, their
interactions with the nuclear matrix may influence chromatin structure
and thus the function of neighboring genes.
The different response of A-T compared to normal cells after ionizing
radiation exposure could partly be attributed to altered telomere
chromatin organization, as is evident from the differences in
nucleosomal periodicity and nucleosomal compaction after ionizing radiation treatment. When nucleosomal compaction was examined in A-T
and normal fibroblasts after ionizing radiation exposure, bulk
chromatin did not show any distinct difference; however, nucleosomal
compaction was influenced in the telomeric region of A-T cells but not
in normal fibroblasts. An explanation for altered nucleosomal
compaction in A-T cells could be that the major portions of telomeric
DNA are attached to the nuclear matrix, whereas only a fraction of the
bulk DNA is associated with the matrix. Therefore, an altered nuclear
matrix could influence specifically the matrix-associated nucleosomes.
The results presented here suggest that the altered telomere chromatin
responds to DNA damage in a different way and thus influences the
nucleosomal compaction of telomeres only in A-T cells. Although it is
clear that ATM influences the interaction of telomeric DNA with the
nuclear matrix and nucleosomal periodicity, it is not clear how it does
so. Information about the interactions of the ATM gene with
other genes is limited. It has been shown that ATM interacts with
c-Abl, p53, and -adaptin (3, 5, 12, 32, 42, 71). However,
it remains to be established if such gene products can influence
telomere interactions with the nuclear matrix and chromosome stability.
| |
ACKNOWLEDGMENTS |
This work was supported by NIH grant NS34746.
Thanks are due to W. E. Wright, H. B. Liebermann, A. S. Balajee, C. R. Geard, W. N. Hittelman, and M. D. Story
for critical discussion of the manuscript. Thanks are also due to
S. G. Sawant, W. Mellado, and R. K. Pandita for technical assistance.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Center for
Radiological Research, College of Physicians & Surgeons, Columbia
University, VC11-213, 630 West 168th St., New York, NY 10032. Phone:
(212) 305-3911. Fax: (212) 305-3229. E-mail:
tkp1{at}columbia.edu.
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Molecular and Cellular Biology, October 1999, p. 6963-6971, Vol. 19, No. 10
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Copyright © 1999, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
