Section of Endocrinology, Diabetes, and Nutrition, Department of Medicine, Boston University School of Medicine, Boston, Massachusetts 02118
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ABSTRACT |
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Fish oils (FOs) have been noted
to reduce growth and proliferation of certain tumor cells, effects
usually attributed to the content of polyunsaturated fatty acids of the
n-3 family, which are thought to modulate cellular signaling
pathways. We investigated the influence of FO on cell cycle
kinetics of cultured Chinese hamster ovary cells. Exponentially growing
cells were labeled with 5-bromo-2'-deoxyuridine (BrdU) and analyzed by
flow cytometry after 5-day treatment with exogenous fat. Bivariate
BrdU-DNA analysis indicated slower progression through S phase and thus
longer S phase duration time in FO- but not corn oil-treated or control cells. We hypothesize that FO treatment might interfere with
spatial/temporal organization of replication origins. Therefore, we
mapped the well-characterized replication origin ori- downstream of
the dihydrofolate reductase gene with the nascent strand length assay. Three DNA marker segments with known positions relative to this origin
were amplified by PCR. By quantitatively assessing DNA length of the
fragments in all fractions containing these markers, the location of
ori-
was established. In control or corn oil-treated cells, the
location of ori-
was consistent with previous studies. However, in
FO-treated cells, DNA replication appears to start from a new site
located farther upstream from ori-
, suggesting a different
replication initiation pattern. This study suggests novel mechanism(s)
by which fats affect cell proliferation and DNA replication in
mammalian cells.
cell cycle kinetics; n-3 fatty acids; dietary fat; cancer; deoxyribonucleic acid replication; Chinese hamster ovary cells; dihydrofolate reductase
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INTRODUCTION |
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THE RELATIONSHIP between dietary fat and cancer remains controversial. Although epidemiological studies previously suggested that high fat intake is positively associated with certain cancers such as colon (50, 105), prostate (37, 41, 61) and breast (39, 40, 81) cancers, recent studies have cast doubt on this relationship (27, 35, 43, 53). Confounding factors and alternative explanations have been proposed to account for the apparent association between dietary fat and cancer (103, 104). Furthermore, prospective studies to reduce dietary fat intake have also produced conflicting results, especially in regard to the incidence of breast cancer in women (13, 24, 45, 93).
On the other hand, studies in experimental animals have been more consistent in showing a positive association between dietary fat and cancer (19, 20, 42, 79). Fats derived from marine animals (fish oil, FO) have been especially investigated. These oils are enriched with polyunsaturated fatty acids (PUFAs) of the n-3 family and are believed to have protective effects at the initiation and promotion stages of carcinogenesis (74, 75). Several investigators, including our group, have also demonstrated in animal feeding studies that diets enriched with FO can reduce the growth rates of implanted tumors (54, 57, 58). These protective effects of FO are generally attributed to its n-3 PUFA content through several hypothetical mechanisms, such as modification of membrane lipids and subsequent changes in signaling pathways, membrane fluidity, prostaglandin synthesis, and ornithine decarboxylase activity (6, 7, 32, 38, 48, 72, 94).
To further delineate the molecular mechanisms that mediate the effect
of FO on cell functions, studies have used cell culture techniques in
which either FO or individual fatty acids are added to the growth media
(6, 16, 26, 48). Recently, Collett et al.
(21) demonstrated differential effects for the n-3 and n-6 PUFAs docosahexaenoic acid (DHA) and linoleic acid (LA),
respectively, on oncogenic Ras activation in cultured adult mouse colon
(YAMC-ras) cells. Other studies in cultured human breast cancer cells
also demonstrated differential effects for n-3 and n-6 PUFAs on the activity of the nuclear receptor peroxisome proliferator-activated receptor (PPAR)- and related these effects to cell proliferation (59, 80). By showing close similarity to in vivo feeding
experiments, these studies illustrate the validity of cell cultures in
evaluating possible mechanisms by which dietary fats influence
carcinogenesis and cell proliferation.
In previous experiments with tumor-bearing rats, we showed (54) that feeding with diets enriched with FO reduces the rate of tumor growth, consistent with findings of several other investigators (57, 58). We characterized the cell cycle kinetics of this tumor model in FO-fed and control rats by use of 5-bromo-2'-deoxyuridine (BrdU) pulse labeling and flow cytometric analysis. The most noticeable result from that study was a delay in S phase completion, which accounted for a slower rate of cell cycle progression, in the cells from FO-fed rats. In particular, although the fraction of cells estimated to enter the S phase was unaffected, the duration of S phase significantly increased after 6 wk of feeding with a diet enriched with FO. Because S phase progression is independent of extracellular regulation of the cell cycle, which is mostly achieved at the level of G1 restriction point (2, 47), we argued that this effect of FO is distinct from its putative modulatory effects on cell surface signaling such as those of the mitogen-activated protein (MAP) kinase pathways (6, 23, 32, 70). Thus we questioned whether FO feeding could alter the DNA replication machinery, in particular the temporal/spatial organization of DNA replication origins that is the primary determinant of S phase duration (11, 22, 30, 83).
Therefore, to address this possibility, we initially tested the
feasibility of reproducing a similar effect on S phase progression in a
cell culture system. We chose the Chinese hamster ovary (CHO) cell line
in view of the existing knowledge of DNA replication origins in this
cell line. Here we characterize the cell cycle kinetics in
exponentially growing CHO cells that have been treated with FO. After 5 days of treatment, FO significantly lengthens the duration of S phase
of exponentially growing CHO cells, a finding consistent with our
previous in vivo feeding studies (54). To determine
whether this change in S phase progression is associated with altered
firing at the level of DNA replication origins, we used the nascent DNA
strand length assay to map the replication origin ori- of the
dihydrofolate reductase (DHFR) gene locus. The region downstream from
the DHFR gene in CHO cells is the most thoroughly mapped high-frequency
initiation region in mammalian chromosomes. We show that in control and
corn oil (CO)-treated CHO cells ori-
activity localizes to the
expected chromosomal site ~17 kb downstream from the DHFR gene. On
the other hand, FO-treated CHO cells, which are characterized by a
longer S phase duration, exhibit an altered location of this
replication origin. These results suggest that FO, and possibly n-3
PUFAs, may affect cell proliferation at the level of organization of
replication origins. Although the mechanism of such an effect is
not yet understood, the possibility that FO can influence the
chromosomal location and/or activity of replication origins in
mammalian cells represents a novel finding. Understanding of the
interaction between PUFAs and the DNA replication machinery will
further clarify the potential role of dietary fats in cell
proliferation and cancer.
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MATERIALS AND METHODS |
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Cell Culture and Oil Treatment
Chinese hamster ovary cells (CHO-K1) were obtained from American Type Culture Collection (Rockville, MD) and routinely grown in 5% CO2 at 37°C in Dulbecco's modified Eagle's medium supplemented with penicillin (100 U/ml), streptomycin (100 µg/ml), and 10% fetal calf serum. As described in detail elsewhere (36), FO and CO were emulsified with 5% (wt/wt) egg phosphatidylcholine (Sigma) and 0.03% (wt/wt) butylated hydroxytoluene (as antioxidant) in phosphate-buffered saline (PBS) at a final oil concentration of 15 mg/ml. For treatment with extrinsic fat, cells were plated in regular growth medium for 24 h, after which the medium was replaced with fresh oil-enriched medium at a final oil concentration of 50 µg/ml. The control dish received egg phosphatidylcholine alone at an equal volume. The oil-enriched medium was changed daily to decrease the reactions of fatty acid oxidation on cells. After 5 days in culture, cells were harvested by trypsinization and reseeded into 100-mm cell culture dishes, and the oil treatment was continued. In all subsequent experiments, exponentially growing control and oil-treated cells were harvested 24 h after reseeding.Experimental Part I: Cell Cycle Kinetics
Growth characteristics.
Estimates of the actual growth rates of the control and oil-treated CHO
cells were determined from changes in cell number as function of time.
After 5-day oil treatment, cells were reseeded and incubated at a
concentration of 5 × 105 per plate. For three
consecutive days, the cells were sampled and counted each day by use of
a hemocytometer and Trypan blue (GIBCO). The first-order growth rate
constant (Kg) was determined by regression
analysis according to the equation
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BrdU pulse labeling and staining procedure for kinetic analysis
of cell cycle.
To exponentially growing cells in oil-enriched and control media, we
added BrdU (Boehringer Mannheim) at a final concentration of 25 µM.
After a labeling pulse of 30 min at 37°C, cells were washed with
identical fresh medium and then incubated in the corresponding medium
for 1, 3, and 5 h, respectively. To obtain a good average, six
plates were prepared for each time point. After the indicated BrdU-free
chase time, cells were harvested, fixed in 70% cold ethanol, and
stored at 20°C until further processing. The staining procedure to
determine the DNA and BrdU content was described previously
(96). Briefly, ~1-2 × 106 fixed
cells were washed and resuspended in 2 ml of PBS and then incubated in
the dark with 2 ml of 4 N HCl containing 0.5% (vol/vol) Triton X-100
(Sigma) for 30 min at room temperature. This procedure produces
single-stranded DNA molecules, improving the staining with anti-BrdU
antibodies. Subsequently, cells were centrifuged at 500 g
for 5 min and neutralized with 2 ml of 0.1 M sodium tetraborate (pH
8.5). Cells were washed twice with PBS containing 0.5% (vol/vol) Tween
20. After pelleting, cells were resuspended in 50 µl of PBS-Tween 20 and incubated in the dark with 20 µl of fluorescein isothiocyanate
(FITC)-labeled monoclonal anti-BrdU antibodies (Becton-Dickinson,
Mountain View, CA) for 30 min at room temperature. For staining of
total DNA, cells were then washed twice with PBS-Tween 20 and
resuspended in 1 ml of PBS containing 10 µg of propidium iodide
(Calbiochem, San Diego, CA). Samples were ready for bivariate DNA-BrdU
flow cytometric analysis after 15 min.
Flow cytometry. Double-stained cells were analyzed with a FACScan flow cytometer (Becton-Dickinson) at an excitation wavelength of 488 nm and a laser power of 15 mW. Red fluorescence from propidium iodide was collected through a 585-nm band-pass filter as total DNA content. Green fluorescence from FITC-labeled anti-BrdU antibodies was collected through a 530-nm band-pass filter. The red fluorescence was calibrated by adjustment of the G0/G1 peak to a fixed channel number, and the green fluorescence was calibrated by use of FITC-labeled latex beads (Polysciences, Warrington, PA). A total of 105 cells were monitored. Flow cytometric data were accumulated at the highest resolution for each parameter (1,024 channels).
Kinetic analysis of flow cytometric data. Two separate procedures were used to determine the kinetic characteristics of exponentially growing CHO cells. In the first procedure, double-stained cells were subjected to an univariate DNA analysis (red fluorescence) only. The distribution of cells in each phase of the cell cycle was determined by mathematical analysis of the DNA histograms (see, e.g., Fig. 2) with the computer program ModFit (Verity Software House, Topsham, ME).
In the second procedure, cell proliferation kinetic parameters were derived from BrdU pulse labeling and bivariate BrdU-DNA analysis by flow cytometry as previously described by White and colleagues in several publications (88, 98-102). Briefly, bivariate BrdU-DNA contour plots were obtained with IsoContour software (Verity Software House) and the cells were first separated according to their DNA content (x-axis) and their BrdU uptake (y-axis). The BrdU-labeled cells were further separated into labeled undivided (flu) and labeled divided (fld) subgroups, respectively (see Fig. 3), and the function
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Experimental Part II: Mapping of DHFR Replication Origin ori-
in Treated CHO Cells
Cell labeling and isolation of nascent DNA. Control and oil-treated, exponentially growing CHO cells were labeled with [3H]deoxycytidine (3H-dC, 10 µCi/ml) in the presence of 50 µM BrdU for 15 min at 37°C (91). The cells were washed three times with cold PBS and lysed in 7 ml of 0.5% sodium dodecyl sulfate, 1 M NaCl, 10 mM EDTA, and 50 mM Tris · HCl (pH 8.0). After incubation with proteinase K (200 µg/ml), DNA was isolated by phenol-chloroform (1:1) extraction and spooling on glass rods under 70% ethanol. The spooled DNA was rinsed in 95% ethanol.
Size fractionation of DNA by gradient centrifugation. The labeled and isolated DNA was dissolved in 4 ml of TE buffer [10 mM Tris · HCl (pH 8.0), 1 mM EDTA] by reversing the direction of spooling. NaOH (1 N) was added to give a final concentration of 0.2 N NaOH. The DNA was fractionated by sedimentation through a 5-20% (wt/vol) linear sucrose gradient containing 0.2 N NaOH and 2 mM EDTA for 18 h at 15°C in a Beckman SW28 rotor at 24,000 rpm. Gradients were collected in 12 fractions, and portions were taken to assay for 3H radioactivity. Fractions 2-11 were each neutralized with 2 N HCl in the presence of 0.1 M Tris · HCl (pH 7.5) and precipitated with ethanol. Fractions 1 and 12 were discarded. For convenience purposes fractions 2-11 were renumbered as fractions 1-10.
Immunoprecipitation of nascent DNA chains. BrdU-containing nascent DNA strands were purified by two rounds of immunoprecipitation with anti-BrdU monoclonal antibodies (90, 91). Briefly, DNA from each fraction was dissolved in 0.5 ml of TE buffer and denatured for 3 min at 95°C. After rapid cooling with an ice bath, each fraction was adjusted to 10 mM sodium phosphate (pH 7.0), 0.14 M NaCl, and 0.05% Triton X-100 (immunoprecipitation buffer) and incubated with 80 µl of mouse anti-BrdU monoclonal antibody (Becton-Dickinson) for 20 min at room temperature with slow agitation. Rabbit anti-mouse IgG (80 µl; 25 µg/ml) was then added to precipitate BrdU-DNA-antibody complexes. Immunoprecipitates were collected by centrifugation for 5 min and washed once with 0.5 ml of immunoprecipitation buffer. Pellets were deproteinized by overnight incubation at 37°C in 200 µl of 50 mM Tris · HCl (pH 8.0), 10 mM EDTA, and 0.5% sodium dodecyl sulfate containing 250 µg/ml proteinase K, followed by phenol-chloroform extraction (1:1) and ethanol precipitation. DNA pellets were dissolved in 100 µl of TE buffer, subjected to a second round of immunoprecipitation, and then precipitated with ethanol in the presence of 20 µg of Escherichia coli tRNA (Sigma). These nascent DNA fractions were dissolved in 50 µl of TE buffer and used for PCR amplification.
PCR amplification conditions. Oligonucleotide primers and probes for the marker segments A, B, and C, homologous to the region of the DHFR replication origin in CHO cells (15), were chemically synthesized (Baron Biotech, Milford, CT). Nucleotide positions of each primer pair downstream of the XbaI restriction site were as follows (15, 91): segment A, 5': 81 to 100 (biotin-5'-GTG CTA GAA GTA GAT GAG AG-3'), 3': 381 to 400 (5'-AAT CCA GCA TGC GAA CAG TT-3'); segment B, 5': 2677 to 2696 (biotin-5'-TTC TCA GTG AGT CCA CTT CT-3'), 3': 2976 to 2995 (5'-CCT GGT AGG GAC TTC AGA AA-3'); segment C, 5': 4569 to 4588 (biotin-5'-AGT ATT GTA GGT ATG TGC CC-3'), 3': 4955 to 4974 (5'-GTT GTG CTT TAG TGA TAG GG-3'). PCR amplification was performed at a final concentration of 1× PCR buffer containing 50 µM dNTPs, each 5' and 3' primer at 0.1 µM, and 1 unit of AmpliTaq DNA polymerase (Perkin-Elmer, Norwalk, CT) in a total volume of 50 µl. All amplification and hybridization reactions were performed in a DNA Thermal Cycler 480 (Perkin-Elmer). The amplification temperature profile involved a denaturation step at 95°C for 0.5 min, a primer-annealing step at 58°C for 0.5 min, and an extension step at 72°C for 1 min and 30 cycles of amplification.
Quantification of PCR products by electrochemiluminescence-based detection system. An automated QPCR 5000 system (Perkin-Elmer) was used to quantify the amplified PCR products. This system is designed to detect electrochemiluminescence (ECL) generated by the reaction between tris(2,2'-bipyridine)ruthenium(II)-chelate (TBR) and tripropylamine (TPA). TBR is attached to the PCR probe, whereas the reactant TPA is present in the QPCR assay buffer. Typically, 5 µl of PCR product was mixed with 35 µl of PCR buffer [10 mM Tris · HCl (pH 8.3), 50 mM KCl] and 10 µl of 1 pmol/µl TBR-labeled probe. For hybridization, the amplified PCR products were denatured at 95°C for 3 min and hybridized with the probe at 60°C for 5 min. Twenty microliters of 2 mg/ml streptavidin-coated magnetic beads, which bind the biotin-labeled PCR sequences, was then added to separate the ECL-signal quantitatively. After 20 min at 55°C, the products of each hybridization reaction were transferred to a separate QPCR System 5000 tube containing TPA in QPCR assay buffer to measure the ECL signal. The signal measured by this system corresponds proportionally to the amount of hybridized PCR product. The TBR probes used in the hybridization reaction are as follows: segment A, 145 to 165 (TBR-5'-CTA CAA TCC TTC CTC TCT CTT-3'); segment B, 2787 to 2807 (TBR-5'-GCT GAA CTT TAT CAG TGC AGT-3'); segment C, 4643 to 4662 (TBR-5'-TG TGT AGC ACA GTC TAG TCT-3').
Statistical Analysis
Whenever applicable, data are reported as means ± SE. Differences were initially evaluated by one-way analysis of variance (ANOVA) to determine the overall statistical significance of treatment. Differences between means were subsequently evaluated by a standard t-test only whenever ANOVA showed a statistically significant difference at PFor analysis of the effect of FO on ori- mapping characteristics,
95% confidence intervals for relevant ECL signal ratios were
calculated from untreated (control) cells, based on a replicate number
(n) of 4. Subsequently, the corresponding range of
experimental values in the treated cells was evaluated in reference to
the 95% confidence interval thus defined. For statistical evaluation of the overall significance of the effect of FO on ori-
mapping, the
null hypothesis that differences in the ECL ratios among treatment groups were due to chance was tested by
2-analysis. In
this analysis, two possible outcomes were considered, i.e., whether a
specific FO-treated ECL ratio was similar to or different from the
expected ratio defined by the data from untreated cells. All
statistical analyses were performed on a desktop computer with the
Microsoft Excel 97 software package.
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RESULTS |
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FO Prolongs S Phase Duration in Exponentially Growing CHO Cells
Initial experiments established the necessary conditions for CHO cells to maintain exponential growth characteristics after treatment with oil-enriched growth media. CHO cells grew exponentially after the 5-day oil treatment and subsequent reseeding. Figure 1 shows the average number of cells in each treatment group measured on three consecutive days after reseeding in the corresponding medium. On day 2, significantly fewer cells were counted in the FO treatment group compared with either control or CO-treated cells (P < 0.05 for FO vs. either control or CO). Estimates of the first-order growth rate constant Kg were derived from the exponential part of the growth curve by regression analysis with Eq. 1 and expressed as actual doubling time Td as defined by Eq. 2 (see Table 3 for numeric values of Td). From the analysis of the growth curves, cells treated with FO proliferated at a reduced rate with a slightly longer doubling time (32.7 h) compared with control cells (29.8 h) and CO-treated cells (30.6 h). Thus FO treatment reduced the overall proliferation rate of CHO cells by ~10%.
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For estimation of the cell cycle kinetics, we first analyzed DNA
histograms in each treatment group. The one-dimensional histograms represent the total DNA stained by propidium iodide and allow determination of the distribution of cells in each phase of the cell
cycle. In Fig. 2 we show representative
DNA histograms for control (Fig. 2A), CO-treated (Fig.
2B) and FO-treated cells (Fig. 2C). For each
group, the percentages of cells in G0/G1, S,
and G2/M phase are indicated as obtained by the software
program ModFit. The percentages for each cell population averaged over
six independent experiments are summarized in Table
1. The 5-day treatment with FO or CO did
not disturb the diploid nature of the CHO cell line. In addition, flow
cytometric analysis showed no evidence for cell death or
apoptosis. Cells treated with FO had ~5-6% more cells in S phase (P < 0.005) and slightly fewer cells in
G1 phase compared with controls and CO-treated cells.
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To determine kinetic parameters of the cell cycle, we analyzed cells of
the three treatment groups by bivariate flow cytometry. The cells were
stained for total DNA content (with propidium iodide) and for BrdU
content (with FITC-BrdU antibodies). For kinetic analysis, cells were
harvested and fixed at three different time points after a BrdU pulse
for 30 min. Figure 3 represents a typical example of bivariate BrdU-DNA contour plots recorded at the earliest (Fig. 3A) and the last (Fig. 3B) time point. In
Fig. 3A we show that populations of cells in
G0/G1 and G2/M phase are separated according to their DNA content and cells in S phase are distinguished according to their BrdU content at 1 h after pulse labeling. As displayed in Fig. 3B, BrdU-labeled cells have separated into
labeled, divided (fld) and labeled, undivided
(flu) fractions after 5 h. The percentage of cells in
each fraction was determined by use of the software program IsoContour
and is summarized in Table 2. At the 5-h
time point, significantly fewer FO-treated cells had divided compared
with either control or CO-treated cells. In the FO group only 12% of
cells had completed cell division by that time and appeared in the
G1 phase population, compared with 20% and 16% in control
and CO-treated groups, respectively (P < 0.05).
Because each division of labeled cell results in two labeled daughter
cells, the contribution of divided and undivided fractions to the total
number of labeled cells is expressed by the function (Eq. 3). This function characterizes the fraction of cells traversing a
single S phase and is directly proportional to the rate of
G1/S phase transition. Numeric values of
are listed in
Table 2. In each of the treatment groups, ~28% of exponentially growing CHO cells underwent DNA replication during the S phase. This
finding implies that the rate of G1-to-S phase transition in the FO-treated group was similar to that of either control or
CO-treated cells.
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Further analysis of BrdU-DNA plots allows estimation of the S phase
duration time TS as well as the mean times spent
in G1, G2/M, and the total cell cycle. Two
analytical methods are presented, both of which are derived from RM
measurements, as defined by Eq. 6. The results of relative
movement at 1, 3, and 5 h after BrdU pulse labeling are summarized
in Fig. 4. One hour after BrdU labeling,
the RM values were significantly higher in FO-treated cells
(P < 0.01 for FO vs. either control or CO). However,
in cells examined 5 h after pulsing, FO-treated cells had
significantly lower values of RM compared with the other treatment
groups (P < 0.005 for FO vs. either control or CO).
Thus the slope of the linear regression function RM(t) was
significantly smaller after FO treatment as shown in Fig. 4. The actual
numeric values of RM are given in Table 2 for the 5 h time point
[RM(5), Eq. 6] and for the initial time point
t = 0 (RMo, Eq. 8).
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The results of determining the S phase duration times
TS, which are based on the linear fit of
RM(5) and RMo, as summarized in Table
3, show that the S phase duration time
was 8.2 ± 0.2 h for FO-treated cells, which is significantly
longer compared with 6.7 ± 0.2 h and 6.6 ± 0.2 h
for control and CO-treated cells, respectively (P < 0.005 for FO vs. either control or CO). Table 3 also summarizes the
results of TS derived from the cubic fit of the
5-h BrdU/DNA data according to Eqs. 9-11. It is noted
that estimates of TS derived from this method
were consistently smaller than those obtained from the linear model.
However, the difference between treatment groups persisted, with
FO-treated cells having significantly longer S phase duration times.
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Knowing the S phase duration time TS, which is
equivalent to the DNA replication time, we calculated the potential
doubling time Tpot for the three treatment
groups according to Eqs. 4 and 5. Corresponding
numeric values are listed in Table 3, which also contains the estimates
of the total cell cycle time (Tc), the mean
times spent in G1 and G2/M phase, and the
actual doubling time (Td) derived from the
growth curves (Fig. 1). These results show that FO treatment, in
addition to the effect on S phase, also significantly increased the
mean time spent in G2/M phase. On the other hand, estimates
of the total cell cycle time and potential doubling time, derived from
the cubic fit model, were more variable within each treatment group and
therefore did not achieve statistical significance between the
treatment groups. We also note that the estimates of total cell cycle
time were in close agreement with and almost equivalent to the
corresponding estimates of potential doubling time. This finding
indicates that all the cells were cycling (growth fraction equal to 1),
with no significant increases in the nonproliferating cell pool
resulting from oil treatments. Furthermore, estimates of the actual
doubling times were longer than those of the potential doubling times, indicating significant cell loss from this CHO cell line. However, comparison of the ratio of Tpot to
Td was similar in all the treatment groups. Thus
the degree of cell loss, which amounted to 43-50% under the
conditions of this experiment, was not affected by the oil treatment,
as noted by the parameter in Table 3 (P = not significant). On the basis of these results, we deduce that most of the
difference in the observed proliferation potential in FO-treated cells
compared with the control and CO-treated cells is accounted for by the
longer S phase duration. Factors that reduce the proliferation potential by halting cells at the G1 restriction point,
thus reducing the growth fraction, or increasing the rate of cell death
do not seem to account for a significant part of the effect of FO in this kinetic model.
We conclude from the kinetic cell cycle studies that FO significantly slows the progression of proliferating cells through the S phase, and possibly through mitosis. However, the net effect on actual cellular growth is smaller and more difficult to measure. This apparent discrepancy between the effect on S phase and total growth may be explained by compensatory changes in other cell cycle regulatory mechanisms. These possibilities were not addressed by the current study.
Effect of FO on Replication Origin ori-
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Preliminary experiments were initially performed and repeated with
untreated (control) CHO cells to determine the reproducibility of the
fractionation and PCR conditions for delineating the putative ori-
replication origin, as previously documented in exponentially growing
CHO cells (91). Subsequently, two independent experiments, consisting of three treatment groups each, were performed. The relative
intensities of the ECL signal generated by each of the three hybridized
probes, expressed as the average value from the two experiments where
all three treatments were conducted simultaneously, are presented in
Fig. 6 as a function of fraction number.
In the control and CO-treated cells, segment B continues to
be represented in the smallest DNA fractions (highest fraction numbers
in Fig. 6B), whereas segments A and C
are diminished significantly in lighter DNA fractions, starting
noticeably in fraction 7 (Fig. 6, A and
C). These results are very similar to those reported previously by Vassilev et al. (91). It is noted that
segment C, which also diminishes starting in fraction
7, is undetectable in fractions 9 and 10. In
contrast to control and CO-treated cells, cells treated with FO
continue to exhibit significant amounts of segment A even in
the highest fraction numbers containing the shortest DNA fragments
(Fig. 6A). Segment B decreases significantly in
fractions 9 and 10 in the FO-treated cells (Fig.
6B), whereas segment C behaves similarly in
FO-treated cells as well as in the two control groups (Fig.
6C). These results indicate that in the chromosome region of
control and CO-treated cells, DNA replication is initiated closest to
segment B, i.e., closest to the replication origin ori-
as expected from previous studies (60, 91). In FO-treated
cells, however, this activity is suppressed. Instead, the initiation of
DNA replication in the FO-treated cells appears to favor a location
closer to segment A.
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To approximate the location of a replication origin in reference to a
marker, one determines the smallest length of nascent DNA that includes
the marker in question. In the control cells, where the replication
origin appears to lie closer to segment B, this is best
illustrated by examining the ratio of B to A. Because longer segments of nascent DNA contain both markers, the ratio
B/A is expected to be ~1.0 in fractions
containing larger fragments (low fraction numbers). For smaller nascent
DNA fragments, the marker farther away from the replication origin has
not been replicated yet, so that no signal can be detected and thus,
theoretically, the ratio tends to go to infinity. In actual data,
however, DNA fractions contain a range of lengths and therefore, the
change in fragment ratios is more gradual. Vassilev et al.
(91) previously showed that the transition in
hybridization ratios (equivalent to ECL signal in Fig. 6) is likely to
occur at values between 1.0 and 3.0. In Fig.
7A, we show the experimental
values of the ratio B/A derived for the three
treatment groups. In the control and CO groups,
B/A achieves a value >3.0 in fraction
7 (control) and fraction 8 (CO), respectively. In the
FO group, B/A varies between 1.0 and 0.1, with
A/B achieving a value of 3 in fraction 9 (see Fig. 7B). Plots of the ratios
C/A (Fig. 7C) and
C/B (Fig. 7D) show variations between
1.0 and 10
4 in each of three treatment groups because the
ECL signal for segment C is very similar for all groups and
is almost undetectable for the smallest DNA fragments. ECL signal
strength ratios A/C and B/C
are also presented in Fig. 7. As noted above and in Fig. 6C, segment C did not amplify in fractions
9 and 10, a finding that could result either from the
distance of this marker relative to the actual initiation point or from
reduced PCR efficiency for this segment. Therefore, values of the
ratios A/C and B/C in
fractions 9 and 10 are inappropriate because
segment C was not quantitatively different from 0. However,
despite this limitation, we note that B/C shows
an increase from 1.0 to 3-4 in fractions 7 and
8 in the CO and control cells, but not in the FO-treated cells. On the other hand, A/C tends to increase
in fraction 8 in FO cells but remains equivalent to 1 in the
other two groups.
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It is noteworthy that differences in ECL signal strength ratios in the
FO group compared with the other two groups were significantly larger
in fractions 9 and 10, with a coefficient of
variation in these measurements ranging between 10 and 25%. In
fractions 7 and 8, however, differences related
to the FO treatment were more subtle. As noted in
DISCUSSION, the actual size of nascent DNA may have a
significant impact on the relative importance of these fractions in
origin mapping. In the absence of experimental information about DNA
fraction sizes in the current study, it is important to further
evaluate the smaller differences in ECL ratios noted in fractions
7 and 8. In the control cells, four separate sets of
ECL measurements were available (2 from preliminary experiments in
untreated exponentially growing CHO cells and 2 in the control groups
of the main study). Thus it is possible to define 95% confidence
intervals for specific ECL ratios in untreated CHO cells
(n = 4). The results of this analysis are presented in
Fig. 8 (95% confidence intervals
depicted in boxes) for the key ratios B/A,
B/C, and A/C in
fractions 7 and 8. The range of experimental
values obtained in the corresponding fractions in CO- and FO-treated
cells, as well as reference values from Vassilev et al.
(91), are also depicted in Fig. 8. We note that the 95%
confidence ranges for B/A and
B/C in fractions 7 and 8 from untreated CHO cells were similar in the current study to the
previously reported values (91). It is also noted in Fig. 8 that, in the FO-treated cells, measured B/A and
B/C ratios were consistently excluded from the
95% confidence intervals defined by data from the control cells. In
contrast, the corresponding values from the CO-treated cells overlapped
with these hybridization ratio ranges. Figure 8 also shows that
A/C from FO-treated cells, but not those from
CO-treated cells, were significantly outside the 95% confidence range
of untreated cells in fraction 8. These results, together
with the larger differences already noted in fractions 9 and
10 (Fig. 7), imply a possible difference in the initiation
of DNA replication in FO-treated CHO cells in reference to the expected
location of ori-.
|
Together, the results presented in Figs. 6-8 show a consistent
pattern of hybridization ratios for the FO group in each of the two
independent experiments where all three treatments were carried out
simultaneously. On the other hand, CO and control cells were consistent
with the expected location of ori- in both experiments. These
findings led us to propose that the DHFR replication origin ori-
,
which is normally in close proximity to marker segment B, is
suppressed in FO-treated cells (P = 0.083, by
2-analysis based on differences in a single ECL ratio).
Instead, the initiation site of DNA replication near segment
B in FO-treated cells appears to be closer to marker segment
A, which is located upstream from segment B (see Fig.
5). However, it is important to note that in view of the limitations in
the mapping method used here, especially when ori-
is no longer in
proximity to segment B (see below), further experiments with
more detailed mapping markers are needed to confirm the results of this study.
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DISCUSSION |
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This study primarily addresses the mechanism by which FO modulates
cell proliferation. Sufficient evidence in the literature indicates
that FO, possibly because of its high content of n-3 PUFAs, has a
protective effect against carcinogenesis. The potential mechanisms that
account for this effect are numerous and have been addressed by many
studies in the past 10-15 years. Here, we have focused on the S
phase in view of previous in vivo animal experiments in our laboratory,
which have shown that FO feeding lengthens the DNA replication time in
a breast cancer tumor model. This is an unexpected result because the
cell cycle is primarily regulated at specific checkpoints that control
exit and entry of cells from one phase of the cell division cycle into
the next phase (47, 67). Progression through S phase is
generally regarded to be independent of extrinsic regulation
(52). We previously hypothesized that changes in the S
phase duration can be explained by alteration in the spatial and/or
temporal organization of replication origins. However, these
replication origins, which are known to be DNA sequence specific in
yeast and lower eukaryotic cells, are less well understood in mammalian
cells. Furthermore, because of experimental difficulties in higher
eukaryotic cells, it was important to address the possibility of an
effect on the DNA replication machinery in a system in which
replication origins have already been characterized. Thus we have
chosen the CHO cell line, in which the replication origin ori- has
been described in detail by several investigators using various techniques.
Our initial approach to this problem was to examine the cell cycle kinetics of the CHO cell line to determine the feasibility of reproducing an effect on S phase progression similar to that previously observed in animal feeding studies. Measurement of BrdU uptake in newly synthesized DNA and the rate of progression of this label in S phase allow estimation of the DNA replication time and the generation rate of new cells (defined by Tpot) according to the analytical methods of White and his group (88, 97, 102). In the current study, we report estimates of the DNA synthesis and potential doubling times derived from two separate analytical methods. As noted by White et al. (98), the linear RM(t) model tends to overestimate these parameters for sampling times close to TS, whereas the cubic fit method is more accurate under these conditions. Data summarized in Table 3 are consistent with these expectations, showing consistently smaller TS values in the cubic fit model. However, both techniques point to the lengthening of S phase duration as the main cell cycle kinetic effect of FO treatment. We note that despite theoretical simplifications, these methods provide significant information about cell cycle regulatory sites, as demonstrated by the current study. Such information would not be available from cell cycle distribution percentages as usually achieved from simple DNA histograms. For example, the slight increase in percentage of cells in S phase in FO-treated cells could possibly be explained by either an increase in the actual number of proliferating cells or an increase in S phase duration relative to the total cell cycle time. Use of bivariate BrdU-DNA analysis in the current study points toward the latter explanation, thus suggesting a reduction in the proliferation potential with FO treatment. The alternative possibility that an increased S phase population implies a larger growth fraction is inconsistent with the finding of reduced growth rate in the FO-treated cells.
Proliferating cells respond to extracellular factors through an intricate network of signal transduction pathways that start at receptor sites within the plasma membrane. In mammalian cells, growth factor stimulation triggers a cascade of events that lead to the expression of early-response genes, many of which are transcription factors for delayed-response genes, such as cyclins D and E, cdks 2, 4, and 6, and E2Fs. Through regulated protein phosphorylation/dephosphorylation events, cdk/cyclin D and cdk/cyclin E complexes control the passage of cells across the G1 restriction point and activate the transcription of S phase cyclins and subsequent degradation of the S phase inhibitor. The latter event allows the S phase cdk/cyclin complexes to phosphorylate the regulatory sites of DNA prereplication origins, which are already assembled on replication origins during G1. Therefore, the net regulatory event arising from the extracellular environment is a decision about the transition of G1 cells into DNA synthesizing S phase cells (22, 95). Within the S phase, checkpoint regulation of DNA replication is primarily a protective mechanism that averts the replication of damaged DNA (1).
We examined the effects of FO on the proliferation kinetics of CHO cells to determine whether fats with a high n-3 fatty acid content can actually inhibit growth, as has been previously claimed (38, 74, 76, 82). Dietary fats have long been suspected to be involved in cancer causation (18, 34, 49, 84) and cellular growth (26, 38), although controversy still prevails (34, 84, 103). Because lipid molecules are important components of cell membranes and signaling pathways, proposals have been made about possible mechanisms by which dietary fats alter cellular function (6, 9, 56, 68, 86, 87). The majority of the proposed mechanisms focus on the actions of phospholipase C and second messengers diacylglycerol and inositol trisphosphate. Activation of this system initiates a series of events that lead to the activation of protein kinases, release of intracellular calcium, and subsequently activation of MAP kinase and its translocation into the nucleus (6, 32, 64, 73). Inside the nucleus, an active MAP kinase induces the transcription of early growth-response genes (c-fos, c-jun, c-myc), which are transcription factors necessary for the expression of G1 cyclins (17, 25, 26, 73), as noted above. Thus, at least theoretically, dietary fats have the ability to modulate cell proliferation inasmuch as they influence signal transduction pathways. A large volume of literature has addressed the salutary properties of n-3 fatty acids in marine oils in regard to cell proliferation and cancer (23, 38, 76). By altering the lipid composition of cell membranes in favor of n-3 fatty acids (e.g., linoleic over arachidonic acid), FO reduces the activity of protein kinase C and the subsequent events, outlined above, that activate the early- and delayed-response genes (25, 26, 33, 46, 73, 76, 86). In addition, FO feeding alters the composition of eicosanoid metabolites, for example, reducing the concentration of prostaglandin E2, which then modulates the activity of several protein kinase systems. The experimental evidence supporting this hypothesis is well documented in various cellular and tissue systems (6, 9, 36, 87).
The finding of prolonged S phase duration is unexpected given the effects of FO known so far and the established links between the extracellular environment, signal transduction pathways, and nuclear events. The current understanding of the DNA replication machinery, its assembly, and function in eukaryotic cells is primarily derived from studies on the various replication proteins and is reviewed in detail by Waga and Stillman (95). Key events include the identification of specific DNA origins, the unwinding of DNA and primosome assembly, the formation of RNA primers, and the activation of polymerases. Binding of the replication protein A (RPA) to prereplication centers, which subsequently serve as replication foci, is the initial step believed to occur in G1. Subsequent instructions to the assembled DNA replication machinery are mainly derived from S phase cdk/cyclin complexes. Once DNA replication begins at a fully assembled replication fork (origin of bidirectional replication), it proceeds very efficiently until all genomic DNA is replicated. During the progression of S phase, further regulation is achieved by a checkpoint mechanism that protects the cell against excessive DNA damage (1, 69). The components of this signal transduction pathway have been best characterized in yeast and are thought to involve several proteins (known as Rad proteins) and the protein kinase Chk1. Damage to DNA by ionizing radiation causes phosphorylation of Chk1 by a mechanism that requires Rad proteins and leads to inhibition of Cdc25. This results in a decrease in Cdc2 tyrosine-15 dephosphorylation. Furthermore, Chk1 induces the export of Cdc25 from the nucleus, thus reducing the Cdc2/cyclin B complex. This signaling cascade leads to inhibition of mitosis. Other S phase regulatory proteins (e.g., Cds1) are known in yeast. These are activated by hydroxyurea in a mechanism that also involves Cdc25, as well as on Cdc2, through an effect on the two protein kinases Wee1 and Mik1 (1, 12, 14, 47, 78). Although FO treatment of cultured cells could possibly lead to DNA damage through induction of free radicals by PUFAs, the results presented here are unlikely to be explained on the basis of this S phase checkpoint regulation. Cells treated with FO do not show evidence of DNA damage by flow cytometry (Figs. 2 and 3); they appear to start DNA synthesis normally and can still complete DNA replication, albeit at a slower rate; and they go through mitosis without evidence of an interruption. An additional support of these arguments is given by the CO treatment group, which was also exposed to PUFAs and thus to possible free radical formation. However, these cells proceeded through the cell cycle unaffected and in a similar manner as the control cells.
Approximately 3 × 109 base pairs comprise genomic DNA
in mammalian cells, which is normally replicated in ~8 h, at a fork
rate of only 100 bp/s. If each replication fork were active for the total duration of S phase, ~1,000 initiation origins would be needed
to replicate the whole genome. In fact, autoradiographic studies
suggest that between 10,000 and 100,000 replicons participate in the
replication process. Thus replication at each origin is active only
during a fraction of the S phase. This implies that the pattern of
spatial and temporal distribution of replicons is a major factor in
determining the time needed for completion of DNA replication.
Therefore, to explain the alteration in S phase kinetics in FO-treated
cells, we hypothesize that these cells have an altered pattern of
firing of replication origins. This alteration could be spatial (i.e.,
the location and/or number of active replication origins could be
affected), temporal (i.e., the timing of firing might be affected), or
a combination of both. More than 30 years ago, Huberman and Riggs
(51) demonstrated that the total duration time of S phase
is a characteristic parameter of each cell type. They showed that DNA
replication is initiated simultaneously at multiple locations.
Furthermore, it was clear from this early work that the duration of S
phase is determined by the spatial and temporal frequency of firing of
the replication origins rather than by the rate of elongation of newly
synthesized DNA (51). Since this pioneering work, major
advances in understanding of DNA replication as well as in the
structural organization of the replication origins have been made. In
yeast and lower eukaryotic cells, DNA replication starts at chromosomal
sites with specific DNA sequences. However, in higher eukaryotic cells
the identification of sequence-defined DNA replication origins has been
more elusive (28-30, 65). Studies using
two-dimensional electrophoresis in higher eukaryotic cells suggest that
DNA replication begins in a broad zone that is unlikely to be defined
by specific DNA sequences. However, subsequent studies with PCR-based
techniques using nascent DNA strand analyses were able to localize DNA
initiation origins to discrete sites. In CHO cells, a replication
initiation zone, which comprises ~55 kb and is located between the
DHFR gene and the 2BE2121 gene, has been thoroughly investigated by
several research groups using a variety of techniques (4, 15, 31, 55, 60, 63, 91, 92). Three major initiation sites have been
characterized, two of which are high-frequency start sites (called
ori- and ori-
) and one a low-frequency start site that was
discovered recently (ori-
') (60). Ori-
is located
~17 kb downstream from the 3' end of the DHFR gene, and the active initiation zone has been consistently determined as 2 kb; ori-
' lies
~5 kb further downstream from ori-
; and ori-
is found at ~40
kb downstream from the DHFR gene (Fig. 5). Despite this
characterization of the DHFR replication origin locations, the exact
significance of specific DNA sequences remains incompletely understood.
Kalejta et al. (55) demonstrated that, in CHO cells,
restoration of the 3' end missing sequence of a DHFR gene knock-out
without the ori-
locus did support replication normally but was
suppressed in variants lacking the 3' end. These investigators
concluded that it is the 3' end of the DHFR gene, rather than a
specific DNA sequence, that is required for initiating DNA replication at ori-
in the DHFR gene locus in CHO cells. On the other hand, Altman and Fanning (4) were able to initiate replication
in random ectopic chromosomal sites by transfecting a 5.8-kb fragment containing ori-
into CHO cells lacking the endogenous DHFR locus. These results suggested that specific DNA sequences within the DHFR
locus are required for efficient initiation activity. Therefore, although it is generally accepted that specific DNA sequences play an
important role in the recognition of DNA replication origins in higher
eukaryotic cells, it is apparent that other factors are also involved
as well. These factors that determine site specificity of metazoan
cells were reviewed recently by DePamphilis (30). In
addition to DNA sequences, both nuclear structure and chromatin structure appear to play an important, but incompletely understood, role in this process. The requirement for an intact nucleus could relate to the regulation of concentration and accessibility of replication factors to chromosomal DNA substrates, whereas the nuclear
matrix may provide physical attachment sites for the replication complexes (30, 62, 71). Differences in chromatin
organization throughout the genome may also account for the variable
activity of replication origins and the decision about which sites are actually used. For example, mouse fibroblast cell lines overexpressing the linker histone H1 have an altered chromatin organization
characterized by an increase in the internucleosomal spacing
(44). In a separate study, Lu et al. (66)
investigated the effects of histone H1 on DNA replication with the
Xenopus egg extract system. Presence of H1 on sperm
chromatin reduced both the rate and extent of DNA replication.
Together, these studies support the possibility that chromatin
organization within the nucleus can affect the accessibility of
replication factors to a DNA binding site.
In the current study, we used the nascent DNA strand length assay, as
originally described by Vassilev et al. (91). Although more recent studies have used multiple DNA reference segments to better
localize the initiation regions and detect lower-frequency sites, we
chose the more simplified method because our primary goal was to
determine major differences between the treatment groups. We chose
three reference DNA sequences covering a range of ~5 kb in the zone
where the high-frequency replication origin ori- is expected. One
reference sequence overlapped with the actual ori-
initiation site,
whereas the two others were flanking ori-
. Although we have followed
the same experimental protocol of Vassilev et al. (91),
our study is limited by the fact that we did not measure the actual DNA
fragment lengths in the fractions subjected to PCR
amplification. Therefore, although the expected DNA size in
fraction 10 is in the range 500-800 bp
(91), inclusion of smaller DNA lengths in this fraction
could potentially contribute Okazaki fragments from sites distant from
the replication origin. However, this possibility cannot account for
the lack of activity of segment B in the FO-treated cells in
DNA fractions 7 and 8, which are unlikely to
contain Okazaki fragments. As noted in Fig. 8, the experimental ranges
of ratios B/A and B/C in
fractions 7 and 8 in the FO-treated cells fall
outside the 95% confidence intervals defined by data from the control
group. Furthermore, our results in the non-FO-treated cells are
indistinguishable from those of the previous report (Ref.
91; see Fig. 8) and are consistent with the known location
of this replication origin in control in CHO cells (60,
91). Another technical constraint in our study is the absence of
segment C amplification in two uppermost DNA fractions, thus
limiting the usefulness of hybridization ratios
A/C and B/C in localizing
ori-
within the mapped region. Whether this limitation is related to
the choice of segment or to reduced PCR efficiency is not clear in the
current study. However, despite these limitations, our data point to
the possibility that treatment of CHO cells with FO inhibits the firing
of ori-
at the original location [P = 0.083;
Ho (null hypothesis) based on equivalence of
B/A in fractions 7 or
8]. Whether this origin is actually shifted toward
marker segment A in FO-treated cells requires further
experimental evaluation. It should be noted that the accuracy in
determining the exact location of a replication origin is dependent on
the choice of the reference DNA sequence. Choosing one marker in close
proximity to the putative center with two other markers on each side
optimizes the accuracy. Although this arrangement was held in the
control and CO-treated cells, the location of the replication origin
ori-
appears to have shifted in FO-treated cells from segment
B toward segment A, so that two of the three marker
segments are now on the same side. Therefore, our method is sensitive
enough to suggest that FO suppresses firing at the original ori-
location and suggests an upstream shift. Further studies with multiple
chromosomal markers and nascent DNA strand assays will be needed to
confirm the current findings and better localize this replication
origin and also to determine the activity at ori-
' and ori-
in
FO-treated cells.
Currently, only a few experimental conditions have been shown to lengthen S phase duration. As noted above, somatic linker histone, H1, reduces the rate and extent of DNA replication in Xenopus egg extract (66). Treatment of CHO cells with a polyamine analog, to generate a state of functional polyamine deficiency, has also been shown to prolong the duration of S phase (3). Because both histone H1 and the polyamines function in the normal packing of DNA inside the nucleus, it is possible to postulate that changes in chromatin structure can regulate the accessibility of replication factors to potential replication origins, thus altering the time required for completion of DNA replication. In another experimental manipulation, Rat-1 fibroblasts that were induced to overexpress human cyclin D1 and E had a shorter G1 phase and a compensatory longer S phase but the mean cell cycle length was unchanged (77). However, stimulation of peripheral blood lymphocytes with dexamethasone has been associated with a prolonged S phase duration while at the same time the expression of cyclin E and Cdk2 are decreased (5).
We conclude that addition of FO to culture media alters the proliferation kinetics of CHO cells mostly at the level of S phase progression. Current signal transduction pathways do not explain the mechanism of this action, which seems to involve a change in the spatial organization of replication origins. We speculate that this effect could be related to a change within the nucleus of chromatin structure and/or the association between replicating DNA and structural components such as the nuclear matrix. FO could affect these changes either directly, by altering the fluidity of the nuclear membrane, or indirectly, through mechanisms that might involve polyamine metabolism or histone acetylation. Alternative hypotheses that could possibly involve novel signal transduction pathways within the nuclear membrane, as recently suggested for prostaglandin E2 (10), also must be addressed. Regardless of the exact pathways, the result of this study, pointing to a possible change in the spatial location of a well-defined replication origin, may also prove to be important for the study of the process of DNA replication in mammalian cells.
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ACKNOWLEDGEMENTS |
---|
This work was supported by National Cancer Institute Grant CA-45768 to N. W. Istfan.
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FOOTNOTES |
---|
Address for reprint requests and other correspondence: N. W. Istfan, Boston Univ. School of Medicine, Section of Endocrinology, Diabetes and Nutrition, 88 East Newton St., Evans 201, Boston, MA 02118 (E-mail: nsteph3{at}cs.com).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
May 29, 2002;10.1152/ajpcell.00614.2001
Received 26 December 2001; accepted in final form 22 May 2002.
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