Fish oil slows S phase progression and may cause upstream shift of DHFR replication origin ori-beta in CHO cells

Nawfal W. Istfan, Zhi-Yi Chen, and Sybille Rex

Section of Endocrinology, Diabetes, and Nutrition, Department of Medicine, Boston University School of Medicine, Boston, Massachusetts 02118


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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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-beta 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-beta was established. In control or corn oil-treated cells, the location of ori-beta was consistent with previous studies. However, in FO-treated cells, DNA replication appears to start from a new site located farther upstream from ori-beta , 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


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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)-gamma 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-beta 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-beta 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.


    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
C(t)=C<SUB>o</SUB><IT> · e</IT><SUP>(<IT>K</IT><SUB>g</SUB><IT> · t</IT>)</SUP> (1)
where C is the total number of viable cells and Co is the initial cell number at the time point of plating. From the growth rate constant of exponentially growing cells, the actual doubling time Td can be calculated as
T<SUB>d</SUB><IT>=</IT>ln 2<IT>/K</IT><SUB>g</SUB> (2)

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 nu  was calculated, as previously defined by White et al. (102), according to the following equation
&ngr;=ln <FR><NU>1<IT>+</IT>f<SUB>lu</SUB></NU><DE>1<IT>−</IT>(f<SUB>ld</SUB><IT>/</IT>2)</DE></FR> (3)
This function nu  is related to the potential doubling time (Tpot), the DNA synthesis time (TS), and the cell production rate (c) by the following two equations (102)
T<SUB>pot</SUB><IT>=</IT>ln 2<FR><NU><IT>T</IT><SUB>S</SUB></NU><DE><IT>&ngr;</IT></DE></FR> (4)

T<SUB>pot</SUB><IT>=</IT><FR><NU>ln 2</NU><DE><IT>c</IT></DE></FR> (5)
TS, which represents the duration time of S phase, is derived from measurements of the relative movement. The parameter relative movement (RM) is defined by the position of BrdU-labeled undivided cells on the DNA axis relative to the position of cell populations in G1 and G2/M phase and is determined by the following equation (8)
RM = <FR><NU>F<SUB>L</SUB><IT>−</IT>F<SUB>G<SUB>1</SUB></SUB></NU><DE>F<SUB>G<SUB>2</SUB></SUB><IT>−</IT>F<SUB>G<SUB>1</SUB></SUB></DE></FR> (6)
where FL, FG1, and FG2 represent the mean red fluorescence (DNA content) of labeled undivided cells and of cells in G0/G1 and G2/M phases, respectively.

Two analytical procedures were used to determine TS. In the first method, the linear relationship RM(t) defines TS as the time required for the relative movement to equal 1.0 (i.e., the mean DNA content of flu is equivalent to that of G2) according to the following equation (97)
RM(<IT>t</IT>)<IT>=</IT>RM<SUB>O</SUB><IT>+</IT><FR><NU>1</NU><DE>2<IT>T</IT><SUB>S</SUB></DE></FR><IT>t</IT> (7)
when the sampling time t is TG2+M <=  t <=  TS. RMo is the initial relative movement at time 0 after the BrdU pulse and is calculated as (88, 97)
RM<SUB>O</SUB> = <FR><NU>(S − Z)</NU><DE>Z(S+G<SUB>2</SUB>)</DE></FR> (8)

Z = ln <FR><NU>(1 + S + G<SUB>2</SUB>)</NU><DE>(1 + G<SUB>2</SUB>)</DE></FR>
In this equation, S and G2 represent the fraction of cells in S and G2/M phases, respectively, as determined from the one-dimensional DNA histograms.

In the second analytical procedure, the cell production rate (c) is determined from the solution of a cubic equation relating RM(t), nu , and flu, as previously described by White et al. (98), according to the equation
<IT>c</IT> = <FENCE>&ngr; − 2 cos <FENCE><FR><NU>&thgr; − 2&pgr;</NU><DE>3</DE></FR></FENCE> − 1</FENCE>/<IT>t</IT> (9)
where
&thgr;=arccos <FENCE>1 − <FR><NU>3&ngr;f<SUB>lu</SUB>(<IT>t</IT>)[1<IT>−</IT>RM(<IT>t</IT>)]</NU><DE>1<IT>+</IT>f<SUB>lu</SUB>(<IT>t</IT>)</DE></FR></FENCE> (10)
After estimation of the cell production rate, the DNA synthesis time (TS) and the time spent in G2/M phase (TG2+M) are derived from the following two relationships
T<SUB>S</SUB> = &ngr;/<IT>c</IT> (11)
and
<IT>T</IT><SUB>G2+M</SUB><IT>=t+</IT><FR><NU>ln [1 − f<SUB>ld</SUB>(<IT>t</IT>)<IT>/</IT>2]</NU><DE><IT>c</IT></DE></FR> (12)
Assuming exponential growth characteristics and a growth fraction of 1, the total cell cycle time, Tc, is determined according to the equation of Steel (85)
T<SUB>c</SUB> = <FR><NU>ln 2(<IT>T</IT><SUB>G2+M</SUB>)</NU><DE>ln (1+G<SUB>2</SUB>)</DE></FR> (13)
where G2 is the percentage of cells in G2/M phase as determined from DNA histograms. Consequently, the time spent in G1 is estimated as the difference between Tc and the sum of TS plus TG2+M. Finally, an estimate of cell loss is derived from comparison of Tpot and Td according to the following equation for the cell loss factor (Phi )
&PHgr;=1−(T<SUB>pot</SUB><IT>/T</IT><SUB>d</SUB>) (14)

Experimental Part II: Mapping of DHFR Replication Origin ori-beta in Treated CHO Cells

We used a method initially described by Vassilev and colleagues (89-91) to determine the position of the DHFR replication origin ori-beta in exponentially growing, treated CHO cells relative to three marker segments A, B, and C, as defined in detail elsewhere (89, 91). Briefly, these segments were selected within a 5-kb region flanking the replication origin ori-beta , which is located 17 kb downstream of the 3' end of the DHFR gene. In exponentially growing cells, nascent DNA was labeled with BrdU and then purified and separated by length into several gravimetric fractions with a sucrose gradient. Those DNA sequences that contained the marker segments were amplified by a standard PCR technique and quantified by an automated system as described below. Measurements of the abundance of each marker in the various fractions of nascent DNA allowed an estimation of the position of the replication origin relative to the marker segments. The initiation site is located closest to that segment that remains represented in the fraction of the shortest newly synthesized DNA.

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 P <=  0.05.

For analysis of the effect of FO on ori-beta 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-beta mapping, the null hypothesis that differences in the ECL ratios among treatment groups were due to chance was tested by chi 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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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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Fig. 1.   Growth curves of Chinese hamster ovary (CHO) cells treated for 5 days with control growth medium (circle) or medium enriched in corn oil (triangles) or fish oil (squares). After reseeding, cells were counted daily for 3 consecutive days. Data are mean values pooled from 3 experiments; error bars represent SE. Lines represent the best fit for exponential growth (Eq. 1) by use of regression analysis. The corresponding values of the actual doubling times (Td) are given in Table 3.

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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Fig. 2.   Representative DNA histograms for control (A) and corn oil (B)- and fish oil (C)-treated CHO cells. The total DNA content of cells (x-axis) was obtained by staining with propidium iodide. Cells were analyzed by flow cytometry. The percentage of cells in each cell cycle phase was determined with the program ModFit. Average values from 6 experiments are summarized in Table 1. The first, large peak represents population of cells (y-axis) in G0/G1 phase, the second, small peak shows population of cells in G2/M phase, and the gray area between both peaks represents cells in S phase. CV, coefficient of variation and a measure of resolution for the G0/G1 peak.


                              
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Table 1.   Cell cycle phase distribution derived from DNA histograms

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 nu  (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 nu  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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Fig. 3.   General example for bivariate analysis of 5-bromo-2'-deoxyuridine (BrdU)-DNA contour plots obtained by flow cytometry. After a BrdU pulse for 30 min, treated cells were incubated for 1 (A) and 5 (B) h with the corresponding BrdU-free medium. The x-axis represents the linear intensity of red fluorescence obtained from propidium iodide (total DNA content), and the y-axis represents the logarithmic intensity of green fluorescence obtained from FITC-conjugated anti-BrdU antibodies (BrdU-labeled population). Cells are separated into G0/G1 phase and G2/M phase according to their DNA content, and into labeled undivided (flu) and labeled divided (fld) subgroups according to the DNA content of BrdU-labeled cells. Estimates of the percentage of cells in both subgroups as well as the relative position of cells in G0/G1 and G2/M phase (according to their DNA content) were derived with IsoContour software. As a result of DNA synthesis, BrdU-labeled cells (S phase, A) move toward higher DNA content (flu, B) as demonstrated.


                              
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Table 2.   Kinetic parameters derived from bivariate BrdU-DNA flow cytometry

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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Fig. 4.   Plot of the relative movement of labeled undivided cells as function of time shown for control (circle) and corn oil (CO; triangles)- and fish oil (FO; squares)-treated cells. The relative movement of labeled cells in each treatment group was measured 1, 3, and 5 h after the BrdU pulse and calculated according to Eq. 5. Data are averaged over 6 experiments and given as mean values ± SE. The lines represent the best linear regression curve (r2 > 0.98 for each treatment group). Numeric values of the relative movement at 5 h [RM(5)] are displayed in Table 2. S phase duration times TS are calculated from Eqs. 7 and 11 and are shown in Table 3. *P < 0.05 or better for fish oil compared with either control or corn oil.

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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Table 3.   Estimates of actual doubling time, potential doubling time, and DNA replication time and other cell cycle times

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 Phi  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-beta

The finding of an altered S phase kinetics in the FO treatment group led us to hypothesize a change in the temporal and/or spatial initiation pattern of replication origins. In CHO cells a replication initiation zone located ~17 kb downstream from the 3' end of the DHFR gene was previously established and characterized thoroughly by a variety of methods (4, 15, 28, 30, 31, 55, 60, 63, 91). We tested our hypothesis by mapping the activation of the replication origin ori-beta within the DHFR replication zone in treated CHO cells. We used a nascent strand length assay following a protocol initially described by Vassilev et al. (89-91) with modifications in the detection method of the PCR products. Nascent DNA was labeled with 3H-dC and BrdU, sedimented, and fractionated according to size by use of an alkaline sucrose gradient. 3H radioactivity in each fraction was measured to ensure uniform labeling of the nascent DNA, because the total activity varied proportionally to size. Nascent BrdU-containing DNA was then separated from parent DNA by repeated precipitation with anti-BrdU antibodies. The percent recovery was checked by counting 3H activity, which exceeded 50% in all the samples being used in further measurements. After purification, nascent DNA was used to amplify the marker segments defined by sequences A, B, and C, which span a region of ~5 kb flanking the replication site ori-beta within the DHFR replication initiation zone (Fig. 5). Separate experiments were conducted to determine the appropriate PCR conditions to generate distinct DNA bands for each of the primers used in the amplification procedure and to verify the specificity of the probes (data not shown). These preliminary experiments confirmed earlier results from Vassilev et al. (91) about the specificity of the primers and the applicability of PCR conditions in the separate sucrose gradient fractions. As noted in MATERIALS AND METHODS, a 5-20% linear sucrose gradient was subdivided into 12 fractions, with the first and last fractions discarded. In this range, the size of DNA fragments is expected to average ~500-800 bp in the smallest fraction (labeled as fraction 10 in the current study) and ~10 kb in fraction 3 (91). Fractions 1-10 as defined above, containing purified nascent DNA, were mixed with primers A, B, and C and subjected to PCR amplification. The PCR products in each fraction were quantified by specific TBR probes for each reference segment. An ECL signal was generated on reacting TBR in the PCR product with TPA in the assay buffer. The signal was quantified by use of an automated system and is proportional to the amount of hybridized PCR product.


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Fig. 5.   Schematic representation of the organization of the replication initiation zone between the dihydrofolate reductase (DHFR) and 2BE2121 genes (55 kb) in CHO cells. The high-frequency initiation sites (ori-beta and ori-gamma ) as well as the low-frequency site (ori-beta ') are indicated. Furthermore, the position of the 3 marker segments relative to ori-beta and ori-beta ' are shown in greater detail.

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-beta 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-beta 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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Fig. 6.   Summary of quantitative PCR analysis for the 3 marker segments AB, and C (A-C, respectively) flanking the DHFR replication origin ori-beta in treated CHO cells. Nascent BrdU-labeled DNA was purified by 2 rounds of immunoprecipitation and separated by use of a sucrose gradient. Fractions are numbered starting with the largest DNA fragments (fraction 1). Segments A, B, and C were amplified by PCR in each fraction and subsequently quantified by an electrochemiluminescence (ECL) signal. In each experiment, ECL was highest for the largest DNA fragments (fractions 1 and 2). Here, the mean ECL value of 2 experiments is shown for each fraction and treatment group. Data are normalized to the average ECL signal in fractions 1 and 2 (measured counts - blank) and expressed as a percentage. Each fraction shows control, fish oil-treated, and corn oil-treated cells.

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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Fig. 7.   Plot of individual ratios of ECL signals for marker segments A, B, and C from Fig. 6. Ratios B/A (A), A/B (B), C/A (C), C/B (D), A/C (E), and B/C (F) for control, corn oil-treated, and fish oil-treated cells are shown. A/C and B/C in fractions 9 and 10 were deleted from the figure (missing data denoted by asterisk) because estimates of the PCR product of this segment were not different from zero in 2 uppermost DNA fractions (see text).

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-beta .


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Fig. 8.   Plot of the ratios B/A, B/C, and A/C in fractions 7 and 8 in FO- and CO-treated CHO cells in reference to the expected ranges of these ratios in untreated cells. Boxes indicate the 95% confidence intervals (n = 4) derived from untreated control cells in the current study. Data shown as open circles with range lines represent the corresponding values reported in Ref. 91 from 2 independent experiments in exponentially growing CHO cells similar to the control cells of the current study. Mean values for FO- and CO-treated cells, respectively, from 2 independent experiments in the current study are also shown, with lines representing the range of values.

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-beta in both experiments. These findings led us to propose that the DHFR replication origin ori-beta , which is normally in close proximity to marker segment B, is suppressed in FO-treated cells (P = 0.083, by chi 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-beta 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.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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-beta 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-beta and ori-gamma ) and one a low-frequency start site that was discovered recently (ori-beta ') (60). Ori-beta 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-beta ' lies ~5 kb further downstream from ori-beta ; and ori-gamma 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-beta 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-beta 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-beta 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-beta is expected. One reference sequence overlapped with the actual ori-beta initiation site, whereas the two others were flanking ori-beta . 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-beta 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-beta 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-beta 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-beta 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-beta ' and ori-gamma 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.


    ACKNOWLEDGEMENTS

This work was supported by National Cancer Institute Grant CA-45768 to N. W. Istfan.


    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.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Aguda, BD. A quantitative analysis of the kinetics of the G2 DNA damage checkpoint system. Proc Natl Acad Sci USA 96: 11352-11357, 1999[Abstract/Free Full Text].

2.   Aguda, BD, and Tang Y. The kinetic origins of the restriction point in the mammalian cell cycle. Cell Prolif 32: 321-335, 1999[ISI][Medline].

3.   Alm, K, Berntsson PSH, Kramer DL, Porter CW, and Oredsson SM. Treatment of cells with the polyamine analog N1,N11-diethylnorspermine retards S phase progression within one cell cycle. Eur J Biochem 267: 4157-4164, 2000[Abstract/Free Full Text].

4.   Altman, AL, and Fanning E. The chinese hamster dihydrofolate reductase replication origin beta is active at multiple ectopic chromosomal locations and requires specific DNA sequence elements for activity. Mol Cell Biol 21: 1098-1110, 2001[Abstract/Free Full Text].

5.   Baghdassarian, N, Catallo R, Mahly MA, Ffrench P, Chizat F, Bryon PA, and Ffrench M. Glucocorticoids induce G1 as well as S-phase lengthening in normal human stimulated lymphocytes: differential effects on cell cycle regulatory proteins. Exp Cell Res 240: 263-273, 1998[ISI][Medline].

6.   Bandyopadhyay, GK, Hwang S-i, Imagawa W, and Nandi S. Role of polyunsaturated fatty acids as signal transducers: amplification of signals from growth factor receptors by fatty acids in mammary epithelial cells. Prostaglandins Leukot Essent Fatty Acids 48: 71-78, 1993[ISI][Medline].

7.   Bartram, HP, Gostner A, Scheppach W, Reddy BS, Dusel G, Richter F, and Richter A. Effects of fish oil on rectal cell proliferation, mucosal fatty acids and prostaglandin E2 release in healthy subjects. Gastroenterology 105: 1317-1322, 1993[ISI][Medline].

8.   Begg, AC, McNally NJ, Shrieve DC, and Kaercher H. A method to measure the duration of DNA synthesis and the potential doubling time from a single sample. Cytometry 6: 620-626, 1985[ISI][Medline].

9.   Bell, RM, and Burns DJ. Lipid activation of protein kinase C. J Biol Chem 266: 4661-4664, 1991[Free Full Text].

10.   Bhattacharya, M, Peri K, Ribeiro-da-Silva A, Almazan G, Shichi H, Hou X, Varma DR, and Chemtob S. Localization of functional prostaglandin E2 receptors EP3 and EP4 in the nuclear envelope. J Biol Chem 274: 15719-15724, 1999[Abstract/Free Full Text].

11.   Blow, JJ. S phase and its regulation. In: Cell Cycle Control, edited by Hutchison C, and Glover DM.. Oxford, UK: Oxford Univ. Press, 1995.

12.   Bousset, K, and Diffley JFX The Cdc7 protein kinase is required for origin firing during S phase. Genes Dev 12: 480-490, 1998[Abstract/Free Full Text].

13.   Braga, C, La Vecchia C, Franceschi S, Negri E, Parpinel M, Decarli A, Giacosa A, and Trichopoulos D. Olive oil, other seasoning fats, and the risk of colorectal carcinoma. Cancer 82: 448-453, 1998[ISI][Medline].

14.   Brondello, JM, Boddy MN, Furnari B, and Russell P. Basis for the checkpoint signal specificity that regulates Chk1 and Cds1 protein kinases. Mol Cell Biol 19: 4262-4269, 1999[Abstract/Free Full Text].

15.   Caddle, MS, Lussier RH, and Heintz NH. Intramolecular DNA triplexes, bent DNA and DNA unwinding elements in the initiation region of an amplified dihydrofolate reductase replicon. J Mol Biol 211: 19-33, 1990[ISI][Medline].

16.   Chen, ZY, and Istfan NW. Docosahexaenoic acid is a potent inducer of apoptosis in HT-29 colon cancer cells. Prostaglandins Leukot Essent Fatty Acids 63: 301-308, 2000[ISI][Medline].

17.   Chinery, R, Coffey RJ, Graves-Deal R, Kirkland SC, Sanchez SC, Zackert WE, Oates JA, and Morrow JD. Prostaglandin J2 and 15-deoxy-Delta 12,14-prostaglandin J2 induce proliferation of cyclooxygenase-depleted colorectal cells. Cancer Res 59: 2739-2746, 1999[Abstract/Free Full Text].

18.   Clifford, C, and Kramer B. Diet as risk and therapy for cancer. Med Clin North Am 77: 725-744, 1993[ISI][Medline].

19.   Clinton, SK, Imrey PB, Mangian HJ, Nandkumar S, and Visek WJ. The combined effects of dietary fat, protein, and energy intake on azoxymethane-induced intestinal and renal carcinogenesis. Cancer Res 52: 857-865, 1992[Abstract].

20.   Clinton, SK, Li PS, Mulloy AL, Imrey PB, Nandkumar S, and Visek WJ. The combined effects of dietary fat and estrogen on survival, 7,12-dimethylbenz(a)anthracene-induced breast cancer and prolactin metabolism in rats. J Nutr 125: 1192-1204, 1995[ISI][Medline].

21.   Collett, ED, Davidson LA, Fan YY, Lupton JR, and Chapkin RS. n-6 and n-3 polyunsaturated fatty acids differentially modulate oncogenic Ras activation in colonocytes. Am J Physiol Cell Physiol 280: C1066-C1075, 2001[Abstract/Free Full Text].

22.   Coverley, D, and Laskey RA. Regulation of eukaryotic DNA replication. Annu Rev Biochem 63: 745-776, 1994[ISI][Medline].

23.   Cowing, BE, and Saker KE. Polyunsaturated fatty acids and epidermal growth factor receptor/mitogen-activated protein kinase signaling in mammary cancer. J Nutr 131: 1125-1128, 2001[Abstract/Free Full Text].

24.   Crighton, IL, Dowsett M, Hunter M, Shaw C, and Smith IE. The effect of a low-fat diet on hormone levels in healthy pre- and postmenopausal women: relevance for breast cancer. Eur J Cancer 28A: 2024-2027, 1992.

25.   Danesch, U, Weber PC, and Sellmayer A. Arachidonic acid increases c-fos and Egr-1 mRNA in 3T3 fibroblasts by formation of prostaglandin E2 and activation of protein kinase C. J Biol Chem 269: 27258-27263, 1994[Abstract/Free Full Text].

26.   Danesch, U, Weber PC, and Sellmayer A. Differential effects of n-6 and n-3 polyunsaturated fatty acids on cell growth and early gene expression in Swiss 3T3 fibroblasts. J Cell Physiol 168: 618-624, 1996[ISI][Medline].

27.   Davis, BM. High fiber diet and colorectal adenomas. N Engl J Med 343: 736-738, 2000[Free Full Text].

28.   DePamphilis, ML. Eukaryotic DNA replication: anatomy of an origin. Annu Rev Biochem 62: 29-63, 1993[ISI][Medline].

29.   DePamphilis, ML. Origins of DNA replication that function in eukaryotic cells. Curr Opin Cell Biol 5: 434-441, 1993[Medline].

30.   DePamphilis, ML. Replication origins in metazoan chromosomes: fact or fiction? Bioessays 21: 5-16, 1999[ISI][Medline].

31.   Dijkwel, PA, and Hamlin JL. The chinese hamster dihydrofolate reductase origin consists of multiple potential nascent-strand start sites. Mol Cell Biol 15: 3023-3031, 1995[Abstract].

32.   DiMarzo, V. Arachidonic acid and eicosanoids as targets and effectors in second messenger interactions. Prostaglandins Leukot Essent Fatty Acids 53: 239-254, 1995[ISI][Medline].

33.   Divecha, N, and Irvine RF. Phospholipid signaling. Cell 80: 269-278, 1995[ISI][Medline].

34.   Dwyer, JT. Diet and nutritional strategies for cancer risk reduction. Focus on the 21st century. Cancer 72: 1024-1031, 1993[ISI][Medline].

35.   Erickson, KL. Is there a relation between dietary linoleic acid and cancer of the breast, colon, or prostate? Am J Clin Nutr 68: 5-7, 1998[Free Full Text].

36.   Fox, PL, and DiCorleto PE. Fish oils inhibit endothelial cell production of platelet-derived growth factor-like protein. Science 241: 453-456, 1988[ISI][Medline].

37.   Fradet, Y, Meyer F, Bairati I, Shadmani R, and Moore L. Dietary fat and prostate cancer progression and survival. Eur Urol 35: 388-391, 1999[ISI][Medline].

38.   Fritsche, KL, and Johnston PV. Effect of dietary alpha -linolenic acid on growth, metastasis, fatty acid profile and prostaglandin production of two murine mammary adenocarcinomas. J Nutr 120: 1601-1609, 1990[ISI][Medline].

39.   Gaard, M, Tretli S, and Loken EB. Dietary fat and the risk of breast cancer: a prospective study of 25,892 Norwegian women. Int J Cancer 63: 13-17, 1995[ISI][Medline].

40.   Ganz, PA, and Schag AC. Nutrition and breast cancer. Oncology (Huntingt) 7: 71-75, 1993.

41.   Giovannucci, E, Rimm EB, Colditz GA, Stampfer MJ, Ascherio A, Chute CC, and Willett WC. A prospective study of dietary fat and risk of prostate cancer. J Natl Cancer Inst 85: 1571-1579, 1993[Abstract].

42.   Gonzalez, MJ, Schemmel RA, Gray JI, Dugan L, Sheffield LG, and Welsch CW. Effect of dietary fat on growth of MCF-7 and MDA-MB231 human breast carcinomas in athymic nude mice: relationship between carcinoma growth and lipid peroxidation product levels. Carcinogenesis 12: 1231-1235, 1991[Abstract].

43.   Graham, S, Zielezny M, Marshall J, Priore R, Freudenheim J, Brasure J, Haughey B, Nasca P, and Zdeb M. Diet in the epidemiology of postmenopausal breast cancer in the New York State Cohort. Am J Epidemiol 136: 1327-1337, 1992[Abstract].

44.   Gunjan, A, Alexander BT, Sittman DB, and Brown DT. Effects of H1 histone variant overexpression on chromatin structure. J Biol Chem 274: 37950-37956, 1999[Abstract/Free Full Text].

45.   Gurr, MI. Diet and the prevention of cancer. No evidence has linked ovarian cancer with high intakes of fat and meat. BMJ 319: 187-188, 1999.

46.   Handler, JA, Danilowicz RM, and Eling TE. Mitogenic signaling by epidermal growth factor (EGF), but not platelet-derived growth factor, requires arachidonic acid metabolism in BALB/c 3T3 cells. J Biol Chem 265: 3669-3673, 1990[Abstract/Free Full Text].

47.   Hartwell, LH, and Weinert TA. Checkpoints: controls that ensure the order of cell cycle events. Science 246: 629-634, 1989[ISI][Medline].

48.   Hashimoto, M, Hossain MS, Yamasaki H, Yazawa K, and Masumura S. Effects of eicosapentaenoic acid and docosahexaenoic acid on plasma membrane fluidity of aortic endothelial cells. Lipids 34: 1297-1304, 1999[ISI][Medline].

49.   Holmes, MD, Hunter DJ, Colditz GA, Stampfer MJ, Hankinson SE, Speizer FE, Rosner B, and Willett WC. Association of dietary intake of fat and fatty acids with risk of breast cancer. JAMA 281: 914-920, 1999[Abstract/Free Full Text].

50.   Howe, GR, Aronson KJ, Benito E, Castelleto R, Cornee J, Duffy S, Gallagher RP, Iscovich JM, Deng-ao J, Kaaks R, Kune GA, Kune S, Lee HP, Lee M, Miller AB, Peters RK, Potter JD, Riboli E, Slattery ML, Trichopoulos D, Tuyns A, Tzonou A, Watson LF, Whittemore AS, and Shu Z. The relationship between dietary fat intake and risk of colorectal cancer: evidence from the combined analysis of 13 case-control studies. Cancer Causes Control 8: 215-228, 1997[ISI][Medline].

51.   Huberman, JA, and Riggs AD. On the mechanism of DNA replication in mammalian chromosomes. J Mol Biol 32: 327-341, 1968[ISI][Medline].

52.   Hunt, T, and Kirschner MW. Cell multiplication. Curr Opin Cell Biol 5: 163-165, 1992.

53.   Hunter, DJ. Cohort studies of fat intake and the risk of breast cancer---a pooled analysis. N Engl J Med 334: 356-361, 1996[Abstract/Free Full Text].

54.   Istfan, NW, Wan JM, Bistrian BR, and Chen ZY. DNA replication time accounts for tumor growth variation induced by dietary fat in a breast carcinoma model. Cancer Lett 86: 177-186, 1994[ISI][Medline].

55.   Kalejta, RF, Li X, Mesner LD, Dijkwel PA, Lin HB, and Hamlin JL. Distal sequences, but not ori-beta /OBR-1, are essential for initiation of DNA replication in the Chinese hamster DHFR origin. Mol Cell 2: 797-806, 1998[ISI][Medline].

56.   Karin, M. Signal transduction from cell surface to nucleus in development and disease. FASEB J 6: 2581-2590, 1992[Abstract/Free Full Text].

57.   Karmali, RA, Marsh J, and Fuchs C. Effect of omega-3 fatty acids on growth of a rat mammary tumor. J Natl Cancer Inst 73: 457-461, 1984[ISI][Medline].

58.   Katz, EB, and Boylan ES. Effect of the quality of dietary fat on tumor growth and metastasis from a rat mammary adenocarcinoma. Nutr Cancer 12: 343-350, 1989[ISI][Medline].

59.   Keller, H, Dreyer C, Medin J, Mahfoudi A, Ozato K, and Wahli W. Fatty acids and retinoids control lipid metabolism through activation of peroxisome proliferator-activated receptor-retinoid X receptor heterodimers. Proc Natl Acad Sci USA 90: 2160-2164, 1993[Abstract].

60.   Kobayashi, T, Rein T, and DePamphilis ML. Identification of primary initiation sites for DNA replication in the hamster dihydrofolate reductase gene initiation zone. Mol Cell Biol 18: 3266-3277, 1998[Abstract/Free Full Text].

61.   Kolonel, LN, Nomura AM, and Cooney RV. Dietary fat and prostate cancer. J Natl Cancer Inst 91: 414-428, 1999[Abstract/Free Full Text].

62.   Laskey, RA, and Madine M. Roles of nuclear structure in DNA replication. In: DNA Replication in Eukaryotic Cells, edited by DePamphilis ML.. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press, 1996, p. 119-130.

63.   Leu, TH, and Hamlin JL. High resolution mapping of replication fork movement through the amplified dihydrofolate reductase domain in CHO cells by in-gel renaturation analysis. Mol Cell Biol 9: 523-531, 1989[ISI][Medline].

64.   Lin, LL, Wartmann M, Lin AY, Knopf JL, Seth A, and Davis RJ. cPLA2 is phosphorylated and activated by MAP kinase. Cell 72: 269-278, 1993[ISI][Medline].

65.   Liskens, MHK, and Huberman JA. The two faces of higher eukaryotic DNA replication origins. Cell 62: 845-847, 1990[ISI][Medline].

66.   Lu, ZH, Sittman DB, Brown DT, Munhi R, and Leno GH. Histone H1 modulates DNA replication through multiple pathways in Xenopus egg extract. J Cell Sci 110: 2745-2758, 1997[Abstract/Free Full Text].

67.   Murray, AW. Creative blocks: cell-cycle checkpoints and feedback controls. Nature 359: 599-604, 1992[ISI][Medline].

68.   Neri, LM, Capitani S, Borgatti P, and Martelli AM. Lipid signaling and cell responses at the nuclear level. Histol Histopathol 14: 321-335, 1999[ISI][Medline].

69.   Paulovich, AG, and Hartwell LH. A checkpoint regulates the rate of progression through S phase in S. cerevisiae in response to DNA damage. Cell 82: 841-847, 1995[ISI][Medline].

70.   Pearson, G, Robinson F, Gibson TB, Xu B, Karandikar M, Berman K, and Cobb MH. Mitogen-activated protein (MAP) kinase pathways: regulation and physiological functions. Endocr Rev 22: 153-183, 2001[Abstract/Free Full Text].

71.   Pemov, A, Bavykin S, and Hamlin JL. Attachment to the nuclear matrix mediates specific alterations in chromatin structure. Proc Natl Acad Sci USA 95: 14757-14762, 1998[Abstract/Free Full Text].

72.   Rao, CV, and Reddy BS. Modulating effect of amount and types of dietary fat on ornithine decarboxylase, tyrosine protein kinase and prostaglandins production during colon carcinogenesis in male F344 rats. Carcinogenesis 14: 1327-1333, 1993[Abstract].

73.   Rao, GN, Baas AS, Glasgow WC, Eling TE, Runge MS, and Alexander RW. Activation of mitogen-activated protein kinases by arachidonic acid and its metabolites in vascular smooth muscle cells. J Biol Chem 269: 32586-32591, 1994[Abstract/Free Full Text].

74.   Reddy, BS, Burill C, and Rigotty J. Effect of diets high in omega-3 and omega-6 fatty acids on initiation and postinitiation stages of colon carcinogenesis. Cancer Res 51: 487-491, 1991[Abstract].

75.   Reddy, BS, and Maruyama H. Effect of dietary fish oil on azoxymethane-induced colon carcinogenesis in male 344 rats. Cancer Res 46: 3367-3370, 1986[Abstract].

76.   Reddy, BS, and Sugie S. Effect of different levels of omega-3 and omega-6 fatty acids on azoxymethane-induced colon carcinogenesis in F344 rats. Cancer Res 48: 6642-6647, 1988[Abstract].

77.   Resnitzky, D, Gossen M, Bujrd H, and Reed SI. Acceleration of the G1/S phase transition by expression of cyclins D1 and E with an inducible system. Mol Cell Biol 14: 1669-1679, 1994[Abstract].

78.   Roberts, BT, Ying CY, Gautier J, and Maller JL. DNA replication in vertebrates requires a homolog of the Cdc7 protein kinase. Proc Natl Acad Sci USA 96: 2800-2804, 1999[Abstract/Free Full Text].

79.   Rose, DP, Connolly JM, and Heschter CI. Effects of dietary fat on human breast cancer growth and lung metastasis in nude mice. J Natl Cancer Inst 83: 1491-1495, 1991[Abstract].

80.   Rumi, MAK, Sato H, Ishihara S, Kawashima K, Hamamoto S, Kazumori H, Okuyama T, Fukuda R, Nagasue N, and Kinoshita Y. Peroxisome proliferator-activated receptor gamma  ligand-induced growth inhibition of human hepatocellular carcinoma. Br J Cancer 84: 1640-1647, 2001[ISI][Medline].

81.   Sasaki, S, Horacsek M, and Kesteloot H. An ecological study of the relationship between dietary fat intake and breast cancer mortality. Prev Med 22: 187-202, 1993[ISI][Medline].

82.   Sauer, LA, and Dauchy RT. The effect of omega-6 and omega-3 fatty acids on 3H-thymidine incorporation in hepatoma 7288CTC perfused in situ. Br J Cancer 66: 297-303, 1992[ISI][Medline].

83.   Shirahige, K, Hori Y, Shiraishi K, Yamashita M, Takahashi K, Obuse C, Tsurimoto T, and Yoshikawa H. Regulation of DNA-replication origins during cell-cycle progression. Nature 395: 618-621, 1998[ISI][Medline].

84.   Simonsen, N, van't Veer P, Strain JJ, Martin-Moreno JM, Huttunen JK, Navajas JFC, Martin BC, Thamm M, Kardinaal AFM, Kok FJ, and Kohlmeier L. Adipose tissue omega-3 and omega-6 fatty acid content and breast cancer in the EURAMIC study. Am J Epidemiol 147: 342-352, 1998[Abstract].

85.   Steel, GG. Growth Kinetics of Tumors: Cell Population Kinetics in Relation to the Growth and Treatment of Cancer. Oxford, UK: Clarendon, 1977.

86.   Stulnig, TM, Berger M, Sigmund T, Raederstorff D, Stockinger H, and Waldhaeusl W. Polyunsaturated fatty acids inhibit T cell signal transduction by modification of detergent-insoluble membrane domains. J Cell Biol 143: 637-644, 1998[Abstract/Free Full Text].

87.   Sumida, C, Graber R, and Nunez E. Role of fatty acids in signal transduction: modulators and messengers. Prostaglandins Leukot Essent Fatty Acids 48: 117-122, 1993[ISI][Medline].

88.   Terry, NHA, White RA, Meistrich ML, and Calkins DP. Evaluation of flow cytometric methods for determining population potential doubling times using cultured cells. Cytometry 12: 234-241, 1991[ISI][Medline].

89.   Vassilev, L, and Johnson EM. Mapping initiation sites of DNA replication in vivo using polymerase chain reaction amplification of nascent strand segments. Nucleic Acids Res 17: 7693-7705, 1989[Abstract].

90.   Vassilev, L, and Johnson EM. An initiation zone of chromosomal DNA replication located upstream of the c-myc gene in proliferating HeLa cells. Mol Cell Biol 10: 4899-4904, 1990[ISI][Medline].

91.   Vassilev, LT, Burhans WC, and DePamphilis ML. Mapping an origin of DNA replication at a single-copy locus in exponentially proliferating mammalian cells. Mol Cell Biol 10: 4685-4689, 1990[ISI][Medline].

92.   Vassilev, LT, and DePamphilis ML. Guide to identification of origins of DNA replication in eukaryotic cell chromosomes. Crit Rev Biochem Mol Biol 27: 445-472, 1992[Abstract].

93.   Veierod, MB, Laake P, and Thelle DS. Dietary fat intake and risk of prostate cancer: a prospective study of 25,708 Norwegian men. Int J Cancer 73: 634-638, 1997[ISI][Medline].

94.   Vognild, E, Elvevoll EO, Brox J, Olsen RL, Barstad H, Aursand M, and Osterud B. Effects of dietary marine oils and olive oil on fatty acid composition, platelet membrane fluidity, platelet responses and serum lipids in healthy humans. Lipids 33: 427-436, 1998[ISI][Medline].

95.   Waga, S, and Stillman B. The DNA replication fork in eukaryotic cells. Annu Rev Biochem 67: 721-751, 1998[ISI][Medline].

96.   Wan, JM, Fogt F, Bistrian BR, and Istfan NW. Evaluation of antitumor effect of tumor necrosis factor in terms of protein metabolism and cell cycle kinetics. Am J Physiol Cell Physiol 265: C365-C374, 1993[Abstract/Free Full Text].

97.   White, RA, and Meistrich ML. A comment on "A method to measure the duration of DNA synthesis and the potential doubling time from a single sample." Cytometry 7: 486-490, 1986[ISI][Medline].

98.   White, RA, Meistrich ML, Pollack A, and Terry NHA Simultaneous estimation of TG2+M, TS, and Tpot using single sample dynamic tumor data from bivariate DNA-thymidine analogue cytometry. Cytometry 41: 1-8, 2000[ISI][Medline].

99.   White, RA, and Terry NHA A quantitative method for evaluating bivariate flow cytometric data obtained using monoclonal antibodies to bromodeoxyuridine. Cytometry 13: 490-495, 1992[ISI][Medline].

100.   White, RA, Terry NHA, Baggerly KA, and Meistrich ML. Measuring cell proliferation by relative movement. I. Introduction and in vitro studies. Cell Prolif 24: 257-270, 1991[ISI][Medline].

101.   White, RA, Terry NHA, and Meistrich ML. New methods for calculating kinetic properties of cells in vitro using pulse labelling with bromodeoxyuridine. Cell Tissue Kinet 23: 561-573, 1990[ISI][Medline].

102.   White, RA, Terry NHA, Meistrich ML, and Calkins DP. Improved method for computing potential doubling time from flow cytometric data. Cytometry 11: 314-317, 1990[ISI][Medline].

103.   Willett, WC. Dietary fat intake and cancer risk: a controversial and instructive story. Semin Cancer Biol 8: 245-253, 1998[ISI][Medline].

104.   Willett, WC. Diet and cancer: one view at the start of the millennium. Cancer Epidemiol Biomarkers Prev 10: 3-8, 2001[Abstract/Free Full Text].

105.   Zhao, LP, Kushi LH, Klein RD, and Prentice RL. Quantitative review of studies of dietary fat and rat colon carcinoma. Nutr Cancer 15: 169-177, 1991[ISI][Medline].


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