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Molecular and Cellular Biology, January 2007, p. 426-437, Vol. 27, No. 2
0270-7306/07/$08.00+0     doi:10.1128/MCB.01382-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

An Origin of DNA Replication in the Promoter Region of the Human Fragile X Mental Retardation (FMR1) Gene{triangledown} ,{dagger}

Steven J. Gray ,{ddagger},§ Jeannine Gerhardt,{ddagger} Walter Doerfler, Lawrence E. Small, and Ellen Fanning*

Department of Biological Sciences and Vanderbilt-Ingram Cancer Center, Vanderbilt University, Nashville, Tennessee 37235

Received 27 July 2006/ Returned for modification 21 August 2006/ Accepted 30 October 2006


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ABSTRACT
 
Fragile X syndrome, the most common form of inherited mental retardation in males, arises when the normally stable 5 to 50 CGG repeats in the 5' untranslated region of the fragile X mental retardation protein 1 (FMR1) gene expand to over 200, leading to DNA methylation and silencing of the FMR1 promoter. Although the events that trigger local CGG expansion remain unknown, the stability of trinucleotide repeat tracts is affected by their position relative to an origin of DNA replication in model systems. Origins of DNA replication in the FMR1 locus have not yet been described. Here, we report an origin of replication adjacent to the FMR1 promoter and CGG repeats that was identified by scanning a 35-kb region. Prereplication proteins Orc3p and Mcm4p bind to chromatin in the FMR1 initiation region in vivo. The position of the FMR1 origin relative to the CGG repeats is consistent with a role in repeat maintenance. The FMR1 origin is active in transformed cell lines, fibroblasts from healthy individuals, fibroblasts from patients with fragile X syndrome, and fetal cells as early as 8 weeks old. The potential role of the FMR1 origin in CGG tract instability is discussed.


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INTRODUCTION
 
Fragile X syndrome, the most common form of inherited mental retardation in males, is characterized by expansion of a CGG repeat tract in the 5' untranslated region of the human fragile X mental retardation protein 1 (FMR1) gene on Xq27.3 (54, 77, 80). The repeat number expands from 5 to 50 in healthy males to over 200 in affected patients, accompanied by methylation of CpG dinucleotides and silencing of FMR1 gene expression, giving rise to the disease (54, 77, 80). DNA methylation of CpG dinucleotides in the FMR1 promoter is a necessary step in the inactivation of transcription, since FMR1 transcription can be reactivated in fragile X syndrome-affected cells upon treatment with 5-aza-2-deoxycytidine (28, 58). Moreover, rare individuals, whose CGG repeats are expanded but not methylated, still express FMR1 and display normal intelligence (24, 59, 71). The cause of the CGG expansion in the patient's maternal lineage is not known, but cis-acting elements are thought to play an important role (9, 39, 45, 46).

More than 20 genetic diseases are caused by the expansion of trinucleotide repeats, including myotonic dystrophy and Huntington's disease (9, 19, 56). Trinucleotide repeat diseases begin with a "premutation" state when the repeat number becomes unstable for unknown reasons. Through successive generations, as the repeats expand, the symptoms of disease appear and increase in severity (9, 10, 19, 45, 56). Expansion of trinucleotide repeats can thus be divided into two categories: (i) the initial trigger that leads to instability and (ii) the mechanism for expansion once the repeat tract is lengthened and unstable. The mechanism that first initiates repeat instability may be different from the mechanism that propagates repeat lengthening once the repeat tract becomes unstable. The expanded repeat length itself may be enough to create instability, as observed when expanded CGG tracts are integrated at different loci in Saccharomyces cerevisiae and Escherichia coli (6, 29, 39, 69). Instability is thought to occur because expanded repeats form extensive stable secondary structures, such as hairpins and triplex and quadruplex DNA, that are difficult to process correctly during replication, recombination, or repair (48, 57, 63, 69). In contrast, the initial transition from short, stable repeat tracts to longer, unstable ones remains poorly understood.

Some evidence has implicated replicon organization as a potential factor in the initial expansion of a stable repeat tract. Instability of a wild-type-length CAG repeat tract placed within ~350 bp of a simian virus 40 replication origin was observed during viral DNA replication in monkey cells (8). In these experiments, the distance between the repeat tract and the origin, as well as the orientation of the repeats relative to the origin, affected the stability of the repeats. Similar experiments using CGG repeats demonstrated that replication fork dynamics, repeat length, and CpG methylation can affect repeat stability (51). Orientation-dependent repeat instability was also observed when CGG repeats were placed in the E. coli or S. cerevisiae genome near an origin of DNA replication (6, 63, 84). CGG repeats in the lagging strand template favored contractions, while CGG repeats in the newly synthesized Okazaki fragment favored expansions (6, 63, 84). Consistent with the importance of replication direction, trinucleotide repeats can be induced to expand or contract in S. cerevisiae by mutating replication proteins involved in lagging strand, but not leading strand, DNA synthesis (39, 57, 62, 67, 68). These observations point to a potential role for origins of DNA replication in initiating and/or exacerbating trinucleotide repeat length instability.

In this study, we sought to identify origins of DNA replication at sites surrounding the CGG repeats in the human FMR1 locus and to monitor their initiation activity in cells from healthy individuals and individuals with fragile X syndrome. An approximately 35-kb region surrounding the FMR1 promoter and the repeats was investigated. Initiation activity localized in the FMR1 promoter region, and the activity of the FMR1 origin was equivalent to that of two previously characterized origins of DNA replication analyzed in parallel as controls. The FMR1 origin was active in untransformed fibroblasts derived from healthy males and females and from fragile X syndrome-affected males. The potential role of the FMR1 origin in CGG tract instability is discussed.


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MATERIALS AND METHODS
 
Cells and culture conditions. Cells used for analysis of origin activity at the FMR1 locus are described in Table 1. HCT116 and HeLa were grown in Dulbecco modified Eagle medium with 10% heat-inactivated fetal bovine serum (FBS) and 5% CO2 at 37°C. All untransformed fibroblasts were grown in Dulbecco modified Eagle medium with 15% FBS and 5% CO2 at 37°C, except GM05381 (grown with 20% FBS) and HAF (grown in 10% heat-inactivated FBS).


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  		TABLE 1. Cells used for analysis of origin activity at the FMR1 locus

Southern blot analysis of repeat tract length. Genomic DNA was isolated from cells as described previously (3). The isolated genomic DNA was then digested overnight with 10 µg RNase A at 4°C and purified by phenol-chloroform extraction followed by ethanol precipitation. Approximately 50 µg of genomic DNA was digested with PstI (New England Biolabs catalog no. R0140S) and electrophoresed through a 1% agarose gel. The DNA was blotted onto a Hybond N+ or Hybond XL membrane (Amersham catalog no. RPN203B or RPN203S, respectively) and cross-linked using a Stratagene UV Stratalinker 1800. The blot was prehybridized in 15 ml Church buffer (0.5 M Na2HPO4, 1 mM EDTA, 7% sodium dodecyl sulfate [SDS], 1% bovine serum albumin) at 65°C for several hours. Fifty nanograms of a 1-kb PstI fragment from the FMR1 locus that includes the CGG repeats (Fig. 1B) was radiolabeled with 50 µCi [{alpha}-32P]dCTP (3,000 Ci/mmol; Perkin-Elmer LAS catalog no. BLU513H250UC) using the High Prime labeling reagent (Roche catalog no. 11585592001). The radiolabeled probe (~50 ng at 1 x 106 to 2 x 106 dpm/ng) was heat denatured in the presence of 750 µg salmon sperm DNA, added to the prehybridization solution, and hybridized overnight at 65°C. The blot was washed as follows: twice with 2x SSC (1x SSC is 150 mM NaCl plus 15 mM sodium citrate) plus 0.1% SDS for 15 min each time at 25°C, once with 1x SSC-0.1% SDS for 15 min at 25°C, and four times with 0.1x SSC-0.1% SDS for 5 min each time at 65°C. The washed blot was exposed to a phosphorimager screen to detect the radioactive signal. PstI fragment lengths were verified with at least two independent Southern blots for each genomic DNA sample.


Figure 1
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  		FIG. 1. Diagram of the FMR1 gene locus. (A) A 40-kb region containing part of the FMR1 coding sequence and upstream region (GenBank accession no. L29074) indicates the start site for transcription of the FMR1 gene (bent arrow), with the CGG repeats indicated by a gray triangle. Above the scale is the FMR1 open reading frame, with the first 12 of 17 exons shown as vertical bars. Below the scale are the locations of the outlying primer sets (black boxes) used for quantitative PCR (see Table S1 in the supplemental material). (B) A detailed diagram of the FMR1 promoter region, with hatch marks every 100 bp. Black dots indicate the locations of CpG dinucleotides in the sequence. The FMR1 promoter (box with vertical bars), CGG repeat tract (gray box), primary start site for transcription (bent arrow), and exon 1 (white box) are indicated. PCR amplicons used for quantitative PCR (see Table S1 in the supplemental material) are shown below the scale as horizontal bars, and pertinent restriction sites are shown above the scale.

Reverse transcriptase PCR. Actively growing cells (~1 x 106 to 3 x 106) were collected by trypsinization, and total RNA was extracted using the QIAGEN RNeasy Mini kit (catalog no. 74104) according to the manufacturer's instructions. cDNA was synthesized using the Invitrogen SuperScript III reverse transcriptase following the manufacturer's instructions, using 1 µg RNA (determined by absorbance at 260 nm) and 500 ng oligo(dT)15 in a 20-µl reaction mixture at 50°C. Samples (0.5 µl) of the reaction product were then used as a template for real-time quantitative PCR. The FMR1 primer set was designed to cross the exon 13/14 junction (see Table S1 in the supplemental material) so that it specifically amplified cDNA from spliced mRNA, but not genomic DNA which contained the intervening intron. The standard used for this quantitative PCR was serial dilutions of the target PCR amplicon from HCT116 cells. Raw values for real-time PCR quantitation of cDNA for all samples are provided in Table S3 in the supplemental material.

Isolation of nascent-strand-enriched DNA. Nascent DNA was isolated from asynchronously growing cells as described previously (41). Briefly, ~7 x 107 HCT116 cells, ~3 x 107 HeLa S3 cells, or ~1 x 107 to 3 x 107 untransformed fibroblasts were harvested by trypsinization and loaded onto a 1.25% alkaline agarose gel. After a 15- to 20-min alkaline lysis of the cells in the well, the DNA was electrophoresed for 16 h at 30 V. Single-stranded DNA of 0.5 to 1 kb or 1 to 2 kb was cut out of the gel and purified with a QIAGEN gel extraction kit (catalog no. 28706) following the manufacturer's instructions. The DNA was eluted in 10 mM Tris-HCl (pH 8.5) and used directly for quantitative PCR. Between two and four independent preparations of nascent DNA were isolated from each cell type.

Quantitative PCR primer optimization. PCR primers (purified by polyacrylamide gel electrophoresis; Integrated DNA Technologies) were designed to have similar annealing temperatures and to create amplicons approximately 180 to 200 bp long. Each primer was used to amplify 1.2 ng of genomic DNA under quantitative PCR conditions (see Table S4 in the supplemental material) with 2 to 5 mM Mg2+ (final concentration) to empirically optimize the Mg2+ concentration. The annealing temperature of the primers was also varied to obtain a single clean PCR product. Primers were considered optimized when the PCR product melted completely at a single temperature (specific for each product) and when the PCR product gave a single band of the appropriate size when run on an agarose gel. A list of all primers used is provided in Table S1 in the supplemental material. PCR products, melting curve analysis, and examples of standard curves are provided in Fig. S1 in the supplemental material.

Quantitative real-time PCR. Fourteen primer sets for the FMR1 locus (Fig. 1), three primer sets for the lamin B2 origin (21) (see Fig. 4A), and three primer sets for the MCM4 origin (37) (see Fig. 4B) were used to quantitate target sequences in nascent-strand DNA samples. The LightCycler FastStart DNA master SYBR green I kit (Roche catalog no. 12239264001) was used following the manufacturer's instructions. All reactions were carried out on a Roche diagnostic real-time PCR LightCycler. Magnesium concentration and annealing temperature (Ta) were optimized for each primer (see "Quantitative PCR primer optimization" above; also see Table S1 and Fig. S1 in the supplemental material). For each reaction, the cycling parameters were as follows: 10 min at 95°C; 5 cycles of 95°C for 15 s, Ta plus 4°C for 5 s, and 72°C for 15 s; 5 cycles of 95°C for 15 s, Ta plus 2°C for 5 s, and 72°C for 15 s; and 40 cycles of 95°C for 15 s, Ta for 5 s, and 72°C for 15 s. Some CG-rich amplicons used a melting temperature of 96°C or 97°C. At the end of each run, a melting curve analysis was performed in which the PCR products were annealed at 72°C and the temperature was gradually raised to 99°C. In all cases, the PCR products melted in a narrow temperature range, indicating a pure PCR product without detectable nonspecific amplification (see Fig. S1 in the supplemental material).


Figure 4
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  		FIG. 4. Replication initiates at the FMR1 locus in human cell lines. (A and B) Gene exons (thick lines), introns (thin lines), transcriptional start sites (bent arrows), and primer set locations (black boxes) (see Table S1 in the supplemental material) are shown below the scales. The location of the origin of bidirectional replication (OBR) has been mapped approximately in the MCM4/PRKDC locus (37) (A) and at high resolution in the lamin B2 locus (2) (B). Locations in base pairs are shown on the scale relative to GenBank accession no. U63630 for the MCM4 locus (A) and to the OBR for the lamin B2 locus (B). (C) DNA sequence abundance at the indicated regions of the FMR1 locus in four independent HCT116 nascent DNA samples (n = 4). Initiation activity calculations are described in Materials and Methods. Black bars indicate 0.5- to 1-kb nascent DNA, and white bars indicate 1- to 2-kb nascent DNA. For internal standards in each preparation of nascent DNA, initiation activity was measured at the human MCM4 (gray bars) and lamin B2 loci (checkered bars). The sample that was not tested (*) is indicated. (D) Abundance of the indicated FMR1 sequences in 0.5- to 1-kb nascent DNA from two independent HeLa S3 cell samples (n = 2). The bars in panels C and D indicate the average values plus standard deviations (error bars) for the indicated number of nascent DNA preparations (n).

To quantitate the abundance of specific target sequences in nascent DNA, standard curves of amplification were done using twofold serial dilutions of XhoI/NarI-digested HCT116 genomic DNA (50 ng to 0.1 ng). This range of genomic DNA included copy numbers of target sequence equivalent to those in the nascent DNA samples. Genomic HCT116 DNA digested with NarI only was used as a standard for lamin B2 amplicons, since XhoI cuts within two of the amplicons used. Amplification profiles obtained from target sequences in nascent DNA samples were compared to the standard curve of cut HCT116 genomic DNA to quantitate the abundance of each target sequence in the nascent DNA sample.

Each PCR was done in duplicate, and the average values from the two reactions with every nascent DNA sample with every primer set are given in Table S4 in the supplemental material. For each nascent DNA preparation, relative nascent DNA abundance (initiation activity) was calculated by dividing the amount of target sequence in nascent DNA at each primer set by the amount of target sequence in nascent DNA at the outlying primer set(s) at that locus. Specifically, all values for the control MCM4 primer sets were divided by the EX6b value, all values for control lamin B2 primer sets were divided by the LB2C1 value, and all values for FMR1 primer sets were divided by the average of the values from primer sets 10, 3, and 20 (see Table S4 in the supplemental material for formulas and see Fig. 4C for an example). Initiation activities for like primer sets in different nascent DNA preparations of the same cell type were averaged, and standard deviation was calculated on the basis of the number (n) of nascent DNA preparations with that cell type.

Chromatin immunoprecipitation (ChIP). A total of 1 x 108 HCT116 cells were washed with phosphate-buffered saline (PBS) and treated with 1% formaldehyde in prewarmed medium for 5 min at 37°C. Nuclei were prepared by a modified version of the protocol of Mendez and Stillman (44). Briefly, cells were harvested, washed with PBS, and resuspended in 4 ml hypotonic buffer A (10 mM HEPES, pH 7.9, 10 mM KCl, 1.5 mM MgCl2, 0.34 M sucrose, 10% glycerol, 1 mM dithiothreitol, protease inhibitor). Cells were lysed by adding 0.04% Triton X-100 and incubated for 10 min on ice. Samples were centrifuged (4 min, 1,300 x g, 4°C). Nuclei were washed in ice-cold buffer A supplemented with 200 mM NaCl. After centrifugation (1,300 x g, 5 min, 4°C), fixed nuclei were washed with PBS, resuspended in 2.7 ml LSB (10 mM HEPES, pH 7.9, 10 mM KCl, 1.5 mM MgCl2) and lysed by adding 300 µl of 20% Sarkosyl. The chromatin was transferred onto a 40-ml sucrose cushion (LSB plus 100 mM sucrose) and centrifuged (10 min, 4°C, 2,500 x g). Supernatant was removed, and the chromatin was resuspended in 4 ml 10 mM Tris-1 mM EDTA and sonicated (Branson sonifier 250-D, output setting 5, 50 1 s pulses, twice). For partial DNA digests, 3 mM CaCl2 and 10 U micrococcal nuclease (Roche) per 1 mg chromatin were added and incubated for 10 min at 37°C. The reaction was stopped by adding 20 mM EDTA. For immunoprecipitation, 1/10 volume of 11x NET (550 mM Tris-HCl, pH 7.4, 1.65 M NaCl, 5.5 mM EDTA, 5.5% NP-40) was added to the extract followed by 10 µg affinity-purified polyclonal antibodies (HsOrc3p and HsMcm4p) or 10 µg rabbit immunoglobulin G (IgG) for a control. The immunoprecipitation and purification of coprecipitated DNA were performed as described previously (66).

Real-time PCR analysis was performed according to the manufacturer's instructions (Roche) using the same parameters and primer pairs described above and in Table S1 in the supplemental material. "Enrichment" of immunoprecipitated DNA is defined as the amount of target sequence detected in the specific Orc3p or Mcm4p immunoprecipitate minus the amount of target sequence detected by a nonspecific IgG immunoprecipitate, divided by the amount of target sequence detected in 30 ng of DNA purified from the preimmunoprecipitation chromatin preparation (65).


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RESULTS
 
CGG repeat tract length and FMR1 transcription. A panel of human tumor cell lines and primary cultures from healthy individuals and individuals with fragile X syndrome (Table 1) was initially characterized by Southern blotting to estimate the length of the CGG repeat tract in the FMR1 locus. A PstI fragment containing the FMR1 promoter region and the normal 5 to 50 CGG repeats has a length of ~1 kb (Fig. 1), whereas a full-mutation allele containing >200 repeats has a length of over 1.5 kb (59, 85). Blots of PstI-digested genomic DNA from HeLa S3, HCT116, HAF, and GM05381 cells all displayed a PstI fragment of approximately 1 kb that hybridized with FMR1 promoter sequences, indicating a normal repeat length of ~30 repeats as previously reported for HCT116 cells (17) (Fig. 2A, lanes 1 to 5 and 9). The fragile X genomic DNA (GM07072, GM05848, and GM04026 cells) displayed PstI fragments of 2 to 4 kb (Fig. 2A, lanes 6 to 8 and 10). The PstI fragment lengths for GM05848 and GM04026 cells were not uniform, suggesting incomplete digestion, repeat instability, and/or mosaicism in the length of the repeat tract in the cell population. Despite the heterogeneity, these results confirm CGG repeat tract expansions in the fragile X genomic DNA.


Figure 2
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  		FIG. 2. CGG tract length and FMR1 transcription. (A) Total genomic DNA from the indicated cell cultures was digested with PstI and analyzed by Southern blotting, using the radiolabeled 1-kb PstI promoter fragment (Fig. 1B) as the probe. Lanes 1 to 5 and 9, unaffected cells with normal-length CGG repeats; lanes 6 to 8 and 10, fragile X syndrome-affected cells with expanded repeat lengths. (B) One microgram of total RNA from the indicated cells was electrophoresed through an agarose gel and stained with ethidium bromide. Lanes M, molecular size markers (in base pairs). (C) Equal amounts of total RNA from the indicated cells were reverse transcribed to cDNA by reverse transcriptase PCR, and equal amounts of cDNA were used as template in quantitative, real-time PCR with the primer set FMR1 13/14 (see Table S1 in the supplemental material). The bars indicate the average values plus standard deviations (error bars) for three independent sample preparations for each cell type.

The expanded CGG tract observed in the cells from individuals with fragile X syndrome (Fig. 2A, lanes 6 to 8 and 10) predicts that FMR1 expression should be silenced (54, 77). To monitor FMR1 gene expression in the fragile X patient cells, we measured the level of FMR1 transcription by quantitative reverse transcriptase PCR. Equal quantities of RNA from all cells tested were used for cDNA synthesis, as determined by absorbance at 260 nm and visualization by agarose gel chromatography (Fig. 2B; see Table S3 in the supplemental material). FMR1 mRNA was detected at similar levels in HCT116 cells and in the untransformed fibroblasts from healthy males and females (Fig. 2C). FMR1 mRNA levels were suppressed by 2 to 3 orders of magnitude in the fragile X cells with expanded CGG tracts relative to the levels in healthy cells (Fig. 2C), supporting the prediction that FMR1 transcription is silenced (54, 77, 80).

Characteristics of primer amplification in the FMR1 locus. Primers were developed to amplify target sequences over a ~35-kb region of the FMR1 locus (see Table S1 in the supplemental material). By changing magnesium concentrations and temperature, primer usage was optimized to specifically amplify each target sequence (see Fig. S1 in the supplemental material). However, initial characterization of primer efficiency on genomic DNA revealed that primers located near the CGG repeat tract displayed lower efficiencies of amplification than those located farther from the repeats (see Table S2 in the supplemental material; also data not shown). These preliminary results raised the question of whether the CGG repeats might suppress the amplification efficiency of flanking sequences, even though the amplicon did not directly contain the repeats. To test this possibility, we reasoned that if the amplicon template were on a separate DNA molecule, its efficiency of amplification should be greater than when it was flanked by the repeat tract. To separate the repeats from the amplification templates, genomic HCT116 DNA was cut with XhoI (which cuts immediately downstream of the repeats), with NarI (which cuts immediately upstream of the repeats), or with both NarI and XhoI (Fig. 1B). PCR amplification was carried out with five primer sets on each of these genomic DNAs. The NarI/XhoI-digested DNA was used to generate the standard curve for these experiments (Fig. 3). PCR amplification using undigested genomic DNA as the template with primer sets 1c and 11 was clearly lower than that with the other three primer sets (Fig. 3A). When XhoI-digested DNA was used as the template, separating the primer set 11 sequence from the CGG tract, amplification with primer set 11 increased to a level comparable to that seen with the other three CGG-distal primer sets (Fig. 3B). Conversely, when NarI-digested DNA was the template, separating primer set 1c from the CGG tract, amplification with primer set 1c increased (Fig. 3C). These results demonstrate that the physical linkage of the CGG tract with the template DNA reduced the amplification efficiency of the flanking target sequences and that eliminating the linkage with the CGG repeats alleviated the reduction in amplification efficiency. On the basis of these results, HCT116 genomic DNA digested with XhoI and NarI was chosen as the calibration standard to quantitate the amplification of target sequences in nascent DNA samples.


Figure 3
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  		FIG. 3. Flanking (CGG)n repeats reduce PCR amplification efficiency. Genomic HCT116 DNA either undigested (A) or digested with NarI (B) or with XhoI (C) was used as the template for real-time PCR amplification with the indicated primer sets in the FMR1 promoter region. Amplification was calibrated using known amounts of NarI/XhoI-digested genomic DNA to generate the standard curves. The arrows indicate the locations of XhoI (B) and NarI (C) restriction sites relative to primer locations and the CGG repeat tract.

Detection of the FMR1 origin of DNA replication in transformed cells. In order to measure potential initiation activity near the CGG repeats of the FMR1 locus, a fraction enriched in short, newly synthesized, single-stranded DNA was prepared from HCT116 cells, which are derived from a male individual and therefore have only one copy of the FMR1 locus per cell. As internal controls for each preparation of nascent DNA, real-time quantitative PCR was used to measure relative levels of target sequences at the well-characterized MCM4 and lamin B2 origins of DNA replication (2, 11, 21, 33, 35, 37, 55, 70, 75, 79) (Fig. 4A and B). The MCM4 and lamin B2 origins showed easily detectable initiation activity in HCT116 cells at origin-proximal sequences, well above that at distal sequences (Fig. 4C).

Real-time quantitative PCR was then used with multiple FMR1 primer sets to measure relative levels of target sequences in the same preparations of nascent DNA at sites across the FMR1 locus. In HCT116 nascent DNA, a peak of abundance was detected at primer set 1.4 adjacent to the FMR1 promoter relative to outlying primer sets, using both the 0.5- to 1-kb and the 1- to 2-kb nascent DNA preparations as templates (Fig. 4C). The level of initiation activity at the FMR1 promoter was comparable to that at the MCM4 and lamin B2 origins in the same preparations of nascent DNA. The pattern of initiation activity at the FMR1 locus in HCT116 nascent DNA was identical whether XhoI/NarI-digested genomic DNA (Fig. 4C) or a XhoI/NarI-digested plasmid, containing the cloned HCT116 FMR1 sequence, was used as the real-time PCR calibration standard (data not shown).

Measurements of the PCR efficiency of the HCT116 nascent DNA compared to that of the XhoI/NarI-cut genomic HCT116 DNA standard showed that these two templates had nearly identical efficiencies with primer sets EX6b, UPR4, 3, 1.4, and 20, validating the peaks of nascent DNA abundance at the MCM4 and FMR1 origins (see Table S2 in the supplemental material). However, the amplification efficiency of primer sets within the FMR1 promoter (1.2, 1c, and 1d) was lower than with the cut genomic DNA used as a quantitation standard (see Table S2 in the supplemental material), resulting in an underestimate of the 1.2, 1c, and 1d target sequences in the nascent DNA. Thus, the peak of FMR1 initiation activity detected with primer set 1.4 may actually extend into the promoter, closer to the CGG repeats (Fig. 1).

To determine whether replication initiated at the FMR1 origin in other cells, HeLa S3 nascent DNA was also tested. Since the peak of nascent DNA with primer set 1.4 was observed with both HCT116 nascent DNA size classes, only the 0.5- to 1-kb nascent DNA fraction was tested. The initiation activity at MCM4 and lamin B2 origin-proximal sequences in two HeLa S3 nascent DNA preparations was lower than that in HCT116 nascent DNA preparations, but it was still greater than that at distal sequences (Fig. 4D). Initiation activity was detected with several closely spaced primer sets in the FMR1 promoter region (Fig. 4D). Even though the level of initiation activity in the FMR1 origin was lower in HeLa S3 cells than in HCT116 cells, it was similar to the initiation activities seen at the MCM4 and lamin B2 control origins in the same preparations of nascent DNA. Relative to the control origins, the FMR1 initiation activity was equivalent in that in HCT116 and HeLa S3 cells.

Orc and Mcm proteins associate with the FMR1 origin in vivo. During the G1 phase of the cell cycle, licensed replication origins are bound by prereplication proteins, such as Orc1-6p and Mcm2-7p (1, 34, 37, 65, 79). If an origin of DNA replication exists in or near the promoter of the FMR1 gene, as suggested by nascent DNA abundance assays (Fig. 4), prereplication complexes would be expected to assemble in this region. To test this prediction, we used a ChIP assay. For a control, we first confirmed enrichment of Orc3p and Mcm4p at the MCM4 origin (UPR4 primer pair) relative to a distal control region (Ex6 primer pair) (Fig. 5A). In the FMR1 locus, the greatest enrichment of Orc3p was detected with primer set 1d at the FMR1 promoter region and slightly lower enrichment with primer set 1.5 (Fig. 5B). Orc3p bound to origin-distal chromatin was detected with the outlying primer sets 3, 10, 16, and 30, but at significantly lower levels. With primer pairs 5 and 11, no Orc3p binding to chromatin was observed. Consistent with these findings, the greatest binding of Mcm4p in the FMR1 locus was detected with primer set 1d, with lower enrichment with the other primer sets (Fig. 5C). The specific binding of Orc3p and Mcm4p within the FMR1 promoter is consistent with the existence of an origin of DNA replication in this region.


Figure 5
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  		FIG. 5. Orc3p and Mcm4p localize to the FMR1 promoter in vivo. A ChIP assay was done on three independently prepared samples from asynchronously growing HCT116 cells for Orc3p (A and B) and Mcm4p (A and C). (A) "Enrichment" for Orc3p and Mcm4p, as described in Materials and Methods, was tested using primer sets at the MCM4 origin (Fig. 4B; see Table S1 in the supplemental material). Enrichment was measured at the MCM4 origin (primer set UPR4) for Orc3p and Mcm4p relative to the outlying primer set EX6b. Enrichment for Orc3p (B) and Mcm4p (C) was tested using primer sets at the FMR1 locus (Fig. 1; see Table S1 in the supplemental material), with peaks of enrichment detected at primer set 1d in the FMR1 promoter. The bars indicate the average values plus standard deviations (error bars) for multiple independent immunoprecipitates.

FMR1 origin activity in untransformed fibroblasts. The activity of the FMR1 origin was measured in nascent DNA isolated from untransformed human fibroblasts to test whether the initiation site detected in the HCT116 tumor cell line is also utilized in cells from healthy humans. The peak initiation activity detected at the FMR1 origin in either HAF or GM05381 nascent DNA was equivalent to that at the MCM4 and lamin B2 control origins (Fig. 6A and B), suggesting that the FMR1 origin is fully active in these healthy male cells. The FMR1 origin in healthy female cells (GM08400) showed initiation activity at closely spaced primer sets 1.5, 1.4, and 1c in the promoter region (Fig. 6C). The level of FMR1 initiation activity in the female cells was nearly as high as that of the MCM4 control origin.


Figure 6
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  		FIG. 6. Replication initiates at the FMR1 origin in healthy male and female fibroblasts. As described in the legend to Fig. 4 and in Materials and Methods, initiation activity was tested with primer sets at the FMR1 locus in independent nascent DNA preparations from HAF male cells (n = 2) (A), GM05381 male cells (n = 4) (B), GM08400 female cells (n = 2) (C), GM00011 male fetal cells (n = 2) (D), GM01603 male fetal cells (n = 2) (E), and GM06170 male fetal cells (n = 2) (F). Initiation activity was tested in the same nascent DNA preparations using primer sets from the MCM4 (gray bars) and lamin B2 (checkered bars) control origins. The bars indicate the average values plus standard deviations (error bars) for the indicated number of nascent DNA preparations (n).

To determine whether a change in FMR1 origin usage could be observed during fetal development, we tested the activity of the origin in untransformed fetal fibroblasts. The FMR1 origin was active in cells from three male individuals at 8, 12, and 20 fetal weeks (GM00011 in Fig. 6D, GM01603 in Fig. 6E, and GM06170 in Fig. 6F). The FMR1 origin activity in these cells was similar to that in adult male fibroblasts (Fig. 6A and B). However, initiation activity at the MCM4 and lamin B2 origins was lower.

FMR1 origin activity in fragile X syndrome-affected cells. FMR1 origin activity was tested in cells from five individuals affected with fragile X syndrome to determine whether the site or level of activity differed from that observed in healthy cells. Importantly, the same site of initiation of DNA replication in the FMR1 locus was used in cells from all five affected individuals (Fig. 7). The levels of initiation activity at the FMR1 origin in the two postnatal patient cell lines, GM05848 and GM04026, were similar (Fig. 7A and B) to that observed in healthy male fibroblasts (Fig. 6A and B). However, initiation activity at the MCM4 and lamin B2 origins was lower in the fragile X cells than in the healthy male cells. These comparisons suggest that the FMR1 origin is at least as active in the adult fragile X cells as in the healthy male cells and perhaps more active.


Figure 7
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  		FIG. 7. Replication initiates at the FMR1 origin in fragile X cells. As described in the legend to Fig. 4 and in Materials and Methods, initiation activity was tested with primer sets at the FMR1 locus in independent nascent DNA preparations from GM05848 male patient cells (n = 2) (A), GM04026 male patient cells (n = 2) (B), GM07730 male fetal patient cells (n = 3) (C), GM05282 male fetal patient cells (n = 3) (D), and GM07072 male fetal patient cells (n = 3) (E). Initiation activity was tested in the same nascent DNA preparations using primer sets from the MCM4 (gray bars) and lamin B2 (checkered bars) control origins. The bars indicate the average values plus standard deviations (error bars) for the indicated number of nascent DNA preparations (n).

The FMR1 origin activity in fetal cells from two individuals with fragile X syndrome, one 20 weeks old (GM07730) and one 21 weeks old (GM05282), was approximately equivalent to that in the postnatal fragile X patient cells (Fig. 7C and D). As in the postnatal patient cells, the MCM4 and lamin B2 control origin activities were low. Unexpectedly, in three independent preparations of nascent-strand-enriched DNA from fibroblasts from a 22-week-old fetus with fragile X syndrome, GM07072, initiation activity at the FMR1 origin was reproducibly lower than in any of the other cells tested (Fig. 7E). In contrast, GM07072 cells displayed strong initiation activity at the MCM4 and lamin B2 origins, comparable to that observed in healthy cells (compare Fig. 7E to Fig. 6). These controls argue against the possibility that the low activity of the FMR1 origin in these nascent DNA fractions was due to poor nascent strand enrichment or slow cell growth. We conclude that FMR1 origin activity relative to that of the control origins was reduced in the fragile X GM07072 fibroblasts, but this reduction is not a general feature of fetal fragile X cells.


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DISCUSSION
 
Detection of a novel origin of DNA replication at the FMR1 promoter. We have mapped a peak of replication initiation in the promoter region of the human FMR1 gene that displays initiation activity comparable to those of known origins tested in the same nascent DNA preparations (Fig. 4 and 6). The colocalization of Orc and Mcm proteins (Fig. 5) with the peak of initiation activity further supports the identification of the FMR1 origin of DNA replication. The lack of nascent DNA enrichment at primer sets distal from the FMR1 origin suggests that this is the primary initiation site within a 35-kb region flanking the FMR1 promoter and CGG repeats (Fig. 4C and data not shown). These features of the FMR1 origin identify it as a new member of a class of origins that reside near CpG islands and the promoters of housekeeping genes (5, 12, 15, 22, 42, 78).

The low efficiency of PCR amplification at target sequences flanking the CGG repeat tract limited the region of accurate PCR quantitation of target sequences in genomic DNA but was alleviated by cleaving the template away from the repeats (Fig. 3). Since the same limitation applies to quantitation of CGG-proximal target sequences in the single-stranded nascent DNA fraction (see Table S2 in the supplemental material), the abundance of target sequences detected by primer sets 1.2, 1c, and 1d is somewhat underestimated. Hence, the peak of initiation activity may include these target sequences in the promoter (Fig. 4C and D and 6 and 7). Our results illustrate the effects that unequal PCR efficiencies can have on the accuracy of quantitation, warranting caution in interpreting peaks of nascent DNA abundance near sequences such as trinucleotide repeats.

Differential regulation of the FMR1 promoter and FMR1 origin in fragile X patient cells. The FMR1 promoter region has a complex architecture of bent DNA, CG-rich tracts, and a (CAAAC)6 tract and occupancy by multiple transcription factors (18, 28, 36, 40, 72, 73) and replication factors (Fig. 5). The proximity of the promoter to the origin suggests that features of the promoter may influence FMR1 origin activity, but this remains to be determined. Methylation of CpG dinucleotides (18, 54, 80), remodeling of chromatin (18, 28), and loss of transcription factor binding (72, 73) have been reported in the promoter of somatic cells from individuals with fragile X syndrome. These alterations result in silencing of FMR1 gene expression in patient cells (Fig. 2B), yet the FMR1 origin activity is maintained in fragile X cells at levels comparable to those in healthy male cells (Fig. 6 and 7). These findings imply that FMR1 origin activity is regulated differently than that of the adjacent promoter.

The transcriptionally silent FMR1 locus in patients with fragile X syndrome replicates later in S phase than does the active FMR1 locus in unaffected individuals (25-27, 74, 87). Since our data show that the FMR1 origin was active in GM04026 cells (Fig. 7B), and these cells were previously shown to have delayed replication in the fragile X locus (26), transcriptional silencing appears to be correlated with a delay in replication timing at the FMR1 origin, but not FMR1 origin usage.

The position and usage of the FMR1 origin have implications for CGG repeat stability. A replication fork emanating from the FMR1 origin would replicate the CGG repeats such that the CGG sequence is in the lagging strand template (Fig. 8, top). The CGG repeats form stable DNA secondary structures, such as hairpins, more readily than CCG repeats (39, 45-48, 88). When hairpin-forming trinucleotide repeats are located in the lagging strand template, as in the FMR1 locus, contractions are favored over expansions (9, 39, 45, 57, 61-63, 81, 84). Thus, the location of the FMR1 origin relative to the CGG repeat tract suggests that it might normally favor contractions when replicating unstable repeats.


Figure 8
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  		FIG. 8. Model of replication fork movement originating from the FMR1 origin. Black lines represent the template strand, and gray arrows indicate the daughter leading and lagging strands. A dashed line indicates the transition point between leading and lagging strand synthesis. The lagging strand template thus contains the CGG repeats within 600 bp downstream of the initiation site. CGG more readily forms secondary structures than CCG, and DNA secondary structure formation in the lagging strand template is implicated in repeat instability, favoring contractions (39, 45, 46).

Since the direction of DNA replication can also play a role in expansion of the CGG repeats (9, 39, 45, 57, 61-63, 81, 84), one can hypothesize that a change in the pattern of origin usage in the FMR1 locus to an origin downstream of the repeats, such that the CGG repeats are in the nascent Okazaki fragments, could lead to repeat expansion (46). The expansions seen in patients with fragile X syndrome occur through maternal transmission, and the repeat instability associated with transmission of the fragile X allele is mostly specific to gametogenesis and/or early embryogenesis (9, 43, 49). During early development, the pattern of origin usage in humans is not known, but in model systems such as Xenopus and Drosophila, origin usage is relatively nonspecific during early development (32, 38, 64, 83). A different pattern of origin usage during early human development might provide a time window in which the repeats would escape the stable or potentially contraction-favoring pattern of replication offered by the FMR1 origin reported here, and therefore become more susceptible to repeat expansion. One of three fetal patient samples used in this study, GM07072, was derived from a 22-week-old fetus and showed reduced usage of the FMR1 origin, in relation to the MCM4 and lamin B2 control origins (Fig. 7E). However, this reduction of FMR1 origin activity was not detected in other fetal cells from healthy individuals or individuals with fragile X syndrome. Thus, if a development-specific change in FMR1 origin usage is generally involved in the initial expansion of CGG repeats, it would need to occur before 8 weeks, the age of the earliest healthy fetal cells that we tested (GM00011 [Fig. 6D]).

Other mechanisms known to cause transient changes in the pattern of origin usage are depletion of nucleotide pools during DNA replication (4, 14), global changes in histone acetylation (35), and agents that induce replication stress (16, 82). Moreover, developmentally regulated chromatin reorganization at the mouse IgH, mouse HoxB, and chicken beta-globin loci are accompanied by changes in the pattern of origin usage (13, 23, 52, 53). If a dormant origin on the downstream side of the CGG repeat tract became transiently activated, its position would tend to favor repeat expansion.

Since the position of the FMR1 origin would be predicted to favor CGG tract contractions in FMR1 patients (10, 39, 45, 46, 57, 62), why are they not more commonly observed in somatic cells of patients? An emerging role for CpG methylation in the stabilization of CGG repeats may give clues to this question. Methylation stabilizes CGG repeats in E. coli (50) and in primate cells in culture (51). The timing of CGG repeat instability, prior to germ line segregation in early human development, coincides with the period in which the DNA lacks epigenetic modifications such as CpG methylation (9, 43). After this time, the repeats are stable and methylated, except in the case of the male testes and sperm, which are unmethylated and display contractions (43, 76). An exception occurs in some high-functioning fragile X patients who display mosaicism for repeat length and methylation (24, 71). Hypermethylated fragile X cells display homogeneous and stable repeat lengths, but unmethylated repeats are heterogeneous for repeat length and stability (9, 20, 85, 86). Since the position of the FMR1 origin favors contraction events, the hypomethylation and consequent instability of the expanded repeats in spermatocytes may provide a mechanism by which the repeats contract in these cells. This could occur through normal DNA replication from the FMR1 origin. Postnatal somatic cells in the same individual would have hypermethylated, stable repeats that would resist contraction during normal DNA replication.

In summary, the results presented here indicate that an origin of DNA replication exists in the promoter region of the FMR1 gene and that this origin is active in somatic cells with either normal-length or expanded CGG repeats, regardless of FMR1 transcription. The discovery and characterization of the FMR1 origin opens the possibility of exploring the relationship of a trinucleotide repeat sequence with its native origin of DNA replication.


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ACKNOWLEDGMENTS
 
We thank C. Pearson and J. Cleary for stimulating discussions in the early phase of this project. We also thank G. Liu, D. Schaarschmidt, and R. Knippers for advice and reagents, X. Qin for help optimizing PCR conditions, and L. J. Zwiebel for LightCycler access. We acknowledge the Coriell Institute for Medical Research, T. R. Peters, P. Wright, and M. Ikizler for providing cells used in this study.

The financial support of NIH (GM 52948 and training grant 5 T32 CA09385-20), the Howard Hughes Medical Institute, Vanderbilt University, and Deutsche Akademische Austauschdienst (Kurzzeit-Dozentur to W.D.) is gratefully acknowledged.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biological Sciences, Vanderbilt University, VU Station B 351634, Nashville, TN 37235-1634. Phone: (615) 343-5677. Fax: (615) 343-6707. E-mail: ellen.fanning{at}vanderbilt.edu. Back

{triangledown} Published ahead of print on 13 November 2006. Back

{dagger} Supplemental material for this article may be found at http://mcb.asm.org/. Back

{ddagger} Co-first authors. Back

§ Present address: Gene Therapy Center, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599. Back

Permanent address: Institute for Clinical and Molecular Virology, Universität Erlangen, Schlossgarten 4, 91054 Erlangen, Germany. Back


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Molecular and Cellular Biology, January 2007, p. 426-437, Vol. 27, No. 2
0270-7306/07/$08.00+0     doi:10.1128/MCB.01382-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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