Defective replication initiation results in locus specific chromosome breakage and a ribosomal RNA deficiency in yeast

Joseph C Sanchez; Elizabeth X Kwan; Thomas J Pohl; Haley M Amemiya; M K Raghuraman; Bonita J Brewer

doi:10.1371/journal.pgen.1007041

. 2017 Oct 16;13(10):e1007041. doi: 10.1371/journal.pgen.1007041

Defective replication initiation results in locus specific chromosome breakage and a ribosomal RNA deficiency in yeast

Joseph C Sanchez ^1,², Elizabeth X Kwan ², Thomas J Pohl ^1,², Haley M Amemiya ², M K Raghuraman ², Bonita J Brewer ^1,^2,^*

Editor: Julian E Sale³

PMCID: PMC5658192 PMID: 29036220

Abstract

A form of dwarfism known as Meier-Gorlin syndrome (MGS) is caused by recessive mutations in one of six different genes (ORC1, ORC4, ORC6, CDC6, CDT1, and MCM5). These genes encode components of the pre-replication complex, which assembles at origins of replication prior to S phase. Also, variants in two additional replication initiation genes have joined the list of causative mutations for MGS (Geminin and CDC45). The identity of the causative MGS genetic variants strongly suggests that some aspect of replication is amiss in MGS patients; however, little evidence has been obtained regarding what aspect of chromosome replication is faulty. Since the site of one of the missense mutations in the human ORC4 alleles is conserved between humans and yeast, we sought to determine in what way this single amino acid change affects the process of chromosome replication, by introducing the comparable mutation into yeast (orc4^Y232C). We find that yeast cells with the orc4^Y232C allele have a prolonged S-phase, due to compromised replication initiation at the ribosomal DNA (rDNA) locus located on chromosome XII. The inability to initiate replication at the rDNA locus results in chromosome breakage and a severely reduced rDNA copy number in the survivors, presumably helping to ensure complete replication of chromosome XII. Although reducing rDNA copy number may help ensure complete chromosome replication, orc4^Y232C cells struggle to meet the high demand for ribosomal RNA synthesis. This finding provides additional evidence linking two essential cellular pathways—DNA replication and ribosome biogenesis.

Author summary

The origin recognition complex (ORC) is essential for licensing replication origins during M/G1 for their firing in the subsequent S phase. Individuals with a rare form of dwarfism called Meier-Gorlin syndrome (MGS) have mutations in proteins required for origin activation, including various subunits of ORC. To better understand the molecular and cellular consequences of these mutations, we introduced an equivalent MGS mutation in ORC4 into yeast. We find that origin activity in the ribosomal DNA (rDNA) repeats is severely compromised in yeast that harbor the MGS allele. Consequently, cells that have reduced their rDNA copy number from ~150 to fewer than 30 copies overtake the culture. Although the loss of rDNA repeats helps ensure the complete replication of chromosome XII during S phase, cells with fewer rDNA repeats struggle to meet the high demand for ribosomal RNA.

Introduction

The faithful and timely duplication of a cell’s genome is required every round of division. During eukaryotic S phase, DNA replication initiates at multiple sites along each chromosome called origins of replication. Eukaryotic replication initiation has been best characterized in the budding yeast Saccharomyces cerevisiae, where chromosomal origins were first identified by their ability to maintain recombinant plasmids after transformation into yeast [1]. The majority of these Autonomous Replication Sequences or ARS elements correspond to the ~300 chromosomal origins of replication that are scattered across the genome and share a core consensus sequence called the ACS (ARS consensus sequence) [2]. Subsequent biochemical and genetic work in yeast identified many of the essential genes for replication initiation [3–5]. Features that define origins in higher eukaryotes differ significantly from yeast ARSs, but the proteins that carry out origin recognition and initiation are strikingly conserved in sequence and structure across eukaryotes (S1 Fig) [6–9].

In budding yeast, a six-membered protein complex called the Origin Recognition Complex (Orc1-6) binds ARSs throughout the cell cycle [10]. To become competent (or licensed) for initiation, additional proteins are recruited by ORC during the M and G1 phases [11]. The first licensing factor to bind is Cdc6, which facilitates the recruitment of the Mcm2-7 helicase component through an interaction with Cdt1 [9,12–16]. Collectively, this protein complex is known as the Pre-Replication Complex (Pre-RC) and, once assembled on an origin, licenses it to initiate DNA replication or “fire” in the subsequent S phase. During the onset of S phase CDK- and DDK-dependent phosphorylation events complete assembly of the replisomes, including two helicase complexes, allowing replication to proceed bi-directionally from the origin of replication [17–19].

As chromosome replication is essential for cell division, there has been a tacit assumption that mutations that impair the function of proteins involved in DNA replication would be incompatible with metazoan life. Yet, researchers reported in 2011 that amino acid substitutions in proteins involved in the initiation of DNA replication, proteins first identified in S. cerevisiae, are responsible for a form of proportionate dwarfism called Meier-Gorlin syndrome (MGS) [20,21]. Individuals with MGS have phenotypes that include short stature, small external ears and missing or underdeveloped kneecaps [22,23]—phenotypes not obviously associated with chromosome replication defects.

The specific genetic variants found in patients with MGS include homozygous or compound heterozygous alterations in six different Pre-RC genes (ORC1, ORC4, ORC6, CDT1, CDC6, and MCM5) [20,21,24]. Recent work has identified de novo autosomal dominant mutations in Geminin (encoded by GMNN), an inhibitor of DNA replication that is unique to higher eukaryotes [25]. Additionally, biallelic mutations in CDC45, which is required for both origin initiation and elongation during S phase, have been found to be causative for some cases of MGS [26]. Considering the known roles of these proteins in origin initiation, a reasonable hypothesis is that these mutations are adversely affecting DNA replication and thus reducing cell proliferation so that individuals harboring these variants are uncommonly small. Consistent with this hypothesis, previous work using Epstein-Barr virus replication as an assay has found that immortalized fibroblasts and cultured lymphoblastoid cells derived from MGS patients are diminished in their ability to initiate replication [27,28]; however, defects in replication initiation did not always correlate with slowed S phase in these cells [27]. Additionally, MGS mutations have been shown to affect aspects of cell biology other than DNA replication, such as centrosome duplication and cilia formation [27,29]. The defective cilia formation phenotype observed in MGS cells is thought to contribute to some of the developmental abnormalities associated with this condition [27]. Although the proteins linked with MGS have been studied extensively in yeast and other eukaryotes, it is not clear how MGS mutations might affect chromosome replication to give rise to the phenotypes observed in humans. Therefore, understanding how MGS mutations affect chromosome replication may shed light on how they contribute to the phenotypes in humans.

In this study, we have replaced the genomic copy of the budding yeast ORC4 with a mutated version (orc4^Y232C) bearing a tyrosine-to-cysteine change that is orthologous to the Tyr174Cys mutation reported in human patients (S2A and S2B Fig) [21]. We find that yeast cells bearing this orc4^Y232C allele have a longer cell cycle time that is mostly accounted for by a lengthened S phase. Additionally, we find that in orc4^Y232C cells more than 85% of the earliest firing origins are unaltered in their time and/or efficiency; however, the origins present in each copy of the ribosomal DNA (rDNA) are severely compromised in their ability to fire and the number of copies of the rDNA repeat drops from ~150 to as few as 10. Previous work with ORC1 and ORC2 temperature sensitive mutants also revealed shrinkage of the rDNA; however, that work did not provide a model for how the copy number reduction occurred and proposed a checkpoint control of genome-wide replication initiation as an explanation for loss of rDNA repeats [30]. Our findings with the orc4^Y232C allele reveal that the mechanism for rDNA copy number loss is chromosome XII breakage as a consequence of the “random replication gap” problem and that insufficient replication initiation outside of the rDNA locus is not likely the cause for rDNA shrinkage. Furthermore, we show that the reduction in rDNA copy number, by restricting rRNA synthesis, constrains the translational capacity, possibly explaining the slow growth observed in orc4^Y232C cells. While it remains to be seen whether these phenotypes are also common to Meier-Gorlin patient cells, our characterization of the orc4^Y232C allele in S. cerevisiae highlights an unsuspected pathway linking replication dysfunction and growth control.

Results

The slow growth of yeast with the orc4^Y232C allele is due to a delay in completing S phase

A missense mutation in ORC4 has been shown to be causative for some instances of MGS in humans [21]. This specific mutation results in an amino acid substitution (Tyrosine to Cysteine) at position 174 of the human Orc4 protein and occurs in a region with homology to the AAA+ (ATPases Associated with diverse cellular Activities) related domain of the S. cerevisiae Orc4p [21]. The initial work investigating the equivalent MGS mutation (orc4^Y232C) in S. cerevisiae revealed a slow-growth phenotype [21]. In that experiment, the investigators constructed a strain that had the orc4^Y232C allele on a plasmid rescuing the inviable chromosomal deletion of ORC4. Because the orc4^Y232C allele was not in its native location, it was not clear to what extent the slow growth was due to plasmid loss or poor expression from the plasmid versus other, more wide-spread defects in chromosome replication or segregation. To explore in more detail the consequences of the orc4^Y232C mutation, we replaced the wild type yeast ORC4 allele with the MGS equivalent allele orc4^Y232C at its native chromosomal locus. Cells with the chromosomal orc4^Y232C allele grow more slowly than wild type (population doubling time of 2.7 hr. vs. 2.4 hr; Fig 1A). The more marked difference in growth rates reported for the strains analyzed by Guernsey et al. [21] is probably due to loss of the plasmid bearing the sole source of Orc4p in their experiments.

To explore the slow growth phenotype, we performed flow cytometry on cells synchronously proceeding through the cell cycle after having been arrested at START by treatment with alpha-factor (Fig 1B). Initiation of S phase appears to be slightly delayed in the mutant but after replicating much of its genomic DNA the mutant shows a more pronounced delay in reaching a full G2 DNA content and progressing back into G1. We reasoned that a delay in S phase entry could result if the orc4^Y232C mutation altered early origin firing. To examine this possibility we used an assay that specifically examines early origin activation across the genome [31]. This assay uses microarray hybridization to measure the levels of single stranded DNA exposed at replication forks. Both the peak position and peak amplitude of ssDNA formed at genomic loci are informative. While we do not fully understand the molecular processes that give rise to peaks of different amplitudes—e.g., number of cells that have activated a particular origin vs. amount of ssDNA revealed at different forks—the results from different replicates of the experiment are highly reproducible. We find that origins that are known to fire early and are efficient produce the peaks of greatest magnitude, while later firing and less efficient origins produce smaller or no peaks in this assay [31].

We carried out the ssDNA assay on the wild type ORC4 and orc4^Y232C mutant and a representative comparison of the two is shown in Fig 1C (to view comparisons of all chromosomes see S3 Fig). We observed differences in origin usage in ORC4 compared to orc4^Y232C and could discern three classes of origins. Some origins had comparable ssDNA peaks in both strains (cf. origins at 420 kb, 490 kb, 890 kb). Some origins that were active in ORC4 showed significant reduction in ssDNA accumulation in orc4^Y232C (cf. 200 kb, 390 kb, 830 kb). Perhaps most surprisingly, the converse was also true: some origins that are late or inefficiently initiated in ORC4 cells had significant ssDNA peaks in the orc4^Y232C mutant, indicating earlier or more efficient initiation than in wild type (cf. 15 kb, 170 kb, 670 kb). To assess quantitatively the magnitude in change to the early replication program of origin firing, we quantified the area under the peaks observed in the ssDNA assays for ORC4 and orc4^Y232. Peak areas in ORC4 vs. orc4^Y232C were highly discordant (Fig 1D; R² = ~0.14). These results highlight substantial differences in the early origin firing of origins in orc4^Y232 cells: of the 213 origins represented in the scatter plot (Fig 1D), 31 are significantly depressed or delayed in activity in the orc4^Y232C mutant (x-axis: outlined in orange) while another 24 origins are now detected as firing early (y-axis: outlined in orange). However, overall, the number of early firing origins was similar between wild type and orc4^Y232 cells. While the ssDNA assay limits us to monitoring only early-firing origins we anticipate that even if no more origin activation occurred (as is the case for cells with a deletion of CLB5, the major S phase cyclin), the completion of S phase would only be delayed by an additional 15 minutes [32].

The flow cytometry data of a synchronous culture of orc4^Y232C cells showed more than a 15 minute delay in completion of S phase and cell division (Fig 1B). For example, by 85 minutes wild type cells had divided and were undergoing a second round of DNA synthesis. The mutant cells do not show a comparable DNA content profile until 120 minutes after release. We hypothesized that this 35 minute delay could be due to cells encountering problems in late S phase and that the gradual shift to 2C DNA content results from slow progress in completing genomic DNA replication. Interestingly, patient-derived lymphoblastoid cell lines harboring an ORC4-MGS mutation also exhibit a delayed S phase transit [27]. An alternative hypothesis for the gradual shift to 2C DNA content that is observed in orc4^Y252C cells may be that they have completed genome replication on time and are continuing to replicate mitochondrial DNA but are delaying the G2/M transition for other reasons. However, the observation that cells with the orc4^Y252C mutation are unable to form colonies on plates with 200 mM hydroxyurea (HU; Fig 1E), a drug that inhibits ribonucleotide reductase and thereby slows the progress of replication forks, supports the hypothesis that the cell cycle delay stems from a chromosome replication defect.

The orc4^Y232C mutation results in reduced copy number and origin activity at the rDNA locus

When comparing the flow cytometry profiles of the orc4^Y232C cells relative to their wild type ORC4 control cells, we noticed a consistent shift to an approximately 10% lower DNA content for the orc4^Y232C cells (Fig 2A) regardless of their cell cycle phase. Considering that a haploid yeast genome is ~13 Mb in size, this difference in peak location means that ~1.3 Mb of sequence is missing from the mutant. Since the ORC4 and orc4^Y232C strains analyzed in this experiment are haploid, the difference in DNA content cannot be ascribed to chromosome aneuploidy. As an alternative possibility, we investigated whether loss of repetitive sequences could explain the shift in the mutant’s flow cytometry histogram. Candidate repetitive DNA species included mitochondrial DNA (mtDNA; ~85 kb, ~50 copies per cell), the native 2-micron plasmid (6.3 kb, ~50 copies per cells), and ribosomal DNA (rDNA; 9.1 kb, 100–200 copies per cell) [33–35]. The 2-micron plasmid and mitochondrial genomes are autonomous elements and copy number variation can occur through failure of replication and/or segregation [33,36]. In contrast, the rDNA locus is a tandemly repeated array located on chromosome XII and copy number changes occur through homologous inter- or intra-chromosome recombination [37,38]. To determine which, if any, of these repeated sequences might account for the missing genomic DNA content of the orc4^Y232C strain we performed quantitative Southern blotting of the mutant and wild type strains. We found that the mutant cells retained most (~88%) of their mtDNA (whose replication is ORC-independent), but retained only ~21% of the 2-micron plasmid (a selfish DNA element that is dependent on ORC for maintenance [39]) and ~23% of their rDNA copies relative to the wild type cells (Fig 2B). The small decrease in mtDNA cannot account for the shift observed in the flow cytometry profiles; however, together the loss of 2-micron and rDNA repeats is sufficient to account for the missing DNA content. Reduction of the 2-micron plasmid would have no significant impact on the cell’s health as cells completely cured of their 2-micron plasmids actually enjoy a slight selective growth advantage [40]. However, the reduction in the rDNA locus could have a more significant impact on cell physiology.

The rDNA locus comprises over half of the physical length of chromosome XII and accounts for ~10% of the nuclear genome (Fig 2C) [41]. Each 9.1 kb repeat contains the template for making ribosomal RNAs (rRNAs), the main structural component of ribosomes [42]. Based on the quantitative Southern blot analysis, we estimate that the rDNA copy number was reduced from ~150 copies in wild type cells to ~30 copies in the mutant. CHEF gel analysis of whole yeast chromosomes confirmed the expected size reduction of chromosome XII in the mutant (Fig 2D). Thus, the deficit of ~120 copies of rDNA in the mutant would account for ~1.1 Mb of the missing sequence, with the ~40 fewer copies of 2-micron plasmid (~252 kb) accounting for the remainder of the missing ~1.3 Mb.

Since the shrinkage of chromosome XII in orc4^Y232C cells was such a striking phenomenon, we tested six additional isolates of the mutant to determine if this size reduction was a consistent phenotype of the orc4^Y232C mutation. Immediately after selection for loss of the ORC4 allele (~20 generations), all six isolates had a smaller chromosome XII than wild type, with an rDNA locus of ~30–100 repeats (Fig 2E). To determine the stability of the rDNA locus in the different orc4^Y232C isolates over time we monitored the size of their chromosome XII after long-term growth of the strains. We found that after ~100 generations, all six populations had arrived at the same rDNA copy number of ~30 repeats (S4 Fig), suggesting that the orc4^Y232C mutation imposes strong selection for cells that have reduced their rDNA copy number. To determine the dynamics of rDNA repeat loss we analyzed samples collected during the ~100 generations of growth for one isolate (e) (Fig 2F). Rather than a steady decrease in rDNA copy number during propagation of the culture, we observed a decreasing abundance of the original chromosome XII size with concomitant increased abundance of the final size, with no obvious populations with intermediate sizes. These results support the hypothesis that a sub-population of cells with the final rDNA copy number already existed in the initial culture or appeared shortly thereafter, and had a selective advantage over cells with the longer rDNA. The descendants of the decreased-rDNA variants eventually took over the population, approaching fixation by ~80 generations of growth (Fig 2F; indicated by the lane marked with an asterisk).

Next, we asked if the selection for a loss of rDNA repeats was a phenotype specific to the orc4^Y232C allele. To test this hypothesis, we introduced a different MGS-like mutation into budding yeast. Some cases of MGS have been reported to be caused by mutations in CDC45, a core component of the CMG complex, the replicative helicase that travels with the replisome during S phase [26,43]. A specific missense mutation (P463L) occurs at an evolutionary conserved position of the human Cdc45 protein [26]; therefore, we introduced the equivalent MGS mutations into budding yeast (cdc45^P542L). We analyzed the chromosome XII size of five cdc45^P542L isolates immediately after selection for loss of the CDC45 allele and found that all the isolates had shortened chromosome XII (S5 Fig). Although we observe a contraction of the rDNA locus in cdc45^P542L yeast cells, it is important to note that shortening of this locus in the cdc45^P542L and orc4^Y232C mutants analyzed may occur through different mechanisms. However, since both Cdc45p and Orc4p are required for DNA replication, we sought to determine if defects during DNA replication may be responsible for this unusual phenotype.

Because each rDNA repeat contains a potential origin of replication (rDNA ARS) [44], we asked if compromised replication initiation at the rDNA ARS could be responsible for the delay in completion of S phase and ultimately in the reduction of rDNA copy number. Replication of the rDNA locus is a bit unusual: 1) Each repeat contains a potential origin but only a subset of them—usually those rDNA ARSs downstream of transcriptionally active repeats—serve as origins in any given S phase [45]. 2) Because of the high transcriptional activity, replication is almost entirely unidirectional—enforced by the replication fork barrier (RFB) that blocks forks from entering the 3’ end of the 35S transcription unit in a direction that opposes transcription [44]. Compared to bidirectional replication elsewhere in the genome, the unidirectional replication in the rDNA would require twice the number of initiation events for the same territory to be replicated in the same amount of time. 3) The rDNA locus completes its replication late in S phase [46]. We reasoned that if origin initiation at the rDNA locus were less efficient in orc4^Y232C cells it could explain the delay in completion of S phase and the reduction in rDNA copy number because cells that had suffered a reduction in rDNA length would complete replication more quickly and their descendants would take over the culture.

To test the efficiency of rDNA origin firing, we carried out 2D gel electrophoresis of genomic DNA from cells in logarithmic growth [47]. By focusing on the NheI fragment that contains the rDNA ARS at its center we can detect the fraction of rDNA repeats that gives rise to bubble intermediates (active origin) relative to the fraction of repeats in which the origin region is passively replicated by a fork moving through (inactive origin) (Fig 3A). The wild type cells showed the expected frequency of repeats with an active origin (roughly 1 in 5 repeats has an active origin; Fig 3B). In contrast, orc4^Y232C cells had a greatly reduced “bubble arc” relative to the intensities of the “Y arc”, the RFB pause site and the non-replicating “1N” spot. After adjusting for copy number differences in rDNA repeats between the two strains we conclude that origin activation within the rDNA array is a very rare event in the orc4^Y232C mutant cells.

Fig 3 — (A) (Top) Organization and *NheI* restriction map of a yeast rDNA repeat. Shown are the rDNA origin of replication (*rARS*) and replication fork barrier (RFB) located in the intergenic space between the 5S and 37S genes (white arrows). Replication intermediates present in the 4.7 kb *NheI* fragment are resolved using 2D-gel electrophoresis. (Bottom) Cartoon image of a 2D-gel illustrating how different replication intermediates are resolved. Relative origin efficiency is obtained by comparing intensities of replication intermediates with similar mass (dashed-line box). (B) Origin activity is reduced at the rDNA locus in *orc4*^Y232C cells. The 4.7 kb *NheI* fragment was excised from rDNA repeats from logarithmically growing cells and examined by 2D-gel electrophoresis. The Southern blot was probed for sequences specific to *rARS*. Bubble intermediates (white arrow) are reduced in *orc4*^Y232C cells compared to *ORC4* cells.

What might be the cause for the reduced origin activity of rDNA origins in the mutant cells? One possibility is that the mutant allele results in a less stable protein, therefore limiting the amount of Orc4^Y232C protein available to form Pre-RCs on rDNA origins. Such is indeed the case in orc2-1 mutant cells, where protein destabilization leads to reduced pre-RC formation; simply increasing the dosage of the mutant protein by providing an additional copy of the mutant gene on a plasmid alleviates the mutant phenotype [48]. Accordingly, we asked whether an additional copy of the orc4^Y232C allele on a centromere plasmid would provide any level of rescue of the rDNA locus contraction. An orc4^Y232C clone with ~30 rDNA repeats was transformed with plasmids containing either the ORC4 or orc4^Y232C allele and chromosome XII size was measured after ~30 generations of growth. The plasmid copy of ORC4 resulted in a significant increase in the rDNA locus while the same plasmid with the orc4^Y232C allele provided no rescue (S6 Fig). These results suggest that it may not be protein levels that are contributing to reduced rDNA origin firing, but rather, the rDNA origin is less efficiently activated in cells with the mutant ORC complex.

To test whether inefficient rDNA origin firing was responsible for contraction of the rDNA locus we reasoned that a compromised rDNA ARS would exacerbate the phenotype. A naturally occurring rDNA variant, obtained from the Robert Mortimer vineyard strain RM11-1a, was introduced into the laboratory background BY4741 [49]. This variant, which has a T to C transition in a highly-conserved residue of the rDNA ACS (S7 Fig), is known to reduce but not eliminate rARS activity [49]. Cells with this weakened rDNA variant in an otherwise wild type background show no change in growth rate (compare Fig 1A and S8 Fig). For convenience, we shall hereafter refer to the lab (BY4741) version of the rDNA as rDNA^BY and the vineyard strain (RM11-1a) variant rDNA as rDNA^RM.

Plating assays of the wild type and orc4^Y232C strains are shown in Fig 4A. While growth of the orc4^Y232C rDNA^BY strain was comparable to that of ORC4 strains with either rDNA version at 30°C and 37°C, growth of the orc4^Y232C rDNA^RM strain was reduced at 30°C and was undetectable at 37°C. Furthermore, the slow growth phenotype of the orc4^Y232C rDNA^RM strain is recapitulated in liquid medium at 30°C (S8 Fig): relative to the wild type strain, the growth rate of orc4^Y232C rDNA^RM is reduced by 54 minutes as compared to 18 minutes for orc4^Y232C rDNA^BY. We analyzed the size of the rDNA locus by measuring the BamHI genomic fragment from chromosome XII that contains the intact rDNA locus plus adjacent single copy sequences (Fig 4B) by CHEF gel electrophoresis and found that the rDNA locus had undergone an even greater reduction in the orc4^Y232C rDNA^RM strain compared to the orc4^Y232C rDNA^BY strain (Fig 4C)—from 30 copies to ~10 copies. An rDNA locus of this size (~91 kb) is at the upper limit of a replicon that could reasonably be expected to be replicated by a fork established at the nearest upstream origin in the flanking unique sequences [50,51]. 2D gel analysis (Fig 4D) confirmed that there is no detectable origin initiation within the rDNA locus itself. Unlike the cells containing the BY version of the rDNA ARS, all the initial orc4^Y232C isolates that contained the RM version of the rDNA ARS showed this drastic reduction in rDNA content and did not require an extended period of growth to achieve the steady-state reduction in rDNA copy number (Fig 4E). Cells released from an alpha-factor arrest into a synchronous S phase showed a similar late-S/G1 delay but an additional and exaggerated delay in entry into S phase (~40 minutes in orc4^Y232C rDNA^RM compared to ~20 minutes in ORC4 rDNA^RM; Fig 4F; compare with Fig 1B).

If the prolonged G1 to S delay in orc4^Y232C rDNA^RM cells were caused by insufficient origin activation across the genome, we would anticipate the total number of early active origins to be lower in this strain. To test this hypothesis, we carried out the same ssDNA assay as before on both ORC4 rDNA^RM and orc4^Y232C rDNA^RM cells. The full set of comparisons for all the chromosomes between ORC4 rDNA^RM and orc4^Y232C rDNA^RM cells are shown in S9 Fig and quantification of the area under peaks and pairwise comparisons between the strains are shown in S10 Fig. Similar to our previous observation, we found differences in the activation of early origins between ORC4 rDNA^RM and orc4^Y232C rDNA^RM cells; however, the total number of active origins was similar between the two. Additionally, pairwise comparisons of the two ORC4 or two orc4^Y232C strains with different rDNAs showed extremely good concordance, indicating that the rDNA ARS genotype does not affect early genome wide replication initiation dynamics. These observations suggest that the prolonged G1 to S transition observed in the cell cycle analysis was not due simply to inadequate genome-wide origin activation in orc4^Y232C rDNA^RM cells but suggest that some aspect of defective rDNA replication contributes to both the entry into and completion of S phase.

The initiation defect in orc4^Y232C rDNA^RM leads to chromosome XII breakage

Why are orc4^Y232C rDNA^RM cells slower to complete S phase and enter cell division? Are they experiencing difficulty in replicating their chromosome XII due to reduced origin activity at the rDNA locus, and if so, what might the consequences be? In the process of measuring rDNA copy number in the different isolates of orc4^Y232C rDNA^RM we noticed an additional minor band (Fig 4E; indicated by an asterisk) that was present only in the orc4^Y232C rDNA^RM samples and that migrated faster than the predominant chromosome XII band. We ruled out the possibility that the presence of the additional band was due to cross hybridization of the probe we were using because we did not observe its presence in the lane loaded with DNA from ORC4 rDNA^RM cells. Therefore, we hypothesized that the additional band might be a result of breakage of chromosome XII, specifically at the rDNA locus. As previously mentioned, we estimated that the rDNA copy number of orc4^Y232C rDNA^RM cells is ~10 copies, which should result in a reduction in the overall size of chromosome XII to ~1.13 Mb (Fig 4G). If chromosome XII were breaking at the rDNA locus, the two resulting sizes should be ~450 kb (left of the rDNA locus) and ~590 kb (right of the rDNA locus) and we should be able to detect these two different products using probes specific to different locations along the chromosome (Fig 4G). When hybridizing the Southern blot with a probe specific to the right of the rDNA locus (R-rDNA), in addition to the full-length chromosome XII, we also detected a band that migrated with a similar mobility to chromosomes V and VIII (~580 kb) in the lane loaded with orc4^Y232C rDNA^RM DNA (Fig 4H). However, when probing the same Southern blot for a sequence to the left of the rDNA locus (L-rDNA), the minor band previously detected was not seen; instead, a new minor band appeared at a size of ~450 kb. Based on the locations of the probes used and the different sizes of minor bands detected, we believe that chromosome XII is breaking at the rDNA locus in a small population of orc4^Y232C rDNA^RM cells. To determine if chromosome breakage is a general feature observed at other chromosomes in orc4^Y232C rDNA^RM cells, we next probed the same Southern blot for a sequence on the second largest yeast chromosome (chromosome IV) and observed only a single band in both lanes (Fig 4H), indicating that chromosome breakage is specific to the rDNA locus in orc4^Y232C rDNA^RM cells. Lastly, to determine if the chromosome breakage observed in orc4^Y232C rDNA^RM cells was a transient or recurrent event, we assayed for chromosome breakage again after ~60 generations of growth and observed only the major band corresponding to the intact chromosome XII (Fig 4H). This result suggests that the locus specific breakage in orc4^Y232C rDNA^RM cells occurs during a very narrow window, shortly after loss of the ORC4 allele.

The further loss of rDNA repeats in orc4^Y232C rDNA^RM cells leads to a reduction in ribosomal components and an increased sensitivity to the ribosome inhibiting drug cycloheximide

The combination of the orc4^Y232C mutation and the rDNA^RM locus reduces the growth rate of cells further compared to that of the orc4^Y232C rDNA^BY strain. While reducing the size of the rDNA array could permit completion of genome replication, it may come at a cost—namely, in the ability to make enough ribosomes to support robust growth. Just what is the lower limit of rDNA repeat number a yeast cell can tolerate and still support a normal ribosome population? When the rDNA copy number in yeast is artificially reduced to ~40 copies there are no obvious negative effects on growth rate and the cells increase the density of Pol I RNA polymerases per rDNA repeat to produce levels of rRNA similar to cells with ~150 copies of rDNA [52]. However, only a finite number of polymerases can be loaded onto a single rDNA repeat before space becomes limited. Based on the estimates of Pol I density and polymerization rates, yeast cells with ~30 copies of rDNA fall just short of being capable of producing levels of rRNA needed to support a normal growth rate [52]. By these calculations, orc4^Y232C cells with only ~10 copies of rDNA^RM could be further compromised in their ability to make rRNA. Therefore, we next tested our hypothesis that in addition to replication defects, the slow growth and prolonged G1 phase observed in orc4^Y232C strains is due to an inability to meet the demand for ribosome production.

To test whether orc4^Y232C cells with a reduced rDNA copy number also have decreased levels of rRNA, we separated the total nucleic acid content from cells by gel electrophoresis (S11 Fig) and then independently transferred to hybridization membrane the two different parts of the gel containing either genomic DNA (Southern blot) or rRNA (northern blot). We found that both orc4^Y232C mutants with a reduced rDNA copy number showed a reduction in 25S rRNA levels compared to their respective wild type counterparts (Fig 5A). However, orc4^Y232C rDNA^RM cells were more severely affected, being able to make only approximately half as much 25S rRNA as wild type cells. If the drastic reduction of rRNA in orc4^Y232C rDNA^RM cells was compromising ribosome production, we proposed that ribosomal proteins would also be reduced in orc4^Y232C rDNA^RM cells. Since the per-cell fluorescence output of GFP is directly proportional to its concentration in living cells [53], we reasoned that we could determine the relative abundance of ribosomal proteins per cell by GFP labeling a ribosomal protein and then measuring the cells’ relative fluorescence using flow cytometry. Ribosomal protein levels are tightly regulated, with excess proteins being targeted for rapid destruction [54], so the abundance of ribosomal proteins is a good proxy for the abundance of ribosomes. We constructed strains of the wild type ORC4 and orc4^Y232C mutant in the rDNA^BY and rDNA^RM backgrounds harboring a GFP-tagged version of the single-copy ribosomal protein Rpl10 (Rpl10-GFP) and measured their relative fluorescence during exponential growth by flow cytometry. The histogram of orc4^Y232C rDNA^RM cells is shifted to the left (a decrease of nearly one log) compared to wild type (Fig 5B). To determine if the shift were statistically significant, we recorded the fluorescence value for each of the 20,000 events recorded for each sample and performed a Wilcoxon Rank-Sum Test. The decrease in fluorescence we observed in orc4^Y232C rDNA^RM cells is statistically significant (p-value < 2.2e-16). We therefore conclude that there are fewer Rpl10-GFP molecules per cell in the orc4^Y232C rDNA^RM strain compared to wild type. The same decrease in fluorescence was not observed in orc4^Y232C rDNA^BY cells (S12 Fig), consistent with the relatively modest reduction of 25S rRNA measured in this strain (Fig 5A).

Fig 5 — (A) Quantitative hybrid Southern/northern blot analysis of the 25S ribosomal RNA. Total nucleic acids (DNA+RNA) from the four strains were separated by gel electrophoresis (See S11 Fig). The top portion of the gel, containing high-molecular weight DNA, was blotted as a Southern blot and probed with *ACT1* sequence (as a single copy control). The lower portion of the gel was blotted as a northern blot and probed with *25S rRNA* sequence to measure rRNA abundance. Values representing the relative abundance of 25S rRNA in the *orc4*^Y232C strains were calculated by first normalizing hybridization signals of *25S rRNA* to *ACT1* in each mutant strain and then relative to its respective wild type parent. (B) Relative fluorescence of *ORC4 rDNA*^RM (blue) and *orc4*^Y232C *rDNA*^RM (red) cells harboring a GFP tagged version of the single-copy ribosomal protein Rpl10. Cells were grown to mid-log phase and relative fluorescence was measured by flow cytometry. (C) Sensitivity of *orc4*^Y232C *rDNA*^RM cells to cycloheximide. Serial dilutions (1:3) of cells were plated on YPD medium with or without cycloheximide (100 ng/ml) and incubated at 30°C for 3 days.

To investigate whether the reduction of ribosomal components in orc4^Y232C rDNA^RM cells might negatively impact their growth when translation is slightly restricted, we tested their ability to grow in the presence of a low concentration of the translation inhibiting drug cycloheximide that is tolerated by wild type ORC4 cells. We observed that orc4^Y232C rDNA^RM cells could form colonies on the plate without cycloheximide; however, their growth was severely inhibited on the plate containing the drug (Fig 5C). This observation supports the notion that the slower growth of orc4^Y232C rDNA^RM cells and their delay in G1 is due to a lower translation capacity, rooted in their inability to make sufficient numbers of ribosomes.

Discussion

In an attempt to uncover the link between potential chromosome replication defects and the phenotypes that characterize Meier-Gorlin syndrome we sought to identify the cellular and molecular defects conferred by the ORC4-MGS mutation (orc4^Y232C) using yeast as a model system. We find that haploid yeast cells harboring orc4^Y232C at the endogenous chromosomal locus grow slowly, with altered S phase kinetics. Cells have reduced DNA content resulting from the loss of most copies of the nuclear 2-micron plasmid and from shrinkage of the chromosomal rDNA locus from the normal ~150 repeats to 30 repeats. Introduction of a single nucleotide variant in the rDNA origin (rDNA^RM) reduces its function as an origin in ORC4 cells and results in a further shrinkage of the rDNA locus to ~10 copies in orc4^Y232C cells. Replication initiation within the rDNA locus is greatly diminished in both rDNA types; in orc4^Y232C cells with rDNA^RM, replication of the rDNA locus is thus mostly dependent on a single fork initiated at one of the adjacent, upstream origins in the unique flanking sequences. We propose that this situation is responsible for the S/G2 delay and for the growth advantage enjoyed by cells that have a shortened rDNA locus. The presence of the rDNA^RM origin exacerbates all phenotypes associated with the orc4^Y232C mutant: slower growth, further reduction in rDNA origin firing, shorter rDNA locus, chromosome XII breakage specifically within the rDNA, and a more rapid selective sweep of the short rDNA variants through the cell population. We also find that the rDNA^RM strain with the orc4^Y232C mutation experiences a drastic decrease of ribosomal components, which may account for the additional ~30 minute G1 delay before cells enter S phase.

Across the yeast genome, replication initiation is also altered by the orc4^Y232C mutation: some unique chromosomal origins share with the rDNA origin a reduction or delay in their activation but others are advanced in their replication. While we have only examined the origins that fire early in S phase, it seems unlikely, given the density of unique origins and the proportion that are affected by the orc4^Y232C allele, that there would be other stretches of the genome that would require a single replication fork to travel ~90 kb—the distance required in the truncated rDNA^RM locus. Thus, even though replication dynamics are altered genome-wide in the orc4^Y232C mutant, we conclude that difficulties in rDNA replication, resulting in a loss of rDNA repeats and ultimately a ribosome deficiency, are at the heart of the phenotypes of yeast harboring this allele.

Our work demonstrates a strong selection for reducing rDNA copy number when ORC function is compromised. A link between impaired ORC function and variation in rDNA copy number has been noted in previous studies. Ide et al. showed that temperature sensitive mutations in two other ORC complex proteins, Orc1 and Orc2, result in the shrinkage of the rDNA locus when cells are grown at the restrictive, or semi-restrictive temperatures [30]. In addition, they demonstrated that the shortened rDNA locus was responsible for suppressing the temperature sensitivity of these mutations. Finally, they demonstrated that replication difficulties in the rDNA triggered a Rad53 checkpoint response and that rDNA reduction attenuates this checkpoint response. Based on these observations, the authors proposed that the rDNA locus plays an important role in monitoring when origin initiation across the genome is compromised. The specific molecular event that was responsible for activation of the checkpoint response was not addressed in that study, nor was it clear why rDNA shrinkage would attenuate the checkpoint response. Our work provides new insights into the observations made by Ide et al. and suggests an alternative interpretation of their results. One limitation of the Ide et al. study is that although their model assumes reduced origin firing across the genome in the orc mutants, origin activity was only examined for one chromosomal origin (ARS1) other than rARS. In our broader assessment of origin activity, we find that ORC4 and orc4^Y232C cells have a very similar number of early active origins, arguing that there is unlikely to be replication defects genome-wide. Rather, it appears that the rDNA itself is the “weak link” suffering from replication gaps. Our results suggest that as in our orc4^Y232C cells, chromosome breakage at the rDNA locus is likely responsible for triggering the Rad53 response observed by Ide et al.; we presume that as in the orc4^Y232C strain, the orc1 and orc2 cells with reduced rDNA would circumvent the rDNA replication gap problem and would therefore attenuate the checkpoint signal. Finally, although Ide, et al. demonstrated shrinkage of the rDNA locus partially suppressed the temperature sensitive mutations in Orc1 and Orc2 they did not explore how this rDNA shrinkage might affect rRNA synthesis. We find that having fewer rDNA repeats may allow for complete replication of this locus in orc4^Y232C cells, but at a cost. Losing too many rDNA repeats, as in the case of orc4^Y232C rDNA^RM cells, limits the amount of rRNA that can be transcribed. Therefore, orc4^Y232C cells must walk a tightrope, as it were—too many copies of rDNA and they suffer chromosome XII instability, but too few copies and they are unable to meet the demand for ribosomal RNAs.

A second study expanded on the idea that the ~150 rDNA origins compete for limiting replication initiation factors with the ~300 unique origins across the yeast genome [55]. The authors of that study discovered that this competition is regulated oppositely by two histone deacetylases—Sir2 and Rpd3. When initiation in the rDNA is increased, replication at some unique genomic origins is reduced, and vice versa. In their experiments, ORC was not the limiting factor. Instead, they showed that they could increase initiation at unique origins in a sir2Δ strain by overexpressing three of the initiation factors known to be in limiting supply (Sld7, Sld3 and Cdc45). We initially entertained the possibility that reduction of rDNA copy number in the orc4^Y232C mutant strain might restore a more favorable replication initiation balance between the rDNA and unique origins. However, three observations make this explanation unlikely. First, we did not find an overall reduction in unique chromosomal origin firing in the orc4^Y232C mutant—the number of early origins that failed to fire was similar to the number of new origins that appeared in the early firing class. Second, if competition between rDNA origins and unique origins were causing the growth defect in orc4^Y232C, reducing the efficiency of the rDNA^BY origin by replacing the locus with rDNA^RM should have improved growth of the orc4^Y232C strain—instead, growth was further restricted and the only locus to suffer from failed replication initiation was the rDNA. Third, introducing a second copy of orc4^Y232C on a centromere plasmid did not produce any rescue in the size of the rDNA locus—suggesting that the Orc4^Y232C protein was not in limiting supply. Together, these observations led us to hypothesize that the orc4^Y232C mutation has altered the protein’s function.

What aspect of ORC function has been altered by the orc4^Y232C mutation? One possibility is that this single amino acid substitution has changed the DNA sequence recognized by the ORC complex. Origins in budding yeast share a similar core sequence called the ACS (ARS Consensus Sequence, an approximately 17 bp AT-rich sequence necessary but not sufficient for origin function) and variation in this sequence has been shown to impact origin usage [2]. Analysis of the different groups of origins did not reveal any simple pattern(s) of polymorphism within the ACS that distinguished wild type specific origins (i.e., origins that showed reduced activity in the mutant) from mutant specific origins (i.e., origins that had ssDNA peaks in the mutant but not in wild type), although we cannot rule out the possibility that such sequence differences do exist.

A second possibility is that there is some aspect of chromatin structure or nuclear architecture that is influencing origin choice in the orc4^Y232C mutant. One class of origins whose activity was influenced by the orc4^Y232C allele was those near centromeres: 11 of the 31 wild type specific origins were located within 10 kb of a centromere (S13 Fig). To determine if this apparently skewed distribution of wild type specific origins was significantly different than would be expected to occur by chance, we performed a permutation test. We randomly labeled 31 of the of the 213 origins used to generate the scatter plots (Fig 1D and S10 Fig) as wild type specific and asked whether at least 11 of those origins were within 10 kb of a centromere. In 10,000 trials of this test, we found no occurrence of 11 or more affected origins being within 10 kb of a centromere (p < 10⁻⁴). Relevant to this discussion is the observation that we and others have made that indicates that centromeres promote early firing time of origins in their vicinity [56–58]. The early firing of centromere proximal origins is thought to occur as a consequence of the kinetochore protein Ctf19 recruiting the S-phase kinase, DDK, to phosphorylate components of the pre-RC for replication initiation [57]. Whether the ORC complex with the orc4^Y232C variant is deficient in this interaction is unknown.

Lastly, nucleosome occupancy or transcription factor binding around the origin may be preventing ORC containing Orc4^Y232C protein from binding to some origins. Moving forward, it will be important to determine if the centromere itself, chromatin state, or specific proteins that make up the kinetochore are influencing these changes.

ORC is known to play important roles in cellular processes other than DNA replication; however, its role maintaining a healthy ribosome population has yet to be characterized. Here we show that properly functioning ORC is necessary for maintaining adequate ribosomal RNA levels through its role in rDNA replication. But when the function of Orc4 is compromised, leading to a loss of rDNA repeats and the capacity to make rRNA, how might a ribosome deficiency cause slow growth in cells? The obvious answer would be that cells cannot meet the demand for protein production—but how might certain processes such as translation and protein degradation be affected? Does the translation of certain housekeeping mRNAs take priority over non-essential ones or do cells manage to deal with a ribosome deficit by speeding up the rate of translation of all mRNAs equally? Ribosome profiling experiments to identify and quantify the mRNAs that are being actively translated by ribosomes will prove helpful in addressing these questions [59]. Additionally, if cells are struggling to keep up with the protein synthesis demands, it is possible that they modulate their protein turnover rates in response. Coupling mass spectrometry based proteomics with metabolic pulse-labeling or cycloheximide treatment of cells will help shed light on the rate of protein turnover and the general state of the proteome in cells with a ribosomal RNA deficiency [60].

Is our proposed model for loss of rDNA repeats relevant to higher eukaryotes? Just as in yeast, the human rDNA is present as tandemly repeated arrays; however, there are some notable differences. In yeast the rDNA locus is located on a single chromosome with two transcription units separately producing the 5S rRNA transcript and 35S rRNA precursor, which is processed into the 18S, 5.8S and 25S rRNAs [42]. In humans, rDNA clusters are located on six different chromosomes [61]. A locus near the end of the long arm of Chromosome 1 contains 50–200 copies of a 2.2 kb repeat that produces the 5S rRNA [61]. Loci at the ends of the short arms of Chromosomes 13, 14, 15, 21, and 22 contain 43 kb repeats coding for a 47S transcript that gets processed into the 5.8S, 18S and 28S rRNAs [61]. The number of repeats on each chromosome varies and individuals show a wide distribution of copy numbers at these five loci, ranging from between 10 to more than 100 repeats per locus [62]. While most chromosomal origins in humans are thought not to be defined by primary DNA sequence, replication initiation events in the 43 kb rDNA repeats are confined to the non-transcribed spacer [63]. What might be the consequences of impaired replication at the rDNA locus in higher eukaryotes? Bloom’s syndrome (BLM) is a rare autosomal recessive disorder characterized by short stature, immunodeficiency, and predisposition for cancer [64]. Individuals with BLM have mutations in the gene BLM, which encodes a member of the RecQ family of DNA helicases that acts during DNA replication [64]. Cells derived from individuals with BLM exhibit a high frequency of sister chromatid exchange and genomic instability [64]. Particularly, BLM cells exhibit elevated levels of instability at the rDNA locus [65]; however, the exact cause for this instability is unknown. Deletion of the BLM orthologue SGS1 in budding yeast also results in increased instability at the rDNA locus [66]. Additionally, cancers in both human and mouse have also been shown to exhibit elevated levels of instability at the rDNA locus [67,68]. Stultz et al. found an increase in rDNA rearrangements in the majority of tumor samples they analyzed from lung and colorectal cancers, leading them to the conclusion that the rDNA locus is a recombination hotspot in some cancers [67]. Lastly, mice deficient for MCM2 (a component of the Pre-RC) develop lymphomas and exhibit elevated levels of DSBs at the 45S rDNA repeats in their genomes [68].

What could be the consequence(s) of increased instability at the rDNA locus in cancer? Recently, Xu et al. analyzed rDNA copy number in both human and mouse cancer genomes and contrary to their initial prediction, they found that rDNA copy number is reduced in the cancer state [69]. One explanation the authors propose for this unexpected result is that having fewer rDNA repeats may allow for more efficient DNA replication and thus greater cell proliferation in cancer. Given the fact that the rDNA locus is highly sensitive to replication stress in higher eukaryotes and its copy number can change rapidly during the disease state, a similar problem with rDNA replication—due to reduced origin initiation—could lead to a reduction in rDNA copy number and possibly limit the amount of rRNA and thus reduce ribosome levels in higher eukaryotes.

Meier-Gorlin syndrome phenotypes include a number of skeletal defects that have similarities to other, better-understood syndromes, all associated with deficiencies in ribosome biogenesis [70]. For example, Treacher Collins syndrome (TCS) can be caused by one of three different genetic mutations that are likely to affect ribosome production [71,72]. One of the TCS mutations is in TCOF1, a gene that encodes a protein called Treacle that is involved in rDNA transcription through its interaction with upstream binding factor (UBF) [73]. Treacle also interacts physically with human Nop56p, a component of the rRNA modifying box C/D small nucleolor ribonucleoprotein complex [74]. The other two genes, POLR1C and POLR1D, are subunits of RNA Polymerases I and III [72]. Work with mouse models of this syndrome suggest that ribosome deficiencies are reducing the migration of neural crest cells that are essential for proper craniofacial development [75]. Whether derived from neural crest cells or mesenchymal cells, chondrocytes also have a high demand for ribosomes as they divide, enlarge and produce collagen and other bone-matrix proteins [70]. In addition to TCS, Postaxial Acrofacial Dystosis (POADS), Diamond Blackfan Anemia (DBA), Roberts Syndrome (RBS), Schwachman-Diamond Syndrome (SDS), and Cartilage-Hair Hyploplasia (CHH) are distinct ribosomopathies with a common set of skeletal malformations [70]. Their genetic mutations affect ribosome production in different ways—a mutation that reduces uracil biosynthesis (POADS) [76,77], mutations in individual ribosomal protein genes (DBA) (reviewed in[78,79]), a mutation in a gene for cohesion (ESCO2) that in yeast (ECO1) reduces 18S and 28S production (RBS) [80,81], a mutation in SBDS (yeast SDO1) that reduces maturation of the 60S ribosomal subunit (SDS) [82–84], and a mutation in RMPR, the RNA component of RNase MRP, a snoRNA involved in rRNA processing (CHH) [85,86].

A ribosomopathy that is associated with a deficiency of rDNA has not yet been identified; however, one might speculate on the phenotypes that may be manifested with such a disorder [87].

Further work will be necessary to determine whether the phenotypes in orc4^Y232C yeast are consistent with cells derived from individuals with MGS. Additionally, determining if the slow growth phenotype observed in orc4^Y232C yeast is due primarily to defects in chromosome replication or a ribosome deficiency will prove necessary in elucidating how mutations in proteins necessary for origin initiation may inadvertently affect cellular processes in addition to DNA replication.

Materials and methods

Yeast strains and plasmids

A complete list of yeast strains and plasmids can be found in the S1 Table. BY4741 was used as wild type (ORC4) for this study and all strains were derived from this background. The rDNA locus from the Robert Mortimer vineyard strain RM11 was introduced into the BY4741 background by standard backcrossing (10 times) to create ORC4 rDNA^RM [49]. Subsequently, the MGS-like variant orc4^Y323C was introduced into either ORC4 rDNA^BY or ORC4 rDNA^RM by two-step gene replacement [88]. A plasmid containing URA3 and the orc4^Y232C allele was integrated at the ORC4 locus; correct integrants were confirmed by PCR and Southern analysis. We selected for loss of the integrated sequences through homologous recombination by selecting against the URA3 gene on plates containing 5-FOA. To screen for clones that had lost the wild type ORC4 allele and had kept the orc4^Y232C allele we performed PCR using an allele specific oligonucleotide as one of the PCR primers. The MGS-like allele cdc45^P542L was introduced into BY4741 using CRISPR-Cas9 following the steps described by Laughery et al [89]. For all experiments, yeast cultures were grown at 30°C in synthetic complete medium supplemented with 2% glucose unless stated otherwise.

Structural analysis of protein models

Protein structure visualization was performed using UCSF Chimera [90].

Flow cytometry for cell cycle analysis

Cell cycle progression was examined using flow cytometry. Early log phase cells (OD₆₆₀ ~0.25) were arrested in G1 by the addition of alpha factor at a final concentration of 3 μM. When >90% of the population was un-budded (time equivalent to ~1.5 population doublings), cells were synchronously released into S-phase by the addition of Pronase (Calbiochem) at a final concentration of 0.3 mg/ml. After the release into S phase, cells were harvested at 5-minute intervals, mixed with sodium azide at a final concentration of 0.1% and fixed with 70% ethanol. Cells were prepared for flow cytometry as previously described [32]. Flow cytometry was performed on a BD FACSCanto II and data were analyzed using FlowJo software.

Contour-clamped homogeneous electric field (CHEF) gel analysis

Stationary phase cells were embedded in agarose plugs and prepared using standard procedures [91]. CHEF gel analysis of whole yeast chromosomes was performed using a BioRad CHEF-DR II Pulsed Field Electrophoresis System. Whole chromosomes were resolved in 0.8% LE agarose gels with a switch time ramped from 300–900 seconds at 100 volts for 68 hours in 0.5X TBE at 14°C. BamHI digested chromosomal DNA fragments were resolved in 1.0% LE agarose gels with a switch time ramped from 47–175 seconds at 165 volts for 62 hours in 0.5X TBE at 14°C. Gels were stained with ethidium bromide and photographed. Southern blotting and hybridization were performed using standard procedures.

Two-Dimensional (2D) agarose gel analysis

Mid-log phase cells were harvested and embedded in 0.5% low melt agarose (SeaPlaque) in 50 mM EDTA and prepared as previously described [92] with slight modifications (see http://fangman-brewer.genetics.washington.edu/plug.html). DNA was subsequently digested in-gel using NheI (NEB). 2D gel electrophoresis was used to visualize the relative abundance of replication intermediates and was performed as described by [47]. Gels were blotted and hybridized with a probe specific to the rDNA origin of replication.

Quantitative Southern blotting

DNA was harvested from stationary phase cells via the “Smash-and-grab” DNA isolation protocol [93] and subsequently digested with EcoRV (NEB). Digested DNA was separated by electrophoresis in a 0.7% Agarose LE (GeneMate) gel and then blotted following standard Southern blotting protocols. Blots were then sequentially hybridized with probes specific to ACT1 (single copy control), mtDNA, 2-micron plasmid, and then rDNA. The hybridization signal was analyzed using the BioRad Personal Molecular Imager and Quantity One software. Hybridization signals of repetitive sequences were first normalized to ACT1 and then relative to wild type.

Long term growth experiments

Our initial replacement of ORC4 with the mutant orc4^Y232C allele was performed using a “pop-in/pop-out” strategy ([88]; see Yeast strains and plasmids section above). To track long-term rDNA copy number changes following replacement of wild type with mutant orc4^Y232C, we picked fresh “pop-out” candidates and inoculated liquid cultures for growth to saturation (~30 generations). After confirming the loss of ORC4 by allele specific PCR, plugs for CHEF gel analysis were made for the orc4^Y323C positive isolates (a-f; Fig 2D). From the initial overnight cultures, a 1/100 dilution was made into 5 ml of fresh medium and allowed to grow to saturation (~7 generations). Growing of cells to saturation and then diluting back into fresh medium was performed for a total of 10 times which accounted for a total of ~100 generations of growth. In each round, cells were harvested when the culture reached saturation and embedded in agarose plugs for CHEF gel analysis.

Quantitative hybrid Southern/northern blotting

The relative abundance of 25S ribosomal RNA was measured by quantitative hybrid Southern/northern blotting. The total nucleic acid content of cells was extracted using a version of the “Smash-and-grab” DNA isolation protocol [93] with the modification that cell walls were enzymatically disrupted using Zymolyase - 20T (Amsbio) instead of glass beads. Nucleic acids were resuspended in TE pH 8 and then separated by electrophoresis in a 1.5% LE agarose gel with ethidium bromide (0.3 μg/ml). After the ribosomal RNA was separated away from genomic DNA, the gel was photographed and then cut to separate the two portions containing genomic DNA and rRNA. Subsequently, the two different portions of the gel were blotted following standard Southern (genomic DNA) and northern (rRNA) blotting protocols. The Southern blot was probed for ACT1 as a single copy control and the northern blot was probed for 25S rRNA sequence. Because the amount of rRNA on the blot could be in excess of the probe, hybridization of the northern blot was limited to 2 hours to ensure that the hybridization signal was proportional to the amount of target sequence. The hybridization signals were analyzed using a Bio-Rad Personal Molecular Imager and Quantity One software. The 25S rRNA hybridization signal was first normalized to ACT1 and then relative to wild type.

Flow cytometry analysis for cells expressing GFP tagged Rpl10

The GFP fluorescence of living cells was measured using flow cytometry. Strains were grown overnight in synthetic complete medium at 30°C. Fresh cultures were made by diluting these overnight cultures back to a starting OD₆₆₀ of ~0.05. When cultures reached an OD₆₆₀ of ~0.6, cells were diluted and sonicated, then analyzed directly using a BD Accuri C6 flow cytometer. Flow cytometry data was exported and analyzed using FloJo software.

Mapping of single stranded DNA

A detailed protocol for this assay has previously been published [94]. Cells growing in log phase (OD₆₆₀ ~0.25) were arrested in G1 by the addition of alpha factor and then synchronously released into S phase in the presence of 200 mM HU. Samples were collected every 15 minutes after release into S phase and cells were embedded in agarose plugs and then spheroplasted. The ssDNA from either S phase or G1 control samples were then differentially labeled with either Cy5- or Cy3-dUTP by in-gel random-primed labeling using exo- Klenow polymerase (NEB) without denaturation of template. The differently labeled DNAs were then collected and co-hybridized to Agilent G4493A yeast 4x44K ChIP to chip DNA microarrays according to the manufacturer’s recommendations. The data from scanned microarrays was extracted using Agilent’s Feature Extraction software. The ssDNA microarray data are available at NCBI GEO under accession no. GSE104671.

ssDNA peak area script description

Areas under ssDNA peaks were assessed from Loess-smoothed microarray data (coordinates spaced 500 bp apart) using a custom Python script and a reference list of origins and their locations from OriDB [95] (http://cerevisiae.oridb.org/). Since most of the genome is double-stranded and therefore not a template for ssDNA labeling, the mean genome-wide signal was used as a “threshold” value for each sample. The “ssDNA peak area” for each origin was then calculated as a sum of S/G1 value at the origin’s location and the sequentially-added S/G1 values from adjacent data points until three “below threshold" values were reached on each side of the origin. The script does not distinguish overlapping origin peaks and therefore overlapping early firing origins in close proximity were manually curated and excluded from the analysis. This Python script is available as a supplemental file (S1 File).

Supporting information

S1 Fig. Structural comparison of H. sapiens and S. cerevisiae ORC subunits.

Comparison of the structural models of H. sapiens (Hs; top panel) and S. cerevisiae (Sc; middle panel) origin recognition complex subunits (Orc1, 2, 3, 4, and 5; PDB IDs 5UJM and 5UDB, respectively). The bottom panel shows a superimposition of the Hs and Sc complexes. Data are from [8] and [9].

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S2 Fig. Position of H. sapiens Orc4-Tyr174 and S. cerevisiae Orc4-Tyr232.

(A).

Superimposition of the Hs and Sc Orc4 subunits highlights the high degree of structural similarity between the proteins from the two species. The right panel focuses on the Tyrosine mutated in human MGS patients (Tyr174) and the corresponding Tyrosine in yeast (Tyr232) with the side chains of these amino acids displayed in red.

(B).

Structural models of ScOrc1-6 and ScCdc6 in complex with a double stranded DNA molecule. The right panel focuses on Tyr232 of ScOrc4, with the side chain of this amino acid depicted as red spheres.

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S3 Fig. Comparison of ssDNA profiles for ORC4 and orc4^Y232C chromosomes.

Genome wide ssDNA profiles for ORC4 (black) and orc4^Y232C (red) are shown for cells after exposure to HU for 30 min. The relative ratio of ssDNA (S/G1) is plotted against chromosome coordinates (kb). A yellow circle denotes centromere locations and the positions of verified origins of replication are marked by orange triangles. The rDNA locus and adjacent flanking sequence on Chr XII (cf. 440–490 kb) were omitted due to insufficient probe coverage on the microarray slide.

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S4 Fig. The rDNA copy number in orc4^Y232C cells stabilizes at ~30 copies.

Variation in rDNA copy number was analyzed in the six isolates (a-f) of orc4^Y232C after growth for ~100 generations. Change in Chr XII size was measured by CHEF gel electrophoresis. Left, ethidium bromide stained image; right, Southern blot image following hybridization with a Chr XII-specific single-copy sequence. By ~100 generations the size of Chr XII had stabilized at ~30 copies of rDNA for most of the population in all six isolates of orc4^Y232C.

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S5 Fig. CHEF gel analysis of variation in rDNA copy number of five isolates (a-e) of cdc45^P542L.

Left, ethidium bromide stained image; right, Southern blot hybridization for Chr XII. All five isolates had a smaller Chr XII than CDC45 due to loss of rDNA repeats.

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S6 Fig. The orc4^Y232C rDNA copy number phenotype is not rescued by an additional copy of orc4^Y232C.

An isolate of orc4^Y232C with ~30 copies of rDNA (lane #2) was transformed with a centromere plasmid (pRS415) containing a copy of either ORC4 or orc4^Y232C. rDNA copy number was analyzed in three isolates from each transformation. An increase in rDNA copy number was observed in cells transformed with the plasmid containing ORC4; however, no increase in rDNA copy number was observed in cells transformed with the plasmid containing orc4^Y232C.

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S7 Fig. Sequence comparison of the ACS of BY4741 and RM11-1a rDNA origin, rARS.

Red arrow indicates the polymorphism in the ACS.

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S8 Fig. The slow growth phenotype is exacerbated in orc4^Y232C rDNA^RM cells.

Growth curves of ORC4 rDNA^RM and orc4^Y232C rDNA^RM cells generated by measuring the optical density over time of mid-log cultures in synthetic complete medium at 30°C. The mutant (white circle) shows a substantial growth defect with a doubling-time 54 minutes (27%) longer than wild-type cells (black circle).

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S9 Fig. Comparison of ssDNA profiles for ORC4 rDNA^RM and orc4^Y232C rDNA^RM chromosomes.

Genome wide ssDNA profiles for ORC4 rDNA^RM (black) and orc4^Y232C rDNA^RM (red) are shown for cells after exposure to HU for 30 min. The relative ratio of ssDNA (S/G1) is plotted against chromosome coordinates (kb). A yellow circle denotes centromere locations and the positions of verified origins of replication are marked by orange triangles. The rDNA locus and adjacent flanking sequence on Chr XII (cf. 440–490 kb) were omitted due to insufficient probe coverage on the microarray slide.

(TIF)

Click here for additional data file.^{(1.5MB, tif)}

S10 Fig. Pairwise comparisons of the relative area under the peak at each origin measured in ssDNA profiles.

The relative areas under the peaks for the four strains (ORC4 rDNA^BY, ORC4 rDNA^RM, orc4^Y232C rDNA^BY, and orc4^Y232C rDNA^RM) were measured and pair-wise comparisons of those values at each origin between the different strains are shown in the three scatter plots.

(TIF)

Click here for additional data file.^{(285.8KB, tif)}

S11 Fig. The total nucleic acid content of cells is resolved by gel electrophoresis.

An ethidium bromide stained gel image of the total nucleic acid content from exponentially growing cells separated by electrophoresis. The blue dashed line indicates where the gel was cut so that the two different parts of the gel containing either genomic DNA or rRNA could be separately treated for Southern or northern transfer to hybridization membranes. The hybridization images and quantifications are shown in Fig 5A.

(TIF)

Click here for additional data file.^{(369.2KB, tif)}

S12 Fig. Relative fluorescence of ORC4 rDNA^BY and orc4^Y232C rDNA^BY cells harboring Rpl10-GFP.

Relative fluorescence of ORC4 rDNA^BY (blue) and orc4^Y232C rDNA^BY (red) cells harboring a GFP tagged version of the single-copy ribosomal protein Rpl10. Cells were grown to mid-log phase and relative fluorescence was measured by flow cytometry.

(TIF)

Click here for additional data file.^{(253.6KB, tif)}

S13 Fig. Origins proximal to centromeres are less active in orc4^Y232C cells.

(A) The locations of ORC4 (blue circles) and orc4^Y232C (red circles) specific origins are shown across the sixteen yeast chromosomes aligned by their centromeres at x = 0. Centromere locations are marked by a yellow circle. (B) The locations of only the ORC4 specific origins are shown relative to the locations of centromeres (yellow line). (C) The locations of only the orc4^Y232C specific origins are shown relative to the locations of centromeres (yellow line).

(TIF)

Click here for additional data file.^{(684.6KB, tif)}

S1 Table. Yeast strains and plasmids used in this study.

(PDF)

Click here for additional data file.^{(78.2KB, pdf)}

S1 File. Python script used to calculate the area under ssDNA peaks.

(PY)

Click here for additional data file.^{(14KB, py)}

Acknowledgments

We wish to thank the members of the Brewer/Raghuraman and Dunham labs for their support and helpful discussions. We thank Gina Alvino and Kelsey Lynch for their critical review of this manuscript. We would also like to thank Monica Sanchez for her insightful discussions and expert technical assistance. We extend our gratitude to current and former Brewer/Raghuraman lab members Mackenzie Croy, Shanti Neff-Baro, and Alex Mason who have worked on various aspects of this project not included in this manuscript. We are grateful to Marissa LeBlanc and Christopher McMaster for their kind gifts of plasmid and yeast strains.

Data Availability

The microarray data from the single stranded assays is available from the NCBI GEO database (accession number GSE104671). All other relevant data are within the paper and its Supporting Information files.

Funding Statement

BJB and MKR were supported by grants GM18926 and 1R35GM122497 to BJB from the National Institute of General Medical Sciences (http://www.nigms.nih.gov/). JCS was supported by the National Science Foundation Graduate Research Fellowship Program under Grant Nos. DG#-0718124 and DGE-1256082 (https://www.nsfgrfp.org/) and by a National Institutes of Health Genome Training Grant T32 (HG00035) (https://www.genome.gov/). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

References

1.Stinchcomb DT, Struhl K, Davis RW. Isolation and characterisation of a yeast chromosomal replicator. Nature. 1979;282: 39–43. doi: 10.1038/282039a0 [DOI] [PubMed] [Google Scholar]
2.Liachko I, Youngblood RA, Keich U, Dunham MJ. High-resolution mapping, characterization, and optimization of autonomously replicating sequences in yeast. Genome Res. 2013;23: 698–704. doi: 10.1101/gr.144659.112 [DOI] [PMC free article] [PubMed] [Google Scholar]
3.Bell SP, Stillman B. ATP-dependent recognition of eukaryotic origins of DNA replication by a multiprotein complex. Nature. 1992;357: 128–34. doi: 10.1038/357128a0 [DOI] [PubMed] [Google Scholar]
4.Liang C, Weinreich M, Stillman B. ORC and Cdc6p interact and determine the frequency of initiation of DNA replication in the genome. Cell. 1995;81: 667–76. doi: 10.1016/0092-8674(95)90528-6 [DOI] [PubMed] [Google Scholar]
5.Tanaka S, Diffley JFX. Interdependent nuclear accumulation of budding yeast Cdt1 and Mcm2-7 during G1 phase. Nat Cell Biol. 2002;4: 198–207. doi: 10.1038/ncb757 [DOI] [PubMed] [Google Scholar]
6.Cvetic C, Walter JC. Eukaryotic origins of DNA replication: Could you please be more specific? Semin Cell Dev Biol. 2005;16: 343–353. doi: 10.1016/j.semcdb.2005.02.009 [DOI] [PubMed] [Google Scholar]
7.Li H, Stillman B. The Origin Recognition Complex: A Biochemical and Structural View. Subcellular Biochemistry. 2012. pp. 37–58. doi: 10.1007/978-94-007-4572-8_3 [DOI] [PMC free article] [PubMed] [Google Scholar]
8.Tocilj A, On KF, Yuan Z, Sun J, Elkayam E, Li H, et al. Structure of the active form of human origin recognition complex and its ATPase motor module. Elife. 2017;6: e20818 doi: 10.7554/eLife.20818 [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Yuan Z, Riera A, Bai L, Sun J, Nandi S, Spanos C, et al. Structural basis of Mcm2-7 replicative helicase loading by ORC-Cdc6 and Cdt1. Nat Struct Mol Biol. 2017; doi: 10.1038/nsmb.3372 [DOI] [PMC free article] [PubMed] [Google Scholar]
10.Liang C, Stillman B. Persistent initiation of DNA replication and chromatin-bound MCM proteins during the cell cycle in cdc6 mutants. Genes Dev. 1997;11: 3375–86. doi: 10.1101/gad.11.24.3375 [DOI] [PMC free article] [PubMed] [Google Scholar]
11.Fragkos M, Ganier O, Coulombe P, Méchali M. DNA replication origin activation in space and time. Nat Rev Mol Cell Biol. Nature Publishing Group; 2015;16: 360–74. doi: 10.1038/nrm4002 [DOI] [PubMed] [Google Scholar]
12.Randell JCW, Bowers JL, Rodríguez HK, Bell SP. Sequential ATP hydrolysis by Cdc6 and ORC directs loading of the Mcm2-7 helicase. Mol Cell. 2006;21: 29–39. doi: 10.1016/j.molcel.2005.11.023 [DOI] [PubMed] [Google Scholar]
13.Cocker JH, Piatti S, Santocanale C, Nasmyth K, Diffley JF. An essential role for the Cdc6 protein in forming the pre-replicative complexes of budding yeast. Nature. 1996;379: 180–2. doi: 10.1038/379180a0 [DOI] [PubMed] [Google Scholar]
14.Diffley JF, Cocker JH, Dowell SJ, Rowley A. Two steps in the assembly of complexes at yeast replication origins in vivo. Cell. 1994;78: 303–16. [DOI] [PubMed] [Google Scholar]
15.Speck C, Chen Z, Li H, Stillman B. ATPase-dependent cooperative binding of ORC and Cdc6 to origin DNA. Nat Struct Mol Biol. 2005;12: 965–71. doi: 10.1038/nsmb1002 [DOI] [PMC free article] [PubMed] [Google Scholar]
16.Chen S, de Vries MA, Bell SP. Orc6 is required for dynamic recruitment of Cdt1 during repeated Mcm2-7 loading. Genes Dev. 2007;21: 2897–907. doi: 10.1101/gad.1596807 [DOI] [PMC free article] [PubMed] [Google Scholar]
17.Sclafani RA, Holzen TM. Cell cycle regulation of DNA replication. Annu Rev Genet. 2007;41: 237–80. doi: 10.1146/annurev.genet.41.110306.130308 [DOI] [PMC free article] [PubMed] [Google Scholar]
18.Stillman B. Origin recognition and the chromosome cycle. FEBS Lett. 2005;579: 877–84. doi: 10.1016/j.febslet.2004.12.011 [DOI] [PubMed] [Google Scholar]
19.Ilves I, Petojevic T, Pesavento JJ, Botchan MR. Activation of the MCM2-7 Helicase by Association with Cdc45 and GINS Proteins. Mol Cell. Elsevier Ltd; 2010;37: 247–258. doi: 10.1016/j.molcel.2009.12.030 [DOI] [PMC free article] [PubMed] [Google Scholar]
20.Bicknell LS, Bongers EMHF, Leitch A, Brown S, Schoots J, Harley ME, et al. Mutations in the pre-replication complex cause Meier-Gorlin syndrome. Nat Genet. 2011;43: 356–9. doi: 10.1038/ng.775 [DOI] [PMC free article] [PubMed] [Google Scholar]
21.Guernsey DL, Matsuoka M, Jiang H, Evans S, Macgillivray C, Nightingale M, et al. Mutations in origin recognition complex gene ORC4 cause Meier-Gorlin syndrome. Nat Genet. 2011;43: 360–364. doi: 10.1038/ng.777 [DOI] [PubMed] [Google Scholar]
22.Gorlin RJ, Cervenka J, Moller K, Horrobin M, Witkop CJ. Malformation syndromes. A selected miscellany. Birth Defects Orig Artic Ser. 1975;11: 39–50. [PubMed] [Google Scholar]
23.Meier Z, Poschiavo, Rothschild M. Case of arthrogryposis multiplex congenita with mandibulofacial dysostosis (Franceschetti syndrome). Helv Paediatr Acta. 1959;14: 213–6. [PubMed] [Google Scholar]
24.Vetro A, Savasta S, Russo Raucci A, Cerqua C, Sartori G, Limongelli I, et al. MCM5: a new actor in the link between DNA replication and Meier-Gorlin syndrome. Eur J Hum Genet. Nature Publishing Group; 2017; 1–5. doi: 10.1038/ejhg.2017.5 [DOI] [PMC free article] [PubMed] [Google Scholar]
25.Burrage LC, Charng WL, Eldomery MK, Willer JR, Davis EE, Lugtenberg D, et al. De Novo GMNN Mutations Cause Autosomal-Dominant Primordial Dwarfism Associated with Meier-Gorlin Syndrome. Am J Hum Genet. The American Society of Human Genetics; 2015;97: 904–913. doi: 10.1016/j.ajhg.2015.11.006 [DOI] [PMC free article] [PubMed] [Google Scholar]
26.Fenwick AL, Kliszczak M, Cooper F, Murray J, Sanchez-Pulido L, Twigg SRF, et al. Mutations in CDC45, Encoding an Essential Component of the Pre-initiation Complex, Cause Meier-Gorlin Syndrome and Craniosynostosis. Am J Hum Genet. American Society of Human Genetics; 2016;99: 125–138. doi: 10.1016/j.ajhg.2016.05.019 [DOI] [PMC free article] [PubMed] [Google Scholar]
27.Stiff T, Alagoz M, Alcantara D, Outwin E, Brunner HG, Bongers EMHF, et al. Deficiency in origin licensing proteins impairs cilia formation: implications for the aetiology of Meier-Gorlin syndrome. PLoS Genet. 2013;9: e1003360 doi: 10.1371/journal.pgen.1003360 [DOI] [PMC free article] [PubMed] [Google Scholar]
28.Bicknell LS, Walker S, Klingseisen A, Stiff T, Leitch A, Kerzendorfer C, et al. Mutations in ORC1, encoding the largest subunit of the origin recognition complex, cause microcephalic primordial dwarfism resembling Meier-Gorlin syndrome. Nat Genet. 2011;43: 350–355. doi: 10.1038/ng.776 [DOI] [PubMed] [Google Scholar]
29.Hossain M, Stillman B. Meier-Gorlin syndrome mutations disrupt an Orc1 CDK inhibitory domain and cause centrosome reduplication. Genes Dev. 2012;26: 1797–810. doi: 10.1101/gad.197178.112 [DOI] [PMC free article] [PubMed] [Google Scholar]
30.Ide S, Watanabe K, Watanabe H, Shirahige K, Kobayashi T, Maki H. Abnormality in initiation program of DNA replication is monitored by the highly repetitive rRNA gene array on chromosome XII in budding yeast. Mol Cell Biol. 2007;27: 568–78. doi: 10.1128/MCB.00731-06 [DOI] [PMC free article] [PubMed] [Google Scholar]
31.Feng W, Collingwood D, Boeck ME, Fox LA, Alvino GM, Fangman WL, et al. Genomic mapping of single-stranded DNA in hydroxyurea-challenged yeasts identifies origins of replication. Nat Cell Biol. 2006;8: 148–55. doi: 10.1038/ncb1358 [DOI] [PMC free article] [PubMed] [Google Scholar]
32.McCune HJ, Danielson LS, Alvino GM, Collingwood D, Delrow JJ, Fangman WL, et al. The temporal program of chromosome replication: genomewide replication in clb5Δ Saccharomyces cerevisiae. Genetics. 2008;180: 1833–47. doi: 10.1534/genetics.108.094359 [DOI] [PMC free article] [PubMed] [Google Scholar]
33.Liu Y-T, Fan H-F, Kachroo AH, Jayaram M, Rowley PA, Sau S, et al. The Partitioning and Copy Number Control Systems of the Selfish Yeast Plasmid: An Optimized Molecular Design for Stable Persistence in Host Cells. Microbiol Spectr. 2014;2: 1–32. doi: 10.1128/microbiolspec.PLAS-0003-2013 [DOI] [PMC free article] [PubMed] [Google Scholar]
34.Williamson D. The curious history of yeast mitochondrial DNA. Nat Rev Genet. 2002;3: 475–81. doi: 10.1038/nrg814 [DOI] [PubMed] [Google Scholar]
35.Schweizer E, MacKechnie C, Halvorson HO. The redundancy of ribosomal and transfer RNA genes in Saccharomyces cerevisiae. J Mol Biol. 1969;40: 261–277. doi: 10.1016/0022-2836(69)90474-4 [DOI] [PubMed] [Google Scholar]
36.Chen XJ, Butow R a. The organization and inheritance of the mitochondrial genome. Nat Rev Genet. 2005;6: 815–825. doi: 10.1038/nrg1708 [DOI] [PubMed] [Google Scholar]
37.Petes TD. Yeast ribosomal DNA genes are located on chromosome XII. Proc Natl Acad Sci U S A. 1979;76: 410–4. doi: 10.1073/pnas.76.1.410 [DOI] [PMC free article] [PubMed] [Google Scholar]
38.Kobayashi T. Strategies to maintain the stability of the ribosomal RNA gene repeats—collaboration of recombination, cohesion, and condensation. Genes Genet Syst. 2006;81: 155–161. JST.JSTAGE/ggs/81.155 [pii] [DOI] [PubMed] [Google Scholar]
39.Micklem G, Rowley A, Harwood J, Nasmyth K, Diffley JF. Yeast origin recognition complex is involved in DNA replication and transcriptional silencing. Nature. 1993;366: 87–89. doi: 10.1038/366087a0 [DOI] [PubMed] [Google Scholar]
40.Futcher AB, Cox BS. Maintenance of the 2 microns circle plasmid in populations of Saccharomyces cerevisiae. J Bacteriol. 1983;154: 612–622. [DOI] [PMC free article] [PubMed] [Google Scholar]
41.Warner JR. The economics of ribosome biosynthesis in yeast. Trends Biochem Sci. 1999;24: 437–440. doi: 10.1016/S0968-0004(99)01460-7 [DOI] [PubMed] [Google Scholar]
42.Woolford JL, Baserga SJ. Ribosome biogenesis in the yeast Saccharomyces cerevisiae. Genetics. 2013;195: 643–681. doi: 10.1534/genetics.113.153197 [DOI] [PMC free article] [PubMed] [Google Scholar]
43.Simon AC, Sannino V, Costanzo V, Pellegrini L. Structure of human Cdc45 and implications for CMG helicase function. Nat Commun. Nature Publishing Group; 2016;7: 11638 doi: 10.1038/ncomms11638 [DOI] [PMC free article] [PubMed] [Google Scholar]
44.Brewer BJ, Fangman WL. A replication fork barrier at the 3′ end of yeast ribosomal RNA genes. Cell. 1988;55: 637–643. doi: 10.1016/0092-8674(88)90222-X [DOI] [PubMed] [Google Scholar]
45.Muller M, Lucchini R, Sogo JM. Replication of yeast rDNA initiates downstream of transcriptionally active genes. Mol Cell. 2000;5: 767–777. doi: 10.1016/S1097-2765(00)80317-2 [DOI] [PubMed] [Google Scholar]
46.Foss EJ, Lao U, Dalrymple E, Adrianse RL, Loe T, Bedalov A. SIR2 suppresses replication gaps and genome instability by balancing replication between repetitive and unique sequences. Proc Natl Acad Sci U S A. 2017;114: 552–557. doi: 10.1073/pnas.1614781114 [DOI] [PMC free article] [PubMed] [Google Scholar]
47.Brewer BJ, Fangman WL. The localization of replication origins on ARS plasmids in S. cerevisiae. Cell. 1987;51: 463–471. doi: 10.1016/0092-8674(87)90642-8 [DOI] [PubMed] [Google Scholar]
48.Shimada K, Pasero P, Gasser SM. ORC and the intra-S-phase checkpoint: A threshold regulates Rad53p activation in S phase. Genes Dev. 2002;16: 3236–3252. doi: 10.1101/gad.239802 [DOI] [PMC free article] [PubMed] [Google Scholar]
49.Kwan EX, Foss EJ, Tsuchiyama S, Alvino GM, Kruglyak L, Kaeberlein M, et al. A Natural Polymorphism in rDNA Replication Origins Links Origin Activation with Calorie Restriction and Lifespan. PLoS Genet. 2013;9: e1003329 doi: 10.1371/journal.pgen.1003329 [DOI] [PMC free article] [PubMed] [Google Scholar]
50.Miller CA, Umek RM, Kowalski D. The inefficient replication origin from yeast ribosomal DNA is naturally impaired in the ARS consensus sequence and in DNA unwinding. Nucleic Acids Res. 1999;27: 3921–3930. doi: 10.1093/nar/27.19.3921 [DOI] [PMC free article] [PubMed] [Google Scholar]
51.Rivin CJ, Fangman WL. Replication fork rate and origin activation during the S phase of Saccharomyces cerevisiae. J Cell Biol. 1980;85: 108–15. [DOI] [PMC free article] [PubMed] [Google Scholar]
52.French SL, Osheim YN, Cioci F, Nomura M, Beyer AL. In exponentially growing Saccharomyces cerevisiae cells, rRNA synthesis is determined by the summed RNA polymerase I loading rate rather than by the number of active genes. Mol Cell Biol. 2003;23: 1558–68. doi: 10.1128/MCB.23.5.1558-1568.2003 [DOI] [PMC free article] [PubMed] [Google Scholar]
53.Lo C-A, Kays I, Emran F, Lin T-J, Cvetkovska V, Chen BE. Quantification of Protein Levels in Single Living Cells. Cell Rep. The Authors; 2015;13: 2634–44. doi: 10.1016/j.celrep.2015.11.048 [DOI] [PubMed] [Google Scholar]
54.Sung M-K, Reitsma JM, Sweredoski MJ, Hess S, Deshaies RJ. Ribosomal proteins produced in excess are degraded by the ubiquitin-proteasome system. Mol Biol Cell. 2016;27: 2642–52. doi: 10.1091/mbc.E16-05-0290 [DOI] [PMC free article] [PubMed] [Google Scholar]
55.Yoshida K, Bacal J, Desmarais D, Padioleau I, Tsaponina O, Chabes A, et al. The Histone Deacetylases Sir2 and Rpd3 Act on Ribosomal DNA to Control the Replication Program in Budding Yeast. Mol Cell. 2014;54: 691–697. doi: 10.1016/j.molcel.2014.04.032 [DOI] [PubMed] [Google Scholar]
56.Blitzblau HG, Chan CS, Hochwagen A, Bell SP. Separation of DNA Replication from the Assembly of Break-Competent Meiotic Chromosomes. Hawley RS, editor. PLoS Genet. 2012;8: e1002643 doi: 10.1371/journal.pgen.1002643 [DOI] [PMC free article] [PubMed] [Google Scholar]
57.Natsume T, Müller CA, Katou Y, Retkute R, Gierliński M, Araki H, et al. Kinetochores coordinate pericentromeric cohesion and early DNA replication by Cdc7-Dbf4 kinase recruitment. Mol Cell. 2013;50: 661–74. doi: 10.1016/j.molcel.2013.05.011 [DOI] [PMC free article] [PubMed] [Google Scholar]
58.Pohl TJ, Brewer BJ, Raghuraman MK. Functional Centromeres Determine the Activation Time of Pericentric Origins of DNA Replication in Saccharomyces cerevisiae. Newlon CS, editor. PLoS Genet. 2012;8: e1002677 doi: 10.1371/journal.pgen.1002677 [DOI] [PMC free article] [PubMed] [Google Scholar]
59.Brar G a, Weissman JS. Ribosome profiling reveals the what, when, where and how of protein synthesis. Nat Rev Mol Cell Biol. Nature Publishing Group; 2015;16: 651–64. doi: 10.1038/nrm4069 [DOI] [PMC free article] [PubMed] [Google Scholar]
60.Larance M, Lamond AI. Multidimensional proteomics for cell biology. Nat Rev Mol Cell Biol. Nature Publishing Group; 2015;16: 269–80. doi: 10.1038/nrm3970 [DOI] [PubMed] [Google Scholar]
61.Stults DM, Killen MW, Pierce HH, Pierce AJ. Genomic architecture and inheritance of human ribosomal RNA gene clusters. Genome Res. 2008;18: 13–18. doi: 10.1101/gr.6858507 [DOI] [PMC free article] [PubMed] [Google Scholar]
62.Gibbons JG, Branco AT, Godinho SA, Yu S, Lemos B. Concerted copy number variation balances ribosomal DNA dosage in human and mouse genomes. Proc Natl Acad Sci U S A. 2015;112: 2485–90. doi: 10.1073/pnas.1416878112 [DOI] [PMC free article] [PubMed] [Google Scholar]
63.Little RD, Platt TH, Schildkraut CL. Initiation and termination of DNA replication in human rRNA genes. Mol Cell Biol. 1993;13: 6600–13. doi: 10.1128/MCB.13.10.6600 [DOI] [PMC free article] [PubMed] [Google Scholar]
64.Cunniff C, Bassetti JA, Ellis NA. Bloom’s syndrome: Clinical spectrum, molecular pathogenesis, and cancer predisposition. Molecular Syndromology. 2017. pp. 4–23. doi: 10.1159/000452082 [DOI] [PMC free article] [PubMed] [Google Scholar]
65.Killen MW, Stults DM, Adachi N, Hanakahi L, Pierce AJ. Loss of Bloom syndrome protein destabilizes human gene cluster architecture. Hum Mol Genet. 2009;18: 3417–3428. doi: 10.1093/hmg/ddp282 [DOI] [PubMed] [Google Scholar]
66.Weitao T, Budd M, Campbell JL. Evidence that yeast SGS1, DNA2, SRS2, and FOB1 interact to maintain rDNA stability. Mutat Res. 2003;532: 157–72. doi: 10.1016/j.mrfmmm.2003.08.015 [DOI] [PubMed] [Google Scholar]
67.Stults DM, Killen MW, Williamson EP, Hourigan JS, Vargas HD, Arnold SM, et al. Human rRNA gene clusters are recombinational hotspots in cancer. Cancer Research. 2009. doi: 10.1158/0008-5472.CAN-09-2680 [DOI] [PubMed] [Google Scholar]
68.Pruitt SC, Qin M, Wang J, Kunnev D, Freeland A. A Signature of Genomic Instability Resulting from Deficient Replication Licensing. Symington LS, editor. PLOS Genet. 2017;13: e1006547 doi: 10.1371/journal.pgen.1006547 [DOI] [PMC free article] [PubMed] [Google Scholar]
69.Xu B, Li H, Perry JM, Singh VP, Unruh J, Yu Z, et al. Ribosomal DNA copy number loss and sequence variation in cancer. Eng C, editor. PLOS Genet. 2017;13: e1006771 doi: 10.1371/journal.pgen.1006771 [DOI] [PMC free article] [PubMed] [Google Scholar]
70.Trainor PA, Merrill AE. Ribosome biogenesis in skeletal development and the pathogenesis of skeletal disorders. Biochim Biophys Acta. 2014;1842: 769–78. doi: 10.1016/j.bbadis.2013.11.010 [DOI] [PMC free article] [PubMed] [Google Scholar]
71.The Treacher Collins Syndrome Collaborative Group. Positional cloning of a gene involved in the pathogenesis of Treacher Collins syndrome. The Treacher Collins Syndrome Collaborative Group. Nat Genet. 1996;12: 130–6. doi: 10.1038/ng0296-130 [DOI] [PubMed] [Google Scholar]
72.Dauwerse JG, Dixon J, Seland S, Ruivenkamp C a L, van Haeringen A, Hoefsloot LH, et al. Mutations in genes encoding subunits of RNA polymerases I and III cause Treacher Collins syndrome. Nat Genet. Nature Publishing Group; 2011;43: 20–2. doi: 10.1038/ng.724 [DOI] [PubMed] [Google Scholar]
73.Valdez BC, Henning D, So RB, Dixon J, Dixon MJ. The Treacher Collins syndrome (TCOF1) gene product is involved in ribosomal DNA gene transcription by interacting with upstream binding factor. Proc Natl Acad Sci U S A. 2004;101: 10709–14. doi: 10.1073/pnas.0402492101 [DOI] [PMC free article] [PubMed] [Google Scholar]
74.Hayano T, Yanagida M, Yamauchi Y, Shinkawa T, Isobe T, Takahashi N. Proteomic analysis of human Nop56p-associated pre-ribosomal ribonucleoprotein complexes. Possible link between Nop56p and the nucleolar protein treacle responsible for Treacher Collins syndrome. J Biol Chem. 2003;278: 34309–19. doi: 10.1074/jbc.M304304200 [DOI] [PubMed] [Google Scholar]
75.Dixon J, Jones NC, Sandell LL, Jayasinghe SM, Crane J, Rey J-P, et al. Tcof1/Treacle is required for neural crest cell formation and proliferation deficiencies that cause craniofacial abnormalities. Proc Natl Acad Sci U S A. 2006;103: 13403–8. doi: 10.1073/pnas.0603730103 [DOI] [PMC free article] [PubMed] [Google Scholar]
76.Ng SB, Buckingham KJ, Lee C, Bigham AW, Tabor HK, Dent KM, et al. Exome sequencing identifies the cause of a mendelian disorder. Nat Genet. Nature Publishing Group; 2010;42: 30–5. doi: 10.1038/ng.499 [DOI] [PMC free article] [PubMed] [Google Scholar]
77.Rainger J, Bengani H, Campbell L, Anderson E, Sokhi K, Lam W, et al. Miller (Genee-Wiedemann) syndrome represents a clinically and biochemically distinct subgroup of postaxial acrofacial dysostosis associated with partial deficiency of DHODH. Hum Mol Genet. 2012;21: 3969–83. doi: 10.1093/hmg/dds218 [DOI] [PubMed] [Google Scholar]
78.Lipton JM, Ellis SR. Diamond-Blackfan anemia: diagnosis, treatment, and molecular pathogenesis. Hematol Oncol Clin North Am. 2009;23: 261–82. doi: 10.1016/j.hoc.2009.01.004 [DOI] [PMC free article] [PubMed] [Google Scholar]
79.Narla A, Ebert BL. Ribosomopathies: human disorders of ribosome dysfunction. Blood. 2010;115: 3196–3205. doi: 10.1182/blood-2009-10-178129 [DOI] [PMC free article] [PubMed] [Google Scholar]
80.Vega H, Waisfisz Q, Gordillo M, Sakai N, Yanagihara I, Yamada M, et al. Roberts syndrome is caused by mutations in ESCO2, a human homolog of yeast ECO1 that is essential for the establishment of sister chromatid cohesion. Nat Genet. 2005;37: 468–70. doi: 10.1038/ng1548 [DOI] [PubMed] [Google Scholar]
81.Bose T, Lee KK, Lu S, Xu B, Harris B, Slaughter B, et al. Cohesin proteins promote ribosomal RNA production and protein translation in yeast and human cells. PLoS Genet. 2012;8: e1002749 doi: 10.1371/journal.pgen.1002749 [DOI] [PMC free article] [PubMed] [Google Scholar]
82.Boocock GRB, Morrison JA, Popovic M, Richards N, Ellis L, Durie PR, et al. Mutations in SBDS are associated with Shwachman-Diamond syndrome. Nat Genet. 2003;33: 97–101. doi: 10.1038/ng1062 [DOI] [PubMed] [Google Scholar]
83.Menne TF, Goyenechea B, Sánchez-Puig N, Wong CC, Tonkin LM, Ancliff PJ, et al. The Shwachman-Bodian-Diamond syndrome protein mediates translational activation of ribosomes in yeast. Nat Genet. 2007;39: 486–95. doi: 10.1038/ng1994 [DOI] [PubMed] [Google Scholar]
84.Moore JB, Farrar JE, Arceci RJ, Liu JM, Ellis SR. Distinct ribosome maturation defects in yeast models of Diamond-Blackfan anemia and Shwachman-Diamond syndrome. Haematologica. 2010;95: 57–64. doi: 10.3324/haematol.2009.012450 [DOI] [PMC free article] [PubMed] [Google Scholar]
85.Ridanpää M, van Eenennaam H, Pelin K, Chadwick R, Johnson C, Yuan B, et al. Mutations in the RNA component of RNase MRP cause a pleiotropic human disease, cartilage-hair hypoplasia. Cell. 2001;104: 195–203. doi: 10.1016/S0092-8674(01)00205-7 [DOI] [PubMed] [Google Scholar]
86.Welting TJM, van Venrooij WJ, Pruijn GJM. Mutual interactions between subunits of the human RNase MRP ribonucleoprotein complex. Nucleic Acids Res. 2004;32: 2138–46. doi: 10.1093/nar/gkh539 [DOI] [PMC free article] [PubMed] [Google Scholar]
87.Bergman S, Rodman TC, Sirlin JL. Measurement of Human Ribosomal DNA in Relation to a Possible Deficiency. Humangenetik. 1972;16: 337–340. [DOI] [PubMed] [Google Scholar]
88.Duff K, Huxley C. Targeting mutations to YACs by homologous recombination. Methods Mol Biol. 1996;54: 187–98. [DOI] [PubMed] [Google Scholar]
89.Laughery MF, Hunter T, Brown A, Hoopes J, Ostbye T, Shumaker T, et al. New vectors for simple and streamlined CRISPR-Cas9 genome editing in Saccharomyces cerevisiae. Yeast. 2015;32: 711–20. doi: 10.1002/yea.3098 [DOI] [PMC free article] [PubMed] [Google Scholar]
90.Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, et al. UCSF Chimera—a visualization system for exploratory research and analysis. J Comput Chem. 2004;25: 1605–12. doi: 10.1002/jcc.20084 [DOI] [PubMed] [Google Scholar]
91.Kwan EX, Wang XS, Amemiya HM, Brewer BJ, Raghuraman MK. rDNA Copy Number Variants Are Frequent Passenger Mutations in Saccharomyces cerevisiae Deletion Collections and de Novo Transformants. G3 (Bethesda). 2016;6: 2829–38. doi: 10.1534/g3.116.030296 [DOI] [PMC free article] [PubMed] [Google Scholar]
92.Iadonato SP, Gnirke A. RARE-cleavage analysis of YACs. Methods Mol Biol. 1996;54: 75–85. doi: 10.1017/CBO9781107415324.004 [DOI] [PubMed] [Google Scholar]
93.Hoffman CS, Winston F. A ten-minute DNA preparation from yeast efficiently releases autonomous plasmids for transformaion of Escherichia coli. Gene. 1987;57: 267–272. doi: 10.1016/0378-1119(87)90131-4 [DOI] [PubMed] [Google Scholar]
94.Peng J, Raghuraman MK, Feng W. Analysis of ssDNA Gaps and DSBs in Genetically Unstable Yeast Cultures. Methods Mol Biol. 2014;1170: 501–515. doi: 10.1007/978-1-4939-0888-2_27 [DOI] [PMC free article] [PubMed] [Google Scholar]
95.Siow CC, Nieduszynska SR, Müller CA, Nieduszynski CA. OriDB, the DNA replication origin database updated and extended. Nucleic Acids Res. 2012;40: D682–6. doi: 10.1093/nar/gkr1091 [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

S1 Fig. Structural comparison of H. sapiens and S. cerevisiae ORC subunits.

(TIF)

Click here for additional data file.^{(3.8MB, tif)}

S2 Fig. Position of H. sapiens Orc4-Tyr174 and S. cerevisiae Orc4-Tyr232.

(A).

(B).

Structural models of ScOrc1-6 and ScCdc6 in complex with a double stranded DNA molecule. The right panel focuses on Tyr232 of ScOrc4, with the side chain of this amino acid depicted as red spheres.

(TIF)

Click here for additional data file.^{(4MB, tif)}

S3 Fig. Comparison of ssDNA profiles for ORC4 and orc4^Y232C chromosomes.

(TIF)

Click here for additional data file.^{(1.4MB, tif)}

S4 Fig. The rDNA copy number in orc4^Y232C cells stabilizes at ~30 copies.

(TIF)

Click here for additional data file.^{(828.1KB, tif)}

S5 Fig. CHEF gel analysis of variation in rDNA copy number of five isolates (a-e) of cdc45^P542L.

Left, ethidium bromide stained image; right, Southern blot hybridization for Chr XII. All five isolates had a smaller Chr XII than CDC45 due to loss of rDNA repeats.

(TIF)

Click here for additional data file.^{(202.2KB, tif)}

S6 Fig. The orc4^Y232C rDNA copy number phenotype is not rescued by an additional copy of orc4^Y232C.

(TIF)

Click here for additional data file.^{(591.1KB, tif)}

S7 Fig. Sequence comparison of the ACS of BY4741 and RM11-1a rDNA origin, rARS.

Red arrow indicates the polymorphism in the ACS.

(TIF)

Click here for additional data file.^{(126.7KB, tif)}

S8 Fig. The slow growth phenotype is exacerbated in orc4^Y232C rDNA^RM cells.

(TIF)

Click here for additional data file.^{(36.7KB, tif)}

S9 Fig. Comparison of ssDNA profiles for ORC4 rDNA^RM and orc4^Y232C rDNA^RM chromosomes.

(TIF)

Click here for additional data file.^{(1.5MB, tif)}

S10 Fig. Pairwise comparisons of the relative area under the peak at each origin measured in ssDNA profiles.

(TIF)

Click here for additional data file.^{(285.8KB, tif)}

S11 Fig. The total nucleic acid content of cells is resolved by gel electrophoresis.

(TIF)

Click here for additional data file.^{(369.2KB, tif)}

S12 Fig. Relative fluorescence of ORC4 rDNA^BY and orc4^Y232C rDNA^BY cells harboring Rpl10-GFP.

(TIF)

Click here for additional data file.^{(253.6KB, tif)}

S13 Fig. Origins proximal to centromeres are less active in orc4^Y232C cells.

(TIF)

Click here for additional data file.^{(684.6KB, tif)}

S1 Table. Yeast strains and plasmids used in this study.

(PDF)

Click here for additional data file.^{(78.2KB, pdf)}

S1 File. Python script used to calculate the area under ssDNA peaks.

(PY)

Click here for additional data file.^{(14KB, py)}

Data Availability Statement

[pgen.1007041.ref001] 1.Stinchcomb DT, Struhl K, Davis RW. Isolation and characterisation of a yeast chromosomal replicator. Nature. 1979;282: 39–43. doi: 10.1038/282039a0 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref002] 2.Liachko I, Youngblood RA, Keich U, Dunham MJ. High-resolution mapping, characterization, and optimization of autonomously replicating sequences in yeast. Genome Res. 2013;23: 698–704. doi: 10.1101/gr.144659.112 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref003] 3.Bell SP, Stillman B. ATP-dependent recognition of eukaryotic origins of DNA replication by a multiprotein complex. Nature. 1992;357: 128–34. doi: 10.1038/357128a0 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref004] 4.Liang C, Weinreich M, Stillman B. ORC and Cdc6p interact and determine the frequency of initiation of DNA replication in the genome. Cell. 1995;81: 667–76. doi: 10.1016/0092-8674(95)90528-6 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref005] 5.Tanaka S, Diffley JFX. Interdependent nuclear accumulation of budding yeast Cdt1 and Mcm2-7 during G1 phase. Nat Cell Biol. 2002;4: 198–207. doi: 10.1038/ncb757 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref006] 6.Cvetic C, Walter JC. Eukaryotic origins of DNA replication: Could you please be more specific? Semin Cell Dev Biol. 2005;16: 343–353. doi: 10.1016/j.semcdb.2005.02.009 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref007] 7.Li H, Stillman B. The Origin Recognition Complex: A Biochemical and Structural View. Subcellular Biochemistry. 2012. pp. 37–58. doi: 10.1007/978-94-007-4572-8_3 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref008] 8.Tocilj A, On KF, Yuan Z, Sun J, Elkayam E, Li H, et al. Structure of the active form of human origin recognition complex and its ATPase motor module. Elife. 2017;6: e20818 doi: 10.7554/eLife.20818 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref009] 9.Yuan Z, Riera A, Bai L, Sun J, Nandi S, Spanos C, et al. Structural basis of Mcm2-7 replicative helicase loading by ORC-Cdc6 and Cdt1. Nat Struct Mol Biol. 2017; doi: 10.1038/nsmb.3372 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref010] 10.Liang C, Stillman B. Persistent initiation of DNA replication and chromatin-bound MCM proteins during the cell cycle in cdc6 mutants. Genes Dev. 1997;11: 3375–86. doi: 10.1101/gad.11.24.3375 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref011] 11.Fragkos M, Ganier O, Coulombe P, Méchali M. DNA replication origin activation in space and time. Nat Rev Mol Cell Biol. Nature Publishing Group; 2015;16: 360–74. doi: 10.1038/nrm4002 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref012] 12.Randell JCW, Bowers JL, Rodríguez HK, Bell SP. Sequential ATP hydrolysis by Cdc6 and ORC directs loading of the Mcm2-7 helicase. Mol Cell. 2006;21: 29–39. doi: 10.1016/j.molcel.2005.11.023 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref013] 13.Cocker JH, Piatti S, Santocanale C, Nasmyth K, Diffley JF. An essential role for the Cdc6 protein in forming the pre-replicative complexes of budding yeast. Nature. 1996;379: 180–2. doi: 10.1038/379180a0 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref014] 14.Diffley JF, Cocker JH, Dowell SJ, Rowley A. Two steps in the assembly of complexes at yeast replication origins in vivo. Cell. 1994;78: 303–16. [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref015] 15.Speck C, Chen Z, Li H, Stillman B. ATPase-dependent cooperative binding of ORC and Cdc6 to origin DNA. Nat Struct Mol Biol. 2005;12: 965–71. doi: 10.1038/nsmb1002 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref016] 16.Chen S, de Vries MA, Bell SP. Orc6 is required for dynamic recruitment of Cdt1 during repeated Mcm2-7 loading. Genes Dev. 2007;21: 2897–907. doi: 10.1101/gad.1596807 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref017] 17.Sclafani RA, Holzen TM. Cell cycle regulation of DNA replication. Annu Rev Genet. 2007;41: 237–80. doi: 10.1146/annurev.genet.41.110306.130308 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref018] 18.Stillman B. Origin recognition and the chromosome cycle. FEBS Lett. 2005;579: 877–84. doi: 10.1016/j.febslet.2004.12.011 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref019] 19.Ilves I, Petojevic T, Pesavento JJ, Botchan MR. Activation of the MCM2-7 Helicase by Association with Cdc45 and GINS Proteins. Mol Cell. Elsevier Ltd; 2010;37: 247–258. doi: 10.1016/j.molcel.2009.12.030 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref020] 20.Bicknell LS, Bongers EMHF, Leitch A, Brown S, Schoots J, Harley ME, et al. Mutations in the pre-replication complex cause Meier-Gorlin syndrome. Nat Genet. 2011;43: 356–9. doi: 10.1038/ng.775 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref021] 21.Guernsey DL, Matsuoka M, Jiang H, Evans S, Macgillivray C, Nightingale M, et al. Mutations in origin recognition complex gene ORC4 cause Meier-Gorlin syndrome. Nat Genet. 2011;43: 360–364. doi: 10.1038/ng.777 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref022] 22.Gorlin RJ, Cervenka J, Moller K, Horrobin M, Witkop CJ. Malformation syndromes. A selected miscellany. Birth Defects Orig Artic Ser. 1975;11: 39–50. [PubMed] [Google Scholar]

[pgen.1007041.ref023] 23.Meier Z, Poschiavo, Rothschild M. Case of arthrogryposis multiplex congenita with mandibulofacial dysostosis (Franceschetti syndrome). Helv Paediatr Acta. 1959;14: 213–6. [PubMed] [Google Scholar]

[pgen.1007041.ref024] 24.Vetro A, Savasta S, Russo Raucci A, Cerqua C, Sartori G, Limongelli I, et al. MCM5: a new actor in the link between DNA replication and Meier-Gorlin syndrome. Eur J Hum Genet. Nature Publishing Group; 2017; 1–5. doi: 10.1038/ejhg.2017.5 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref025] 25.Burrage LC, Charng WL, Eldomery MK, Willer JR, Davis EE, Lugtenberg D, et al. De Novo GMNN Mutations Cause Autosomal-Dominant Primordial Dwarfism Associated with Meier-Gorlin Syndrome. Am J Hum Genet. The American Society of Human Genetics; 2015;97: 904–913. doi: 10.1016/j.ajhg.2015.11.006 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref026] 26.Fenwick AL, Kliszczak M, Cooper F, Murray J, Sanchez-Pulido L, Twigg SRF, et al. Mutations in CDC45, Encoding an Essential Component of the Pre-initiation Complex, Cause Meier-Gorlin Syndrome and Craniosynostosis. Am J Hum Genet. American Society of Human Genetics; 2016;99: 125–138. doi: 10.1016/j.ajhg.2016.05.019 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref027] 27.Stiff T, Alagoz M, Alcantara D, Outwin E, Brunner HG, Bongers EMHF, et al. Deficiency in origin licensing proteins impairs cilia formation: implications for the aetiology of Meier-Gorlin syndrome. PLoS Genet. 2013;9: e1003360 doi: 10.1371/journal.pgen.1003360 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref028] 28.Bicknell LS, Walker S, Klingseisen A, Stiff T, Leitch A, Kerzendorfer C, et al. Mutations in ORC1, encoding the largest subunit of the origin recognition complex, cause microcephalic primordial dwarfism resembling Meier-Gorlin syndrome. Nat Genet. 2011;43: 350–355. doi: 10.1038/ng.776 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref029] 29.Hossain M, Stillman B. Meier-Gorlin syndrome mutations disrupt an Orc1 CDK inhibitory domain and cause centrosome reduplication. Genes Dev. 2012;26: 1797–810. doi: 10.1101/gad.197178.112 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref030] 30.Ide S, Watanabe K, Watanabe H, Shirahige K, Kobayashi T, Maki H. Abnormality in initiation program of DNA replication is monitored by the highly repetitive rRNA gene array on chromosome XII in budding yeast. Mol Cell Biol. 2007;27: 568–78. doi: 10.1128/MCB.00731-06 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref031] 31.Feng W, Collingwood D, Boeck ME, Fox LA, Alvino GM, Fangman WL, et al. Genomic mapping of single-stranded DNA in hydroxyurea-challenged yeasts identifies origins of replication. Nat Cell Biol. 2006;8: 148–55. doi: 10.1038/ncb1358 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref032] 32.McCune HJ, Danielson LS, Alvino GM, Collingwood D, Delrow JJ, Fangman WL, et al. The temporal program of chromosome replication: genomewide replication in clb5Δ Saccharomyces cerevisiae. Genetics. 2008;180: 1833–47. doi: 10.1534/genetics.108.094359 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref033] 33.Liu Y-T, Fan H-F, Kachroo AH, Jayaram M, Rowley PA, Sau S, et al. The Partitioning and Copy Number Control Systems of the Selfish Yeast Plasmid: An Optimized Molecular Design for Stable Persistence in Host Cells. Microbiol Spectr. 2014;2: 1–32. doi: 10.1128/microbiolspec.PLAS-0003-2013 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref034] 34.Williamson D. The curious history of yeast mitochondrial DNA. Nat Rev Genet. 2002;3: 475–81. doi: 10.1038/nrg814 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref035] 35.Schweizer E, MacKechnie C, Halvorson HO. The redundancy of ribosomal and transfer RNA genes in Saccharomyces cerevisiae. J Mol Biol. 1969;40: 261–277. doi: 10.1016/0022-2836(69)90474-4 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref036] 36.Chen XJ, Butow R a. The organization and inheritance of the mitochondrial genome. Nat Rev Genet. 2005;6: 815–825. doi: 10.1038/nrg1708 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref037] 37.Petes TD. Yeast ribosomal DNA genes are located on chromosome XII. Proc Natl Acad Sci U S A. 1979;76: 410–4. doi: 10.1073/pnas.76.1.410 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref038] 38.Kobayashi T. Strategies to maintain the stability of the ribosomal RNA gene repeats—collaboration of recombination, cohesion, and condensation. Genes Genet Syst. 2006;81: 155–161. JST.JSTAGE/ggs/81.155 [pii] [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref039] 39.Micklem G, Rowley A, Harwood J, Nasmyth K, Diffley JF. Yeast origin recognition complex is involved in DNA replication and transcriptional silencing. Nature. 1993;366: 87–89. doi: 10.1038/366087a0 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref040] 40.Futcher AB, Cox BS. Maintenance of the 2 microns circle plasmid in populations of Saccharomyces cerevisiae. J Bacteriol. 1983;154: 612–622. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref041] 41.Warner JR. The economics of ribosome biosynthesis in yeast. Trends Biochem Sci. 1999;24: 437–440. doi: 10.1016/S0968-0004(99)01460-7 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref042] 42.Woolford JL, Baserga SJ. Ribosome biogenesis in the yeast Saccharomyces cerevisiae. Genetics. 2013;195: 643–681. doi: 10.1534/genetics.113.153197 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref043] 43.Simon AC, Sannino V, Costanzo V, Pellegrini L. Structure of human Cdc45 and implications for CMG helicase function. Nat Commun. Nature Publishing Group; 2016;7: 11638 doi: 10.1038/ncomms11638 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref044] 44.Brewer BJ, Fangman WL. A replication fork barrier at the 3′ end of yeast ribosomal RNA genes. Cell. 1988;55: 637–643. doi: 10.1016/0092-8674(88)90222-X [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref045] 45.Muller M, Lucchini R, Sogo JM. Replication of yeast rDNA initiates downstream of transcriptionally active genes. Mol Cell. 2000;5: 767–777. doi: 10.1016/S1097-2765(00)80317-2 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref046] 46.Foss EJ, Lao U, Dalrymple E, Adrianse RL, Loe T, Bedalov A. SIR2 suppresses replication gaps and genome instability by balancing replication between repetitive and unique sequences. Proc Natl Acad Sci U S A. 2017;114: 552–557. doi: 10.1073/pnas.1614781114 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref047] 47.Brewer BJ, Fangman WL. The localization of replication origins on ARS plasmids in S. cerevisiae. Cell. 1987;51: 463–471. doi: 10.1016/0092-8674(87)90642-8 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref048] 48.Shimada K, Pasero P, Gasser SM. ORC and the intra-S-phase checkpoint: A threshold regulates Rad53p activation in S phase. Genes Dev. 2002;16: 3236–3252. doi: 10.1101/gad.239802 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref049] 49.Kwan EX, Foss EJ, Tsuchiyama S, Alvino GM, Kruglyak L, Kaeberlein M, et al. A Natural Polymorphism in rDNA Replication Origins Links Origin Activation with Calorie Restriction and Lifespan. PLoS Genet. 2013;9: e1003329 doi: 10.1371/journal.pgen.1003329 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref050] 50.Miller CA, Umek RM, Kowalski D. The inefficient replication origin from yeast ribosomal DNA is naturally impaired in the ARS consensus sequence and in DNA unwinding. Nucleic Acids Res. 1999;27: 3921–3930. doi: 10.1093/nar/27.19.3921 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref051] 51.Rivin CJ, Fangman WL. Replication fork rate and origin activation during the S phase of Saccharomyces cerevisiae. J Cell Biol. 1980;85: 108–15. [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref052] 52.French SL, Osheim YN, Cioci F, Nomura M, Beyer AL. In exponentially growing Saccharomyces cerevisiae cells, rRNA synthesis is determined by the summed RNA polymerase I loading rate rather than by the number of active genes. Mol Cell Biol. 2003;23: 1558–68. doi: 10.1128/MCB.23.5.1558-1568.2003 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref053] 53.Lo C-A, Kays I, Emran F, Lin T-J, Cvetkovska V, Chen BE. Quantification of Protein Levels in Single Living Cells. Cell Rep. The Authors; 2015;13: 2634–44. doi: 10.1016/j.celrep.2015.11.048 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref054] 54.Sung M-K, Reitsma JM, Sweredoski MJ, Hess S, Deshaies RJ. Ribosomal proteins produced in excess are degraded by the ubiquitin-proteasome system. Mol Biol Cell. 2016;27: 2642–52. doi: 10.1091/mbc.E16-05-0290 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref055] 55.Yoshida K, Bacal J, Desmarais D, Padioleau I, Tsaponina O, Chabes A, et al. The Histone Deacetylases Sir2 and Rpd3 Act on Ribosomal DNA to Control the Replication Program in Budding Yeast. Mol Cell. 2014;54: 691–697. doi: 10.1016/j.molcel.2014.04.032 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref056] 56.Blitzblau HG, Chan CS, Hochwagen A, Bell SP. Separation of DNA Replication from the Assembly of Break-Competent Meiotic Chromosomes. Hawley RS, editor. PLoS Genet. 2012;8: e1002643 doi: 10.1371/journal.pgen.1002643 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref057] 57.Natsume T, Müller CA, Katou Y, Retkute R, Gierliński M, Araki H, et al. Kinetochores coordinate pericentromeric cohesion and early DNA replication by Cdc7-Dbf4 kinase recruitment. Mol Cell. 2013;50: 661–74. doi: 10.1016/j.molcel.2013.05.011 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref058] 58.Pohl TJ, Brewer BJ, Raghuraman MK. Functional Centromeres Determine the Activation Time of Pericentric Origins of DNA Replication in Saccharomyces cerevisiae. Newlon CS, editor. PLoS Genet. 2012;8: e1002677 doi: 10.1371/journal.pgen.1002677 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref059] 59.Brar G a, Weissman JS. Ribosome profiling reveals the what, when, where and how of protein synthesis. Nat Rev Mol Cell Biol. Nature Publishing Group; 2015;16: 651–64. doi: 10.1038/nrm4069 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref060] 60.Larance M, Lamond AI. Multidimensional proteomics for cell biology. Nat Rev Mol Cell Biol. Nature Publishing Group; 2015;16: 269–80. doi: 10.1038/nrm3970 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref061] 61.Stults DM, Killen MW, Pierce HH, Pierce AJ. Genomic architecture and inheritance of human ribosomal RNA gene clusters. Genome Res. 2008;18: 13–18. doi: 10.1101/gr.6858507 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref062] 62.Gibbons JG, Branco AT, Godinho SA, Yu S, Lemos B. Concerted copy number variation balances ribosomal DNA dosage in human and mouse genomes. Proc Natl Acad Sci U S A. 2015;112: 2485–90. doi: 10.1073/pnas.1416878112 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref063] 63.Little RD, Platt TH, Schildkraut CL. Initiation and termination of DNA replication in human rRNA genes. Mol Cell Biol. 1993;13: 6600–13. doi: 10.1128/MCB.13.10.6600 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref064] 64.Cunniff C, Bassetti JA, Ellis NA. Bloom’s syndrome: Clinical spectrum, molecular pathogenesis, and cancer predisposition. Molecular Syndromology. 2017. pp. 4–23. doi: 10.1159/000452082 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref065] 65.Killen MW, Stults DM, Adachi N, Hanakahi L, Pierce AJ. Loss of Bloom syndrome protein destabilizes human gene cluster architecture. Hum Mol Genet. 2009;18: 3417–3428. doi: 10.1093/hmg/ddp282 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref066] 66.Weitao T, Budd M, Campbell JL. Evidence that yeast SGS1, DNA2, SRS2, and FOB1 interact to maintain rDNA stability. Mutat Res. 2003;532: 157–72. doi: 10.1016/j.mrfmmm.2003.08.015 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref067] 67.Stults DM, Killen MW, Williamson EP, Hourigan JS, Vargas HD, Arnold SM, et al. Human rRNA gene clusters are recombinational hotspots in cancer. Cancer Research. 2009. doi: 10.1158/0008-5472.CAN-09-2680 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref068] 68.Pruitt SC, Qin M, Wang J, Kunnev D, Freeland A. A Signature of Genomic Instability Resulting from Deficient Replication Licensing. Symington LS, editor. PLOS Genet. 2017;13: e1006547 doi: 10.1371/journal.pgen.1006547 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref069] 69.Xu B, Li H, Perry JM, Singh VP, Unruh J, Yu Z, et al. Ribosomal DNA copy number loss and sequence variation in cancer. Eng C, editor. PLOS Genet. 2017;13: e1006771 doi: 10.1371/journal.pgen.1006771 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref070] 70.Trainor PA, Merrill AE. Ribosome biogenesis in skeletal development and the pathogenesis of skeletal disorders. Biochim Biophys Acta. 2014;1842: 769–78. doi: 10.1016/j.bbadis.2013.11.010 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref071] 71.The Treacher Collins Syndrome Collaborative Group. Positional cloning of a gene involved in the pathogenesis of Treacher Collins syndrome. The Treacher Collins Syndrome Collaborative Group. Nat Genet. 1996;12: 130–6. doi: 10.1038/ng0296-130 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref072] 72.Dauwerse JG, Dixon J, Seland S, Ruivenkamp C a L, van Haeringen A, Hoefsloot LH, et al. Mutations in genes encoding subunits of RNA polymerases I and III cause Treacher Collins syndrome. Nat Genet. Nature Publishing Group; 2011;43: 20–2. doi: 10.1038/ng.724 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref073] 73.Valdez BC, Henning D, So RB, Dixon J, Dixon MJ. The Treacher Collins syndrome (TCOF1) gene product is involved in ribosomal DNA gene transcription by interacting with upstream binding factor. Proc Natl Acad Sci U S A. 2004;101: 10709–14. doi: 10.1073/pnas.0402492101 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref074] 74.Hayano T, Yanagida M, Yamauchi Y, Shinkawa T, Isobe T, Takahashi N. Proteomic analysis of human Nop56p-associated pre-ribosomal ribonucleoprotein complexes. Possible link between Nop56p and the nucleolar protein treacle responsible for Treacher Collins syndrome. J Biol Chem. 2003;278: 34309–19. doi: 10.1074/jbc.M304304200 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref075] 75.Dixon J, Jones NC, Sandell LL, Jayasinghe SM, Crane J, Rey J-P, et al. Tcof1/Treacle is required for neural crest cell formation and proliferation deficiencies that cause craniofacial abnormalities. Proc Natl Acad Sci U S A. 2006;103: 13403–8. doi: 10.1073/pnas.0603730103 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref076] 76.Ng SB, Buckingham KJ, Lee C, Bigham AW, Tabor HK, Dent KM, et al. Exome sequencing identifies the cause of a mendelian disorder. Nat Genet. Nature Publishing Group; 2010;42: 30–5. doi: 10.1038/ng.499 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref077] 77.Rainger J, Bengani H, Campbell L, Anderson E, Sokhi K, Lam W, et al. Miller (Genee-Wiedemann) syndrome represents a clinically and biochemically distinct subgroup of postaxial acrofacial dysostosis associated with partial deficiency of DHODH. Hum Mol Genet. 2012;21: 3969–83. doi: 10.1093/hmg/dds218 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref078] 78.Lipton JM, Ellis SR. Diamond-Blackfan anemia: diagnosis, treatment, and molecular pathogenesis. Hematol Oncol Clin North Am. 2009;23: 261–82. doi: 10.1016/j.hoc.2009.01.004 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref079] 79.Narla A, Ebert BL. Ribosomopathies: human disorders of ribosome dysfunction. Blood. 2010;115: 3196–3205. doi: 10.1182/blood-2009-10-178129 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref080] 80.Vega H, Waisfisz Q, Gordillo M, Sakai N, Yanagihara I, Yamada M, et al. Roberts syndrome is caused by mutations in ESCO2, a human homolog of yeast ECO1 that is essential for the establishment of sister chromatid cohesion. Nat Genet. 2005;37: 468–70. doi: 10.1038/ng1548 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref081] 81.Bose T, Lee KK, Lu S, Xu B, Harris B, Slaughter B, et al. Cohesin proteins promote ribosomal RNA production and protein translation in yeast and human cells. PLoS Genet. 2012;8: e1002749 doi: 10.1371/journal.pgen.1002749 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref082] 82.Boocock GRB, Morrison JA, Popovic M, Richards N, Ellis L, Durie PR, et al. Mutations in SBDS are associated with Shwachman-Diamond syndrome. Nat Genet. 2003;33: 97–101. doi: 10.1038/ng1062 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref083] 83.Menne TF, Goyenechea B, Sánchez-Puig N, Wong CC, Tonkin LM, Ancliff PJ, et al. The Shwachman-Bodian-Diamond syndrome protein mediates translational activation of ribosomes in yeast. Nat Genet. 2007;39: 486–95. doi: 10.1038/ng1994 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref084] 84.Moore JB, Farrar JE, Arceci RJ, Liu JM, Ellis SR. Distinct ribosome maturation defects in yeast models of Diamond-Blackfan anemia and Shwachman-Diamond syndrome. Haematologica. 2010;95: 57–64. doi: 10.3324/haematol.2009.012450 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref085] 85.Ridanpää M, van Eenennaam H, Pelin K, Chadwick R, Johnson C, Yuan B, et al. Mutations in the RNA component of RNase MRP cause a pleiotropic human disease, cartilage-hair hypoplasia. Cell. 2001;104: 195–203. doi: 10.1016/S0092-8674(01)00205-7 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref086] 86.Welting TJM, van Venrooij WJ, Pruijn GJM. Mutual interactions between subunits of the human RNase MRP ribonucleoprotein complex. Nucleic Acids Res. 2004;32: 2138–46. doi: 10.1093/nar/gkh539 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref087] 87.Bergman S, Rodman TC, Sirlin JL. Measurement of Human Ribosomal DNA in Relation to a Possible Deficiency. Humangenetik. 1972;16: 337–340. [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref088] 88.Duff K, Huxley C. Targeting mutations to YACs by homologous recombination. Methods Mol Biol. 1996;54: 187–98. [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref089] 89.Laughery MF, Hunter T, Brown A, Hoopes J, Ostbye T, Shumaker T, et al. New vectors for simple and streamlined CRISPR-Cas9 genome editing in Saccharomyces cerevisiae. Yeast. 2015;32: 711–20. doi: 10.1002/yea.3098 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref090] 90.Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, et al. UCSF Chimera—a visualization system for exploratory research and analysis. J Comput Chem. 2004;25: 1605–12. doi: 10.1002/jcc.20084 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref091] 91.Kwan EX, Wang XS, Amemiya HM, Brewer BJ, Raghuraman MK. rDNA Copy Number Variants Are Frequent Passenger Mutations in Saccharomyces cerevisiae Deletion Collections and de Novo Transformants. G3 (Bethesda). 2016;6: 2829–38. doi: 10.1534/g3.116.030296 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref092] 92.Iadonato SP, Gnirke A. RARE-cleavage analysis of YACs. Methods Mol Biol. 1996;54: 75–85. doi: 10.1017/CBO9781107415324.004 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref093] 93.Hoffman CS, Winston F. A ten-minute DNA preparation from yeast efficiently releases autonomous plasmids for transformaion of Escherichia coli. Gene. 1987;57: 267–272. doi: 10.1016/0378-1119(87)90131-4 [DOI] [PubMed] [Google Scholar]

[pgen.1007041.ref094] 94.Peng J, Raghuraman MK, Feng W. Analysis of ssDNA Gaps and DSBs in Genetically Unstable Yeast Cultures. Methods Mol Biol. 2014;1170: 501–515. doi: 10.1007/978-1-4939-0888-2_27 [DOI] [PMC free article] [PubMed] [Google Scholar]

[pgen.1007041.ref095] 95.Siow CC, Nieduszynska SR, Müller CA, Nieduszynski CA. OriDB, the DNA replication origin database updated and extended. Nucleic Acids Res. 2012;40: D682–6. doi: 10.1093/nar/gkr1091 [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Defective replication initiation results in locus specific chromosome breakage and a ribosomal RNA deficiency in yeast

Joseph C Sanchez

Elizabeth X Kwan

Thomas J Pohl

Haley M Amemiya

M K Raghuraman

Bonita J Brewer

Roles

Abstract

Author summary

Introduction

Results

The slow growth of yeast with the orc4Y232C allele is due to a delay in completing S phase

Fig 1. Growth and cell cycle characterization of orc4Y232C cells.

The orc4Y232C mutation results in reduced copy number and origin activity at the rDNA locus

Fig 2. Reduction of rDNA copy number in orc4Y232C cells.

Fig 3. Origin initiation at the rDNA locus is reduced in orc4Y232C cells.

Fig 4. orc4Y232C cells with a less efficient rARS exhibit exacerbated growth and rDNA phenotypes.

The initiation defect in orc4Y232C rDNARM leads to chromosome XII breakage

The further loss of rDNA repeats in orc4Y232C rDNARM cells leads to a reduction in ribosomal components and an increased sensitivity to the ribosome inhibiting drug cycloheximide

Fig 5. The orc4Y232C rDNARM strain has reduced ribosomal components and is sensitive to the ribosome inhibiting drug cycloheximide.

Discussion

Materials and methods

Yeast strains and plasmids

Structural analysis of protein models

Flow cytometry for cell cycle analysis

Contour-clamped homogeneous electric field (CHEF) gel analysis

Two-Dimensional (2D) agarose gel analysis

Quantitative Southern blotting

Long term growth experiments

Quantitative hybrid Southern/northern blotting

Flow cytometry analysis for cells expressing GFP tagged Rpl10

Mapping of single stranded DNA

ssDNA peak area script description

Supporting information

Acknowledgments

Data Availability

Funding Statement

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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Links to NCBI Databases

The slow growth of yeast with the orc4^Y232C allele is due to a delay in completing S phase

Fig 1. Growth and cell cycle characterization of orc4^Y232C cells.

The orc4^Y232C mutation results in reduced copy number and origin activity at the rDNA locus

Fig 2. Reduction of rDNA copy number in orc4^Y232C cells.

Fig 3. Origin initiation at the rDNA locus is reduced in orc4^Y232C cells.

Fig 4. orc4^Y232C cells with a less efficient rARS exhibit exacerbated growth and rDNA phenotypes.

The initiation defect in orc4^Y232C rDNA^RM leads to chromosome XII breakage

The further loss of rDNA repeats in orc4^Y232C rDNA^RM cells leads to a reduction in ribosomal components and an increased sensitivity to the ribosome inhibiting drug cycloheximide

Fig 5. The orc4^Y232C rDNA^RM strain has reduced ribosomal components and is sensitive to the ribosome inhibiting drug cycloheximide.