A novel cell response triggered by interphase centromere structural instability

Centromeres are specialized chromosome domains that are responsible for chromosome segregation during meiosis and mitosis. They assemble around repetitive DNA sequences in a complex structure that has yet to be fully elucidated. A simplistic view involves the division of this domain into two areas: the central core region or centromeric chromatin (Schueler and Sullivan, 2006) and the flanking heterochromatic regions, which are called pericentromeres. The protein composition of the central core region varies between interphase and mitosis. In this model, constitutive proteins are permanently associated with the centromere even during interphase, whereas facultative proteins are recruited only during mitosis to assemble the kinetochore, which is the site of microtubule attachment. As such, the central core region serves as the assembly platform for the kinetochore. A specific feature of the chromatin structure of the core centromere is that it contains interspersed blocks of nucleosomes that contain histone H3 and a histone H3 variant called centromeric protein (CENP) A in human cells (Blower et al., 2002). In addition to histones, six constitutive proteins named CENP-A, -B, -C, -H, -I, and hMis12 are known as the major components of the interphase centromeric chromatin. However, another set of 11 proteins associated with the CENP-A–containing nucleosomes or with the CENP-H–I complex has recently been described (Foltz et al., 2006; Okada et al., 2006).

Cajal bodies (CBs) are nuclear domains that were discovered in 1903 by the Spanish physiologist Santiago Ramón y Cajal (Gall, 2003). These bodies are concentrates of several proteins and small nuclear ribonucleoproteins (Matera and Shpargel, 2006). Among these proteins, coilin was described in the early 1990s as the major component of CBs (Raska et al., 1991), although its precise biological activity remains elusive. Orthologues of human coilin are known in many vertebrates, including the mouse (Tucker et al., 2000), Xenopus laevis (Tuma et al., 1993), Danio rerio (Tucker et al., 2000), Arabidopsis thaliana (Collier et al., 2006), and Drosophila melanogaster (Liu, J.L., and J.G. Gall, personal communication). Coilin is not strictly essential for mouse embryonic development, although a substantial reduction of viability has been observed in inbred homozygous embryos (Tucker et al., 2001). Coilin contains nuclear and nucleolar localization domains, an arginine-glycine (RG)–rich box, and an autointeraction domain that facilitates CB formation (Hebert and Matera, 2000). The formation of CBs depends, at least in part, on the autointeraction domain and on posttranslational modifications of coilin. Indeed, hyperphosphorylation considerably reduces the coilin autointeraction, which leads to CB disassembly during mitosis (Hebert and Matera, 2000; Shpargel et al., 2003). The biological function of coilin within CBs remains mysterious, and its additional diffuse staining in the nucleoplasm has been proposed to be the mark of still unrevealed CB-independent activity (Matera and Frey, 1998).

Herpes simplex virus type 1 (HSV-1) infection of cultured cells induces the destabilization of centromeres during interphase, preventing the assembly of the kinetochore and the binding of microtubules during mitosis (Everett et al., 1999a). The factor responsible for this centromere destabilization is the viral protein infected cell protein 0 (ICP0). ICP0 is a RING finger nuclear protein with characterized E3 ubiquitin ligase activity (for review see Hagglund and Roizman, 2004). As soon as it enters the nucleus, ICP0 temporarily localizes to centromeres and induces the proteasomal degradation of CENP-A, -B, and -C (Everett et al., 1999a; Lomonte et al., 2001; Lomonte and Morency, 2007). Thus, ICP0-induced degradation of essential constitutive CENPs during interphase is likely to modify the structure of the central core region extensively, thereby preventing the assembly of the kinetochore. As a consequence, cells that express ICP0 just before entering mitosis are stalled in early mitosis and eventually suffer premature cell division without chromosomal segregation, leading to aneuploidy (Lomonte and Everett, 1999). Although the biological significance of ICP0-induced centromere destabilization is unclear, from the cellular viewpoint, ICP0 is of exceptional interest as a unique tool for studying centromere structure and the putative cell mechanisms implicated in centromere architectural maintenance. Indeed, although it is known that the cell uses kinetochore surveillance mechanisms during mitosis, such as the mitotic checkpoints (for review see Cleveland et al., 2003), it remains unknown whether the cell is able to detect centromeric structural defects in interphase.

In this study, using ICP0 as a tool, we reveal a previously unreported and unexpected cell response to human and mouse centromeres that have sustained structural modifications during interphase. This response is characterized by the accumulation at damaged centromeres of two CB proteins, coilin and fibrillarin, and one CB-associated gemini of CB (gems) nuclear domain protein, the survival motor neuron (SMN) protein. We show that this response does not implicate the entire CBs and/or gems. In addition, we confirm the physical association between coilin and centromere higher order type I α-satellite (α-SAT) DNA. Using siRNA against CENPs, we demonstrate that depletion of CENP-B leads to the accumulation of coilin at centromeres. This confirms the existence of a cell response that is triggered by interphase centromere instability, for which we propose the term interphase centromere damage response (iCDR).

After our work on the ICP0-induced degradation of CENPs and the resulting destabilization of centromere structure, we decided to check whether and how cells were able to respond to centromere instability during interphase. Interphase HeLa cells were infected with HSV-1 wild-type (wt) virus and analyzed by immunostaining at 2 and 4 h after infection. As expected from our previous work, ICP0 transiently colocalized with centromeres at 2 h after infection but not at 4 h after infection (i.e., after degradation of the CENPs; compare ICP0 patterns in Fig. 1, b and c; insets; Everett et al., 1999b). At 4 h after infection (Fig. 1 c), ICP0 accumulated in large, visible foci inside the nucleoli as described previously (Morency et al., 2005). We found that at early stages of infection, the nuclear pattern of coilin underwent profound changes. Fig. 1 a shows noninfected cells in which coilin was localized in CBs. Infection by HSV-1 wt for 2 h did not substantially affect the coilin distribution (Fig. 1 b), whereas at 4 h after infection, coilin adopted a multidotted pattern in >90% of the infected cells (Fig. 1 c).

These coilin foci were more abundant and smaller than those that corresponded to CBs (0.2–0.5 μm for coilin foci vs. 1–1.2 μm for CBs in noninfected cells; compare coilin patterns in Fig. 1, a and c). Upon close examination, it was noticed that the nuclear distribution of these coilin dots looked very much like the pattern of immunostained centromeres. Therefore, we stained the latter together with coilin in ICP0-expressing cells; strikingly, most of the coilin foci indeed colocalized with centromeres (Fig. 1 c, yellow foci in the merged and magnified images). As the centromeric localizations of ICP0 and coilin were temporally separated (2 vs. 4 h after infection), the accumulation of coilin at centromeres is unlikely to be a consequence of an interaction with ICP0. Collectively, these data suggested that coilin accumulates at centromeres as a consequence of ICP0 activity on centromeres. To verify this notion, cells were infected with either the vFXE virus, which expresses the nonfunctional ICP0 RING finger mutant FXE, or the ICP0-null mutant dl1403, whose infection is detectable by ICP4 viral protein staining. We found that infection with these viruses did not induce coilin redistribution (Fig. 1, d and e).

ICP0 has E3 ubiquitin ligase activity that is associated with its RING finger domain. This activity is responsible for the induction of degradation via the proteasome of CENPs, which can lead to the destabilization of interphase centromeres and to defects in kinetochore assembly (Everett et al., 1999a). To verify the implication of proteasome activity in ICP0-induced coilin accumulation at centromeres, cells were infected with the HSV-1 wt virus in the presence of the proteasomal inhibitor MG132 or DMSO alone (unpublished data). In the presence of MG132, HSV-1 wt no longer induced the accumulation of coilin at centromeres (Fig. 1 f). These data suggest that there is a strong correlation between ICP0- and proteasomal-induced protein degradation (and probably CENP degradation) and the accumulation of coilin at centromeres.

Finally, to exclude any effect of the infection, we verified that transfected cells that expressed ICP0 (Fig. 1 g) but not those that expressed FXE (not depicted) showed a centromeric accumulation of coilin similar to HSV-1 wt–infected cells. As expected, 100% of the ICP0-expressing cells showed centromeric coilin. Note that during the course of these experiments, coilin colocalization with centromeres was never seen in mitotic cells. In conclusion, the aforementioned results demonstrate (1) the existence of a cell response that is triggered by the ICP0-induced structural damage of interphase centromeres (i.e., the iCDR) and (2) that coilin is implicated in the iCDR. These observations strongly suggest a role for coilin in a mechanism that is dedicated to the detection and/or repair of unstable interphase centromeric structures.

To determine whether coilin physically interacts with the central core region, we performed chromatin immunoprecipitation (ChIP) assays with HSV-1 wt–infected cells (Fig. 2 a) and looked for the association of coilin with centromeric, chromatin-specific higher order type I α-SAT DNA. In each case, we compared the ChIP of noninfected cells with that of cells at 4 h after infection (i.e., the time point at which coilin was found to have accumulated at centromeres; Fig. 1 c). The method and results were validated using an anti–CENP-A antibody as a control. Indeed, as expected, we observed a major decrease in the amount of CENP-A associated with type I α-SAT DNA as a consequence of ICP0-induced CENP-A degradation. We then tested an anticoilin antibody in parallel with the anti–CENP-A and anti-myc antibodies, with the latter being used as a nonspecific control (Fig. 2 b). In each experiment, a single batch of chromatin was split into three aliquots, one for each antibody, and the three ChIPs were performed simultaneously.

The results obtained (Fig. 2 b, left) show that more coilin is associated with α-SAT DNA at 4 h after infection compared with noninfected cells, whereas less CENP-A is present at 4 h after infection. In these experiments, the amount of α-SAT DNA associated with the anti-myc control antibody did not notably change. Considering that the 4-h postinfection CENP-A was undetectable at centromeres by immunofluorescence (IF) and almost completely degraded, as shown by Western blotting (WB; Lomonte et al., 2001), a twofold decrease in the amount of α-SAT DNA retrieved (P < 0.05) with CENP-A can be regarded as representative of a major effect. In this context, the 1.7-fold increase in α-SAT DNA retrieved with coilin (P < 0.05) is suggestive of a significant increase in coilin levels at centromeres, which fits with our aforementioned results (Fig. 1). The DNA of the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was measured to control for binding of the anticoilin and anti–CENP-A antibodies to unrelated DNA, and, as expected, no enrichment of coilin was noted (Fig. 2 b, right). From these data, we conclude that ICP0-induced centromere destabilization results in a physical interaction between coilin and centromeric DNA.

A major issue in the biology of nuclear domains is whether the entire domain is involved in a process or only some components thereof. Thus, there was a need to determine whether components of CBs other than coilin also accumulate at damaged centromeres and, if so, whether whole CB domains are associated with centromeres in ICP0-expressing cells. In addition to coilin, CBs concentrate several small noncoding RNAs as well as an array of proteins, all of which are more or less implicated in transcription (Gall, 2001). Apart from fibrillarin, no other tested CB-associated protein exhibits centromeric accumulation in ICP0-expressing cells (Fig. 3 a, i and ii; Fig. 4, a–e; and Table I). Fibrillarin is one of the most abundant proteins in the fibrillar regions of the nucleolus. It is conserved from yeast to humans and is essential for early development in the mouse, being a catalyst of preribosomal RNA methylation (Omer et al., 2002; Newton et al., 2003). Because CBs also contain RNAs, immuno-RNA FISH assays were performed to analyze the pattern of the major CB-associated U2 small nuclear RNA (snRNA) in ICP0-expressing cells. No centromeric accumulation of U2 was detected (Fig. 3 b, i and ii). In addition, antibodies raised against both the 5′-terminal caps of the snRNAs and the snRNA-binding Sm proteins did not show any centromeric signals in ICP0-expressing cells (Fig. 4, d and e; and Table I). Collectively, these data indicate that only some components of the CBs, such as coilin and fibrillarin, accumulate at damaged centromeres.

In the cell nucleus, gems were originally defined as CB-associated domains on the basis of IF staining with antibodies against the SMN protein (Liu and Dreyfuss, 1996). This protein is the product of the SMN gene, whose loss of function mutations are responsible for the severe inherited disorder spinal muscular atrophy, one of the major genetic causes of infant mortality (Lefebvre et al., 1995). We decided to check the patterns of gem- associated proteins in ICP0-expressing cells. Three gem proteins, SMN, gemin 2, and gemin 3, were tested and showed foci that colocalized with CBs in control cells (Fig. 3 a, iii; arrows for SMN; and Fig. 4, f and g; gemin 2 and 3). In ICP0-expressing cells, SMN but not gemin 2 or 3 was detected in numerous small foci that, similar to coilin and fibrillarin, colocalized with centromeres (Fig. 3 a, iv; arrowheads; Fig. 4, f and g; and Table I). As was the case for coilin, 100% of the ICP0-expressing cells showed centromeric fibrillarin and SMN.

The aforementioned experiments suggest that neither coilin, fibrillarin, nor SMN is degraded in ICP0-expressing cells. To confirm this point, cells were infected with HSV-1 wt in the presence or absence of the proteasome inhibitor MG132, vFXE, or dl1403 viruses at a multiplicity of infection of 10 (100% of the cells infected; Fig. 3 c). Coilin, fibrillarin, and SMN did not sustain ICP0-induced degradation.

These data rule out a putative relocalization of the entire CB and/or gem to damaged centromeres and strongly suggest the existence of a specific subset of nuclear proteins (containing at least coilin, fibrillarin, and SMN) that accumulates in response to centromere damage. Therefore, coilin, fibrillarin, and SMN most likely have activities associated with damaged centromeres in addition to their nuclear domain-associated functions.

We observed the iCDR in additional human cell lines of various origins (e.g., breast carcinoma [T47D], colon carcinoma [SW480, HCT116, and HT29], and skin cancer [HaCaT]) as well as in primary keratinocyte cells. To determine whether this response is conserved in other species, the centromeric accumulations of coilin, fibrillarin, and SMN were analyzed in mouse NIH3T3 cells, which express ICP0. In mouse nuclei, chromosomes are distributed in clusters, and pericentromeric heterochromatin forms chromocenters that can be visualized by DAPI staining. The DNA sequences of the central core and pericentromere regions are based on two different types of DNA repeat, called minor and major satellites, respectively. These two domains are spatially separated and are clearly distinguishable by in situ hybridization (Fig. 5 a; Pietras et al., 1983; Guenatri et al., 2004). We performed immuno-DNA FISH assays on ICP0- expressing cells to detect ICP0 (Fig. 5, d and e), minor satellite DNA, and coilin or fibrillarin (the anti-SMN mAb does not work on mouse cells). In the absence of ICP0, coilin and fibrillarin showed the same patterns in the mouse as in human cells (i.e., present in CBs and/or nucleoli; Fig. 5, b and c). In ICP0-expressing cells, coilin (Fig. 5 d, i–iii) and fibrillarin (Fig. 5 e, i–iii) showed a multidotted pattern that was similar to the pattern in human cells. These dots clearly colocalized with the minor satellite signals and, thus, with the central core region of the centromere. These data show that the iCDR is conserved in mammalian cells, at least between humans and mice, and is not an artifact of a single cell line.

A recent study has described the effect of UV-C irradiation on CB fragmentation (Cioce and Lamond, 2005). Consequently, coilin showed a change of pattern and increased interaction with PA28γ (proteasome activator subunit γ), a protein that is implicated in some aspects of proteasome activity in vitro (Wilk et al., 2000). This interaction resulted in the colocalization of coilin and PA28γ in UV-C–treated cells. Because UV light irradiation induces DNA breaks, the putative participation of coilin in a mechanism that is designed to resolve such damage to DNA has to be considered. Therefore, we investigated whether the response to ICP0-induced damaged centromeres implicated protein complexes involved in DNA break repair. PA28γ did not colocalize with coilin at the centromeres of ICP0-expressing cells (Fig. 4 i and Table I). Likewise, several proteins involved in single- or double-strand DNA break repair pathways (including nucleotide excision, base excision, and double-strand break repair) did not relocalize at the damaged centromeres in ICP0-expressing cells (Fig. 6, a–g; and Table II). Given that irradiation of HeLa cells with γ rays (from 2 to 10 Gy) does not provoke CB fragmentation (unpublished data) and that ICP0 is not known to provoke DNA breaks, these data do not support the accumulation of centromeric coilin, fibrillarin, and SMN as part of a putative DNA damage response–associated mechanism with activity in resolving DNA breaks.

The aforementioned results demonstrate that ICP0 provokes the accumulation of at least three proteins at damaged centromeres. However, one could argue that the accumulations of coilin, fibrillarin, and SMN at centromeres are caused by an as of yet unknown activity of ICP0 and are not a direct result of CENP degradation. Therefore, we induced centromere destabilization independently of ICP0 using siRNAs that target the mRNAs of CENP-A, -B, and -C, the three known CENPs that are degraded in an ICP0-dependent manner. We verified by IF (Fig. 7 a) and WB (Fig. 7 b) the effects of the siRNAs on the stability of the targeted proteins. HeLa cells were transfected with single-type siRNAs or a mixture of two or three siRNAs, and the cells were then immunostained to detect (1) decreases in the amounts of the targeted proteins from centromeres and (2) centromeric accumulations of coilin, fibrillarin, and SMN. Cells with multidotted coilin, fibrillarin, or SMN, which were representative of the accumulations of these proteins at centromeres, were counted in several experiments that were performed independently. A scrambled sequence siRNA never had any effect on the coilin, fibrillarin, or SMN patterns. Similarly, none of the three CENP siRNAs was able to induce multidotted fibrillarin or SMN in a manner similar to ICP0 even when used in combination (unpublished data). Interestingly, coilin showed a multidotted pattern in a small proportion (∼5%) of cells that were treated with the CENP-A or -C siRNA and in a large proportion (∼30%) of cells that were treated with the CENP-B siRNA (Fig. 7 c).

We confirmed by IF that the multidotted coilin colocalized with centromeres (Fig. 7 d). To show a more representative view of the colocalization, the merged images were processed, as in Fig. 1 (c and g) with the colocalization module of the LSM 510 software, so that all of the colocalized pixels appeared black on a white background (Fig. 7 d, right column). CENP-A colocalization with huACA staining was used as a positive control (Fig. 7 d, i). A large proportion of coilin colocalized with centromeres in ICP0-expressing cells (Fig. 7 d, iii) as well as in CENP-B siRNA-treated cells (Fig. 7 d, iv). Much fewer black spots were visible in the few CENP-A (or CENP-C; not depicted) siRNA-treated cells that showed multidotted coilin (Fig. 7 d, v), and almost no black spots were seen in the scrambled sequence siRNA-treated cells (Fig. 7 d, ii). In addition, the proportion of centromeres that colocalized with coilin was determined (Fig. 7 d, right column; numbers in parentheses). Several images were scanned in each experiment (Fig. 7 d, i–v) to determine a mean colocalization coefficient between centromeres and coilin. The coefficient for CENP-A colocalization with centromeres was arbitrarily fixed at 1. The colocalization coefficient in untreated cells or cells transfected with the scrambled sequence siRNA was very low, with a value of 0.046. Cells that were treated with the CENP-A (or CENP-C) siRNA showed a fivefold increase in the colocalization coefficient (0.28). ICP0-expressing cells and CENP-B siRNA-treated cells gave the highest coefficient (0.45) for coilin/centromere colocalization, with a 10-fold increase compared with the normal situation. The specificity of the accumulation of coilin at centromeres that lacked CENP-B and, to a lesser extent, CENP-A and -C renders an off-target effect of the siRNAs highly unlikely. However, we tested another siRNA against CENP-B and obtained similar results (unpublished data). These results confirm that the iCDR is triggered as a direct consequence of the loss of some constitutive CENPs and, thus, by the instability of the centromere structure.

In this study, we took advantage of the effect of the ICP0 protein on the destabilization of centromere structures to investigate whether cells survive and adapt to such dramatic modifications at domains of major importance for their viability. We reveal a novel cellular response, named iCDR, suggesting the existence of mechanisms dedicated to dealing with structural damage to centromeres during interphase (i.e., before the onset of mitosis). We also describe a role for coilin in the iCDR, which implicates two other proteins, fibrillarin and SMN. In addition, the fact that several human cell lines and mouse cells manifest a similar response suggests a general mechanism that is likely to be conserved, at least in mammals.

Importantly, our data from cells knocked down for CENP proteins confirm the existence of a cellular response that is triggered directly by structural modifications to centromeres. Although efficient for coilin, the single and combined siRNAs were, unlike ICP0, ineffective in stimulating fibrillarin and SMN. Therefore, the centromeric accumulations of fibrillarin and SMN are probably induced by more severe damage than that arising from the absence of CENP-A, -B, or -C or by the absence of other CENPs. Interphase centromeres are complex structures organized into multisubunit protein domains that are associated partly with the CENP-H–I complex and partly with the centromere-specific CENP-A–containing nucleosomes in the distal (CENP-A nucleosome distal) and proximal (CENP-A nucleosome- associated complex) layers (Foltz et al., 2006; Okada et al., 2006). In light of our results, these data are informative in two respects. First, to engineer the collapse of the entire centromeric structure using CENP-directed siRNAs, it is necessary to affect more than one protein. Second, it is anticipated that ICP0 induces the proteasomal degradation of more CENPs than the individual CENP-A, -B, and -C proteins. Therefore, a simple explanation for the differential centromeric accumulations of coilin, fibrillarin, and SMN seen in this study may be the levels of damage caused to the different layers of CENPs.

Coilin accumulation at damaged centromeres is induced by CENP-B depletion, which suggests that the absence of CENP-B, unlike the absence of CENP-A and -C, is sufficient to trigger the response. CENP-B is a DNA-binding protein that has been implicated by in vitro studies in the positioning of nucleosomes (Yoda et al., 1998; Tanaka et al., 2005). It is unclear whether there is an absolute requirement for CENP-B for the preservation of centromere structure and function, as knockout mice for CENP-B are viable (Hudson et al., 1998; Kapoor et al., 1998; Perez-Castro et al., 1998) and CENP-B is essential for the de novo formation of centromeres, control of the epigenetic state of centromeric chromatin, and assembly of CENP-A (Masumoto, H., personal communication; Ohzeki et al., 2002). In any case, CENP-B is highly conserved in higher eukaryotes, which does not fit with the absence of major phenotype reported in the cenpB^−/− mice studies (Hudson et al., 1998; Kapoor et al., 1998; Perez-Castro et al., 1998). The fact is that in the particular context of mouse cells, centromeres are stable and functional even if the cenpB gene is missing. One explanation is that in cenpB^−/− mice, the centromere structures could have acquired a CENP-B–independent equilibrium that is epigenetically transmissible. This situation is not quite the same as the one described in our present study, in which we hypothesize the rapid disruption of the equilibrium of the centromere structure by the rapid degradation of CENPs. In this regard, we investigated the coilin and fibrillarin distributions in untreated (not expressing ICP0 and not transfected with CENP siRNAs) cenpB^−/− mouse embryonic fibroblast cells derived from CENP-B knockout mice and did not detect any accumulations of these proteins at centromeres (unpublished data).

We know that the accumulation of coilin and fibrillarin at damaged centromeres occurs in ICP0-expressing mouse NIH3T3 cells (this study) and normal mouse embryonic fibroblast cells (unpublished data). Therefore, the absence of the accumulation of these proteins at the centromeres of steady-state cenpB^−/− mouse embryonic fibroblast cells suggests that the centromeric coilin, fibrillarin, and SMN proteins do not act as part of a putative structural complex that replaces the missing proteins but rather as a response of the cell to dramatic modifications of the centromere structure at a given time point. Centromeres possess a very specific organization of their chromatin compared with noncentromeric chromatin, and, thus, they are likely to concentrate unique cellular processes. Our data showing that ICP0-expressing cells lack centromeric accumulations of proteins implicated in DNA break repair mechanisms do not favor the hypothesis of DNA lesions as a direct consequence of the accumulations of centromeric coilin, fibrillarin, and SMN. Therefore, it is likely that the modification of centromere structure itself is responsible for triggering the iCDR. Interestingly, two out of the three known CENP targets of ICP0, CENP-A and -B, are established chromatin-related proteins. This raises the question as to whether the recruitment of centromeric coilin, fibrillarin, and SMN is initiated by the abnormal protein content of the centromere and/or by the abnormal chromatin structure.

On the basis of the data in the literature, it is difficult to come up with a convincing explanation for the interactions of coilin, fibrillarin, and SMN with damaged centromeres. First, this association was unexpected. Second, there is a general lack of information concerning the architecture of the central core centromere, particularly during interphase. However, coilin, fibrillarin, SMN, and centromeres may be linked by RNAs, especially small noncoding RNAs. Indeed, coilin, fibrillarin, and SMN are components of nuclear bodies, CBs, and gems, whose best-characterized functions remain the maturation of small noncoding RNAs that are implicated in the processing of larger transcripts. The association of small RNAs through the RNA interference mechanism with the epigenetic modifications of centromeric heterochromatin is now well documented, particularly in the yeast Schizosaccharomyces pombe (for reviews see Pidoux and Allshire, 2005; Verdel and Moazed, 2005). Importantly, this RNAi-dependent heterochromatinization is directly linked to transcriptional activity in the pericentromeric heterochromatin. Although the existence of a similar mechanism in higher eukaryotes has not yet been demonstrated, it is clear that centromeric heterochromatin–associated transcriptional activity is conserved at least in humans, maize, rice, and chickens (Saffery et al., 2003; Fukagawa et al., 2004; Topp et al., 2004; Zhang et al., 2005). Moreover, recent studies have shown that transcription and small RNAs can participate in the architecture and function of centromeres in murine cells (Maison et al., 2002; Bouzinba-Segard et al., 2006). From these data, it is tempting to speculate that the presence of coilin, fibrillarin, and SMN at damaged centromeres reflects the involvement of RNAs in maintaining the centromeric chromatin structure. Future studies should provide new information that is relevant to this hypothesis.

At the molecular level, we anticipate a close link between the iCDR and the need to reform a functional centromere structure that has been accidentally damaged during interphase. This response could trigger a mechanism that eventually results in the reformation of a fully functional prekinetochore. This inevitably raises questions as to the consequences of centromere instability for general chromosomal instabilities that result in aneuploidy and cancer development (Jallepalli and Lengauer, 2001). Several types of genetic alteration are responsible for chromosomal instabilities, including those that affect kinetochore functions (Bharadwaj and Yu, 2004). Although there is a clear need for correct kinetochore structures to prevent chromosomal instabilities, not much is known concerning the capacities of interphase centromeres to serve as platforms for kinetochore nucleation. If interphase centromeres are unable to build functional kinetochores as a result of structural problems, it is likely that the mitotic spindle checkpoint will be weakened and its activity bypassed, forcing mitosis and provoking aneuploidy (for review see Rieder and Maiato, 2004). This is precisely what has been described in a recent study of late embryonic development in Drosophila cid/cenp-A–null mutants (Blower et al., 2006). Therefore, it is reasonable to propose the existence of a cellular response that acts as a sensor mechanism to signal the emergence of structural problems to interphase centromeres.

In retrospect, it is not surprising that cells are sensitive to defects at interphase centromeres considering the major roles of these genetic loci. To date, it has been unknown whether cells have developed mechanisms during interphase to check the functionalities of these structures. Our present results clearly show that this is probably the case, and they raise questions as to the existence of pathways that are committed to sensing, signaling, and repairing centromeres before the cell enters mitosis. In this framework, future studies will need to address the importance and functions of proteins, such as coilin, fibrillarin, SMN, and other proteins, in these types of pathways.

HeLa and NIH3T3 cell lines were cultivated in BHK-21 and DME, respectively, which were supplemented with 10% FBS, l-glutamine (1% vol/vol), 10 U/ml penicillin, and 100 μg/ml streptomycin. The pci110 plasmid, which expresses ICP0, has been described previously (Everett et al., 1993). The wt strain HSV-1 syn+ (17+) is the parental strain (referred to as HSV-1 wt in this study). The dl1403 virus, which was deleted of ICP0 (Stow and Stow, 1986), the vFXE virus, which expresses a nonfunctional ICP0 isoform mutated in its RING finger domain, and the standard infection procedures have been described previously (Everett, 1989).

Cells were seeded at 1.5 × 10⁵ cells per well for transfection and at 2.5 × 10⁵ cells per well for infection in 24-well plates that contained round coverslips. 24 h later, the cells were transfected with the pci110 plasmid according to the manufacturer's recommendations (Effectene Transfection Reagent; QIAGEN) or infected. Cells were treated for IF as described previously (Lomonte et al., 2001). With the exception of Fig. 5, in which a CCD camera (CoolSNAP HQ; Roper Scientific) was used for the analysis (Metaview software; Molecular Devices), all of the samples were examined under a confocal microscope (LSM 510 Meta; Carl Zeiss MicroImaging, Inc.). The data from the channels were collected separately to avoid channel overlap, with fourfold averaging at a resolution of 512 × 512 pixels using optical slices of 0.8–1.0-μm thickness. A microscope (Axiovert 200M; Carl Zeiss MicroImaging, Inc.) was used at either 63× (NA 1.25) or 100× (NA 1.3) magnification by oil-immersion objective lenses (Carl Zeiss MicroImaging, Inc.). Datasets were processed using LSM 510 software (Carl Zeiss MicroImaging, Inc.) and exported in preparation for printing using Photoshop (Adobe).

HeLa cells were seeded at 1.5 × 10⁵ cells per well in 24-well plates that contained coverslips. The next day, the cells were transfected with the pci110 plasmid. After 24 h, the cells were rinsed in 1× PBS and fixed with 4% formaldehyde in 1× PBS for 10 min. The cells were rinsed again in 1× PBS and incubated overnight at 4°C in 70% ethanol. The cells were then rehydrated for 5 min in 2× SSC with 50% formamide and hybridized overnight at 37°C with 3 ng of the digoxigenin-labeled snRNA U2 probe (5′-ATACTGATAAGAACAGATACTACACTTGATCTTAGCCATA-3′) diluted in 40 μl of 2× SSC that contained 10% dextran sulfate, 2 mM vanadyl-ribonucleoside complex, 0.02% BSA (Rnase free), 40 μg Escherichia coli tRNA, and 50% formamide. 1 d later, the cells were washed twice for 30 min at 37°C in 2× SSC that contained 50% formamide and were then washed three times at 37°C with 1× PBS that contained 1% FBS. The cells were finally treated for IF to reveal proteins as described previously (Lomonte et al., 2001). An FITC-labeled sheep antidigoxigenin antibody diluted 1:500 was used to reveal the probe.

Cells were first processed for immunostaining as indicated in the previous section and were postfixed with PFA and treated for FISH using the three-dimensional FISH procedure of Solovei et al. (2002) to ensure maximum preservation of the nuclear architecture. Probes were generated from the mouse major and minor satellite DNAs, which were obtained by PCR amplification of genomic DNA from the NIH3T3 cells used in this study using the following primers: for the major satellite MMS 24C, 5′-CATATTCCAGGTCCTTCAGTGTGC-3′; for MMS 24D, 5′-CACTTTAGGACGTGAAATATGGCG-3′; for the minor satellite MS 24C, 5′-ACTCATCTAATATGTTCTACAGTG-3′; and for MS box 1, 5′-AAAACACATTCGTTGGAAAC-GGG-3′. The major and minor satellite DNAs were labeled with either biotin-16-dUTP or digoxigenin-11-dUTP by nick translation (Roche Diagnostics). A standard procedure was used for probe detection using either AlexaFluor555-labeled streptavidin (Invitrogen) or FITC-labeled antidigoxigenin antibodies (Roche Diagnostics). Nuclear DNA was counterstained with Hoechst 33258, and the cells were mounted with Vectashield antifade.

Cells were seeded at 5 × 10⁶ cells per 100-mm Petri dish. The following day, the cells were infected with the HSV-1 virus for 4 h and were subjected to the ChIP assay (Upstate Biotechnology). The samples were incubated with IgG Fab fragments (US Biological) before adding the test antibodies to decrease the nonspecific binding of alphoid DNA repeats to IgG. DNA that was immunoprecipitated with the different antibodies was quantified by PCR using specific primers. The PCR reaction was performed in a LightCycler (Roche Diagnostics) using 1 μl DNA, 0.5 μM of each primer, and 10 μl SYBR green (QIAGEN) in a final volume of 20 μl. The data were analyzed using the LightCycler software (Roche Diagnostics). The α-SAT and GAPDH designations indicate oligonucleotides that are specific for the centromeric type I α-SAT DNA and the gene that encodes the GAPDH enzyme, respectively. The α-SAT oligonucleotides were designed based on the consensus α-SAT sequence (Alexandrov et al., 1993), which was derived from the alignment of >500 type I alphoid sequences. Moreover, a BLAST search using this consensus type I α-SAT showed hits for cosmids from several chromosomes. Therefore, these oligonucleotides amplify DNA fragments from several different chromosomes, and the ChIP data are likely to reflect the association of proteins with several chromosome centromeres. The primers used were as follows: for α-SAT, forward 5′-AATCTGCAAGTGGATATTT-3′ and reverse 5′-CTACAAAAAGAGTGTTTCAAAAC-3′; for GAPDH, forward 5′-CACGTAGCTCAGGCCTCAAGA-3′ and reverse 5′-AGGCTGCGGGCTCAATTTAT-3′. The data shown are the ratios between the values obtained for the immunoprecipitated chromatin and the input DNA and are the mean values from six independent experiments.

HeLa cells were seeded at 4 × 10⁴ cells per well in a 24-well plate that contained round coverslips for IF or at 1 × 10⁶ cells per 60-mm Petri dish for WB. 1 d later, the cells were transfected with 300 ng (for IF) or 7 μg (for WB) siRNA that targets the mRNA of CENP-A, -B, or -C and were incubated for 48 h. A second round of siRNA transfection was performed as described above, and cells were incubated for an additional 48 h before the cells were harvested for IF or WB experiments. For WB, 40-μg aliquots of total protein were loaded per well in an SDS-polyacrylamide gel before electrophoresis, transfer, and detection as described previously (Lomonte et al., 2004). The siRNA sequences used were as follows: for CENP-A, 5′-GGUUGGCUAAAGGAGAUCCTT-3′; for CENP-B, 5′-CUACACCGCCAACUCCAAGTT-3′; and for CENP-C, 5′-GAAGCCUCUCUACAGUUUGTT-3′.

The following antibodies were used at the indicated dilutions for the IF, WB, or ChIP experiments: mAbs anti-p62 (TFIIH at 1:400 for IF), [3–19] anti–CENP-A (1 μg/ml for WB and 2 μg per sample for ChIP; Abcam), [5E6C1] anti–CENP-B (1 μg/ml for WB), anticoilin (1:400 for IF, 1 μg/ml for WB, and 2 μg per sample for ChIP; Sigma-Aldrich), [25–4] anti–DNA-PKcs (1:200 for IF; NeoMarkers); [72B9] antifibrillarin (1:500 for IF), [38F3] antifibrillarin (1:500 for WB; Abcam), anti–gemin 2 (1:200 for IF; Abcam), [12H12] anti–gemin 3 (1:200 for IF; Abcam), anti–HCF-1 (1:400 for IF), [11060] anti-ICP0 (1:1,000 for IF and 1:10,000 for WB), [8.F.137B] anti-ICP4 (1:100 for IF; US Biological), [9E10] anti-myc (2 μg per sample for ChIP; Abcam), [D0-7] anti-p53 (1:500 for IF; DakoCytomation), [5E10] anti–promyelocytic leukaemia (PML; 1:50 for IF), [51RAD01] anti-Rad51 (1:200 for IF; NeoMarkers), [9H8] anti-RPA32 (1:200 for IF; Abcam), Y12 anti-Sm (1:500 for IF; NeoMarkers), anti-SMN (1:400 for IF; 1 μg/ml for WB; BD Biosciences), and anti–2,2,7-trimethylguanosine (TMG at 1:300 for IF; Oncogene Research Products). In addition, the following rabbit polyclonal antibodies were used: antiactin (1 μg/ml for WB; Sigma-Aldrich), 1343 anti-BLM (1:500 for IF), anti–CENP-A (1:200 for IF; Upstate Biotechnology), anti–CENP-B (1:2,000 for IF), r554 anti–CENP-C (1:1,000 for IF and WB), rp80 anticoilin (1:400 for IF), anti-FLASH (1:4,000 for IF), anti-γH2AX (phospho-S139 at 1:500 for IF; Abcam), R190 anti-ICP0 (1:200 for IF), anti-PA28γ (N terminus; 1:200 for IF; Zymed Laboratories), and BL647 antiphospho-RPA32 (S4/S8 at 1:1,000 for IF; Bethyl Laboratories). The human autoimmune serum huACA against CENPs was also used (1:3,000 for IF). For IFs, the secondary antibodies used were as follows: goat anti–rabbit, anti–mouse, and anti–human antibodies coupled to AlexaFluor488, -555, or -647 (1:200; Invitrogen).

We thank the following people for donating materials: Mounira Amor-Guéret for anti-BLM (Institut Curie, Orsay, France), William R. Brinkley for CENP-B^−/− cells (Baylor College of Medicine, Houston, TX), Jérome Cavaillé for mAb 72B9 (Université Paul Sabatier, Toulouse, France), Vincenzo De Laurenzi for anti-FLASH (Universita' di Roma, Rome, Italy), William Earnshaw for huACA, rabbit anti–CENP-B, and r554 (Institute of Cell and Molecular Biology, Edinburgh, Scotland, UK), Jean-Marc Egly for anti-p62 (Institut de Génétique et de Biologie Moléculaire et Cellulaire, Strasbourg, France), Roger Everett for mAb 11060 and R190 (Medical Reseach Council Virology Unit, Glasgow, Scotland, UK), Winship Herr for anti–HCF-1 (Center for Integrative Genomics, Lausanne, Switzerland), Jérome Lamartine for human keratinocyte primary cells (Centre de Génétique Moléculaire et Cellulaire/Université Claude Bernard Lyon 1), Angus Lamond for rp80 anticoilin (Wellcome Trust Biocenter, Dundee, Scotland, UK), Hiroshi Masumoto for mAb 5E6C1 and technical advice (Division of Biological Science, Nagoya University, Nagoya, Japan), and Roel van Driel for mAb 5E10 (University of Amsterdam, Amsterdam, Netherlands). We also thank Joëlle Thomas for her help with real-time PCR and constructive criticism of the manuscript and thank William Earnshaw for insightful comments.

This work was funded by the Centre National de la Recherche Scientifique through the Action Thématique Incitative sur Programme Young Researchers Program, the National Agency for Research (grant ANR-05-MIIM-008-01) through the CENTROLAT program, and the National Institute of Cancer through the EPIPRO program.