Antibody Production

To generate antibodies against two additional domains of C-Nap1, (His)₆-tagged fusion proteins were constructed with the NH₂ terminus (N-term, amino acid [aa] 67–345) as well as the middle region (Middle, aa 1,098–1,248), using the QIAexpress bacterial expression system (QIAGEN). The plasmid pQE10:N-term was constructed by subcloning an RsaI (bp 867)–BclI (bp 1,702) (blunted with Klenow) fragment from C-Nap1 into the pQE31 vector (QIAGEN) digested with SmaI. The plasmid pQE30:Middle was constructed by subcloning the SacI (bp 3,967)–HindIII (bp 4,410) fragment of C-Nap1 into pQE30 digested with SacI and HindIII. Protein expression was performed in the Escherichia coli strain M15(pREP4) using Luria broth (10 mM Tris-HCl, pH 7.5, 10 g Bacto tryptone, 5 g Bacto yeast extract, 5 g NaCl per liter). Recombinant proteins were expressed for 4–6 h with 1 mM isopropyl-β-d-thiogalactoside (IPTG) and purified on nickel columns under denaturing conditions, as described by the manufacturer (QIAGEN). They were then further purified by preparative gel electrophoresis on a 12% SDS-polyacrylamide gel, as described previously (Fry et al. 1998b). For immunizations, purified proteins (350 μg of the [His])₆:N-term and 450 μg of the [His])₆:Middle protein) were injected subcutaneously into New Zealand white rabbits (Elevage Scientifique des Dombes) on days 0, 27, 56, 84, 112, 140, and 160 (N-term) and 0, 28, 56, 84, and 112 (Middle). Immune sera were obtained on days 140, 178, and 182 (N-term) and 97, 125, and 132 (Middle). These antibodies are referred to as N-Ab and M-Ab, respectively. For affinity purification of antibodies, 1 mg of each purified (His)₆:recombinant protein was coupled covalently to 400 mg cyanogen bromide–activated Sepharose 4B (Amersham Pharmacia Biotech), as recommended by the manufacturer, and antibodies were purified on this affinity matrix, as described previously (Harlow and Lane 1998). Purified antibodies were dialyzed extensively against PBS before use.

Cell Culture, Transfections, and Immunochemical Techniques

Human Hs68 fibroblasts, HeLa, U2OS osteosarcoma, and KE37 T-lymphoblastoid cells were grown at 37°C in a 7% CO₂ atmosphere in DME supplemented with 10% heat-inactivated FCS and penicillin-streptomycin (100 IU/ml and 100 μg/ml, respectively). For transient transfection studies, U2OS cells were seeded onto hydrochloric acid–treated glass coverslips at a density of 10⁵ cells per 35-mm dish. They were then transfected with 5 μg of plasmid DNA, using calcium phosphate precipitates, as previously described (Seelos 1997), and fixed 24 h later in cold methanol (−20°C) for 6 min. For microinjection, human diploid Hs68 fibroblasts were grown on gelatin- or polylysine-coated coverslips. Where indicated, Hs68 cells were treated with 1.6 μg/ml aphidicolin, 5 μg/ml nocodazole, 5 μM taxol, 3 μg/ml cytochalasin D, or 10 mM 2,3-butanedione monoxime (BDM) (all drugs were from Sigma-Aldrich).

To prepare cell extracts for immunoblotting, cells were harvested, washed once in PBS/1 mM PMSF, and resuspended in RIPA extraction buffer (50 mM Tris-HCl, pH 8, 1% NP-40, 0.5% deoxycholic acid, 0.1% SDS, 150 mM NaCl, 1 mM PMSF, 1 μg/ml aprotinin, 1 μg/ml leupeptin, 1 μg/ml pepstatin A, 20 mM β-glycerophosphate) to yield 2 × 10⁴ cells/μl. Samples were left for 30 min on ice, passed 10 times through a 27G needle, and centrifuged at 14,000 rpm for 10 min at 4°C. One volume of protein sample buffer was added to the supernatant and the sample was heated to 95°C for 3 min before analysis by SDS-PAGE on a 7.5% gel. Proteins were electrophoretically transferred to nitrocellulose membranes using a Bio-Rad apparatus (Bio-Rad Laboratories), and membranes were incubated for 30 min in blocking buffer (5% low-fat dried milk in PBS/0.1% Tween-20). All antibody incubations were carried out in blocking buffer for one hour at room temperature (N-Ab and C-Ab, 0.5 μg/ml; M-Ab, 1 μg/ml), and bound IgGs were visualized using alkaline phosphatase–conjugated anti–rabbit IgG secondary antibodies (Promega).

Immunofluorescence and Immunoelectron Microscopy

Immunofluorescence microscopy was performed using a ZEISS Axioplan II microscope and 40× or 63× oil immersion objectives. Photographs were taken using a Quantix 1400 (Photometrics, Inc.) or Micromax (Princeton Instruments) CCD cameras and IP-Lab or Metaview (Universal Imaging Corp.) softwares. Primary antibodies used for immunofluorescence were affinity-purified N-Ab (0.5 μg/ml) and M-Ab (1 μg/ml), anti–γ-tubulin IgG (2 μg/ml), GTU-88 anti–γ-tubulin mAb (1:1,000; Sigma-Aldrich), 9E10 anti-myc mAb (undiluted tissue culture supernatant), and GT335 anti-polyglutamylated tubulin mAb (1:2,000). Secondary antibodies were biotinylated donkey anti–rabbit or goat anti–mouse IgG (1:200; Amersham Pharmacia Biotech), followed by Texas red–conjugated streptavidin (1:100; Amersham Pharmacia Biotech), FITC–conjugated goat anti–mouse Fab fragment (1:100; Sigma-Aldrich), FITC-conjugated goat anti–rabbit (1:100; Sigma-Aldrich), and AlexaFluor488– conjugated goat anti–mouse IgG (1:1,000; Molecular Probes).

Preembedding immunoelectron microscopy of whole U2OS cells was performed after fixation with 3% paraformaldehyde/2% sucrose and permeabilized with 0.5% Triton X-100, as previously described (Fry et al. 1998b).

Plasmid Construction

A full-length C-Nap1 construct was generated by subcloning the fragments P5.71, P4.20, and P2.5 (Fry et al. 1998b) into pBlueScript-SK and fusing them at the restriction sites SalI (bp 2,604) and NheI (bp 3,491). For fusion to the myc tag, EcoRV and PmlI restriction sites were introduced upstream of the first methionine ATG codon (bp 672) to create ^kC-Nap1. The NH₂-terminal fragment of C-Nap1 was amplified by PCR with the high fidelity Taq polymerase (Roche) using two oligonucleotides (5′-TTGATATCACGTGCCGCCATGGAGACAAGAAGC-3′ and 5′-ATGGGCTTCCTTCTCCAGCTCC-3′). The amplified fragment was digested with EcoRV–SmaI (bp 1,116) and fused to the SmaI (bp 1,116)–NotI C-Nap1 fragment. The modified full-length ^kC-Nap1 was then digested by EcoRV–NotI and ligated to pBS KS+:myc, which was digested with SmaI and NotI. During this procedure, a 10 aa (MNSCPSPRAA) linker was created between the myc tag and C-Nap1. The myc–C-Nap1 insert was then subcloned in the pBK-CMV vector. T1 (aa 1–362) was obtained by cutting ^KC-Nap1 with EcoRV and BstXI (bp 1,758), T2 (aa 1,982–2,442) was previously described (Fry et al. 1998b), and T3 (aa 243–1,985) was generated using NaeI (bp 1,399) and XhoI (bp 6,624). All fragment were cloned downstream of a myc tag and inserted into pCMV plasmids.

Cell Cycle Analysis of Centrosome Splitting

To determine whether centrosomes were susceptible to splitting at different cell cycle stages, we microinjected anti–C-Nap1 antibodies into cells that had been synchronized in either G1 or G2 phase. For injection of G1 phase cells, Hs68 fibroblasts were arrested in G0 by serum deprivation, and then they were released into the cell cycle by the addition of 20% FCS. For synchronization in G2, cells were released from an aphidicolin-induced block at the G1/S boundary. Synchronized cells, as well as asynchronous populations for control, were injected with either control IgG or anti–C-Nap1 antibodies, and the extent of centrosome splitting was determined by immunofluorescent staining with anti–γ-tubulin antibodies or GT335, as described above. Cultures were subjected to flow cytometric analysis of cell cycle profiles, in parallel. As shown in Fig. 2 A, the G1 population of cells showed a 2N DNA content both at the time of injection and at the time of fixation. The G2 population showed predominantly a 4N DNA content at the time of injection, though some cells had progressed through mitosis by the time of fixation 4 h later. Compared with results for asynchronous cell populations, anti–C-Nap1 antibodies caused centrosome splitting to similar extents, regardless of whether cells were in G1 or in G2 (Fig. 2 B). Furthermore, in both cases, the distances between split centrosomes showed a very broad distribution, with an average distance of about 10 μm (with a range of 2 to >20 μm). This indicates that interference with C-Nap1 function induces the separation of parental centrioles, whether or not they are associated with growing procentrioles.

Figure 2

Cell cycle analysis of centrosome splitting. (A) FACS^® analysis of synchronized Hs68 cells. For cell cycle synchronization, Hs68 cells were serum starved for 36 h and then released into medium containing 20% FCS and 1.6 μg/ml aphidicolin. G1 cells were microinjected 16 h after serum addition. To microinject a G2 population, cells were released for 5 h from a 36 h block in aphidicolin. Aliquots of cells were analyzed by FACS^® before microinjection as well as before fixation 4 h after injection. (B) Centrosome splitting occurs regardless of cell cycle position. Hs68 cells were injected with anti–C-Nap1 antibodies at the indicated cell cycle stages and analyzed 4 h later by immunofluorescence microscopy, as described in the legend to Fig. 1 B. The histogram shows the percent of cells with split centrosomes. For C-Ab, a total of 252, 371, and 309 injected cells were counted for asynchronous, G1, and G2 populations, respectively. Similarly, 150, 318, and 366 cells were counted after injection with control IgG. Rare cells showing obvious chromosome condensation were excluded from the analysis. Results were averaged from two independent experiments. (C) Centrosome splitting does not prevent cell division. Single scattered Hs68 cells were injected with control (n = 50) or anti–C-Nap1 antibodies (n = 58) and analyzed 30 h later by immunofluorescence microscopy. Cell doubling was assessed by counting cell numbers in clusters originating from single injected cells. The histograms indicate the percent of clusters containing 1, 2, 3, or 4 cells. B and C: Black bars refer to cells injected with C-Ab, gray bars to control-injected cells. (D) Double immunofluorescent staining of a representative mitotic cell, injected with anti–C-Nap1 antibody in the preceding interphase. a, DNA; b, γ-tubulin; c, the injected anti–C-Nap1 antibody. Bar, 5 μm.

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It was of interest to determine whether anti–C-Nap1 antibody–induced centrosome splitting adversely affected cell division. To address this question, single cells were injected with C-Ab or control IgG and their progeny were analyzed 30 h later (Lane and Nigg 1996). This assay revealed no significant difference between C-Ab–injected cells and control-injected cells in their ability to divide: in both cases, ~60% of the cells had gone through one division at the time of analysis, and ~20% had even completed a second division (Fig. 2 C). Furthermore, we carefully examined mitotic cells arising after the injection of C-Nap1 antibodies during the preceding interphase and found that such cells had assembled normal-looking bipolar spindles (Fig. 2 D). We conclude from these results that cells harboring split centrosomes are not prevented from undergoing cell division. Although this conclusion may appear surprising at first, it is in complete agreement with previous findings (Fry et al. 1998a).

Overexpression of Wild-type C-Nap1 and Deletion Mutants

To provide further evidence for a role of C-Nap1 in centrosome cohesion, independent of antibody microinjection, we asked whether particular regions of C-Nap1 might cause centrosome splitting by acting as dominant-negative mutants. To this end, three C-Nap1 deletion mutants were designed on the basis of the C-Nap1 sequence, which predicts the existence of small NH₂- and COOH-terminal end domains, separated by a large central coiled-coil structure. The mutants T1 and T2 were thus generated to represent the NH₂- and COOH-terminal end domains, respectively, whereas T3 was made to span the central, putative coiled-coil region (Fig. 3 A). Full-length C-Nap1 and the three C-Nap1 deletion mutants were then expressed in U2OS cells, under the control of a cytomegalovirus (CMV) promoter. The localization of the ectopically expressed proteins was determined by immunofluorescence microscopy, taking advantage of an NH₂-terminal–myc tag. In parallel, the distribution and morphology of centrosomes was examined by staining with antibodies against γ-tubulin, and the extent of centrosome splitting was scored as described above.

Figure 3

Mutational domain analysis of C-Nap1. (A) Schematic view of C-Nap1 domain structure and different C-Nap1 deletion mutant constructs. The dark gray boxes designate the predicted coiled-coil domains in C-Nap1. A semiquantitative assessment of the ability of C-Nap1 mutants to localize to centrosomes is given as follows: −, no centrosomal signal; ++, the ability to localize to centrosomes in ~2/3 of transfected cells; +++, centrosome association in all transfected cells. (B) Illustration of the phenotypes observed after overexpression of three different C-Nap1 deletion mutants: T1 (a and a′), T2 (b and b′), and T3 (c and c′). U2OS cells were analyzed by double immunofluorescence microscopy, by using anti-myc (9E10) mAb to visualize C-Nap1 deletion mutants (a–c) and anti–γ-tubulin (a′–c′). Arrowheads mark the positions of centrosomes. Bar, 10 μm. (C) Overexpression of C-Nap1 deletion mutants induces centrosome splitting. The histogram indicates the percent of transfected cells with split centrosomes. Cells transfected with the full-length C-Nap1 (wt) as well as nontransfected cells (−) are shown for control, and the extent of splitting observed with the protein kinases Nek2 and Nek3 is shown for comparison. A myc tag was present in all constructs. Errors bars indicate standard deviations for data collected from three independent experiments.

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In cells expressing low levels of exogenous full-length C-Nap1, the protein localized correctly, and almost exclusively, to the centrosome, but in cells expressing higher protein levels, one or several compact C-Nap1–positive aggregates became visible (data not shown). This probably reflects a propensity of C-Nap1 to oligomerize. However, under no circumstances did overexpression of full-length C-Nap1 cause any significant centrosome splitting (Fig. 3 C). In contrast, all three C-Nap1 deletion mutants caused substantial centrosome splitting, albeit to a different extent. Although the mutants T2 and T3 were almost as effective in causing centrosome splitting as the active-Nek2 kinase, T1 was only half as effective. Nek3, a Nek2-related kinase analyzed for a control, did not induce any splitting above background. It is noteworthy that the ability of C-Nap1 mutants to cause centrosome splitting was not directly related to their ability to localize to centrosomes (Fig. 3a and Fig. c). Whereas T1 and T2 were efficiently targeted to centrosomes, T3 was almost completely cytoplasmic. Yet, T3 was able to cause a substantial amount of splitting similar to T2. This suggests that C-Nap1 truncation mutants most likely cause centrosome splitting by sequestering one or more centrosomal components in the cytoplasm, thereby preventing them from performing a cohesive function. One protein likely to be titrated by C-Nap1–deletion mutants is C-Nap1 itself. In support of this view, we have observed that endogenous C-Nap1 was displaced from centrosomes when these were split by overexpression of C-Nap1 mutants (data not shown). Experiments aimed at identifying other C-Nap1–interacting proteins are currently in progress.

Centrosome Splitting and Cytoskeletal Dynamics

The mechanisms underlying centrosome cohesion remain largely unknown. On the one hand, it is plausible that centrioles may be held together by a proteinaceous but flexible and dynamic structure. This hypothesis is supported by both electron microscopic evidence (Bornens et al. 1987; Paintrand et al. 1992) and recent live-imaging studies (Piel et al. 2000). On the other hand, it has been argued that centrioles may localize in close proximity primarily as a result of MT dynamics (Jean et al. 1999). To explore a possible critical contribution of the cytoskeleton to the phenomena described here, we asked whether anti–C-Nap1 antibodies were able to induce centrosome splitting in cells in which cytoskeletal dynamics had been disturbed by drugs. MT dynamics were profoundly altered by treatment with either nocodazole or taxol, whereas the functions of the microfilament (MF) system were impaired by treatments with either cytochalasin D or BDM, an inhibitor of myosin ATPase.

We found that depolymerization of MTs by nocodazole roughly doubled the proportion of control-injected cells with split centrosomes, whereas treatment of cells with the MT-stabilizing drug taxol did not significantly alter this proportion (Fig. 4 A, gray bars). These results are in agreement with the observation that MT dynamics can influence the positioning of centrioles within cells (Jean et al. 1999). Most importantly, however, microinjection of anti–C-Nap1 antibodies still induced centrosome splitting even in nocodazole- or taxol-treated cells (Fig. 4 A, black bars). Similarly, although centrosome splitting in control-injected cells was slightly affected by drugs acting on the MF system (Fig. 4 B, gray bars), neither cytochalasin D nor BDM prevented the splitting induced by anti–C-Nap1 antibodies (Fig. 4 B, black bars). As we observed in all our experiments, the distributions of distances between split centrioles were very broad (with a range of 2 to ≥20 μm), with no clear peaks. Quantitative analyses of control cells and drug-treated cells did not allow us to uncover statistically significant differences, indicating that the drug treatments described above exerted at most a minor influence on the average distances between split centrioles (data not shown). Thus, we conclude that anti–C-Nap1 antibody–induced centrosome splitting does not require an intact MT or MF system.

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Figure 4

Anti–C-Nap1 antibody-induced centrosome splitting does not require an intact cytoskeleton. (A) Centrosome splitting caused by C-Ab injection is independent of MTs. The histogram indicates the percent of antibody-injected cells with split centrosomes when treated with nocodazole (5 μg/ml) and taxol (5 μM). Drug treatment was initiated 1–1.5 h before antibody injection and the cells were maintained throughout the injection and the following 4 h of incubation before fixation. In two independent experiments, 213 and 259 cells injected with C-Ab were counted (black bars), along with 183 and 217 cells injected with control IgG (gray bars). (B) Centrosome splitting is independent of the MF network. The histogram indicates the percent of injected cells with split centrosomes after treatment with either cytochalasin D (3 μg/ml) or BDM (10 mM). To avoid cell rounding, BDM was applied only 30 min before microinjection and cytochalasin D was applied right after injection. Cells were incubated for 4 h in the presence of drugs before they were fixed and analyzed by immunofluorescence microscopy. In two independent experiments, 150 and 211 cells injected with C-Ab were counted (black bars), along with 196 and 212 cells injected with control IgG (gray bars). (A and B) The percent of centrosome splitting observed in injected cells in the absence of drugs are indicated by dotted lines (lower line for control IgG and upper line for C-Nap1 antibodies).

We also asked whether centrosome splitting would interfere with the ability of separated centrioles to nucleate MTs. To this end, MT depolymerization and regrowth assays were performed on anti–C-Nap1–injected cells. We observed many cells harboring spit centrosomes in which both separated centrioles were at the centers of MT asters (data not shown). This indicates that centrosome splitting does not prevent MT nucleation, in line with the fact that both separated centrioles are readily stained with anti–γ-tubulin antibodies, implying that they are associated with PCM. Also, these results are in agreement with data showing that both parental centrioles are competent to nucleate MTs through their associated PCM, although they differ in their ability to retain them (Jean et al. 1999; Piel et al. 2000).

The data described so far show that C-Nap1 plays a critical role in mediating centrosome cohesion throughout interphase of the cell cycle. This conclusion is based on both antibody-mediated interference with C-Nap1 function and the analysis of dominant-negative–acting C-Nap1 mutants. Furthermore, we have shown that interference with C-Nap1 function causes centrosome splitting by a mechanism that is largely independent of an intact MT or MF network, and that centrosome splitting does not encumber cell division.

Reversible Dissociation of C-Nap1 from Mitotic Spindle Poles

As an important consequence of the proposed role of C-Nap1 in centrosome cohesion, one would predict that a C-Nap1–containing cohesive structure should be transiently dismantled (or functionally inactivated) during mitosis when parental centrioles separate from each other to form the spindle poles. In line with this prediction, we have observed previously that C-Nap1 staining appeared to fade away upon formation of mitotic spindle poles (Fry et al. 1998b). However, since our former data were obtained using a single antibody reagent directed against the COOH terminus of C-Nap1, C-Ab, it was impossible to exclude the caveat of epitope masking. To rigorously test whether C-Nap1 dissociates from centrosomes during mitosis, we raised two additional antibodies, one directed against the NH₂-terminal end domain, the other against the middle domain (the putative “hinge” region; Fig. 5 A). These reagents are referred to as N-Ab and M-Ab, respectively. As shown by Western blotting, both N-Ab and M-Ab readily recognized a protein of the expected mobility in U2OS total-cell extracts (Fig. 5 A). M-Ab was about as reactive as C-Ab, whereas N-Ab was slightly less reactive. In addition, both N-Ab and M-Ab recognized a few smaller proteins. The same antibodies were also used for Western blots on centrosomes purified from KE37 lymphoid cells (Fig. 5 A). As reported previously, KE37 cells exhibit two forms of C-Nap1 (Fry et al. 1998b). Of these, the slower migrating form was recognized by all three antibodies, but the faster migrating one was only recognized by M-Ab and C-Ab, indicating that it lacks the NH₂ terminus (Fig. 5 A). More importantly, we may also conclude that the additional bands seen in U2OS whole-cell lysates represent noncentrosomal, cross-reactive proteins, as they were not present in purified centrosomes. When tested by immunofluorescence microscopy on methanol-fixed U2OS cells, both new antibodies readily labeled the centrosome, as shown by costaining for γ-tubulin (Fig. 5 B, and data not shown). Furthermore, both N-Ab and M-Ab showed staining of two distinct dots, as reported previously for C-Ab. This suggests that C-Nap1 is unlikely to span the entire distance between the two centrioles, a conclusion strengthened further by immunoelectron microscopy (see below). Most importantly, both N-Ab and M-Ab produced at most very faint staining of mitotic spindle poles (Fig. 5 B, and data not shown), confirming and extending the result obtained previously with C-Ab. These data are important in that they strongly argue against epitope masking, and instead indicate that the bulk of C-Nap1 truly dissociates from centrosomes during mitosis.

Figure 5

Cell cycle–dependent association of C-Nap1 with centrosomes. (A) Total protein was prepared from exponentially growing U2OS cells (lanes 1–3), and centrosomes purified from KE37 cells (lanes 4–6), separated by SDS-PAGE, and probed by immunoblotting with N-Ab (lanes 1 and 4), M-Ab (lanes 2 and 5), or C-Ab (lanes 3 and 6). The positions of molecular weight markers are indicated (in kD). The C-Nap1 fragments used for immunization to produce N-Ab, M-Ab, and C-Ab are illustrated schematically on top. (B) Double-indirect immunofluorescent staining of U2OS cells. Centrosomes were labeled with antibodies against γ-tubulin (c and d), and C-Nap1 with N-Ab (a and b). (a and c) Interphasic cell. (b and d) Mitotic cell. Bar, 5 μm. (C) C-Nap1 reassociates with the centrosome upon exit from mitosis. U2OS cells were stained as indicated. (a, c, and e) Telophase cell. (b, d, and f) Early G1 cell. DNA was stained with Hoechst dye 33258, showing condensed (a) and decondensed chromatin (b). Midbody (c) and postmitotic bridge (d) were stained with antibodies against α-tubulin; C-Nap1 was stained with N-Ab (e and f). Bar, 10 μm. (D) Double-indirect immunofluorescent staining of a dividing HeLa cell. The shape of the dividing cell is shown by differential interference contrast microscopy (a), and chromatin is visualized by staining with Hoechst dye 33258 (b). Centrioles were labeled with γ-tubulin (c and d) and C-Nap1 with N-Ab (e and f). Note that the pictures shown in c and e and d and f were taken in two different focal plains to allow visualization of both centrioles. In this particular cell, the two parental centrioles are separated over a very large distance. Bar, 10 μm.

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We also asked when exactly C-Nap1 reaccumulates at centrosomes and found that this occurs at the very end of cell division (Fig. 5 C). Up to telophase, when chromatin was still condensed (Fig. 5 C, a), only traces of C-Nap1 could be seen on spindle poles (Fig. 5 C, e). However, C-Nap1 staining then increased progressively and approached interphase levels already in very early G1 cells (Fig. 5 C, f and d; in d, note the postmitotic bridge still connecting these cells). Of particular interest, recent live imaging studies have revealed that parental centrioles frequently separate over very large distances in dividing cells (Piel et al. 2000). Fig. 5 D shows an example in a dividing HeLa cell. Clearly, under these conditions, C-Nap1 is associated with both parental centrioles, that is, with the one located in the cell body (d and f), and the one located close to the future cleavage site (c and e).

Split Centrosomes Are Functional

Our data indicate that interference with C-Nap1 function severely affects centrosome structure. Although split centrosomes are commonly seen in a small proportion of untreated cells, the frequency of cells with split centrosomes increases drastically when C-Nap1 function is disrupted. Moreover, this splitting occurs even when the MT or MF networks are disturbed. This argues that, in the absence of a connective structure, split parental centrioles can drift apart passively. Splitting occurs between parental centrioles and each centriole remains associated with the surrounding PCM. Thus, cells harboring split centrosomes are able to nucleate MTs. Furthermore, it is remarkable that centrosome splitting does not interfere with the ability of antibody-injected cells to divide. Spindles formed in the presence of anti–C-Nap1 antibodies appeared normal when stained with anti–γ-tubulin antibodies, suggesting the presence of functional poles. However, live-imaging studies and quantitative analyses of chromosome segregation would be required to determine whether such spindles are fully functional in the error-free segregation of chromosomes (for further discussion see Fry et al. 1998a).

The Centrosome Association of C-Nap1 Is Cell Cycle Regulated

From the proposed function of C-Nap1, we predicted that this protein should be displaced from centrosomes, or functionally inactivated, when parental centrioles are separated at the onset of mitosis. Our earlier studies had indeed suggested that C-Nap1 levels may be diminished at mitotic spindle poles, but it was impossible to exclude epitope masking as an alternative explanation (Fry et al. 1998b). Furthermore, though C-Nap1 protein levels in total cell lysates are detectably diminished during mitosis (Mayor, T., and E.A. Nigg, unpublished results), these results are difficult to interpret in view of a large cytoplasmic C-Nap1 pool. Thus, the use of antibodies directed against distinct parts of C-Nap1 proved critical for examining the fate of C-Nap1 during the cell cycle. We found that antibodies directed against widely separated C-Nap1 domains all produced negligible staining of mitotic spindle poles, strongly arguing against the caveat of epitope masking, and confirming that C-Nap1 truly dissociates from centrosomes during mitosis.

C-Nap1 Shows a Restricted Localization within the Centrosome

Using antibodies raised against the NH₂ terminus, COOH terminus, and middle domain of C-Nap1 for immunoelectron microscopy, we found that all three domains are located close to the proximal tips of centrioles. Although the NH₂-terminal end domain was slightly more distant from the centriole than either the middle or the COOH-terminal domain, we obtained no evidence to indicate that C-Nap1 oligomers could span the entire distance between the two parental centrioles. We conclude, therefore, that additional proteins must participate in a centriole–centriole linkage. The identity of these proteins is presently unknown, although δ-tubulin (Chang and Stearns 2000), as well as a putative calcium-binding protein (Moudjou et al. 1991), represent candidates. Of equal importance, careful examination of centrioles in the process of duplication revealed a conspicuous absence of C-Nap1 from the contact area between parental centrioles and procentrioles. This strongly suggests that the structure connecting a procentriole to its parental centriole is of a different nature than the one proposed to exist between the two parental centrioles.

We thank Bernard Eddé (Collège de France, Paris, France) for a kind gift of the GT335 antibody, Jorgos Pyrowolakis (Max-Planck-Institute for Biochemistry [MPIB], Martinsried, Germany) for assistance with FACS^® analysis, and the Ullrich laboratory (MPIB) for the use of their microinjection setup. We also thank Monika Matzner for technical assistance, and many colleagues in our laboratory and beyond for stimulating discussions.

In its initial stage, this work was supported by the Swiss National Science Foundation (31-50576.97) and the Canton of Geneva.