Tomography of the cell nucleus using confocal microscopy and medium voltage electron microscopy

Critical Reviews in Oncology/Hematology 69 (2009) 127–143

Tomography of the cell nucleus using confocal microscopy and medium voltage electron microscopy Pavel Tchélidzé a , Aurore Chatron-Colliet b , M. Thiry c , Natahlie Lalun d , Hélène Bobichon b , Dominique Ploton b,e,∗ a Institute of Biology Iv. Javakhishvili, Tbilissi State University, Tbilissi, Georgia UMR CNRS 6237, Université de Reims Champagne-Ardenne, 51 rue Cognacq-Jay, 51095 Reims Cédex, France Laboratoire de biologie cellulaire et tissulaire, Université de Liège, Institut d’anatomie (Bât L3), 20 rue de Pitteurs, 4020 Liège, Belgique d Laboratoire de Biologie cellulaire, Génétique et Histologie, UFR de Médecine, 51 rue Cognacq-Jay, 51095 Reims Cédex, France e Laboratoire Pol Bouin, CHRU Maison-Blanche, 45 Rue C. Jay, 51092 Reims Cédex, France b

c

Accepted 18 July 2008

Contents 1.

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Structural organization of the cell nucleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1. Chromatin architecture of the human genome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Chromosome territory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Proximity of distal genes for transcription and translocation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4. Molecular mobility within the cell nucleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5. Organization of nuclear functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Methods for investigating the 3D organization of the cell nucleus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Molecular markers of nuclear domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.1. Nucleic acid synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. DNA, RNA and chromatin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3. In situ hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.4. Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Confocal microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Electron tomography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Examples of application. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. DNA replication sites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. 3D FISH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Nucleolus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.1. rRNA synthesis and processing in the nucleolus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.2. 3D organization of RPI and rDNA genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.3. 3D organization of pKi-67 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Auto-fluorescent fusion proteins (GFP and its derivatives) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Reviewers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix A. Supplementary data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author at: UMR CNRS 6237, UFR de Médecine, 51 Rue Cognacq-Jay, 51095 Reims Cédex, France. Tel.: +33 3 26 91 83 36; fax: +33 3 26 91 35 23. E-mail address: [email protected] (D. Ploton).

1040-8428/$ – see front matter © 2008 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.critrevonc.2008.07.022

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Abstract: Changes in nuclear structures are widely used by pathologists as diagnostic and prognostic indicators in cancer cells. Recent studies have demonstrated that the cell nucleus is probably the most complex organelle in the cell. It contains the genome and is the site of all related activities such as DNA repair, DNA duplication, RNA synthesis, RNA processing and RNA transport. These activities take place within dynamic three-dimensional compartments. The detailed study of these compartments requires an approach termed “cell tomography” based on 3D imaging using confocal microscopy and electron tomography. In this paper, we will first summarize the most recent findings concerning the organization of the cell nucleus. We will then describe markers used to identify molecules specific for various nuclear compartments and their use in tomography of the cell nucleus by confocal microscopy and electron tomography. © 2008 Elsevier Ireland Ltd. All rights reserved. Keywords: Nucleus; Chromatin; Nucleolus; Spatio-temporal organization; 3D visualization; Electron tomography

1. Structural organization of the cell nucleus In tumor cells, many changes in nuclear structures serve as important diagnostic and prognostic indicators for pathologists [1–3]. Although the relationships between nuclear modifications and cancerous status are not fully understood, many studies over recent years have considerably increased our knowledge of the cell nucleus in cancerous cells [4]. The cell nucleus contains the genome and all related activities such as DNA repair, DNA duplication, RNA synthesis, RNA processing and RNA transport. A large number of studies published during the last 10 years have demonstrated that the nucleus is probably the most complex organelle in the cell [5–10]. These studies demonstrated that nuclear processes take place within three-dimensional compartments, which are dynamic at the different levels of their organization, i.e. from constitutive molecules to chromosome territories [11]. We will summarize some of the recent most insightful findings, which will significantly challenge our view of the functional nucleus over the coming years. 1.1. Chromatin architecture of the human genome Chromatin in the nucleus is frequently described as heterochromatin (i.e. condensed and inactive chromatin) and euchromatin (i.e. decondensed and active). This simple view was substantially modified by the generation of the human transcriptome map (HTM) some years ago. It revealed that highly expressed genes are clustered in particular regions present in all chromosomes [12] called RIDGES (regions of increased gene expression). These regions correspond to gene dense and highly expressed chromatin domains [13,14]. These findings support previous studies showing that sucrose sedimentation of chromatin fiber can identify compact and open chromatin [15]. Open chromatin contains the highest gene density and is decondensed but genes in such domains may be inactive. Conversely, compact chromatin contains very few genes, some of which may be active. 1.2. Chromosome territory Within the nucleus, each chromosome occupies a discrete territory called chromosome territory (CT). Although

the position of a CT is not fixed, it is non-random. Thus, gene-rich chromosomes are preferentially positioned in the center of the nucleus, whereas gene-poor chromosomes are located at the periphery of the nucleus [14,16–19]. This organization is probabilistic [13,19] and is strictly defined during the early G1 phase of the cell cycle, during which chromatin mobility is very high [20,21]. It remains unclear whether such positioning may depend on or, conversely, contribute to variations in gene expression [14,22]. Studies examining the position of active genes relative to CT demonstrated that genes located within CT can be transcribed [23]. Additionally, the location of highly active sequences is not restricted to this area but can loop out from the CT into the interchromosomal space [19,24–28]. These loops thus constitute a fuzzy domain around each CT in which different CTs can intermingle [25,29]. 1.3. Proximity of distal genes for transcription and translocation Active RNA polymerase II molecules are found in discrete foci or “transcription factories” in which groups of distal genes may be actively transcribed [30]. Active genes appear to migrate to these sites in order to be transcribed but exit the sites when their initiation and transcription are inhibited [31]. There is substantial evidence demonstrating that chromatin organization is dynamic [32–34], i.e. can change as a function of differentiation or cell activity [35]. Within differentiating cells, co-regulated genes are located close to each other due to the non-random juxtaposition of lineage-specific domains [36–40]. A recent study [41] also demonstrated that several cancer-associated genes change their position within the nucleus during the early stages of breast cancer, although their repositioning was not related to changes in their activity. Recent evidence strongly suggests that the reorganization of chromatin domains results from active movements, which are directly dependent on actin and myosin [42–44]. The proximity of distal genes and chromosome territories has other important implications. Indeed, many studies clearly demonstrate that non-random spatial organization of chromosome territories underlies specific translocation in murine and human cancerous tissues [45–48].

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1.4. Molecular mobility within the cell nucleus The very high concentration of macromolecules in the nucleus (around 100 mg/ml) causes crowding forces, which maintain nuclear sub-compartments [49,50]. However, RNA and proteins are clearly highly dynamic in the nucleus. For example, chromatin-binding proteins rapidly scan the whole genome and link to their binding site for only a few seconds [51,52]. Such characteristics were first demonstrated in the study of RNA pol I subunit assembly for ribosomal genes transcription [53]. This study showed that distinct subunits assemble inefficiently to form active RNA polymerase I on the promoter and that pre-rRNA elongation is around 95 nucleotides per second for a human rDNA gene of 13.3 kb. Similar findings were more recently obtained for RNA polymerase II subunits [54]. Nuclear processes are now considered to be compartmentalized due to the stochastic assembly of freely diffusing proteins around a given gene [55]. This assembly is highly dynamic, allowing for a rapid regulatory mechanism in the control of nuclear activity [35,51,52,56,57]. 1.5. Organization of nuclear functions Nuclear functions occur in three-dimensional nonmembrane-limited domains. These domains are identified by the stochastic association of free molecules from the nucleoplasm to perform dedicated functions [7,8]. For example, DNA synthesis takes place in non-permanent domains whose shape and 3D localization typically vary during the different stages of S-phase [58,59]. Other domains can be clearly identified within each nucleus due to their high level of activity and resulting prominence [60,61]. These domains constitute nuclear organelles such as: nuclear speckles (mRNA synthesis and processing [62]), Cajal bodies (RNA modifications [61]), nucleoli (rRNA synthesis and processing [63,64]), PML bodies [65]. One characteristic of nuclear organelles is that they can be fractionated and their detailed protein composition can be analyzed by mass spectrometry. For example, the nucleolar proteome comprises around 700 proteins [66]. The flux of these proteins in response to metabolic inhibitors has been also analyzed [63,67,68]. Another way to analyze nuclear domains is to investigate their structure using advanced microscopic techniques. Considering that nuclear domains are three-dimensional entities, approaches need to be used, which will provide insight into the 3D organization of both their functions and molecular components. Cell tomography is the method of choice to address these issues. It can be performed by confocal or electron tomography depending on the resolution requirements. In the following section, several examples will be discussed demonstrating the use of cell tomography in the study the cell nucleus. In particular, we will describe some of the markers currently used to specifically identify molecules of interest; we will also compare several image processing tech-

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niques, which are used to interpret the results. We will then describe the potential benefits of new markers aimed at use in correlative microscopy.

2. Methods for investigating the 3D organization of the cell nucleus 2.1. Molecular markers of nuclear domains Each nuclear domain can be identified by the visualization of one or several molecules specifically located within it, such as DNA, RNA or proteins. 2.1.1. Nucleic acid synthesis To localize DNA replication sites, living cells are classically labeled with BrdU for several minutes and then exposed to a fluorescent anti-BrdU antibody [59]. Imaging by confocal microscopy [59,69] and by electron microscopy on ultrathin sections [69] demonstrated that the 3D organization of DNA replication sites varied during the S-phase. Five stages were thereby identified using variables such as the size and 3D localization of replication sites within the nucleus. These results clearly showed that nuclear functions are not randomly organized but are in fact strictly organized both in space and time. Nascent RNA is normally visualized by labeling living cells with BrUTP and using an antibody tagged with a fluorescent marker on fixed cells. These classical methods require either the encapsulation of cells in agarose beads and treatment with ␣-amanitin to label nucleoli [70] or permeabilization of cells [71]. As an alternative approach, we used a transfection reagent as vector for the uptake and incorporation of BrUTP into growing cells [72]. This approach appeared to be very efficient for the rapid labeling of nascent RNA both in the nucleoplasm and nucleolus and for use in 3D studies using confocal microscopy (see Section 3.3.1). 2.1.2. DNA, RNA and chromatin DNA and RNA can be easily and specifically stained with various fluorescent dyes in either fixed or living cells and visualized by confocal microscopy [73–76]. DNA can also be specifically identified using the Feulgen reaction based on DNA hydrolysis with HCl and detection with the fluorescent dye pararosaniline [77]. The equivalent technique used to study cellular ultrastructure is the so-called osmium ammine reaction. This technique has allowed the study of chromatin conformation on ultrathin and thick cryo-sections using stereo-pairs [78,79]. The 3D organization of chromatin in fixed or in living cells can be addressed using a more efficient and elegant approach involving the localization of H2B-GFP in transiently transfected cells or in established cell-lines. This method was developed several years ago [74,80,81].

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2.2. Confocal microscopy

2.1.3. In situ hybridization The identification and localization of one gene or one specific RNA molecule within the nuclear volume requires the use of fluorescence in situ hybridization (FISH). It is far beyond the scope of this paper explain this approach in detail (see cited references for reviews). Classically, DNA probes are nick-translated using biotin-16-dUTP or digoxigenin-11-dUTP and the resulting hybrid molecules are detected using a fluorescent conjugated avidin or antidigoxigenin antibody, respectively [80,82]. A single gene and its transcribed RNA may also be simultaneously visualized [83]. In situ hybridization can be also performed on ultrathin sections to localize genes and RNA at the ultrastructural level. This approach uses nick-translated probes with biotinylated dUTP, which are detected after in situ hybridization with colloidal gold particles coupled to a specific antibody. This approach was particularly useful to confirm the localization of rRNA genes and their products within the nucleolar compartments [84–86]. However, to the best of our knowledge, it has not been used for electron tomography.

The first convincing applications of confocal microscopy in cell biology emerged at the end of the 1980s [95]. The major use of a confocal microscope is to produce images of complex structures located within a well preserved cell using optical sections, i.e. views with a very limited depth of field [96]. For this reason, a confocal microscope can be considered as an optical tomograph aimed at investigating the 3D organization of fluorescently labeled cells. Cell tomography using confocal microscopy has advanced significantly over the last few years mainly through marked improvements in computers and software. Thus, data can now be rapidly and easily processed for complex 3D visualization and 3D quantification. This is particularly beneficial to investigate the 3D complexity of the living or fixed cell. The continued development of new markers and probes for nuclear domains over recent years has led to (and will lead to) significant advances in the study of the nucleus (see above and below).

2.1.4. Proteins Proteins are mostly identified using antibodies (mono or polyclonal) followed by detection with secondary antibodies coupled to a fluorescent marker (confocal microscopy) or to a dense marker such as colloidal gold (electron tomography). Some markers contain both a fluorochrome and a dense marker such as fluoronanogold (FNG), which can be visualized by fluorescence and by electron microscopy ([87,88] and Sections 3.3.2 and 3.3.3). It has been possible to visualize cells expressing GFP fusion proteins (or its derivatives) in transiently transfected cells or established cell-lines for several years. Fusion proteins can be directly observed by confocal microscopy in fixed or living cells or indirectly by using antibodies against GFP labeled with colloidal gold for electron tomography [89]. Finally, some nuclear proteins can be identified by means of cytochemical methods. We developed the Ag-NOR staining method, which identifies argyrophilic nucleolar proteins [90]. This staining gives rise to tiny dots of metallic silver; these can be identified by transmitted light for its quantification in cancer cells as a prognostic marker [91,92] or by reflected light in confocal microscopy for detailed localization [93]. Additionally, silver dots can be identified on thick sections for electron tomography. This approach allowed the description of the 3D organization of metaphasic NORs at a very high resolution [94].

The three-dimensional (3D) organization of structures (from molecules to cells) can be investigated with a resolution of a few nanometers (nm) using electron tomography [97]. The reconstructed volume of the structure (called electron tomogram) is generated using multiple projection views of a sample tilted in a transmission electron microscopy from −60◦ to +60◦ [98]. One rapidly developing application of electron tomography is to image vitrified unstained molecules or supra-molecular complexes in low-dose conditions with a cryo-TEM [99]. Tomograms with a resolution of 4–5 nm can be obtained with this approach, using specimens thinner than 100 nm (suspensions of molecules or ultrathin cryo-sections) [97]. However, electron tomography of complex molecular machinery within organelles (such as the nucleus) requires image sections of several hundred to several thousand nanometers thick and specific markers for identification of the molecules. Specific molecules can be identified using cytochemical or immunocytochemical labeling incorporating electron-dense markers [100]. Indeed, we demonstrated that the end-product of Ag-NOR cytochemical staining, consisting of 5–10 nm silver dots, could be imaged in sections several micrometers thick by using a 300-kV STEM [101]. This approach allowed us to investigate the 3D organization of nucleolar organizer regions (NORs) during metaphase for the first time and to propose a model of their fine structure [94]. Nanogold technology can also be used to specifically

2.3. Electron tomography

Fig. 1. Three-dimensional visualization of DNA replication sites using EdU incorporation. Cells showing one of the five typical patterns of DNA replication (IA, IB, II, IIIA and IIIB) were imaged by confocal microscopy to obtain z-series constituted with around 50 optical sections. Volume and surface rendering were computed for a sliced central sub-volume (figures A1, A2, B1, B2, C1, C2, D1, D2, E1 and E2) and for the whole volume (figures A3, A4, B3, B4, C3, C4, D3, D4, E3 and E4), respectively. Bar: 3 ␮m. Captions for the movies: movies corresponding to figures: A3, A4, B3, B4, C3, C4, D3, D4, E3 and E4 are presented as supplementary data. Each movie shows a total rotation of the computed volume after volume rendering (movies A3–E3) and after surface rendering (movies A4–E4).

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Fig. 2. Localization of BrUTP-labeled rRNAs. Evidence for the migration of BrUTP-labeled rRNAs from the nucleolus to the cytoplasm. ␣-Amanitin-treated HeLa cells were lipofected for 15 min with BrUTP-FuGene-6 complexes and then cultured in medium without BrUTP for 2 min (A and B), 15 min (C and D), 30 min (E and F) and 60 min (G and H) before being fixed. BrUTP-labeled rRNAs were detected by an indirect immunofluorescence method. (A, C, E and G): phase contrast images; (B, D, F and H): single optical section of the same fields obtained in confocal microscopy. Bar: 10 ␮m. With permission, Ref. [72].

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Fig. 3. Three-dimensional structure of nucleolar sub-domains containing the BrUTP labeled rRNAs during their movement within the nucleolus. ␣-Amanitintreated HeLa cells were lipofected for 15 min with BrUTP-FuGene-6 complexes and then cultured in medium without BrUTP for 2 min (A and B), 15 min (C and D), 30 min (E and F) and 60 min (G and H) before being fixed. BrUTP-labeled rRNAs were detected by an indirect immunofluorescence method. (A, C, E and G): single optical section; (B, D, F and H): three-dimensional visualization after three-dimensional reconstruction of the corresponding fields. Insets in (B), (D) and (F) show a higher magnification of three selected areas. Dashed lines indicate the outline of the nucleus. Bar: 5 ␮m (1 ␮m in insets in (B) and (D); 2.5 ␮m in inset in F). With permission, Ref. [72].

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identify proteins with electron-dense labeling [102]. With this new approach, nanogold dots (around 1 nm in diameter), amplified with silver to increase their size to 5–10 nm particles, are used for preembedding immunostaining [103]. We showed that this labeling is a very efficient method for identifying proteins and tracing their 3D organization within thick sections using electron tomography performed with a medium voltage STEM [87,104]. This has been used to study RNA polymerase I and pKi-67 as discussed bellow (Sections 3.3.2 and 3.3.3). Moreover, preliminary results from our group demonstrated that this approach could be very beneficial to study the 3D organization of GFP-tagged proteins identified with anti-GFP antibodies and detected with silver-amplified nanogold particles [89].

3. Examples of application 3.1. DNA replication sites The development of labeling and 3D visualization techniques has led to improved understanding of 3D nuclear organization, particularly in the study of DNA replication sites. As stated above, until recently, replication sites have been identified using incorporation of BrdU within living cells [69,59]. However, this technique requires acid depurination or nuclease treatment to reveal incorporated BrdU molecules using specific antibodies [59]. It is possible that these strong pre-treatments modify the fine structural organization of chromatin and the intensity of labeling [105]. These limitations were recently overcome through the development of a new method based on the incorporation of 5-ethynyl-2 -deoxyuridine (or EdU) instead of BrdU. EdU is easily detected with a fluorescent azide, rather than with an antibody [105], does not require acid or nuclease pre-treatment and preserves chromatin structure. In this section, we will discuss the use of this new approach to investigate the organization of replication sites coupled to high-resolution 3D processing of data obtained by confocal microscopy. Briefly, EdU was added to cell culture for 5–30 min, as recommended (Click-itTM kit, Invitrogen). Cells were washed, fixed with 3% paraformaldehyde and permeabilized with 0.1% Triton X-100. EdU incorporated into S-phase cells was labeled with Click-itTM detection cocktail for 30 min using Alexa Fluor® 488.

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Cells were mounted in Citifluor® and imaged by confocal microscopy using a 1024 ES confocal microscope (BioradZeiss) equipped with a x63, 1.4 NA plan apochromatic objective. z-Series were obtained through the whole nucleus at a pitch of 0.3 ␮m. Each z-series was processed using Amira® 4.1 (Mercury Computer Systems). Volume and surface renderings were processed to produce still images and movies. The five distinct patterns of DNA replication (IA, IB, II, IIIA and IIIB) were identified using this approach (Fig. 1; row A: IA, row B: IB, row C: II, row D: IIIA, row E: IIIB). The labeling reveals numerous discrete tiny dots displaying typical groupings and 3D distribution for each pattern. Surface and volume renderings were computed on the whole (or part of the) data sets to analyze the complexity of this distribution. The same procedure was used for each z-series. A slice 2 ␮m thick was cut parallel to the larger diameter in the center of the nucleus and labeling was visualized using volume and surface rendering (Fig. 1, columns 1 and 2, respectively). Similarly, volume and surface renderings were computed from the whole volume and visualized on still images (Fig. 1, columns 2 and 3, respectively) and movies. Each rendering mode has specific advantages and disadvantages. Surface rendering displays objects with a gray value above a chosen threshold as solid entities. This gives images with a strong 3D appearance. However, data below the threshold are not shown. Volume rendering displays all the objects transparent and with their exact gray level. Thus, all data are shown but the resulting image is complex, requiring analysis as a movie. Each rendering mode thereby has its own benefits and is thus complementary to the other. Images can now be obtained using both approaches within a fraction of a second using a classic personal computer equipped with a good graphic card to process the data. For our purposes, such processing facilitated an investigation of the 3D distribution of replication sites (see still images and movies). Thus, during phase IA (Fig. 1A1–A4), replication sites appeared as discrete spots rarely grouped together and absent from the nuclear periphery. During phase IB (Fig. 1B1–B4), replication sites were organized into large clumps linked with fibers to the periphery of the nucleus. During phase II (Fig. 1C1–C4), replication sites were present in small clumps along the nuclear envelope. There was intense labeling in the nucleolus, but no labeling was visible between the nucleolus and nuclear envelope. During phase IIIA (Fig. 1D1–D4), intense labeling was visible at the

Fig. 4. Ultrastructural localization of RNA polymerase I within A549 cells. Anti-RPI antibodies were revealed with fluoronanogold, followed by silver enhancement. (A) After embedding, ultrathin sections (80 nm) were counterstained and observed with an electron microscope at 100 kV. The main nucleolar components are identified (FC, DFC and GC). A high density of particles is observed within the fibrillar components of the nucleolus. (B) Tomographic study of A549 cells immunolabeled with anti-RNA polymerase I antibodies. Contrast was inverted so that silver particles appear white. A 500-nm thick section observed using a STEM working at 250 kV, is displayed: several independent clusters, 270 nm in diameter, are seen in (B). (C–E) Different projections of the tomogram were calculated after tomographic reconstruction of the cluster framed in (B). At +15◦ (C), five 60-nm coils are evidenced, as indicated by brackets (#1–5). The large circle shows the area where the coils are fused together. The arrows point to twines, 20 nm in thickness. (F) A stereo-pair of the tomogram presented in the same orientation as in (E) was calculated thanks to a surfacic rendering mode. (G–J) Four successive 30-nm thick sections were achieved with a coronal orientation within the tomogram presented in (E). Asterisks (H and I) indicate the internal part of the cluster, devoid of labeling, and arrows (H–J) refer to twines. Bar: 0.5 ␮m (A), 200 nm (B) or 100 nm (C–J). With permission, Ref. [87].

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periphery of the nucleus, within small clumps in the nucleoplasm and in a thick continuous sheath around the nucleolus. Finally, during phase IIIB (Fig. 1E1–E4), labeling was visible as a few medium-sized roundish particles evenly dispersed within the nucleoplasm. Such a labeling method, combined with data processing (surface and volume rendering), will be of great value in the analysis of 3D organization of replication sites within normal and cancer cells, in different conditions of culture or when treated with anti-cancer drugs. 3.2. 3D FISH Fluorescence in situ hybridization is used to precisely identify genes and transcripts within the volume of the nucleus. Using confocal microscopy, this approach has provided extensive insight into the 3D organization and relative distribution of genes within the nucleus of normal and cancer cells. Detailed technical approaches have been described by experts in this field [106–108]. Findings obtained with FISH have been key to demonstrating that genes are not localized at random in the volume of the nucleus. Some of the most important recent findings obtained in this field address: the tissuespecific organization of genes [46], the consequences of non-random organization of the genome in chromosomal translocations in cancer cells [45,47,48], the positioning of active and inactive genes relative to chromosome territories [23], the role of DNA hypomethylation in nuclear organization modifications [109], the importance of gene density in the positioning of chromatin within the nucleus [16,27,28,23,17], gene repositioning during early tumorogenesis [41] and gene positioning within differentiated cells [36,37]. 3.3. Nucleolus 3.3.1. rRNA synthesis and processing in the nucleolus The nucleolus is the nuclear site in which synthesis and processing of pre-ribosomal RNA take place [110]. At the ultrastructural level, four nucleolar sub-compartments can be identified as a function of their electron density and texture: fibrillar centers, dense fibrillar component, granular component and peri- and intra-nucleolar chromatin. Considering that the nucleolus is responsible for synthesis and processing of more than half of all RNA [63], it should be a suitable model to study both the 3D and temporal organization of RNA synthesis and processing. Such a study involves

Fig. 6. The Ki-67 protein was immunolabeled and revealed by steptavidinfluoronanogold. After silver amplification and embedding, a counterstained ultrathin section (80 nm thick) was observed at 80 kV. This conventional electron microscopy picture shows the preservation of the structure, in which FCs associated to DFC (circles) are clearly identified. The labeling appears as numerous dots ∼15 nm in diameter that are mainly localized in the cortical part of the nucleolus. In some areas, the labeling seems to be organized as fibers, 50 nm in diameter (arrows and parallel lines). Bar: 1 ␮m. With permission, Ref. [104].

firstly identifying the sites of pre-rRNA transcription and secondly, localizing the sites of processed rRNA. To this end, we developed a new approach allowing us to rapidly label nascent rRNA using BrUTP complexed with a transfection reagent (fuGene-6® ). This method avoids the permeabilization of living cells [72]. Nascent rRNAs labeled with BrUTP were further identified using an indirect immunocytological method involving an antibody against BrUTP. Their precise localization was determined by confocal microscopy. A major advantage of this approach is the rapid penetration of BrUTP into the cells, which allowed pulse-chased experiments. 3D reconstruction from optical sections obtained by confocal microscopy allowed us to examine the 3D organization of sites where pre-rRNA transcription takes place and where rRNA molecules move in and out of the nucleolus. The migration of BrUTP-labeled rRNAs was observed in ␣-amanitin (an inhibitor of RNA polymerase II)-treated cells which were pulse-labeled for 15 min and then chased for 2, 15, 30 and 90 min (Fig. 2). Visualization of the same cell firstly by phase-contrast microscopy and secondly by confocal microscopy showed that rRNA synthesis takes place in tiny structures within nucleoli. Labeled rRNA molecules then migrated towards the periphery of the nucleolus, into the nucleoplasm and finally into the cytoplasm. Series of optical sections were then processed for 3D reconstruction and surface rendering imaging (Fig. 3). Sites containing nascent pre-rRNA as individual full beads around 0.5 ␮m in diameter

Fig. 5. Spatial distribution of the Ki-67 protein observed by confocal microscopy. After fixation of A549 cells, the Ki-67 antigen was detected by immunolabeling with a specific antibody, revealed by steptavidin-fluoronanogold. The fluorescent dye of the probe (FITC) was visualized with a confocal microscope. (A–E) Gallery of five optical sections taken from a z-series. The labeling is distributed either in cords, approximately 300 nm in width, or in small, perinucleolar dots (A and E). (F) Projection of all the sections contained in the z-series. (G) Orthogonal section of the reconstructed volume, performed at the level of the arrow in Fig. 5C. (H–J): Transversal sections performed at level of the three arrowheads in Fig. 5G. (K–M) Surfacic visualization of pKi-67. A surfacic mode was applied to the optical sections presented in (A–E); (K) Stereo-pair of a back view of the volume presented in (F). (L) Stereo-pair of a front view, corresponding to the area framed in (C). (M) Split view of the stereo-pair shown in (L). Bar: 2 ␮m in (A–J) and 1 ␮m in (K–M). With permission, Ref. [104].

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were organized in the form of several necklaces. In the next stage, the labeling was arranged in several ringlets 0. 8 ␮m in diameter (without fluorescence in their center) comprising large strands. Labeled entities thus constituted fused hollow spheres covered with several threadlike structures 1.5 ␮m in diameter. In the last step, the labelling was arranged as several rims 3–4 ␮m in diameter at the periphery of the nucleolus. We used two approaches to precisely localize pre-rRNA. For the first approach, confocal microscopy revealed colocalization of BrUTP rRNA and RNA polymerase I (RPI) in the same cells [72]. Quantification of pixels with co-labeling showed that nascent pre-rRNA was strictly co-localized with RPI and that processed rRNA migrates towards the periphery of these sites. In the second approach, immunolocalization of BrUTP-labeled rRNA on ultrathin sections clearly identified fibrillar centres as nucleolar sites containing nascent rRNA [87]. 3.3.2. 3D organization of RPI and rDNA genes As described above, fibrillar centers (FC) are sites of prerRNA synthesis and consequently contain rDNA genes. This finding is consistent with previous findings showing highly decondensed DNA (characteristic of rDNA) strictly localized within FC ([110] for a review) and ultrastructural in situ hybridization studies demonstrating rDNA within the cortex of FC [84]. Given that each rDNA gene is covered with around 120 RPI molecules, we hypothesized that electron tomography of immunolabeled RP1 could reveal the 3D organization of transcribed rDNA genes [87]. We immunolabeled RPI molecules before embedding with fluoronanogold, which was further amplified with silver. After embedding, ultrathin sections (100 nm thick) were observed by electron microscopy. Labeling was mostly localized within FCs (Fig. 4A). The presence of RPI outside FCs was interpreted as free RPI subunits diffusing around the rRNA genes within the FCs [53]. Analysis of 500 nm thick sections using a 250 kV scanning and transmission electron microscope (STEM) revealed several discrete clusters of numerous silver/gold particles (with a density of particles 10 times higher than in the nucleoplasm) (Fig. 4B). The thick sections were then tilted within the STEM from −60◦ to +60◦ (2◦ steps) and tomograms were obtained (Fig. 4C–E). Surface rendering (Fig. 4F) and virtual sections (Fig. 4G–J) taken in all direc-

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tions showed that particles were not localized at random. Conversely, RPI molecules were arranged in coils mainly localized in the cortex of FCs and successive RPI molecules were organized into loops and twines [87]. These findings led to the first proposal of a model for the spatial organization of rDNA genes within FC [87]. 3.3.3. 3D organization of pKi-67 The Ki-67 antigen (pKi-67) is present only in nucleoli of cycling cells and not in resting (G0) cells. pKi-67 is widely used as a prognostic marker to estimate the growth fraction in human cancer tissues [111]. Many studies suggest that pKi-67 plays a role in chromatin organization and have frequently reported its localization within the nucleolus. We studied the 3D spatial distribution of pKi-67 by determining its precise localization using confocal microscopy and electron tomography [104]. To compare the 3D reconstructions obtained at different levels of resolution, we used a marker (called Fluoronanogold® or FNG) containing one fluorescent probe and one nanogold particle. This marker can thus be visualized using confocal microscopy and electron microscopy either in different cells labeled in exactly the same conditions or within the same cell (correlative microscopy). Here, we discuss the findings obtained using these complementary approaches. In a first step, immunolabeled pKi-67 was visualized using FNG and confocal microscopy (Fig. 5). In the optical sections (Fig. 5A–F), labeling revealed several thin cords localized mainly around the nucleolus and partly along the nuclear envelope or within the nucleoplasm. In transversal sections (Fig. 5G–J), labeled protrusions appeared to contact the nuclear envelope. All optical sections were processed for surface rendering, presented as stereo-pairs either of the whole nucleus (Fig. 5K) or of one part of the nucleolus (Fig. 5L). This demonstrated an irregular pattern of labeling, arranged as a contorted cord around the nucleolus. This was confirmed in a transversal section obtained from this data (Fig. 5M). Simultaneous labeling of DNA with chromomycin A3 demonstrated that nucleolar pKi-67 is co-localized with DNA [104]. We examined the localization of pKi-67 at a higher resolution. Cells of the same culture were labeled in the same conditions as described above and were studied using electron

Fig. 7. Electron tomography analysis of Ki-67 protein. Thick sections (2 ␮m in thickness), in which pKi-67 molecules were immunodetected prior to embedding, were analyzed with a STEM working at 250 kV. Image contrast was reversed for convenience. (A–E) Gallery of five projections taken from the same tilt series. Using certain view angles, the labeling appears as a cord, 250–300 nm in thickness (bracketed by arrowheads). Contacts with the nuclear envelope are evidenced (arrows). (F) The projections corresponding to an entire tilt series were used to reconstruct a tomogram. The 0◦ projection shows a similar organization to that observed on the initial image (compare C and F), indicating that the reconstruction process introduces no distortion. (G–I): Three 50-nm thick digital sections taken from the top (G), the middle (H) and the bottom (I) of the tomogram. The labeling is distributed in cords (bracketed by arrowheads), 250–300 nm in diameter, disposed around a central, unstained area. The cords consist of fibers, 40–50 nm in diameter (frames in G and H). Arrows point to contacts with the nuclear envelope. (J–L) Electron tomography performed on a 0.5-␮m thick section. (J): STEM observation (250 kV) of a 0.5-␮m thick section. The two framed areas were observed at a higher magnification (K and L). (K) A detailed view of a protrusion in close contact with the nuclear envelope. (L) A view of the central part of the labeling, showing the organization of the perinucleolar cord. In both cases, the labeling appears as a complex network of intertwined fibers, whose compaction is higher in the protrusion (K) than in the perinucleolar cord (L). Bar: 0.5 ␮m (A–I), 2 ␮m (J) and 200 nm (K and L). With permission, Ref. [104].

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microscopy. After silver enhancement, cells were embedded in plastic and ultrathin sections were cut. All typical nucleolar components were clearly recognizable, thereby confirming a good preservation of cellular ultrastructure (Fig. 6). Labeling was predominant on condensed perinucleolar chromatin and was sometimes organized as short threads around 50 nm in diameter. Labeling was not present within fibrillar centers or dense fibrillar components, but was observed within chromatin outside the nucleolus. We then used electron tomography to study pKi-67 in the same cells. Thick sections (0.5–2 ␮m) were tilted in a medium voltage electron microscope operated at 250 kV. Part of the tilt-series is shown in Fig. 7A–F. A tomogram was then obtained and digital sections (50 nm thick) were analyzed to study the complex organization of the labeling. Labelling is organized as different cords 300 nm in diameter comprising thinner fibers 50 nm in diameter (Fig. 7G–L). A more detailed characterization of these fibers was carried out at higher magnification of the tomogram [104]. In summary, this study demonstrated that a thorough 3D study of pKi-67 was possible at different levels of resolution using the same marker: fluoronanogold. We established that pKi-67 co-localizes with perinucleolar heterochromatin and forms cords, 250–300 nm in diameter composed of fibers, 30–50 nm thick. These findings suggest that pKi-67 is involved in the organization of perinucleolar chromatin. 3.4. Auto-fluorescent fusion proteins (GFP and its derivatives) Autofluorescent fusion proteins (GFP and its derivative) are widely used in studies of the nucleus. Their use has demonstrated that proteins and RNA are very dynamic in all nuclear compartments and that the movement of these molecules can control cellular physiology [56,51]. For example, photobleaching experiments demonstrated that GFP-tagged nucleolar proteins are highly dynamic although nucleolar structures are stable [53,64]. Fusion proteins have also been used, without using antibodies, to study nuclear compartments [64]. This application is particularly important because it allows the study of native structures, which are otherwise potentially modified by exposure to fixatives (living cells) or pre-treatment (fixed cells). Moreover, this approach offers the possibility of mapping all nuclear proteins to better understand the nuclear organization. The use of histone H2B-GFP in such studies is a convenient way of analyzing chromatin organization within living cells in 2D [112] or 3D and 4D [113]. Moreover, the localization of genes and chromosome territories identified by FISH in cells with histone H2B-GFP-tagged chromatin allows the precise 3D organization of the nucleus to be determined [80]. The study of fusion proteins within living cells using confocal microscopy allows the detailed study of nuclear reorganization during drug treatments [64]. However, the detailed analysis of 3D time-series will require the devel-

opment of new software to describe, analyze and quantify the numerous structural modifications arising during such treatment as we demonstrated in a preliminary study [114]. Finally, electron tomography of immunolabeled GFP-tagged proteins may represent a convenient way to address the molecular organization of certain nuclear territories at a resolution ranging between 10 and 20 nm [89].

4. Concluding remarks The cell nucleus is a highly complex three-dimensional structure, the study of which requires various complementary approaches. Among the approaches available, tomography based on confocal and electron microscopy appears to be a very useful technique for unraveling the functional complexity of the cell nucleus. These approaches provide extensive insight into fundamental investigations (see references herein) and can also be applied in studies such as those investigating the effects of anti-cancer drugs on nuclear architecture [115]. Moreover, new methods in the field of photon microscopy now allow imaging with a resolution of the order of 30 nm [116]; thus, we may expect our knowledge of spatial organization of the cell nucleus to greatly increase over the next few years. Finally, there is no doubt that new development such as these will prove invaluable in the study of cancer and will be highly beneficial in diagnostic and prognostic applications.

Reviewers Professor Massimo Derenzini, University of Bologna, Department of Experimental Pathology, Via San Giacomo 14, I-40126 Bologna, Italy. Dr. Yves Usson, CNRS/TIMC-IMAG, UMR CNRS 5525, IN3S, Pavillon le Taillefer, Domaine de la Merci, F-38706 La Tronche, France.

Acknowledgements This work received financial support from the “Fonds National de la Recherche Scientifique Médicale” (grant n◦ 3.4540.06), Association de la Recherche contre le Cancer (ARC) (contract N◦ 4497), Ligue contre le Cancer (Departements de l’Aube, de la Marne et des Ardennes) and Région Champagne Ardenne for thesis grant support of ACC.

Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.critrevonc. 2008.07.022.

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Biography Dominique Ploton took his Research Habilitation in 1988. He is a professor of cell biology at the faculty of Medicine in Reims. His scientific expertise concerns the structure of the nucleolus and of the nucleus. More particularly he developed Ag-NOR staining which appeared as a good prognostic marker for human cancerous tissues. More recently he developed the use of 3D reconstruction and visualization to study the 3D organization of the nucleus imaged by confocal and electron microscopy (electron tomography).