“On the move”ments of nuclear components in living cells

“On the move”ments of nuclear components in living cells

Experimental Cell Research 296 (2004) 4 – 11 www.elsevier.com/locate/yexcr Review ‘‘On the move’’ments of nuclear components in living cells Paula A...

247KB Sizes 4 Downloads 31 Views

Experimental Cell Research 296 (2004) 4 – 11 www.elsevier.com/locate/yexcr


‘‘On the move’’ments of nuclear components in living cells Paula A. Bubulya and David L. Spector * Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 11724, USA Received 19 January 2004 Available online 12 April 2004

Abstract The use of GFP fusion proteins has dramatically changed our view of how the cell nucleus is organized and how functions are carried out. In this review we focus on recent advances related to the dynamics of chromatin domains, as well as the dynamics of nuclear proteins and several nuclear organelles. D 2004 Elsevier Inc. All rights reserved. Keywords: Chromatin domains; Gene expression machinery; DNA repair; Nuclear bodies; Live cell microscopy


Movement of chromatin domains

The use of green fluorescent protein (GFP) has led to an explosion of information regarding the localization and dynamics of proteins within living cells. Using this approach, one can determine not only where a protein localizes over time, but also, using photobleaching techniques, how much of the protein is mobile, and the frequency of exchange of mobile proteins between cellular compartments. Such techniques confirmed early on that many cytoplasmic structures are in constant flux (for a review, see Refs. [1,2]). The use of GFP fusion proteins to study processes within the cell nucleus was first initiated in 1997 to examine the dynamics of pre-mRNA splicing factors [3]. Since this initial study, we have learned much about the dynamics and functions of nuclear proteins, the processes in which they participate, and nuclear bodies where they often reside. This review focuses on recently published reports regarding the movements and kinetics of nuclear components in living cells. For related topics, we refer readers to other recent reviews regarding chromatin dynamics [4 – 9], nuclear membrane dynamics [10,11], dynamics of the nuclear pore complex [12], visualizing RNA [13], and computational imaging [14,15].

Subnuclear organization of yeast chromatin is thought to play a major role in transcription repression. One of the bestcharacterized examples of this involves anchoring of telomeres near the nuclear periphery. The dynamic movement of native telomeres was recently studied in living yeast cells by tagging chromosome ends with binding sites for GFP-lac repressor [16]. Yeast telomeres exhibited oscillating movement near the nuclear envelope that changed in frequency and distance depending upon the stage of the cell cycle. In yKu70 mutant strains, the GFP-tagged telomere was released to the internal nuclear space and moved a greater total distance over time within the nuclear interior compared to wild-type cells. Perinuclear anchoring was also disrupted in sir mutants, primarily in S-phase [16]. Restoration of silencing by recruiting Sir4p to telomeres in the absence of yKu restored perinuclear anchoring of GFP-tagged telomeres in S-phase. Based upon this result, the authors concluded that although yKu can anchor telomeres in the absence of Sir proteins, a partially redundant Sir-dependent telomere anchoring mechanism exists independently of yKu anchoring [16]. Interestingly, the dominance of these two mechanisms varies regarding different telomeres and different cell cycle phases. Therefore, it will be interesting to determine how this is regulated and if both telomeres on a particular chromosome are regulated in a similar way. In mammalian cells, telomere position is not constrained to the nuclear periphery; rather, telomeres are located

* Corresponding author. Cold Spring Harbor Laboratory, 1 Bungtown Road, Cold Spring Harbor, NY 11724. E-mail address: [email protected] (D.L. Spector). 0014-4827/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2004.03.018

P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11

throughout the nuclear volume. To observe telomere dynamics in living human U2OS osteosarcoma cells, telomeres were visualized by hybridization of fluorescently labeled protein nucleic acid (PNA) probes and the binding of CFPTRF2 to the telomere repeat sequences [17]. Movement of the majority of telomeres was confined within a radius of less than 1 Am, while a few telomeres exhibited movement over a significant distance (up to approximately 8 Am) during a period of 60 min [17], whereas yeast telomere oscillations rarely exceeded 0.3 Am [18]. Additionally, individual telomeres were observed to temporarily associate and then dissociate from telomere clusters [17]. In U2OS cells, which lack the enzyme telomerase, an alternative mechanism of maintenance of telomere length is used, and cells with this phenotype showed an association of a subset of telomeres with PML bodies that were stable over at least 30 min [17]. Occasionally, a telomere was observed to visit more than one PML body during this time, or PML was observed to accumulate at telomeric DNA. Supposing that all anchored telomeres are transcriptionally silenced, then perhaps as the authors discuss, the regulated release of telomeres from such constraints may allow for the activation of sub-telomeric genes. Interestingly, Everett et al. [19] previously reported a cell cycle-regulated association between centromeres and PML bodies. Together, these studies further support a possible role of specific chromosomal regions in anchoring chromosome territories within the cell nucleus.

DNA repair The process of double strand break (DSB) repair was recently studied in vivo using an elegant approach combining the introduction of the lac operator– repressor system with the incorporation of specific endonuclease cleavage sites into Saccharomyces cerevisiae chromosomes [20]. In S. cerevisiae, induction of DSB causes recruitment of Rad52-YFP from a diffuse nuclear distribution to subnuclear foci. The majority of spontaneous Rad52 foci were observed to form and dissipate rapidly within less than 10 min. To specifically tag DSB sites, a strain of yeast was engineered to contain a HO endonuclease cleavage site adjacent to a tandem array of lac operator sequences. Expression of YFP-lac repressor allowed the direct visualization of DSBs, and with the induction of HO endonuclease, 94% of Rad52-CFP foci colocalized with these sites [20]. Because many DSBs result in the formation of only a few Rad52 foci, it was hypothesized that damaged DNA moves from regions throughout the nucleus into a few repair centers. This idea was supported by results from strains containing a second DSB site labeled with red fluorescent protein (RFP). Both the YFP- and RFP-tagged DSB sites showed significant combined colocalization with a single Rad52 focus (48% of the cells) versus random colocalization (8% of cells) or localization of individual DSBs with different Rad52 foci (11% of the cells) [20]. This result


demonstrates that DSBs are recruited to a common nuclear region for repair, and it will be interesting to pursue further live observations of this process by tracking DSB localization over time during endonuclease induction. UV-induced DNA lesions are corrected in mammalian cells by two nucleotide excision repair (NER) mechanisms: transcription coupled NER (TCR) and global genome repair (GGR). The mechanism for detection of lesions could be either processive scanning of DNA or the stochastic diffusion-based recognition of lesions by repair factors, the latter supported by recent findings [21 –23]. The endonuclease for excision repair complementation group 1 –xeroderma pigmentosum group F (ERRC1 –XPF) as well as NER proteins XPA and TFIIH moves freely in the nucleus with different diffusion rates in the absence of DNA damage [21 – 23]. TFIIH is a multi-subunit dual function protein complex that operates in both NER and transcription initiation. Recently, the integral helicase XPB was tagged with GFP and used to study how TFIIH switches between sites of transcription and repair [22]. FRAP recovery curves in undamaged cells indicate that one population of TFIIH is freely diffusing, 20 – 40% is engaged in RNA pol II transcription initiation, and another fraction is engaged in RNA pol I transcription (see below). Upon UV irradiation, 40% of TFIIH was recruited to sites of UV damage within 2 min. In contrast to its short-term residence at transcription sites, FLIP results show that TFIIH resides at sites of DNA damage for about 4 min [22]. In two related studies, UV exposure immobilized ERCC1-GFP – XPF for about 4 min [21], and GFP-XPA, in an XPC-dependent manner, for 4– 6 min [23], the approximate time estimated to complete a single repair event. The kinetics of TFIIH-GFP in transcription in undamaged nuclear regions, including nucleoli, is unaffected by damage elsewhere that sequesters a significant amount (40%) of TFIIH [22]. Based upon their findings, the authors of these studies suggest that the exchange of TFIIH between transcription and NER occurs by random diffusion and collision, that it does not involve large-scale modifications, and that transcription and repair are not likely to be connected [21 –23]. Together, these results point toward diffusion-based lesion recognition by individual components or small subcomplexes as the primary mode of DNA repair, instead of the recruitment of large preassembled holoenzymes. One clear advantage of this model is that individual molecules and small subcomplexes would presumably have better access to condensed chromatin than large pre-assembled complexes. This model also allows greater flexibility in the use of individual members of a complex for different nuclear functions.

Kinetics of the gene expression machinery in living cells RNA polymerase I transcription The kinetics of assembly and elongation of RNA pol I on ribosomal genes was recently determined in living cells [24].


P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11

GFP-RNA pol I subunits and GFP-RNA pol I transcription factors localized to fibrillar centers (FCs) of nucleoli where they were subjected to photobleaching experiments. The preinitiation factors GFP-UBF1, GFP-UBF-2, and TAFI48GFP all showed rapid and complete fluorescence recovery (within 30– 35 s) after photobleaching (FRAP), as well as a rapid loss of fluorescence after inverse FRAP (iFRAP), suggesting that they associate transiently and continually exchange from the ribosomal genes. Consistent with this finding, Hoogstraten et al. [22] recently showed that nucleolar TFIIH-GFP completely recovered within 20– 30 s after bleaching, and its engagement in RNA pol I transcription (approximately 25 s) was much longer than in RNA pol II transcription (approximately 2 –10 s). In contrast to TFIIH and the above-mentioned preinitiation factors, RNA pol I subunits showed two recovery phases: an initial rapid recovery indicative of the transient association of a subset of RNA pol I molecules at initiation sites, followed by a slower recovery phase consistent with the association of actively elongating RNA pol I molecules on ribosomal genes [24]. This was further supported by iFRAP data, in which there was a rapid loss of fluorescence signal from FCs (transiently associated RNA pol I) followed by a continued decrease in fluorescence signal (elongating RNA pol I). Importantly, while the highly mobile fraction of RNA pol I was insensitive to a drug that terminates elongation, the dissociation of elongating RNA pol I is slowed further by the drug [24]. Based upon the recovery rates calculated in this study, elongating RNA pol I was estimated to complete synthesis of an rDNA gene in 140 s (95 nucleotides per second) with an estimated reinitiation rate of 1.4 s. Preinitiation factors have a residence time of 3 –5 s, suggesting they may be involved in several rounds of initiation. These recent studies are consistent with a previous study showing that proteins involved in different steps of rRNA biogenesis exhibit different kinetics in living cells [25]. Interestingly, kinetic studies now allow us to see individual steps within a single process such as transcription. Additionally, these data indicate that the preinitiation and elongation factors are not present in a common holoenzyme, rather, they reach the ribosomal genes independently and are subsequently assembled. RNA polymerase II transcription The first step in gene activation involves the transcription machinery accessing the chromatin template. One theory is that decondensed euchromatin allows access of this machinery, while condensed heterochromatin prevents transcription factors from reaching binding sites. However, heterochromatin-associated proteins show high turnover, indicating accessibility of heterochromatic nuclear regions [26,27]. Interestingly, Verschure et al. [28] recently showed in living nuclei that regions of condensed heterochromatin are accessible to fluorescently labeled dextrans and proteins of various sizes. Dextrans of 3 and 10 kDa showed reduced

accessibility to nucleoli, but were generally able to access all regions of the nucleus. Dextran (70 kDa) accessibility was more limited in some regions of condensed chromatin and in some regions completely lacking chromatin that probably correspond to nuclear bodies. However, a significant fraction of condensed chromatin domains were accessible to the 70kDa dextran and GFP-RNA polymerase II [28]. These results suggest that not all condensed chromatin is silent merely as a result of steric exclusion of the gene expression machinery, but that transcriptional status is more likely controlled by the chromatin remodeling machinery and the availability of properly modified binding sites such as proper histone-tail modifications (for review, see Ref. [29]). How much time does an active gene spend being transcribed? A study by Kimura et al. [30] focused on this question. Using CHO cells with a temperature-sensitive mutation in the large catalytic subunit of RNA pol II, they rescued transcription by expressing GFP-RNA pol II and studied its kinetics. Photobleaching experiments revealed two populations of RNA pol II. Approximately three-fourths of the RNA pol II showed fast recovery kinetics, representing free GFP-RNA pol II that can diffuse throughout the nucleoplasm and rapidly exchange on a gene promoter. The remaining one-fourth showed slow kinetics, representing GFP-RNA pol II engaged in elongating transcripts [30]. The time calculated to complete half a transcription cycle (i.e., the time it takes for half a gene to be transcribed) was surprisingly long—about 14 –20 min. From these data, it is estimated that a typical ‘‘active’’ gene would not be associated with multiple engaged polymerases; rather, it would only rarely be associated with more than one polymerase at a time [30]. Of course, a limitation of the methods used in this study is that large regions of nucleoplasm are photobleached, and one cannot be absolutely certain about the number and activity of genes or the types of other nuclear structures in the bleached regions. A way around this problem is to use cell lines containing easily visualized sites of RNA pol II transcription. McNally et al. [31] utilized a previously developed cell line containing a tandem array of a mouse mammary tumor virus (MMTV) promoter-driven ras to assess the exchange rate of GFP-RNA pol II at a specific transcription site [32]. While the glucocorticoid receptor (GR) and the GR coactivator GRIP – Tif-2 exchange rapidly on the locus (t1/2 approximately 5 s), initial recovery of RNA pol II occurs rapidly but is followed by a much slower exchange overall. The authors interpret these results to be consistent with the existence of two populations of RNA pol II, the fast one undergoing multiple unsuccessful initiation events, and the slower one engaged in transcription elongation that must be completed before recovery of fluorescence [32]. A system previously developed by Belmont et al. [33 – 35] to study large-scale chromatin decondensation during gene activation utilizes a CHO A03_1 cell line resulting from amplification, through methotrexate selection, of a 256 copy direct repeat of the lac operator sequence in conjunc-

P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11

tion with the DHFR gene. The 90 Mb amplified locus is inducible upon recruitment of the GFP-lac repressor-VP16 acidic activation domain (AAD) fusion protein, which leads to chromatin remodeling, large-scale chromatin decondensation, and transcriptional activation. Following VP16AAD targeting, endogenous HAT and SWI – SNF components were sequentially recruited to the locus [35]. TRRAP, a mammalian homolog of the yeast Tra1 protein and a component of several HAT complexes, was recruited very early to the activated locus (within 5 min). The HAT catalytic subunits GCN5 and PCAF, which preferentially acetylate histone H3, and p300 – CBP were slightly delayed, accumulating at the locus after 20– 30 min. The recruitment of GCN5, pCAF, and p300 –CBP correlated with an increase in histone H3 acetylation, while histone H4 acetylation occurred slightly later (35 – 40 min after gene activation). The surprising result was that TIP60, which preferentially acetylates histone H4, showed a strikingly delayed recruitment, as it reached the locus in 40% of cells after 45 –60 min and accumulated to steady state levels 3 –5 h after activation. Based upon these results, the authors speculate that early H4 acetylation may involve an unidentified HAT or could be a result of p300– CBP activity [35]. An interesting point is that the nature of a transcriptional activator targeted to a promoter can influence the degree of chromatin unwinding. Nye et al. [36] also used the A03_1 cell line to assess the chromatin unwinding ability of the estrogen receptor (ER)-lac repressor fusion protein at the locus described above. Chromatin unfolding was of the same magnitude, but was qualitatively different from that observed with GFP-lac repressor-VP16AAD. Neither ligand nor transcriptional activity was required for chromatin unfolding


activity of the ER targeted to chromatin in this manner. Additionally, competence of the receptor for transcription was not necessary for chromatin decondensation, as a dramatic unfolding of the locus was seen when the ER was recruited via lac repressor-SRC-1, which renders the complex transcriptionally inactive [36]. Experiments such as these have begun to define the stepwise in vivo recruitment of endogenous proteins to a gene locus, and they have allowed the direct visualization of chromatin behavior in living cells. To assess gene expression at the levels of DNA, RNA, and protein, Spector et al. [37,38] has developed a system in which a stably integrated array of transcription units and its RNA and protein products can be visualized simultaneously in living U2OS (2-6-3) cells. This system utilizes the lac repressor – operator interaction to visualize the chromatin, the tetracycline response elements, and activator protein to experimentally regulate transcription and the interaction of the MS2 bacteriophage coat protein with RNA stem loop structures to visualize the transcribed RNA. Additionally, the mRNA encodes for a CFP protein with a peroxisomal targeting signal, and its expression confirms that all posttranscriptional events have occurred properly (Fig. 1). One great advantage of this system is that the locus can be visualized directly in living cells both before and after activation, allowing for the simultaneous observation of chromatin unwinding and the recruitment of the transcription and chromatin-remodeling machineries [38]. In the condensed state, HP1a, h, and g, histone H3 tri-methylated at lysine 9 (H3 tri-meK9) and the histone H3 K9 methyltransferases, Suv39h1, and G9a-L were visualized at the locus. Loss of YFP-HP1a was first detected 30 min after transcriptional activation and levels progressively decreased

Fig. 1. Merged images of a U2OS 2-6-3 cell expressing CFP-lac repressor, MS2 coat-YFP fusion protein, and CFP-SKL. (A) In the ‘‘off’’ state, the chromatin at the locus is condensed, the MS2 binding protein is diffusely distributed throughout the nucleoplasm, and no cytoplasmic protein product is observed. (B) After induction of the transcription units, the locus becomes decondensed, the RNA binding protein is present at the transcription site as well as in particles in the nucleoplasm, and the protein product is observed in cytoplasmic peroxisomes. Scale bar = 10 Am. Images provided by Janicki et al. [38].


P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11

until it was undetectable at 180 min, suggesting that the HP1a-binding sites (histone H3 tri-MeK9) are lost during gene activation. Consistent with this possibility, H3 tri-Me K9 was not detected on an active locus. Interestingly, the histone variant H3.3 [39] became highly concentrated at the locus over a 75-min time period suggesting that histone exchange is part of the mechanism by which chromatin is converted from the inactive to the active state [38]. In the future, this system will prove useful for directly tracking newly synthesized mRNP particles in living cells and visualizing termination of transcription, recruitment of chromatin silencing complexes, and condensation of chromatin as the locus is inactivated.

Kinetics of Cajal body and nucleolar constituents The Cajal bodies Cajal bodies (CBs) are spherical nuclear bodies that contain a variety of components, including nucleolar proteins, snRNPs, coilin, and SMN (for a review, see Refs. [40 –42]). CBs have been implicated in snRNP assembly [43,44], snRNA metabolism [45], and posttranscriptional modification of newly assembled spliceosomal snRNAs [46,47]. They have also been reported to associate with specific gene loci [48,49]. Stable HeLa cell lines expressing either GFP-coilin [50] or GFP-SMN [51] have been used to track CB movements in living cells, which was described as anomalous diffusion [52]. Both studies showed similar types of movements by a subset of CBs in interphase nuclei, for example, the joining of two CBs, the separation of one CB into smaller CBs, and the movement of CBs within close proximity of each other. Stationary CBs were shown to associate with chromatin in an ATP-dependent manner [52]. It was recently reported that different CB components have different residence times in CBs. After photobleaching, YFP-coilin has a half time of fluorescence recovery of 40 s while GFP-SMN has a half time of recovery of more than 1 h in CBs, which could be due to the cytoplasmic localization of the bulk of the protein [51]. Bleaching of cytoplasmic GFP-SMN causes a rapid loss of GFP-SMN from CBs, suggesting a rapid transport of nuclear SMN to the cytosol. With respect to their localization, coilin and SMN also behave much differently during mitosis. Both are found in mitotic CBs in the cytoplasm until telophase, when YFPcoilin rapidly enters daughter nuclei [51]. In contrast, GFPSMN remains in the cytoplasm where it forms a large number of mitotic CBs before a small proportion of GFPSMN enters nuclear CBs in G1 and the majority remains diffuse in the cytoplasm [51]. Further experiments will elucidate how the observed differences in localization and turnover reflect differences in nuclear functions. The exchange of CB components between the nucleoplasm and CBs can also be observed in isolated germinal vesicles (GVs) of Xenopus oocytes [53]. One clear advantage

of using GVs versus mammalian nuclei is that GVs contain up to 100 CBs ranging in size from 2 to 10 Am in diameter, while mammalian nuclei contain only a few CBs, each with a diameter of less than 1 Am. In a study from the Gall laboratory [53], Xenopus oocytes were injected with fluorescent RNA or mRNAs encoding GFP-labeled CB components before GV isolation, allowing accumulation of fluorescently tagged components of interest in the GV nucleoplasm as well as incorporation into CBs. Using this method, GVs containing CBs or nucleoplasm labeled with fluorescein-U7 snRNA, GFP-TBP, and GFP-coilin were isolated and subjected to photobleaching experiments. Multiphasic recovery was reported for all three CB components, with approximate residence times of 14 s, 7.2 min, and 33 min for U7 and coilin, and 15 s, 3.3 min, and 34 min for TBP. Using computer modeling, the authors determined how the three kinetic states could be interconnected along a functional pathway, for example, as stages of macromolecular complex assembly. The data are consistent with the interpretation that the fast phase reflects exchange of RNA or protein between the nucleoplasm and CBs, and make little contribution to the FRAP dynamics [53]. Alternatively, the slower phase was interpreted to account for a significant percentage of the total, and it was suggested to reflect different states of complex formation (e.g., modification or assembly events) along a nonlinear pathway. The nucleolus The nucleolus is the site for rRNA transcription and ribosome assembly. Leary et al. [54] recently reported the nucleocytoplasmic shuttling dynamics and differential nucleolar localization of two major U3-snoRNA containing complexes, the core monoparticle (U3 snoRNA and box C/ D snoRNA-associated proteins) and the pre-ribosomal complex (the core complex and additional pre-ribosome associated proteins). GFP-tagged U3 complex-associated proteins, including fibrillarin, U3-55K, Imp3, Imp4, Mpp10, Rcl1, and Sof1, all shuttled between the nucleus and the cytoplasm in heterokaryon assays [54]. Interestingly, a range of shuttling kinetics among the different proteins suggests that they do not shuttle as a complex (Fig. 2). Additionally, U3 snoRNA and core proteins were enriched in CBs and in nucleolar foci that likely correspond to fibrillar structures, while pre-ribosome-associated proteins were not present in CBs but were enriched in nucleolar regions outside of the nucleolar foci (likely the granular component). Based upon their findings, the authors propose a model whereby U3 monoparticles assemble in the nucleoplasm or CBs, then translocate to the fibrillar components of the nucleolus where they associate at sites of rRNA transcription [54]. Pre-ribosomal associated proteins would then be recruited as the rRNA reaches the granular component, forming functional pre-ribosomes. After subsequent pre-rRNA processing and pre-ribosome assembly, dissociation may allow U3 complex-associated protein recycling in the next round of

P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11


Fig. 2. U3 snoRNP proteins shuttle between the nuclei and the cytoplasm at different rates. (A). Untransfected HeLa cells were fused in heterokaryon assays with HeLa cells transfected with constructs encoding either U3 snoRNP core proteins, GFP-fibrillarin or -U3-55K or the pre-ribosome-associated proteins GFP-MPP10, -Rcl1, or -Sof1. Heterokaryons were analyzed 2 h after fusion. DNA was labeled with DAPI to delineate nuclei. Arrows indicate untransfected nucleoli. Scale bar = 10 Am. (B) Quantitative analyses of the heterokaryon assays to compare the relative shuttling rates of the proteins analyzed in A. FR = fluorescence intensity in the untransfected cell nucleolus ( FU) / fluorescence intensity in the transfected cell nucleolus ( FT) within the same sized area. Images provided by Leary et al. [54].

processing, or it may subject these proteins to modifications involving shuttling between cellular compartments [54]. While much has been learned from studies using fixed cells to elucidate the pathways of rRNA synthesis and processing, little is known about rRNA trafficking in living cells. Ritland-Politz et al. [55] have directly observed endogenous rRNA movement in the nucleus. Complementary oligodeoxynucleotides labeled with caged fluorochromes were introduced into cultured rat myoblasts to follow the movement of endogenous 28S rRNA assembled into 60S ribosomal subunits. Caged fluorochromes do not fluoresce until caging groups (two o-nitrobenzyl groups covalently linked to fluorescein) are released by photolysis.

Upon uncaging of the fluorochrome at nucleolar sites, an average of 63% of the fluorescent signal moved away from the nucleolus in all directions within 30 s in a manner consistent with anomalous diffusion [55]. The fraction of signal remaining in nucleoli after uncaging could include ‘‘anchored’’ 28S rRNA in the process of being synthesized and packaged [55]. The results were similar when the experiment was performed at either 37jC or 23jC, suggesting the bulk of 60S ribosomal subunit movement is independent of metabolic energy. In addition, some of the signal uncaged in the nucleolus or the nucleoplasm sometimes visited other nucleoli, presumably in transit toward the nuclear pore complex.


P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11

Summary In recent years, our understanding of the nucleus has gained much from kinetic studies using GFP-tagged proteins. We have learned that contrary to our earlier images from fixed cells, the nucleus is a highly dynamic environment in which molecules move predominantly by diffusion, in some cases specifically by anomalous diffusion, as they temporarily associate with compartments or domains. Individual components of multi-subunit complexes in general exhibit different diffusion rates, implying a working model of the nucleus in which individual proteins or small subcomplexes involved in a particular process reach their destination(s) independently. Such a model would ultimately allow for multiple regulatory steps, especially for proteins that function in multiple nuclear processes. Acknowledgments The authors would like to thank Susan M. Janicki and Kannanganattu V. Prasanth for critical reading of the manuscript. Research in the Spector laboratory is funded by NIGMS/NIH 42694. References [1] B. Ludin, A. Matus, GFP illuminates the cytoskeleton, Trends Cell Biol. 8 (1998) 72 – 77. [2] J. Lippincott-Schwartz, T.H. Roberts, K. Hirschberg, Secretory protein trafficking and organelle dynamics in living cells, Annu. Rev.Cell Dev. Biol. 16 (2000) 557 – 589. [3] T. Misteli, J.F. Caceres, D.L. Spector, The dynamics of a pre-mRNA splicing factor in living cells, Nature 387 (1997) 523 – 527. [4] S.M. Gasser, Visualizing chromatin dynamics in interphase nuclei, Science 296 (2002) 1412 – 1416. [5] W.A. Bickmore, J.R. Chubb, Chromosome position: now, where was I? Curr. Biol. 13 (2003) R357 – R359. [6] A. Belmont, Dynamics of chromatin, proteins, and bodies within the cell nucleus, Curr. Opin. Cell Biol. 15 (2003) 304 – 310. [7] S.M. Janicki, D.L. Spector, Nuclear choreography: interpretations from living cells, Curr. Opin. Cell Biol. 15 (2003) 149 – 157. [8] L.A. Parada, J.J. Roix, T. Misteli, An uncertainty principle in chromosome positioning, Trends Cell Biol. 13 (2003) 393 – 396. [9] D. Zink, N. Sadoni, E. Stelzer, Visualizing chromatin and chromosomes in living cells, Methods 29 (2003) 42 – 50. [10] D. Gerlich, J. Beaudouin, M. Gebhard, J. Ellenberg, R. Eils, Fourdimensional imaging and quantitative reconstruction to analyse complex spatiotemporal processes in live cells, Nat. Cell Biol. 3 (2001) 852 – 855. [11] R.D. Moir, T.P. Spann, R.I. Lopez-Soler, M. Yoon, A.E. Goldman, S. Khuon, R.D. Goldman, The dynamics of the nuclear lamins during the cell cycle—Relationship between structure and function, J. Struct. Biol. 129 (2000) 324 – 334. [12] S.K. Lyman, L. Gerace, Nuclear pore complexes: dynamics in unexpected places, J. Cell Biol. 154 (2001) 17 – 20. [13] R.W. Dirks, C. Molenaar, H.J. Tanke, Visualizing RNA molecules inside the nucleus of living cells, Methods 29 (2003) 51 – 57. [14] R. Eils, D. Gerlich, W. Tvarusko, D.L. Spector, T. Misteli, Quantitative imaging of pre-mRNA splicing factors in living cells, Mol. Biol. Cell 11 (2000) 413 – 418.

[15] J.R. Swedlow, I. Goldberg, E. Brauner, P.K. Sorger, Informatics and quantitative analysis in biological imaging, Science 300 (2003) 100 – 102. [16] F. Hediger, F.R. Neumann, G. Van Houwe, K. Dubrana, S.M. Gasser, Live imaging of Telomeres. yKu and Sir proteins define redundant telomere-anchoring pathways in yeast, Curr. Biol. 12 (2002) 2076 – 2089. [17] C. Molenaar, K. Wiesmeijer, N.P. Verwoerd, S. Khazen, R. Eils, H.J. Tanke, R.W. Dirks, Visualizing telomere dynamics in living mammalian cells using PNA probes, EMBO J. 22 (2003) 6631 – 6641. [18] P. Heun, T. Laroche, K. Shimada, P. Furrer, S.M. Gasser, Chromosome dynamics in the yeast interphase nucleus, Science 294 (2001) 2181 – 2186. [19] R. Everett, W. Earnshaw, A. Pluta, T. Sternsdorf, A. Ainsztein, M. Carmena, S. Ruchaud, W. Hsu, A. Orr, A dynamic connection between centromeres and ND10 proteins, J. Cell. Sci. 112 (1999) 3443 – 3454. [20] M. Lisby, U.H. Mortensen, R. Rothstein, Colocalization of multiple DNA double-strand breaks at a single Rad52 repair centre, Nat. Cell Biol. 5 (2003) 572 – 577. [21] A.B. Houtsmuller, S. Rademakers, A.L. Nigg, D. Hoogstraten, J.H.J. Hoeijmakers, W. Vermeulen, Action of DNA repair endonuclease ERCC1/XPF in living cells, Science 284 (1999) 958 – 961. [22] D. Hoogstraten, A.L. Nigg, H. Heath, L.H. Mullenders, R. van Driel, J.H. Hoeijmakers, W. Vermeulen, A.B. Houtsmuller, Rapid switching of TFIIH between RNA polymerase I and II transcription and DNA repair in vivo, Mol. Cell 10 (2002) 1163 – 1174. [23] S. Rademakers, M. Volker, D. Hoogstraten, A.L. Nigg, M.J. Mone, A.A. Van Zeeland, J.H. Hoeijmakers, A.B. Houtsmuller, W. Vermeulen, Xeroderma pigmentosum group A protein loads as a separate factor onto DNA lesions, Mol. Cell. Biol. 23 (2003) 5755 – 5767. [24] M. Dundr, U. Hoffmann-Rohrer, Q. Hu, I. Grummt, L.I. Rothblum, R.D. Phair, T. Misteli, A kinetic framework for a mammalian RNA polymerase in vivo, Science 298 (2002) 1623 – 1626. [25] D. Chen, S. Huang, Nucleolar components involved in ribosome biogenesis cycle between the nucleolus and nucleoplasm in interphase cells, J. Cell Biol. 153 (2001) 169 – 176. [26] T. Cheutin, A.J. McNairn, T. Jenuwein, D.M. Gilbert, P.B. Singh, T. Misteli, Maintenance of stable heterochromatin domains by dynamic HP1 binding, Science 299 (2003) 721 – 725. [27] R. Festenstein, S.N. Pagakis, K. Hiragami, D. Lyon, A. Verreault, B. Sekkali, D. Kioussis, Modulation of heterochromatin protein 1 dynamics in primary mammalian cells, Science 299 (2003) 719 – 721. [28] P.J. Verschure, I. van der Kraan, E.M. Manders, D. Hoogstraten, A.B. Houtsmuller, R. van Driel, Condensed chromatin domains in the mammalian nucleus are accessible to large macromolecules, EMBO Rep. 4 (2003) 861 – 866. [29] B.D. Strahl, C.D. Allis, The language of covalent histone modifications, Nature 403 (2000) 41 – 45. [30] H. Kimura, K. Sugaya, P.R. Cook, The transcription cycle of RNA polymerase II in living cells, J. Cell Biol. 159 (2002) 777 – 782. [31] J.G. McNally, W.G. Muller, D. Walker, R. Wolford, G.L. Hager, The glucocorticoid receptor: rapid exchange with regulatory sites in living cells, Science 287 (2000) 1262 – 1265. [32] M. Becker, C. Baumann, S. John, D.A. Walker, M. Vigneron, J.G. McNally, G.L. Hager, Dynamic behavior of transcription factors on a natural promoter in living cells, EMBO Rep. 3 (2002) 1188 – 1194. [33] C.C. Robinett, A. Straight, G. Li, C. Willhelm, G. Sudlow, A. Murray, A.S. Belmont, In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition, J. Cell Biol. 135 (1996) 1685 – 1700. [34] T. Tumbar, G. Sudlow, A.S. Belmont, Large-scale chromatin unfolding and remodeling induced by VP16 acidic activation domain, J. Cell Biol. 145 (1999) 1341 – 1354.

P.A. Bubulya, D.L. Spector / Experimental Cell Research 296 (2004) 4–11 [35] S. Memedula, A.S. Belmont, Sequential recruitment of HAT and SWI/ SNF components to condensed chromatin by VP16, Curr. Biol. 13 (2003) 241 – 246. [36] A.C. Nye, R.R. Rajendran, D.L. Stenoien, M.A. Mancini, B.S. Katzenellenbogen, A.S. Belmont, Alteration of large-scale chromatin structure by estrogen receptor, Mol. Cell. Biol. 22 (2002) 3437 – 3449. [37] T. Tsukamoto, N. Hashiguchi, S.M. Janicki, T. Tumbar, A.S. Belmont, D.L. Spector, Visualization of gene activity in living cells, Nat. Cell Biol. 2 (2000) 871 – 878. [38] S.M. Janicki, T. Tsukamoto, S.E. Salghetti, W.P. Tansey, R. Sachidanandam, K.V. Prasanth, T. Ried, Y. Shav-Tal, E. Bertrand, R.H. Singer, D.L. Spector, From silencing to gene expression: realtime analysis in single cells, Cell 116 (2004) 683 – 698. [39] K. Ahmad, S. Henikoff, The histone variant H3.3 marks active chromatin by replication-independent nucleosome assembly, Mol. Cell 9 (2002) 1191 – 1200. [40] A.G. Matera, Nuclear bodies: multifaceted subdomains of the interchromatin space, Trends Cell Biol. 9 (1999) 302 – 309. [41] S.C. Ogg, A.I. Lamond, Cajal bodies and coilin-moving towards function, J. Cell Biol. 159 (2002) 17 – 21. [42] J.G. Gall, The centennial of the Cajal body, Nat. Rev., Mol. Cell Biol. 4 (2003) 975 – 980. [43] J.E. Sleeman, A.I. Lamond, Newly assembled snRNPs associate with coiled bodies before speckles, suggesting a nuclear snRNP maturation pathway, Curr. Biol. 9 (1999) 1065 – 1074. [44] D. Stanı¨k, S.D. Rader, M. Klingauf, K.M. Neugebauer, Targeting of U4/U6 small nuclear RNP assembly factor SART3/p110 to Cajal bodies, J. Cell Biol. 160 (2003) 505 – 516. [45] C. Verheggen, D.L. Lafontaine, D. Samarsky, J. Mouaikel, J.M. Blanchard, R. Bordonne, E. Bertrand, Mammalian and yeast U3 snoRNPs are matured in specific and related nuclear compartments, EMBO J. 21 (2002) 2736 – 2745. [46] X. Darzacq, B.E. Jady, C. Verheggen, A.M. Kiss, E. Bertrand, T. Kiss, Cajal body-specific small nuclear RNAs: a novel class of 2V-O-meth-











ylation and pseudouridylation guide RNAs, EMBO J. 21 (2002) 2746 – 2756. B.E. Jady, X. Darzacq, K.E. Tucker, A.G. Matera, E. Bertrand, T. Kiss, Modification of Sm small nuclear RNAs occurs in the nucleoplasmic Cajal body following import from the cytoplasm, EMBO J. 22 (2003) 1878 – 1888. E.Y. Jacobs, M.R. Frey, W. Wu, T.C. Ingledue, T.C. Gebuhr, L. Gao, W.F. Marzluff, A.G. Matera, Coiled bodies preferentially associate with U4, U11, and U12 small nuclear RNA genes in interphase HeLa cells but not with U6 and U7 genes, Mol. Biol. Cell 10 (1999) 1653 – 1663. L.S. Shopland, M. Byron, J.L. Stein, J.B. Lian, G.S. Stein, J.B. Lawrence, Replication-dependent histone gene expression is related to Cajal body (CB) association but does not require sustained CB contact, Mol. Biol. Cell 12 (2001) 565 – 576. M. Platani, I. Goldberg, J.R. Swedlow, A.I. Lamond, In vivo analysis of Cajal body movement, separation, and joining in live human cells, J. Cell Biol. 151 (2000) 1561 – 1574. J.E. Sleeman, L. Trinkle-Mulcahy, A.R. Prescott, S.C. Ogg, A.I. Lamond, Cajal body proteins SMN and Coilin show differential dynamic behaviour in vivo, J. Cell. Sci. 116 (2003) 2039 – 2050. M. Platani, I. Goldberg, A.I. Lamond, J.R. Swedlow, Cajal body dynamics and association with chromatin are ATP-dependent, Nat. Cell Biol. 4 (2002) 502 – 508. K.E. Handwerger, C. Murphy, J.G. Gall, Steady-state dynamics of Cajal body components in the Xenopus germinal vesicle, J. Cell Biol. 160 (2003) 495 – 504. D.J. Leary, M.P. Terns, S. Huang, Components of U3 snoRNA-containing complexes shuttle between nuclei and the cytoplasm and differentially localize in nucleoli: implications for assembly and function, Mol. Biol. Cell 15 (2004) 281 – 293. J.C. Ritland-Politz, R.A. Tuft, T. Pederson, Diffusion-based transport of nascent ribosomes in the nucleus, Mol. Biol. Cell 14 (2003) 4805 – 4812.