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Visualizing chromosome dynamics with GFP
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Visualizing chromosome dynamics with GFP
GFP applications
Andrew S. Belmont By allowing easy labeling of chromosomal and nuclear proteins and the tagging of specific chromosomal regions, the use of green-fluorescent protein (GFP) has provided new and special opportunities for directly observing chromosome dynamics in vivo. Here, we review recent applications of this methodology, focusing particularly on examples where new biology has been learned, or at least sighted. In particular, we focus on active bacterial chromosome segregation, yeast mitosis and centromere dynamics, and largescale chromatin structure and dynamics within eukaryotic interphase nuclei.
Andrew S. Belmont Dept of Cell and Structural Biology, University of Illinois, UrbanaChampaign B107 CLSL, 601 South Goodwin Ave., Urbana, IL 61801, USA. e-mail: [email protected]
More than 100 years ago, the dynamic behavior of mitotic chromosomes captured the interest of early microscopists. Identification of chromosomes as the basic units of heredity near the turn of the twentieth century pushed mitotic and meiotic chromosome segregation to center stage, where it has remained ever since as a major cell-biological and genetics research focus. Using transmitted light microscopy without any special staining, mitotic chromosome segregation can be observed directly in living cells over a time scale of seconds to minutes. The ability to visualize this dynamic process has been crucial to dissecting the molecular mechanisms underlying chromosome segregation. Conversely, the inability to easily visualize chromosomes in various organisms has led to serious scientific roadblocks. With regard to chromosome segregation, the inability to visualize bacterial chromosomes easily has meant that progress in understanding this process in prokaryotic cells has remained a ‘black box’for many decades. Even in eukaryotic organisms, the inability to easily visualize mitotic chromosomes, for instance in budding and fission yeast, has hindered the full exploitation of the powerful genetic tools available for these model organisms. More significantly, although it has been possible to visualize with relatively simple methods the segregation in eukaryotic cells of large, condensed mitotic chromosomes, visualization of decondensed, interphase chromosomes has remained technically challenging in all organisms. Basic questions of interphase chromosome structure as a result have relied largely on static images from fixed material and have been plagued by concerns over specimen preparation artifacts induced by fixation, immunostaining and in situ hybridization procedures. More interesting questions relating interphase chromosome structure and dynamics to regulation of transcription, replication, recombination and DNA repair have been difficult to approach owing to this key technical limitation.
In fact, it is not too great a stretch to conclude that this inability to visualize easily chromosome regions within interphase nuclei has contributed greatly to the relative backwardness of the cell biology of many aspects of nuclear structure and function. The introduction of fluorescently labeled nucleotides and proteins into cells has enabled direct observation of in vivo mitotic and interphase chromosome dynamics1. This approach has now been greatly enhanced by the recent applications of greenfluorescent protein (GFP). Not only can specific nuclear proteins be labeled with GFP, but, by using direct repeats of bacterial operator sequences, it is possible to tag specific chromosome sites and visualize these tags using GFP fused with the appropriate repressor protein (Fig. 1)2,3. This approach was first developed with the lac operator–repressor system in mammalian cells4 and yeast5 and then transferred rapidly to bacteria6, Caenorhabditis elegans (A. Serrichio and P. Sternberg, pers. commun.)7 and more recently to Drosophila (A. Belmont et al., unpublished). A similar approach using the tet operator–repressor system has also been highly successful8. Here, we review several recent examples of the scientific impact derived from these new experimental capabilities. Chromosome dynamics in bacteria
In contrast to active segregation of eukaryotic mitotic chromosomes, the textbook picture of bacterial chromosome segregation for several decades was based on the Jacob replicon model in which daughter chromatids attached to centrally located regions on the cell membrane, with passive separation being achieved through cell-wall growth during cell-cycle progression9. This textbook picture now has been supplanted by active chromosome segregation models, based on recent in vivo observations combined with traditional immunofluorescence and fluorescence in situ hybridization (FISH) approaches10,11. Using either GFP fusions to endogenous proteins binding near the chromosome origin region or GFP fused with bacterial repressors binding to operator direct repeats introduced at specific chromosome locations, in vivo dynamics of chromosome origins and other chromosome regions have been visualized. A defined spatial layout of the bacterial chromosome relative to the bacterial cell has been observed. In sporulating Bacillus subtilis, origins were localized at the cell poles, with termini near the
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Fig. 1. Tagging chromosome sites with the lac operator–repressor system3. (a) Detection: a direct repeat of the lac operator can be identified by the in vivo or in vitro binding of lac repressor protein. Originally a 256 copy, 10-kb direct repeat was made, but smaller repeats can also be visualized using this method. In Drosophila, a direct repeat of less than 1 kb has been visualized in diploid nuclei (P. Alvarez and A. Belmont, unpublished). The lac repressor has been engineered to contain a nuclear localization signal (NLS) and the C-terminus has been mutated to prevent lac repressor dimers from forming tetramers, which can bind to two different operators. (b) Transfection: direct repeats of the lac operators can be subcloned into transformation vectors and stable transformants generated by homologous recombination (to date, yeast and bacteria), transposable elements (Drosophila) or nonhomologous recombination (Caenorhabditis elegans, mammalian cells). Here, we show introduction of the lac operator direct repeat into an expression vector for dihydrofolate reductase (DHFR). DHFR serves as the selectable marker for selection of stable transformants in DHFR– Chinese hamster ovary (CHO) cells. Cells expressing a GFP–lac-repressor–NLS fusion protein show small dots of staining where the lac operator containing vector has integrated. A general nuclear background is present owing to nuclear import of the NLS-containing fusion protein. (c) Gene amplification: owing to the high degree of chromatin compaction, even multicopy vector insertions, hundreds of kb in size, appear as small dots in the mammalian cell nucleus by light microscopy. To visualize larger chromosome regions, gene amplification was used to create amplified chromosome regions, ranging in size from chromosome bands to whole chromosome arms. Multiple selections using increasing concentrations of methotrexate, an inhibitor of DHFR, were used to isolate cell lines that have undergone gene amplification. In these cell lines, continual fibers or a series of dots are seen, depending on the size of the coamplified genomic DNA.
cell middle6. Vegetatively growing cells showed a bipolar origin distribution after origin replication, with termini towards the middle and with a preference for polar origin localization prior to replication6,12,13. Labeling origins, termini and quarter positions (90° and 270°) demonstrated in general maximum separation between origin and terminus, with quarter positions lying in between14. E. coli showed similar bipolar distribution of origins and a central location of termini, but, by contrast, http://tcb.trends.com
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both F and P1 plasmids localized to the cell center prior to replication and moved rapidly to the quarter cell positions after replication15. The bacterial chromosome and each of the two plasmids all appear to use different overall mechanisms for active chromosome segregation. FISH results and immunolocalization using proteins mapping to the chromosome origin produced similar results10. Time-lapse imaging on large numbers of B. subtilis cells revealed that approximately two-thirds of origins moved from the pole towards the cell center prior to separation into two origins, with origin separation occurring near the pole in the remaining cells. Strikingly, the majority of movement during separation of newly replicated origins occurred during a short portion of the total cell cycle, approximately 11 minutes out of a 100-minute doubling time, with peak origin velocity during the first 3–6 minutes of origin separation16. Chromosome termini, as well as chromosome quarter positions (90° and 270°), also were observed to separate rapidly, thus demonstrating an active mechanism for partitioning of bacterial chromosomes16. Similarly rapid and abrupt origin partitioning movements were observed using a GFP fusion with SpoOJ, which binds near to the B. subtilis origin14. The recent demonstration that DNA polymerase is located at a discrete spot near the cell center means that these active chromosome partitioning mechanisms must be coupled in some way with spatially localized replication17. By stalling DNA replication at a specific, labeled chromosome site ~100 kb from the replication origin, followed by release of this block, movement away from the replication machinery to the cell poles was observed for newly replicated foci18. A labeled chromosome region lying approximately halfway between origin and terminus moved to the centrally located replication focus prior to duplication and separation. Forces generated by DNA polymerase, therefore, might contribute to chromosome segregation. Fig. 2 shows the spatial arrangement of replisomes containing DNA polymerase III (orange) and labeled chromosomal origins (blue) at different times after cell synchronization and induction and then release of the replication block. Chromosome origins localize to the replisome (d–f) during the replication block but move quickly towards the poles after release of this replication block (g–i). Mitosis in cytologically challenged but genetically blessed eukaryotes
Direct observation of chromosome dynamics in higher eukaryotic cells has been crucial in formulating mechanistic models of mitotic chromosome segregation. The ability to test these models directly, however, and to identify the underlying molecular components that execute these mechanisms has been limited by the functional redundancy for key gene products frequently observed in these organisms. This is compounded by the experimental difficulties in knocking out these gene products, and the relative
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Fig. 2. Movement of newly replicated chromosome origins away from a centrally located replisome. Bacillus subtilis cells were observed at different times after synchronization of the replication cycle. Synchronization was achieved using a temperature-sensitive allele of dnaB134, required for DNA replication initiation. Growth at the nonpermissive temperature allows ongoing rounds of replication to finish while preventing initiation of new rounds of replication. Release of this replication initiation block was followed by a stringent replication arrest, using amino acid starvation, which arrests replication forks at defined chromosome loci, LSTer and RSTer, located ~100–130 kb and ~150–200 kb, respectively, from the origin. The replisome was labeled using a subunit of DNA polymerase III – dnaX – fused with YFP. LSTer was labeled by the nearby insertion of a lac operator direct repeat, binding to lac-repressor–CFP. (a–c) Sixty minutes after synchronization by a block in DNA replication initiation. Two copies of LSTer are located towards the cell poles, away from the centrally located replisome. (d–f) Sixty minutes after synchronization and induction of the replication arrest at LSTer. LSTer is coincident with the replisome. (g–i) Thirty minutes after release from the replication arrest, two copies of LSTer are located away from the replisome. (j–l) A cell undergoing (or about to undergo) multifork replication 30 min after release from stringent replication arrest. The central polymerase is from the initial round of replication, and additional replisomes have assembled at the cell quarter positions. (m–o) Replisomes approximately at the cell quarters in a cell that seems to have finished the initial replication round (no replisome at midcell). Left panels: foci of DNA polymerase (orange) and cell outline (gray). Middle panels: foci of LSTer (cyan) and cell outline (gray). Right panels: merged images from left and middle panels. Bar, 1 µm. (Images reprinted, with permission, from Ref. 18.)
difficulty in identifying novel proteins involved in these processes. By contrast, in genetically tractable organisms such as budding and fission yeast, relatively simple screens have identified a large number of genes involved in chromosome segregation during mitosis and meiosis, but the poor cytology in these organisms has seriously hindered identification of the functional role of each of these gene products. http://tcb.trends.com
Using GFP, it has now been possible to visualize directly in vivo chromosome dynamics during mitosis in budding yeast using marked chromosomal sites5. Chromosome movements relative to the microtubule spindle structure were observed in cells coexpressing GFP–lac-repressor and GFP-α-tubulin19. Budding yeast did not appear to show a conventional metaphase plate alignment; instead chromosomes oscillated along the entire length of the spindle, with chromosomes frequently near one spindle pole at the onset of anaphase chromosome separation. However, a recent study has shown that DNA within several kilobases of the centromere is located within the central 25% of the spindle in the majority of nuclei20, suggesting a time-averaged, metaphase-like alignment for centromere regions but not for chromosome arms. A rapid, polewards chromosome movement during an anaphase A stage was observed, with centromeres leading and telomeres lagging, and with transient chromosome stretching between centromere and telomere19. Comparison of in vivo mitotic chromosome motion in wild-type versus mutant yeast strains revealed unique roles during mitosis for different kinesin-like motors, with Cin8p implicated in the rapid phase of anaphase B, and Kip1p in the slow phase of anaphase B21. Similar mitotic events were observed in fission yeast, with an extensive period during which chromosomes oscillate between poles prior to onset of sister chromatid separation and anaphase A and B movements22. Transient splitting of centromeres was also observed, up to 0.6 µm in distance, just prior to the onset of anaphase chromosome separation, suggesting that, during metaphase, the centromere was positioned in a bioriented manner towards the poles and was under tension. Reinvestigation of centromere dynamics in budding yeast using chromosomes tagged just several kilobases from the actual centromere region demonstrated a similar ‘precocious’ separation of centromere regions early during spindle formation and probably soon after centromere duplication23,24. Similar early splitting of fluorescence signal was observed for GFP-labeled kinetochore proteins bound to wild-type centromeres23. Time-lapse three-dimensional microscopy demonstrated that centromere separation was transient, with dramatic, elastic stretching of centromere regions and movement towards the poles, followed by periods when daughter centromeres rejoin20,24,25. This pre-anaphase stretching occurs predominately in DNA regions within ~9–13 kb flanking the actual centromere sequence and lowers compaction ratios to ~10–20, below the 30–40 ratio expected for a 30-nm fiber and the overall 90–120-fold compaction ratio seen for yeast chromosomes24. Anaphase stretching to below that of nucleosome filaments was observed transiently for the tet-operator-labeled insert DNA during anaphase, but it remains unclear whether
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this extreme stretching is a property of the tet op repeat DNA only or would also be observed for wild-type sequence24. Recent data suggest similar stretching, however, for ~1 kb of DNA between the centromere and a 1.7 kb lac operator repeat20. The existence of this precocious sister centromere separation is consistent with the establishment of properly bioriented sister kinetochores through a tension-generating and tension-sensing mechanism, as established for higher eukaryotic cells where an approximately twofold stretching of GFP–CENP-Blabeled centromeres has been observed26. The magnitude of this stretching and separation in yeast, however, is surprising, with the disruption of chromatin packing to the level of a nucleosome filament and the 5–10-fold stretching much larger proportionally than seen for mammalian centromeres. Precocious separation of sister centromeres does not require Esp1p, a protein required for cohesin degradation. Therefore, separation is independent of cohesin proteolysis25, although prevention of premature daughter chromosome separation requires the cohesin component Scc1p, and therefore cohesin function, to hold the distal chromosome arms together during periods of centromere splitting24. These observations suggest that cohesin complexes near centromeres might separate transiently during centromere splitting. Because splitting falls off sharply beyond a certain distance from the core centromere, this has further supported the possibility of specialized, elastic chromatin regions flanking centromeres25. Comparison of the velocities of centromere-proximal and centromere-distal labeled chromosome sites during anaphase suggests an elastic recoil accompanying separation of sister chromatids, increasing the initial rate of sister chromosome separation for centromere-distal regions20. Interphase chromosomes on the move
As accustomed as we are to dynamic chromosome movements during mitosis, our natural inclination is to view the interphase nucleus and its contents as static. Previously, chromosomes and subnuclear organelles have been difficult to visualize in fixed material, let alone live cells. Moreover, the overall shape of nuclei and their visible contents, such as nucleoli, have not shown rapid movements. Using FISH, distinct chromosome ‘territories’ have been observed. The solid appearance of these chromosome ‘paints’ in some studies has led to models in which chromatin packing within territories would limit diffusion of macromolecular complexes27. Similarly, close contact between different chromosome territories has left the impression that the majority of the nuclear volume might be occupied by chromatin, directly constraining movement of chromosomes themselves. In fact, measurements of dextran intranuclear diffusion in vertebrate tissue-culture cells indicated http://tcb.trends.com
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that no more than 15% of the nuclear volume is inaccessible to small molecules28,29, and electron microscopy has revealed large spaces between largescale chromatin domains30. Stereotypical cell-cycle changes in nucleolar distribution have been described31, as well as large movements of coiled bodies, demonstrated through use of GFP fusion proteins32. A combination of immunostaining and FISH has demonstrated nonrandom localization of DNA sequences in the nucleus; these include associations with the nuclear envelope, nucleoli or heterochromatic blocks (reviewed in Ref. 33). Changes in these nonrandom distributions during the cell cycle, differentiation and changes in gene expression imply intranuclear chromosome movements occurring over a time frame no greater than several hours. Recent data examining the differential localization of a gene-rich human chromosome versus a gene-poor human chromosome by FISH have suggested that changes in interphase chromosomal location of entire chromosomes might occur relatively rapidly34. These studies, however, reveal only steady-state statistical distributions, without revealing actual dynamics. Using fluorescence recovery after photobleaching (FRAP), little or no relative motion was observed in vivo in mammalian cells over several hours for bleached chromosome regions, stained with ethidium bromide and ~1 µm in diameter35. Similarly, in vivo dynamics of chromosome foci labeled by fluorescent nucleotides in general showed little or no motion over periods of several hours, particularly after adjustments for nuclear rotation and spatial translation36. Likewise, centromeres labeled with a GFP–CENP-B fusion protein in mammalian tissueculture cells also in general showed little motion26. Together, these studies examining the aggregate behavior of large chromosome regions within mammalian tissue-culture cells unsynchronized in the cell cycle indicated that long-range motion of interphase chromosomal loci, if present, must be infrequent and confined to a small subset of chromosomal loci and/or cell-cycle stages. These studies also did not address whether rapid, but locally constrained, chromosome movements were occurring. A careful examination of yeast chromosomes labeled near centromeres, however, showed that, over seconds to tens of seconds, there was in fact rapid but localized motion that was best described as diffusion within a constrained radius of ~0.3 µm (Ref. 37). Rapid, localized motion in mammalian cell nuclei on a time scale of seconds or less also occurs within mammalian cells (A.S. Belmont, unpublished). More extended observations, however, have revealed infrequent examples in mammalian cells of sustained, unidirectional motion of centromeres26 and chromosome replication domains labeled by fluorescent nucleotides36. Using artificial chromosome constructs labeled with lac operator repeats, three examples of specific movements now have been
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Fig. 3. Direct observation of a translocation over several microns of a ~90-Mbp heterochromatic, gene-amplified chromosome arm from the nuclear periphery to the nucleolus and back. (a–f) Images represent combination of transmitted- and fluorescence-light images at specific time points, measured from beginning of observation time (t = 0), 4 h after release from an early S-phase block. (a–f) correspond to t = 1, 5, 5.5, 6, 9 and 9.5 hrs, respectively. Movement of the chromosome arm from the nuclear periphery to nucleolus is coupled with chromosome decondensation. The chromosome arm returns to the nuclear periphery in the vicinity of the original starting position. Arrows point to the edge of the nucleus. Bar, 2 µm. (G. Li and A. Belmont, unpublished.)
observed in mammalian cells. A late-replicating, amplified chromosome region, usually located adjacent to the nuclear envelope in G1 and the first half of S phase, was observed to move to the nuclear interior 4–6 hours into S phase. This movement was
tightly correlated with the onset of DNA replication in this chromosome region38. Both the timing of this movement and its correlation with DNA replication were established by statistical analysis of a large number of cells using fixed preparations. An example in which this motion has been observed directly in living cells is shown in Fig. 3. Cells were synchronized in early S phase. Visualization began four hours after release from the early S phase. A motion from the nuclear envelope to the nucleolus and back to the nuclear envelope occurs. A cell clone containing an insertion of 10–20 vector copies containing lac operator inserts showed a single, GFP–lac repressor spot in the nucleus, and this had a high frequency of association with the nuclear envelope during G1 phase. Movement of this spot to the nuclear interior was observed at a distinct stage in early S phase39. Direct in vivo visualization showed that, in the absence of VP16, in most cells the spot frequently moved transiently into the nuclear interior during early G1 but relocated back to the nuclear periphery and remained there until early S phase (Fig. 4, bottom left panel). Targeting the acidic activation domain of VP16 to this spot activated transcription and also induced a permanent relocalization of the spot to the nuclear interior that was established soon after mitosis (Fig. 4, top left panel). This differential nuclear positioning of this chromosome site occurs as a function of cell-cycle progression and the presence or absence of the VP16 activation domain in Fig. 4, right panel. Control
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Fig. 4. Interphase positioning of a DNA chromosome region is established early after mitosis and is modulated by VP16 transcriptional activator39. Left: A specific chromosome site, normally located preferentially adjacent to the nuclear periphery (bottom panel), instead localizes to the nuclear interior when the VP16 acidic activation domain (AAD) is tethered to this site (top panel). Top panel shows example from C4-VP16 cells expressing a GFP–lac-repressor–VP16-AAD fusion protein targeted to a chromosome site (bright spot) containing lac operator repeats. Number in left, bottom corner refers to minutes after cell division. The chromosome site localizes away from nuclear edge early in G1 phase. Bottom panel shows control cells expressing a GFP–lacrepressor protein. The site starts at the nuclear periphery, extends
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linearly into the nuclear interior transiently, then returns as a condensed spot to the nuclear edge. This transient stretching or extension was observed in a significant fraction of cells and was largely restricted to this G1 period. Bar, 5 µm. Image reprinted, with permission, from Ref. 39. Right: summary of cell-cycle changes in nuclear position for a chromosome site containing the lac operator in control cells expressing GFP–lac repressor (left) versus cells expressing the GFP–lacrepressor–VP16-AAD fusion protein (right). Control cells show a peripheral spot localization that moves to the interior early in S phase. Cells expressing the GFP–lac-repressor–VP16 fusion protein show an interior localization established early in G1. Abbreviation: GFP, greenfluorescent protein. (Drawing kindly provided by Tudorita Tumbar.)
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In Drosophila embryos, a higher rate of mobility has been observed over short times, using fluorescently labeled topoisomerase 2 to label a specific satellite repeat, with a much larger radius of constrained motion37. In imaginal disks (J. Radasta and A.S. Belmont, unpublished) and in spermatocytes (J. Vazquez et al., pers. commun.), a much larger range of mobility is observed, over a time scale of minutes, as compared with mammalian cells. During male meiosis I in late G2, this long-range mobility ceases and is replaced by a localized constrained diffusion (J. Vazquez and J. Sedat, pers. commun.). In contrast to the constrained diffusion observed previously near yeast centromeres37, a high degree of mobility was observed for a number of tagged chromosomal sites in yeast. Using FISH, latereplicating origins distant from the yeast telomere were shown to be enriched during G1 in a zone adjacent to the nuclear periphery40. This was in contrast to early-replicating chromosomal origins, which were distributed randomly throughout the nucleus. Live observation, however, revealed large movements (~0.5 µm) over a distance of approximately one-half the radius of the cell nucleus; these occurred during intervals of several seconds approximately once per minute during G1 phase for early-replicating chromosome regions40. Fig. 5 shows an example in which an early-replicating chromosome region moves from near the nuclear center to the periphery in a six-second period. Similar, but somewhat less frequent, movements during G1 phase were seen for the late-replicating origin cluster examined, including movements well into the nuclear interior (S.M. Gasser, pers. commun.). In an interesting parallel with the previously described results in mammalian cells, the late-replicating (a)
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Fig. 5. High mobility of budding yeast interphase chromosomes. Shown are six frames (a–f) of a haploid yeast cell in G1 phase, in which an early-firing autonomously replicating sequence (ARS) on Chr IV (ARS Chr 4-908) is tagged with multiple lac operator repeats, creating a bright focus of green-fluorescent protein (GFP) fluorescence, while the nuclear envelope is visualized with NUP49-GFP40. The images were taken on the Zeiss LSM 510 confocal with a 100× objective. They are time-lapse images, with 1.22 seconds between each image. From the first to the last image, 6.12 seconds have elapsed, and the bright spot representing the origin has moved from the center of the nucleus to the periphery. The nucleus is ~1.9 µm in diameter. (Photomicrograph kindly provided by Susan M. Gasser.)
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origin cluster on chromosome 14 lost its peripherallocalization bias during S phase, becoming distributed throughout the nuclear interior40. The contrasting dynamics of centromere DNA versus other chromosomal regions in yeast again points to different functional properties of different chromosomal loci. In fact, yeast centromeres have been reported to interact with microtubules throughout large portions of the cell cycle41, and, in the published study examining centromere dynamics37, disruption of microtubules with nocodazole led to increased centromere movements, with a reduction in apparent diffusion constraints. Thus, the overall picture emerging shows different rates of chromosome mobility between species, between cell types within a species, within the same cell type for different cell-cycle stages, and within the same cell type and the same cell-cycle stage but different chromosome regions. Aspects of this mobility can be explained by diffusion. Yet, at the heart of these observations, layered on top of this diffusion-like behavior is likely to be interesting regulation occurring at the level of regulated attachments or constraints on the range of motion, with possible hints at directed motion and therefore motor activity in certain examples. Progress over the next few years will be followed with keen interest as the mechanisms of this chromosome mobility and compartmentalization are explored. Sightings of a higher-order
While we are all well aware of the very high compaction present in maximally condensed mitotic chromosomes, the degree of compaction within interphase chromosomes, and the folding motifs underlying this compaction, have been less clear. There has been an implicit assumption in many textbooks that the major form of interphase chromosomes would be as 30-nm chromatin fibers, possibly packaged within loops, with attention focusing on the regulation of the 30-to-10-nm chromatin fiber transition. At least in nuclei of many higher eukaryotes, light and electron microscopy on fixed samples have suggested distinct large-scale chromatin fibers well above the 30 nm chromatin fiber, consistent with FISH experiments measuring compaction ratios 10–30-fold higher than 30-nm fibers33,42 (‘compaction ratio’ refers to the result from dividing the length of B-form DNA by the length of the same DNA folded as a chromatin fiber). The significance of these observations, however, has been questioned due to concerns over potential artifacts introduced by the conditions of specimen preparation. Now, using a GFP–histone fusion protein, in vivo images show remarkably similar large-scale chromatin organization in comparison to previous images of fixed samples, stained with DNA specific dyes, that showed suggestions of fibrillar large-scale chromatin structures at the resolution of light
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Fig. 6. IPL1 is among nine yeast genes identified in a ‘loss of cohesion’ (LOC) screen46. A chromosome marked with green-fluorescent protein (GFP) was used in a direct, primary visual genetic screen of temperature-sensitive (ts) lethals, looking at anaphase segregation of sister chromatids. A tandem lac operator repeat was integrated 12 kb from the centromere of chromosome 4. Temperature-sensitive mutant strains were screened for those with large budded cells containing one GFP signal instead of two. Secondary screens were used to eliminate mutant strains where a single dot was observed because of metaphase delay or arrest. Nine complementation groups defective for sister chromatid separation remained after these secondary screens, including one that involved mutants in the IPL1 gene. Percentages of cells showing one dot per nucleus, two dots in one nucleus and one dot in one nucleus are shown for an ipl1 mutant allele identified in this screen. (Photomicrograph kindly provided by Andrew Murray.)
Acknowledgements This work was supported by grants from NIH to A.S.B. I thank Alan Grossman and Katherine Lemon for providing Fig. 2, Susan M. Gasser for providing Fig. 5 and Andrew Murray for providing Fig. 6 for this paper. I thank Tudorita Tumbar for providing the right-hand panel of Fig. 4. I also thank Julio Vazquez and John Sedat for sharing and allowing citation of unpublished data.
microscopy43. By constructing gene-amplified chromosome regions containing lac operator repeats, selective staining of specific chromosome regions provides a clear in vivo demonstration of distinct, linear large-scale chromatin fibers with compaction ratios much higher than those of 30-nm chromatin fibers and with diameters of ~80 nm estimated by immunoelectron microscopy4. In general, the compaction of these fibers is remarkably constant over periods of several hours, varying no more than 10–20%, with general integrity of overall geometry and orientation within the nucleus being stable over similar periods (G. Li and A. Belmont, unpublished). While the biological significance of this level of chromatin organization remains unclear42, the stability of these structures necessitates an investigation of the impact of this higher-level compaction with regard to accessibility of the large macromolecular complexes involved in DNA function. A reproducible choreography of folding and unfolding of these large-scale structures during the cell cycle has been described, with a major unfolding correlating with onset of DNA replication38. Remarkably, for a late-replicating, heterochromaticlike amplified chromosome arm, DNA replication is observed in late S phase at a stage in which the chromosome arm has a clear linear axis and is only about threefold less compact than during metaphase38. Targeting a transcriptional activator to this heterochromatic chromosome arm resulted in dramatic uncoiling of the condensed chromosome region, in some cases into an extended large-scale chromatin fiber still ~20-fold higher in compaction than a 30-nm chromatin fiber – but now possessing high transcriptional activity44. Unfolding of large-scale chromatin structures http://tcb.trends.com
The development of GFP and methods for localizing specific proteins and DNA sequences has opened a new dimension for investigating chromosome dynamics. Combining these GFP methods with techniques such as fluorescence recovery after photobleaching (FRAP) has already had a large impact on how cell biologists think of nuclear and chromosome structure. In particular, there is now a growing appreciation of how metastable nuclear architecture and chromosome states are created from highly labile molecular interactions29. Future merger of these localization methods with spectroscopic methods, such as FRAP, fluorescence correlation spectroscopy (FCS) and fluorescence resonance energy transfer (FRET), should allow us not only to correlate structural transitions with changes in functional states but eventually also connect both with the underlying biochemistry and macromolecular dynamics. We also envision the harnessing of these GFP methods to carry out novel screens for identifying proteins regulating specific aspects of chromosome dynamics. To date, genetic screens aimed at chromosome dynamics have been largely indirect, based on functional assays such as chromosome mis-segregation or transcriptional activity. GFP methods for visualizing sister chromatid separation already have been used to characterize mutants identified in these indirect screens. Now, two recent papers have applied in vivo visualization of sister centromeres in yeast as direct visual screens for genetic or reverse-genetic approaches. Using failure of sister-chromatid separation as a primary visual screen of 2000 temperature-sensitive mutants, combined with several secondary screens, nine loss of cohesion (LOC) complementation groups that do not segregate sister chromatids at anaphase were identified, with two genes required for sisterchromatid separation and four genes required for sister-chromatid segregation46. Fig. 6 shows representative images from this screen for one of these complementation groups, corresponding to mutations in the Ipl1 gene. By combining functional genomics, to identify 171 genes expressed differentially in meiosis, with a visual, reverse-genetics, screen designed to examine chromosome segregation using GFP-marked centromeres, a novel kinetochore protein, monopolin, was identified that is required for segregation of homologs during meiosis I (Ref. 47). Through direct visual primary and secondary screens designed to specifically examine chromosome structure and dynamics, we envision the application of genetic, reverse-genetic and chemical-genetic approaches to identify proteins regulating chromosome mobility, homologous pairing, nuclear compartmentalization and other aspects of chromosome dynamics over the coming years.
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References 1 Zink, D. and Cremer, T. (1998) Cell nucleus: chromosome dynamics in nuclei of living cells. Curr. Biol. 8, R321–R324 2 Belmont, A.S. and Straight, A.F. (1998) In vivo visualization of chromosomes using lac operator–repressor binding. Trends Cell Biol. 8, 121–124 3 Belmont, A.S. et al. (1998) Visualization of largescale chromatin structure and dynamics using the lac operator/lac repressor reporter system. Methods Cell Biol. 58, 99–124 4 Robinett, C.C. et al. (1996) In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J. Cell Biol. 135, 1685–1700 5 Straight, A.F. et al. (1996) GFP tagging of budding yeast chromosomes reveals that protein–protein interactions can mediate sister chromatid cohesion. Curr. Biol. 6, 1599–1608 6 Webb, C.D. et al. (1997) Bipolar localization of the replication origin regions of chromosomes in vegetative and sporulating cells of B. subtilis. Cell 88, 667–674 7 Carmi, I. et al. (1998) The nuclear hormone receptor SEX-1 is an X-chromosome signal that determines nematode sex. Nature 396, 168–173 8 Michaelis, C. et al. (1997) Cohesins: chromosomal proteins that prevent premature separation of sister chromatids. Cell 91, 35–45 9 Jacob, F. et al. (1963) On the regulation of DNA replication in bacteria. Cold Spring Harbor Symp. Quant. Biol. 28, 329–348 10 Moller-Jensen, J. et al. (2000) Plasmid and chromosome segregation in prokaryotes. Trends Microbiol. 8, 313–320 11 Losick, R. and Shapiro, L. (1999) Changing views on the nature of the bacterial cell: from biochemistry to cytology. J. Bacteriol. 181, 4143–4145 12 Glaser, P. et al. (1997) Dynamic, mitotic-like behavior of a bacterial protein required for accurate chromosome partitioning. Genes Dev. 11, 1160–1168 13 Lin, D.C. et al. (1997) Bipolar localization of a chromosome partition protein in Bacillus subtilis. Proc. Natl. Acad. Sci. U. S. A. 94, 4721–4726 14 Teleman, A.A. et al. (1998) Chromosome arrangement within a bacterium. Curr. Biol. 8, 1102–1109 15 Gordon, G.S. et al. (1997) Chromosome and low copy plasmid segregation in E. coli: visual evidence for distinct mechanisms. Cell 90, 1113–1121 16 Webb, C.D. et al. (1998) Use of time-lapse microscopy to visualize rapid movement of the replication origin region of the chromosome during the cell cycle in Bacillus subtilis. Mol. Microbiol. 28, 883–892
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17 Lemon, K.P. and Grossman, A.D. (1998) Localization of bacterial DNA polymerase: evidence for a factory model of replication. Science 282, 1516–1519 18 Lemon, K.P. and Grossman, A.D. (2000) Movement of replicating DNA through a stationary replisome. Mol. Cell. 6, 1321–1330 19 Straight, A.F. et al. (1997) Mitosis in budding yeast: anaphase A but not metaphase plate. Science 277, 574–578 20 Pearson, C.G. et al. (2001) Budding yeast chromosome structure and dynamics during mitosis. J. Cell Biol. 152, 1255–1266 21 Straight, A.F. et al. (1998) Time-lapse microscopy reveals unique roles for kinesins during anaphase in budding yeast. J. Cell Biol. 143, 687–694 22 Nabeshima, K. et al. (1998) Dynamics of centromeres during metaphase–anaphase transition in fission yeast: Dis1 is implicated in force balance in metaphase bipolar spindle. Mol. Biol. Cell 9, 3211–3225 23 Goshima, G. and Yanagida, M. (2000) Establishing biorientation occurs with precocious separation of the sister kinetochores, but not the arms, in the early spindle of budding yeast. Cell 100, 619–633 24 Tanaka, T. et al. (2000) Cohesin ensures bipolar attachment of microtubules to sister centromeres and resists their precocious separation. Nat. Cell Biol. 2, 492–499 25 He, X. et al. (2000) Transient sister chromatid separation and elastic deformation of chromosomes during mitosis in budding yeast. Cell 101, 763–775 26 Shelby, R.D. et al. (1996) Dynamic elastic behavior of alpha-satellite DNA domains visualized in situ in living human cells. J. Cell Biol. 135, 545–557 27 Cremer, T. et al. (1993) Role of chromosome territories in the functional compartmentalization of the cell nucleus. Cold Spring Harbor Symp. Quant. Biol. 58, 777–792 28 Seksek, O. et al. (1997) Translational diffusion of macromolecule-sized solutes in cytoplasm and nucleus. J. Cell Biol. 138, 131–142 29 Misteli, T. (2001) Protein dynamics: Implications for nuclear architecture and gene expression. Science 291, 843–847 30 Belmont, A.S. (1998) Nuclear ultrastructure: transmission electron microscopy and image analysis. Methods Cell Biol. 53, 99–124 31 Alberts, B. et al. (1994) In Molecular Biology of the Cell, pp. 382–383, Garland Publishing 32 Boudonck, K. et al. (1999) The movement of coiled bodies visualized in living plant cells by the green fluorescent protein. Mol. Biol. Cell. 10, 2297–2307
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33 Mahy, N.L. et al. (2000) Linking large-scale chromatin structure with nuclear function. In Chromatin Structure and Gene Expression (Elgin, S.C.R. and Workman, J.L., eds), Oxford University Press 34 Bridger, J.M. et al. (2000) Re-modelling of nuclear architecture in quiescent and senescent human fibroblasts. Curr. Biol. 10, 149–152 35 Abney, J.R. et al. (1997) Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J. Cell Biol. 137, 1459–1468 36 Bornfleth, H. et al. (1999) Quantitative motion analysis of subchromosomal foci in living cells using four-dimensional microscopy. Biophys. J. 77, 2871–2886 37 Marshall, W.F. et al. (1997) Interphase chromosomes undergo constrained diffusional motion in living cells. Curr. Biol. 7, 930–939 38 Li, G. et al. (1998). Interphase cell cycle dynamics of a late-replicating, heterochromatic homogeneously staining region: precise choreography of condensation/decondensation and nuclear positioning. J. Cell Biol. 140, 975–989 39 Tumbar, T. and Belmont, A.S. (2001) Interphase movements of a DNA chromosome region modulated by VP16 transcriptional activator. Nat. Cell Biol. 3, 134–139 40 Heun, P. et al. (2001) The positioning and dynamics of origins of replication in the budding yeast nucleus. J. Cell Biol. 152, 385–400 41 Jin, Q.W. et al. (2000) Centromere clustering is a major determinant of yeast interphase nuclear organization. J. Cell Sci. 113, 1903–1912 42 Belmont, A.S. et al. (1999) Large-scale chromatin structure and function. Curr. Opin. Cell Biol. 11, 307–311 43 Kanda, T. et al. (1998) Histone–GFP fusion protein enables sensitive analysis of chromosome dynamics in living mammalian cells. Curr. Biol. 8, 377–385 44 Tumbar, T. et al. (1999) Large-scale chromatin unfolding and remodeling induced by VP16 acidic activation domain. J. Cell Biol. 145, 1341–1354 45 Tsukamoto, T. et al. (2000) Visualization of gene activity in living cells. Nat. Cell Biol. 2, 871–878 46 Biggins, S. et al. Genes involved in sister chromatid separation and segregation in the budding yeast Saccharomyces cerevisiae. Mol. Biol. Cell (in press) 47 Toth, A. et al. (2000) Functional genomics identifies monopolin: a kinetochore protein required for segregation of homologs during meiosis I. Cell 103, 1155–1168
GFP applications series This Review by Andrew Belmont is part of a series of articles being featured in Trends in Cell Biology on the uses of GFP in a variety of applications and technologies. See last month’s issue for a review from Fred Wouters, Peter Verveer and Philippe Bastiaens, ‘Imaging biochemistry inside cells’. Next month’s issue will contain a review by Derek Toomre and Dietmar Manstein, ‘Lighting up the cell surface with evanescent wave microscopy’.
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Visualizing chromosome dynamics with GFP
GFP applications
Andrew S. Belmont By allowing easy labeling of chromosomal and nuclear proteins and the tagging of specific chromosomal regions, the use of green-fluorescent protein (GFP) has provided new and special opportunities for directly observing chromosome dynamics in vivo. Here, we review recent applications of this methodology, focusing particularly on examples where new biology has been learned, or at least sighted. In particular, we focus on active bacterial chromosome segregation, yeast mitosis and centromere dynamics, and largescale chromatin structure and dynamics within eukaryotic interphase nuclei.
Andrew S. Belmont Dept of Cell and Structural Biology, University of Illinois, UrbanaChampaign B107 CLSL, 601 South Goodwin Ave., Urbana, IL 61801, USA. e-mail: [email protected]
More than 100 years ago, the dynamic behavior of mitotic chromosomes captured the interest of early microscopists. Identification of chromosomes as the basic units of heredity near the turn of the twentieth century pushed mitotic and meiotic chromosome segregation to center stage, where it has remained ever since as a major cell-biological and genetics research focus. Using transmitted light microscopy without any special staining, mitotic chromosome segregation can be observed directly in living cells over a time scale of seconds to minutes. The ability to visualize this dynamic process has been crucial to dissecting the molecular mechanisms underlying chromosome segregation. Conversely, the inability to easily visualize chromosomes in various organisms has led to serious scientific roadblocks. With regard to chromosome segregation, the inability to visualize bacterial chromosomes easily has meant that progress in understanding this process in prokaryotic cells has remained a ‘black box’for many decades. Even in eukaryotic organisms, the inability to easily visualize mitotic chromosomes, for instance in budding and fission yeast, has hindered the full exploitation of the powerful genetic tools available for these model organisms. More significantly, although it has been possible to visualize with relatively simple methods the segregation in eukaryotic cells of large, condensed mitotic chromosomes, visualization of decondensed, interphase chromosomes has remained technically challenging in all organisms. Basic questions of interphase chromosome structure as a result have relied largely on static images from fixed material and have been plagued by concerns over specimen preparation artifacts induced by fixation, immunostaining and in situ hybridization procedures. More interesting questions relating interphase chromosome structure and dynamics to regulation of transcription, replication, recombination and DNA repair have been difficult to approach owing to this key technical limitation.
In fact, it is not too great a stretch to conclude that this inability to visualize easily chromosome regions within interphase nuclei has contributed greatly to the relative backwardness of the cell biology of many aspects of nuclear structure and function. The introduction of fluorescently labeled nucleotides and proteins into cells has enabled direct observation of in vivo mitotic and interphase chromosome dynamics1. This approach has now been greatly enhanced by the recent applications of greenfluorescent protein (GFP). Not only can specific nuclear proteins be labeled with GFP, but, by using direct repeats of bacterial operator sequences, it is possible to tag specific chromosome sites and visualize these tags using GFP fused with the appropriate repressor protein (Fig. 1)2,3. This approach was first developed with the lac operator–repressor system in mammalian cells4 and yeast5 and then transferred rapidly to bacteria6, Caenorhabditis elegans (A. Serrichio and P. Sternberg, pers. commun.)7 and more recently to Drosophila (A. Belmont et al., unpublished). A similar approach using the tet operator–repressor system has also been highly successful8. Here, we review several recent examples of the scientific impact derived from these new experimental capabilities. Chromosome dynamics in bacteria
In contrast to active segregation of eukaryotic mitotic chromosomes, the textbook picture of bacterial chromosome segregation for several decades was based on the Jacob replicon model in which daughter chromatids attached to centrally located regions on the cell membrane, with passive separation being achieved through cell-wall growth during cell-cycle progression9. This textbook picture now has been supplanted by active chromosome segregation models, based on recent in vivo observations combined with traditional immunofluorescence and fluorescence in situ hybridization (FISH) approaches10,11. Using either GFP fusions to endogenous proteins binding near the chromosome origin region or GFP fused with bacterial repressors binding to operator direct repeats introduced at specific chromosome locations, in vivo dynamics of chromosome origins and other chromosome regions have been visualized. A defined spatial layout of the bacterial chromosome relative to the bacterial cell has been observed. In sporulating Bacillus subtilis, origins were localized at the cell poles, with termini near the
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Fig. 1. Tagging chromosome sites with the lac operator–repressor system3. (a) Detection: a direct repeat of the lac operator can be identified by the in vivo or in vitro binding of lac repressor protein. Originally a 256 copy, 10-kb direct repeat was made, but smaller repeats can also be visualized using this method. In Drosophila, a direct repeat of less than 1 kb has been visualized in diploid nuclei (P. Alvarez and A. Belmont, unpublished). The lac repressor has been engineered to contain a nuclear localization signal (NLS) and the C-terminus has been mutated to prevent lac repressor dimers from forming tetramers, which can bind to two different operators. (b) Transfection: direct repeats of the lac operators can be subcloned into transformation vectors and stable transformants generated by homologous recombination (to date, yeast and bacteria), transposable elements (Drosophila) or nonhomologous recombination (Caenorhabditis elegans, mammalian cells). Here, we show introduction of the lac operator direct repeat into an expression vector for dihydrofolate reductase (DHFR). DHFR serves as the selectable marker for selection of stable transformants in DHFR– Chinese hamster ovary (CHO) cells. Cells expressing a GFP–lac-repressor–NLS fusion protein show small dots of staining where the lac operator containing vector has integrated. A general nuclear background is present owing to nuclear import of the NLS-containing fusion protein. (c) Gene amplification: owing to the high degree of chromatin compaction, even multicopy vector insertions, hundreds of kb in size, appear as small dots in the mammalian cell nucleus by light microscopy. To visualize larger chromosome regions, gene amplification was used to create amplified chromosome regions, ranging in size from chromosome bands to whole chromosome arms. Multiple selections using increasing concentrations of methotrexate, an inhibitor of DHFR, were used to isolate cell lines that have undergone gene amplification. In these cell lines, continual fibers or a series of dots are seen, depending on the size of the coamplified genomic DNA.
cell middle6. Vegetatively growing cells showed a bipolar origin distribution after origin replication, with termini towards the middle and with a preference for polar origin localization prior to replication6,12,13. Labeling origins, termini and quarter positions (90° and 270°) demonstrated in general maximum separation between origin and terminus, with quarter positions lying in between14. E. coli showed similar bipolar distribution of origins and a central location of termini, but, by contrast, http://tcb.trends.com
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both F and P1 plasmids localized to the cell center prior to replication and moved rapidly to the quarter cell positions after replication15. The bacterial chromosome and each of the two plasmids all appear to use different overall mechanisms for active chromosome segregation. FISH results and immunolocalization using proteins mapping to the chromosome origin produced similar results10. Time-lapse imaging on large numbers of B. subtilis cells revealed that approximately two-thirds of origins moved from the pole towards the cell center prior to separation into two origins, with origin separation occurring near the pole in the remaining cells. Strikingly, the majority of movement during separation of newly replicated origins occurred during a short portion of the total cell cycle, approximately 11 minutes out of a 100-minute doubling time, with peak origin velocity during the first 3–6 minutes of origin separation16. Chromosome termini, as well as chromosome quarter positions (90° and 270°), also were observed to separate rapidly, thus demonstrating an active mechanism for partitioning of bacterial chromosomes16. Similarly rapid and abrupt origin partitioning movements were observed using a GFP fusion with SpoOJ, which binds near to the B. subtilis origin14. The recent demonstration that DNA polymerase is located at a discrete spot near the cell center means that these active chromosome partitioning mechanisms must be coupled in some way with spatially localized replication17. By stalling DNA replication at a specific, labeled chromosome site ~100 kb from the replication origin, followed by release of this block, movement away from the replication machinery to the cell poles was observed for newly replicated foci18. A labeled chromosome region lying approximately halfway between origin and terminus moved to the centrally located replication focus prior to duplication and separation. Forces generated by DNA polymerase, therefore, might contribute to chromosome segregation. Fig. 2 shows the spatial arrangement of replisomes containing DNA polymerase III (orange) and labeled chromosomal origins (blue) at different times after cell synchronization and induction and then release of the replication block. Chromosome origins localize to the replisome (d–f) during the replication block but move quickly towards the poles after release of this replication block (g–i). Mitosis in cytologically challenged but genetically blessed eukaryotes
Direct observation of chromosome dynamics in higher eukaryotic cells has been crucial in formulating mechanistic models of mitotic chromosome segregation. The ability to test these models directly, however, and to identify the underlying molecular components that execute these mechanisms has been limited by the functional redundancy for key gene products frequently observed in these organisms. This is compounded by the experimental difficulties in knocking out these gene products, and the relative
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Fig. 2. Movement of newly replicated chromosome origins away from a centrally located replisome. Bacillus subtilis cells were observed at different times after synchronization of the replication cycle. Synchronization was achieved using a temperature-sensitive allele of dnaB134, required for DNA replication initiation. Growth at the nonpermissive temperature allows ongoing rounds of replication to finish while preventing initiation of new rounds of replication. Release of this replication initiation block was followed by a stringent replication arrest, using amino acid starvation, which arrests replication forks at defined chromosome loci, LSTer and RSTer, located ~100–130 kb and ~150–200 kb, respectively, from the origin. The replisome was labeled using a subunit of DNA polymerase III – dnaX – fused with YFP. LSTer was labeled by the nearby insertion of a lac operator direct repeat, binding to lac-repressor–CFP. (a–c) Sixty minutes after synchronization by a block in DNA replication initiation. Two copies of LSTer are located towards the cell poles, away from the centrally located replisome. (d–f) Sixty minutes after synchronization and induction of the replication arrest at LSTer. LSTer is coincident with the replisome. (g–i) Thirty minutes after release from the replication arrest, two copies of LSTer are located away from the replisome. (j–l) A cell undergoing (or about to undergo) multifork replication 30 min after release from stringent replication arrest. The central polymerase is from the initial round of replication, and additional replisomes have assembled at the cell quarter positions. (m–o) Replisomes approximately at the cell quarters in a cell that seems to have finished the initial replication round (no replisome at midcell). Left panels: foci of DNA polymerase (orange) and cell outline (gray). Middle panels: foci of LSTer (cyan) and cell outline (gray). Right panels: merged images from left and middle panels. Bar, 1 µm. (Images reprinted, with permission, from Ref. 18.)
difficulty in identifying novel proteins involved in these processes. By contrast, in genetically tractable organisms such as budding and fission yeast, relatively simple screens have identified a large number of genes involved in chromosome segregation during mitosis and meiosis, but the poor cytology in these organisms has seriously hindered identification of the functional role of each of these gene products. http://tcb.trends.com
Using GFP, it has now been possible to visualize directly in vivo chromosome dynamics during mitosis in budding yeast using marked chromosomal sites5. Chromosome movements relative to the microtubule spindle structure were observed in cells coexpressing GFP–lac-repressor and GFP-α-tubulin19. Budding yeast did not appear to show a conventional metaphase plate alignment; instead chromosomes oscillated along the entire length of the spindle, with chromosomes frequently near one spindle pole at the onset of anaphase chromosome separation. However, a recent study has shown that DNA within several kilobases of the centromere is located within the central 25% of the spindle in the majority of nuclei20, suggesting a time-averaged, metaphase-like alignment for centromere regions but not for chromosome arms. A rapid, polewards chromosome movement during an anaphase A stage was observed, with centromeres leading and telomeres lagging, and with transient chromosome stretching between centromere and telomere19. Comparison of in vivo mitotic chromosome motion in wild-type versus mutant yeast strains revealed unique roles during mitosis for different kinesin-like motors, with Cin8p implicated in the rapid phase of anaphase B, and Kip1p in the slow phase of anaphase B21. Similar mitotic events were observed in fission yeast, with an extensive period during which chromosomes oscillate between poles prior to onset of sister chromatid separation and anaphase A and B movements22. Transient splitting of centromeres was also observed, up to 0.6 µm in distance, just prior to the onset of anaphase chromosome separation, suggesting that, during metaphase, the centromere was positioned in a bioriented manner towards the poles and was under tension. Reinvestigation of centromere dynamics in budding yeast using chromosomes tagged just several kilobases from the actual centromere region demonstrated a similar ‘precocious’ separation of centromere regions early during spindle formation and probably soon after centromere duplication23,24. Similar early splitting of fluorescence signal was observed for GFP-labeled kinetochore proteins bound to wild-type centromeres23. Time-lapse three-dimensional microscopy demonstrated that centromere separation was transient, with dramatic, elastic stretching of centromere regions and movement towards the poles, followed by periods when daughter centromeres rejoin20,24,25. This pre-anaphase stretching occurs predominately in DNA regions within ~9–13 kb flanking the actual centromere sequence and lowers compaction ratios to ~10–20, below the 30–40 ratio expected for a 30-nm fiber and the overall 90–120-fold compaction ratio seen for yeast chromosomes24. Anaphase stretching to below that of nucleosome filaments was observed transiently for the tet-operator-labeled insert DNA during anaphase, but it remains unclear whether
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this extreme stretching is a property of the tet op repeat DNA only or would also be observed for wild-type sequence24. Recent data suggest similar stretching, however, for ~1 kb of DNA between the centromere and a 1.7 kb lac operator repeat20. The existence of this precocious sister centromere separation is consistent with the establishment of properly bioriented sister kinetochores through a tension-generating and tension-sensing mechanism, as established for higher eukaryotic cells where an approximately twofold stretching of GFP–CENP-Blabeled centromeres has been observed26. The magnitude of this stretching and separation in yeast, however, is surprising, with the disruption of chromatin packing to the level of a nucleosome filament and the 5–10-fold stretching much larger proportionally than seen for mammalian centromeres. Precocious separation of sister centromeres does not require Esp1p, a protein required for cohesin degradation. Therefore, separation is independent of cohesin proteolysis25, although prevention of premature daughter chromosome separation requires the cohesin component Scc1p, and therefore cohesin function, to hold the distal chromosome arms together during periods of centromere splitting24. These observations suggest that cohesin complexes near centromeres might separate transiently during centromere splitting. Because splitting falls off sharply beyond a certain distance from the core centromere, this has further supported the possibility of specialized, elastic chromatin regions flanking centromeres25. Comparison of the velocities of centromere-proximal and centromere-distal labeled chromosome sites during anaphase suggests an elastic recoil accompanying separation of sister chromatids, increasing the initial rate of sister chromosome separation for centromere-distal regions20. Interphase chromosomes on the move
As accustomed as we are to dynamic chromosome movements during mitosis, our natural inclination is to view the interphase nucleus and its contents as static. Previously, chromosomes and subnuclear organelles have been difficult to visualize in fixed material, let alone live cells. Moreover, the overall shape of nuclei and their visible contents, such as nucleoli, have not shown rapid movements. Using FISH, distinct chromosome ‘territories’ have been observed. The solid appearance of these chromosome ‘paints’ in some studies has led to models in which chromatin packing within territories would limit diffusion of macromolecular complexes27. Similarly, close contact between different chromosome territories has left the impression that the majority of the nuclear volume might be occupied by chromatin, directly constraining movement of chromosomes themselves. In fact, measurements of dextran intranuclear diffusion in vertebrate tissue-culture cells indicated http://tcb.trends.com
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that no more than 15% of the nuclear volume is inaccessible to small molecules28,29, and electron microscopy has revealed large spaces between largescale chromatin domains30. Stereotypical cell-cycle changes in nucleolar distribution have been described31, as well as large movements of coiled bodies, demonstrated through use of GFP fusion proteins32. A combination of immunostaining and FISH has demonstrated nonrandom localization of DNA sequences in the nucleus; these include associations with the nuclear envelope, nucleoli or heterochromatic blocks (reviewed in Ref. 33). Changes in these nonrandom distributions during the cell cycle, differentiation and changes in gene expression imply intranuclear chromosome movements occurring over a time frame no greater than several hours. Recent data examining the differential localization of a gene-rich human chromosome versus a gene-poor human chromosome by FISH have suggested that changes in interphase chromosomal location of entire chromosomes might occur relatively rapidly34. These studies, however, reveal only steady-state statistical distributions, without revealing actual dynamics. Using fluorescence recovery after photobleaching (FRAP), little or no relative motion was observed in vivo in mammalian cells over several hours for bleached chromosome regions, stained with ethidium bromide and ~1 µm in diameter35. Similarly, in vivo dynamics of chromosome foci labeled by fluorescent nucleotides in general showed little or no motion over periods of several hours, particularly after adjustments for nuclear rotation and spatial translation36. Likewise, centromeres labeled with a GFP–CENP-B fusion protein in mammalian tissueculture cells also in general showed little motion26. Together, these studies examining the aggregate behavior of large chromosome regions within mammalian tissue-culture cells unsynchronized in the cell cycle indicated that long-range motion of interphase chromosomal loci, if present, must be infrequent and confined to a small subset of chromosomal loci and/or cell-cycle stages. These studies also did not address whether rapid, but locally constrained, chromosome movements were occurring. A careful examination of yeast chromosomes labeled near centromeres, however, showed that, over seconds to tens of seconds, there was in fact rapid but localized motion that was best described as diffusion within a constrained radius of ~0.3 µm (Ref. 37). Rapid, localized motion in mammalian cell nuclei on a time scale of seconds or less also occurs within mammalian cells (A.S. Belmont, unpublished). More extended observations, however, have revealed infrequent examples in mammalian cells of sustained, unidirectional motion of centromeres26 and chromosome replication domains labeled by fluorescent nucleotides36. Using artificial chromosome constructs labeled with lac operator repeats, three examples of specific movements now have been
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Fig. 3. Direct observation of a translocation over several microns of a ~90-Mbp heterochromatic, gene-amplified chromosome arm from the nuclear periphery to the nucleolus and back. (a–f) Images represent combination of transmitted- and fluorescence-light images at specific time points, measured from beginning of observation time (t = 0), 4 h after release from an early S-phase block. (a–f) correspond to t = 1, 5, 5.5, 6, 9 and 9.5 hrs, respectively. Movement of the chromosome arm from the nuclear periphery to nucleolus is coupled with chromosome decondensation. The chromosome arm returns to the nuclear periphery in the vicinity of the original starting position. Arrows point to the edge of the nucleus. Bar, 2 µm. (G. Li and A. Belmont, unpublished.)
observed in mammalian cells. A late-replicating, amplified chromosome region, usually located adjacent to the nuclear envelope in G1 and the first half of S phase, was observed to move to the nuclear interior 4–6 hours into S phase. This movement was
tightly correlated with the onset of DNA replication in this chromosome region38. Both the timing of this movement and its correlation with DNA replication were established by statistical analysis of a large number of cells using fixed preparations. An example in which this motion has been observed directly in living cells is shown in Fig. 3. Cells were synchronized in early S phase. Visualization began four hours after release from the early S phase. A motion from the nuclear envelope to the nucleolus and back to the nuclear envelope occurs. A cell clone containing an insertion of 10–20 vector copies containing lac operator inserts showed a single, GFP–lac repressor spot in the nucleus, and this had a high frequency of association with the nuclear envelope during G1 phase. Movement of this spot to the nuclear interior was observed at a distinct stage in early S phase39. Direct in vivo visualization showed that, in the absence of VP16, in most cells the spot frequently moved transiently into the nuclear interior during early G1 but relocated back to the nuclear periphery and remained there until early S phase (Fig. 4, bottom left panel). Targeting the acidic activation domain of VP16 to this spot activated transcription and also induced a permanent relocalization of the spot to the nuclear interior that was established soon after mitosis (Fig. 4, top left panel). This differential nuclear positioning of this chromosome site occurs as a function of cell-cycle progression and the presence or absence of the VP16 activation domain in Fig. 4, right panel. Control
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Fig. 4. Interphase positioning of a DNA chromosome region is established early after mitosis and is modulated by VP16 transcriptional activator39. Left: A specific chromosome site, normally located preferentially adjacent to the nuclear periphery (bottom panel), instead localizes to the nuclear interior when the VP16 acidic activation domain (AAD) is tethered to this site (top panel). Top panel shows example from C4-VP16 cells expressing a GFP–lac-repressor–VP16-AAD fusion protein targeted to a chromosome site (bright spot) containing lac operator repeats. Number in left, bottom corner refers to minutes after cell division. The chromosome site localizes away from nuclear edge early in G1 phase. Bottom panel shows control cells expressing a GFP–lacrepressor protein. The site starts at the nuclear periphery, extends
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linearly into the nuclear interior transiently, then returns as a condensed spot to the nuclear edge. This transient stretching or extension was observed in a significant fraction of cells and was largely restricted to this G1 period. Bar, 5 µm. Image reprinted, with permission, from Ref. 39. Right: summary of cell-cycle changes in nuclear position for a chromosome site containing the lac operator in control cells expressing GFP–lac repressor (left) versus cells expressing the GFP–lacrepressor–VP16-AAD fusion protein (right). Control cells show a peripheral spot localization that moves to the interior early in S phase. Cells expressing the GFP–lac-repressor–VP16 fusion protein show an interior localization established early in G1. Abbreviation: GFP, greenfluorescent protein. (Drawing kindly provided by Tudorita Tumbar.)
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In Drosophila embryos, a higher rate of mobility has been observed over short times, using fluorescently labeled topoisomerase 2 to label a specific satellite repeat, with a much larger radius of constrained motion37. In imaginal disks (J. Radasta and A.S. Belmont, unpublished) and in spermatocytes (J. Vazquez et al., pers. commun.), a much larger range of mobility is observed, over a time scale of minutes, as compared with mammalian cells. During male meiosis I in late G2, this long-range mobility ceases and is replaced by a localized constrained diffusion (J. Vazquez and J. Sedat, pers. commun.). In contrast to the constrained diffusion observed previously near yeast centromeres37, a high degree of mobility was observed for a number of tagged chromosomal sites in yeast. Using FISH, latereplicating origins distant from the yeast telomere were shown to be enriched during G1 in a zone adjacent to the nuclear periphery40. This was in contrast to early-replicating chromosomal origins, which were distributed randomly throughout the nucleus. Live observation, however, revealed large movements (~0.5 µm) over a distance of approximately one-half the radius of the cell nucleus; these occurred during intervals of several seconds approximately once per minute during G1 phase for early-replicating chromosome regions40. Fig. 5 shows an example in which an early-replicating chromosome region moves from near the nuclear center to the periphery in a six-second period. Similar, but somewhat less frequent, movements during G1 phase were seen for the late-replicating origin cluster examined, including movements well into the nuclear interior (S.M. Gasser, pers. commun.). In an interesting parallel with the previously described results in mammalian cells, the late-replicating (a)
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Fig. 5. High mobility of budding yeast interphase chromosomes. Shown are six frames (a–f) of a haploid yeast cell in G1 phase, in which an early-firing autonomously replicating sequence (ARS) on Chr IV (ARS Chr 4-908) is tagged with multiple lac operator repeats, creating a bright focus of green-fluorescent protein (GFP) fluorescence, while the nuclear envelope is visualized with NUP49-GFP40. The images were taken on the Zeiss LSM 510 confocal with a 100× objective. They are time-lapse images, with 1.22 seconds between each image. From the first to the last image, 6.12 seconds have elapsed, and the bright spot representing the origin has moved from the center of the nucleus to the periphery. The nucleus is ~1.9 µm in diameter. (Photomicrograph kindly provided by Susan M. Gasser.)
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origin cluster on chromosome 14 lost its peripherallocalization bias during S phase, becoming distributed throughout the nuclear interior40. The contrasting dynamics of centromere DNA versus other chromosomal regions in yeast again points to different functional properties of different chromosomal loci. In fact, yeast centromeres have been reported to interact with microtubules throughout large portions of the cell cycle41, and, in the published study examining centromere dynamics37, disruption of microtubules with nocodazole led to increased centromere movements, with a reduction in apparent diffusion constraints. Thus, the overall picture emerging shows different rates of chromosome mobility between species, between cell types within a species, within the same cell type for different cell-cycle stages, and within the same cell type and the same cell-cycle stage but different chromosome regions. Aspects of this mobility can be explained by diffusion. Yet, at the heart of these observations, layered on top of this diffusion-like behavior is likely to be interesting regulation occurring at the level of regulated attachments or constraints on the range of motion, with possible hints at directed motion and therefore motor activity in certain examples. Progress over the next few years will be followed with keen interest as the mechanisms of this chromosome mobility and compartmentalization are explored. Sightings of a higher-order
While we are all well aware of the very high compaction present in maximally condensed mitotic chromosomes, the degree of compaction within interphase chromosomes, and the folding motifs underlying this compaction, have been less clear. There has been an implicit assumption in many textbooks that the major form of interphase chromosomes would be as 30-nm chromatin fibers, possibly packaged within loops, with attention focusing on the regulation of the 30-to-10-nm chromatin fiber transition. At least in nuclei of many higher eukaryotes, light and electron microscopy on fixed samples have suggested distinct large-scale chromatin fibers well above the 30 nm chromatin fiber, consistent with FISH experiments measuring compaction ratios 10–30-fold higher than 30-nm fibers33,42 (‘compaction ratio’ refers to the result from dividing the length of B-form DNA by the length of the same DNA folded as a chromatin fiber). The significance of these observations, however, has been questioned due to concerns over potential artifacts introduced by the conditions of specimen preparation. Now, using a GFP–histone fusion protein, in vivo images show remarkably similar large-scale chromatin organization in comparison to previous images of fixed samples, stained with DNA specific dyes, that showed suggestions of fibrillar large-scale chromatin structures at the resolution of light
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after targeting of the transcription factor occurred over several hours44,45 and preceded accumulation of transcript and reporter protein45. Future visions
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Fig. 6. IPL1 is among nine yeast genes identified in a ‘loss of cohesion’ (LOC) screen46. A chromosome marked with green-fluorescent protein (GFP) was used in a direct, primary visual genetic screen of temperature-sensitive (ts) lethals, looking at anaphase segregation of sister chromatids. A tandem lac operator repeat was integrated 12 kb from the centromere of chromosome 4. Temperature-sensitive mutant strains were screened for those with large budded cells containing one GFP signal instead of two. Secondary screens were used to eliminate mutant strains where a single dot was observed because of metaphase delay or arrest. Nine complementation groups defective for sister chromatid separation remained after these secondary screens, including one that involved mutants in the IPL1 gene. Percentages of cells showing one dot per nucleus, two dots in one nucleus and one dot in one nucleus are shown for an ipl1 mutant allele identified in this screen. (Photomicrograph kindly provided by Andrew Murray.)
Acknowledgements This work was supported by grants from NIH to A.S.B. I thank Alan Grossman and Katherine Lemon for providing Fig. 2, Susan M. Gasser for providing Fig. 5 and Andrew Murray for providing Fig. 6 for this paper. I thank Tudorita Tumbar for providing the right-hand panel of Fig. 4. I also thank Julio Vazquez and John Sedat for sharing and allowing citation of unpublished data.
microscopy43. By constructing gene-amplified chromosome regions containing lac operator repeats, selective staining of specific chromosome regions provides a clear in vivo demonstration of distinct, linear large-scale chromatin fibers with compaction ratios much higher than those of 30-nm chromatin fibers and with diameters of ~80 nm estimated by immunoelectron microscopy4. In general, the compaction of these fibers is remarkably constant over periods of several hours, varying no more than 10–20%, with general integrity of overall geometry and orientation within the nucleus being stable over similar periods (G. Li and A. Belmont, unpublished). While the biological significance of this level of chromatin organization remains unclear42, the stability of these structures necessitates an investigation of the impact of this higher-level compaction with regard to accessibility of the large macromolecular complexes involved in DNA function. A reproducible choreography of folding and unfolding of these large-scale structures during the cell cycle has been described, with a major unfolding correlating with onset of DNA replication38. Remarkably, for a late-replicating, heterochromaticlike amplified chromosome arm, DNA replication is observed in late S phase at a stage in which the chromosome arm has a clear linear axis and is only about threefold less compact than during metaphase38. Targeting a transcriptional activator to this heterochromatic chromosome arm resulted in dramatic uncoiling of the condensed chromosome region, in some cases into an extended large-scale chromatin fiber still ~20-fold higher in compaction than a 30-nm chromatin fiber – but now possessing high transcriptional activity44. Unfolding of large-scale chromatin structures http://tcb.trends.com
The development of GFP and methods for localizing specific proteins and DNA sequences has opened a new dimension for investigating chromosome dynamics. Combining these GFP methods with techniques such as fluorescence recovery after photobleaching (FRAP) has already had a large impact on how cell biologists think of nuclear and chromosome structure. In particular, there is now a growing appreciation of how metastable nuclear architecture and chromosome states are created from highly labile molecular interactions29. Future merger of these localization methods with spectroscopic methods, such as FRAP, fluorescence correlation spectroscopy (FCS) and fluorescence resonance energy transfer (FRET), should allow us not only to correlate structural transitions with changes in functional states but eventually also connect both with the underlying biochemistry and macromolecular dynamics. We also envision the harnessing of these GFP methods to carry out novel screens for identifying proteins regulating specific aspects of chromosome dynamics. To date, genetic screens aimed at chromosome dynamics have been largely indirect, based on functional assays such as chromosome mis-segregation or transcriptional activity. GFP methods for visualizing sister chromatid separation already have been used to characterize mutants identified in these indirect screens. Now, two recent papers have applied in vivo visualization of sister centromeres in yeast as direct visual screens for genetic or reverse-genetic approaches. Using failure of sister-chromatid separation as a primary visual screen of 2000 temperature-sensitive mutants, combined with several secondary screens, nine loss of cohesion (LOC) complementation groups that do not segregate sister chromatids at anaphase were identified, with two genes required for sisterchromatid separation and four genes required for sister-chromatid segregation46. Fig. 6 shows representative images from this screen for one of these complementation groups, corresponding to mutations in the Ipl1 gene. By combining functional genomics, to identify 171 genes expressed differentially in meiosis, with a visual, reverse-genetics, screen designed to examine chromosome segregation using GFP-marked centromeres, a novel kinetochore protein, monopolin, was identified that is required for segregation of homologs during meiosis I (Ref. 47). Through direct visual primary and secondary screens designed to specifically examine chromosome structure and dynamics, we envision the application of genetic, reverse-genetic and chemical-genetic approaches to identify proteins regulating chromosome mobility, homologous pairing, nuclear compartmentalization and other aspects of chromosome dynamics over the coming years.
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References 1 Zink, D. and Cremer, T. (1998) Cell nucleus: chromosome dynamics in nuclei of living cells. Curr. Biol. 8, R321–R324 2 Belmont, A.S. and Straight, A.F. (1998) In vivo visualization of chromosomes using lac operator–repressor binding. Trends Cell Biol. 8, 121–124 3 Belmont, A.S. et al. (1998) Visualization of largescale chromatin structure and dynamics using the lac operator/lac repressor reporter system. Methods Cell Biol. 58, 99–124 4 Robinett, C.C. et al. (1996) In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J. Cell Biol. 135, 1685–1700 5 Straight, A.F. et al. (1996) GFP tagging of budding yeast chromosomes reveals that protein–protein interactions can mediate sister chromatid cohesion. Curr. Biol. 6, 1599–1608 6 Webb, C.D. et al. (1997) Bipolar localization of the replication origin regions of chromosomes in vegetative and sporulating cells of B. subtilis. Cell 88, 667–674 7 Carmi, I. et al. (1998) The nuclear hormone receptor SEX-1 is an X-chromosome signal that determines nematode sex. Nature 396, 168–173 8 Michaelis, C. et al. (1997) Cohesins: chromosomal proteins that prevent premature separation of sister chromatids. Cell 91, 35–45 9 Jacob, F. et al. (1963) On the regulation of DNA replication in bacteria. Cold Spring Harbor Symp. Quant. Biol. 28, 329–348 10 Moller-Jensen, J. et al. (2000) Plasmid and chromosome segregation in prokaryotes. Trends Microbiol. 8, 313–320 11 Losick, R. and Shapiro, L. (1999) Changing views on the nature of the bacterial cell: from biochemistry to cytology. J. Bacteriol. 181, 4143–4145 12 Glaser, P. et al. (1997) Dynamic, mitotic-like behavior of a bacterial protein required for accurate chromosome partitioning. Genes Dev. 11, 1160–1168 13 Lin, D.C. et al. (1997) Bipolar localization of a chromosome partition protein in Bacillus subtilis. Proc. Natl. Acad. Sci. U. S. A. 94, 4721–4726 14 Teleman, A.A. et al. (1998) Chromosome arrangement within a bacterium. Curr. Biol. 8, 1102–1109 15 Gordon, G.S. et al. (1997) Chromosome and low copy plasmid segregation in E. coli: visual evidence for distinct mechanisms. Cell 90, 1113–1121 16 Webb, C.D. et al. (1998) Use of time-lapse microscopy to visualize rapid movement of the replication origin region of the chromosome during the cell cycle in Bacillus subtilis. Mol. Microbiol. 28, 883–892
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17 Lemon, K.P. and Grossman, A.D. (1998) Localization of bacterial DNA polymerase: evidence for a factory model of replication. Science 282, 1516–1519 18 Lemon, K.P. and Grossman, A.D. (2000) Movement of replicating DNA through a stationary replisome. Mol. Cell. 6, 1321–1330 19 Straight, A.F. et al. (1997) Mitosis in budding yeast: anaphase A but not metaphase plate. Science 277, 574–578 20 Pearson, C.G. et al. (2001) Budding yeast chromosome structure and dynamics during mitosis. J. Cell Biol. 152, 1255–1266 21 Straight, A.F. et al. (1998) Time-lapse microscopy reveals unique roles for kinesins during anaphase in budding yeast. J. Cell Biol. 143, 687–694 22 Nabeshima, K. et al. (1998) Dynamics of centromeres during metaphase–anaphase transition in fission yeast: Dis1 is implicated in force balance in metaphase bipolar spindle. Mol. Biol. Cell 9, 3211–3225 23 Goshima, G. and Yanagida, M. (2000) Establishing biorientation occurs with precocious separation of the sister kinetochores, but not the arms, in the early spindle of budding yeast. Cell 100, 619–633 24 Tanaka, T. et al. (2000) Cohesin ensures bipolar attachment of microtubules to sister centromeres and resists their precocious separation. Nat. Cell Biol. 2, 492–499 25 He, X. et al. (2000) Transient sister chromatid separation and elastic deformation of chromosomes during mitosis in budding yeast. Cell 101, 763–775 26 Shelby, R.D. et al. (1996) Dynamic elastic behavior of alpha-satellite DNA domains visualized in situ in living human cells. J. Cell Biol. 135, 545–557 27 Cremer, T. et al. (1993) Role of chromosome territories in the functional compartmentalization of the cell nucleus. Cold Spring Harbor Symp. Quant. Biol. 58, 777–792 28 Seksek, O. et al. (1997) Translational diffusion of macromolecule-sized solutes in cytoplasm and nucleus. J. Cell Biol. 138, 131–142 29 Misteli, T. (2001) Protein dynamics: Implications for nuclear architecture and gene expression. Science 291, 843–847 30 Belmont, A.S. (1998) Nuclear ultrastructure: transmission electron microscopy and image analysis. Methods Cell Biol. 53, 99–124 31 Alberts, B. et al. (1994) In Molecular Biology of the Cell, pp. 382–383, Garland Publishing 32 Boudonck, K. et al. (1999) The movement of coiled bodies visualized in living plant cells by the green fluorescent protein. Mol. Biol. Cell. 10, 2297–2307
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33 Mahy, N.L. et al. (2000) Linking large-scale chromatin structure with nuclear function. In Chromatin Structure and Gene Expression (Elgin, S.C.R. and Workman, J.L., eds), Oxford University Press 34 Bridger, J.M. et al. (2000) Re-modelling of nuclear architecture in quiescent and senescent human fibroblasts. Curr. Biol. 10, 149–152 35 Abney, J.R. et al. (1997) Chromatin dynamics in interphase nuclei and its implications for nuclear structure. J. Cell Biol. 137, 1459–1468 36 Bornfleth, H. et al. (1999) Quantitative motion analysis of subchromosomal foci in living cells using four-dimensional microscopy. Biophys. J. 77, 2871–2886 37 Marshall, W.F. et al. (1997) Interphase chromosomes undergo constrained diffusional motion in living cells. Curr. Biol. 7, 930–939 38 Li, G. et al. (1998). Interphase cell cycle dynamics of a late-replicating, heterochromatic homogeneously staining region: precise choreography of condensation/decondensation and nuclear positioning. J. Cell Biol. 140, 975–989 39 Tumbar, T. and Belmont, A.S. (2001) Interphase movements of a DNA chromosome region modulated by VP16 transcriptional activator. Nat. Cell Biol. 3, 134–139 40 Heun, P. et al. (2001) The positioning and dynamics of origins of replication in the budding yeast nucleus. J. Cell Biol. 152, 385–400 41 Jin, Q.W. et al. (2000) Centromere clustering is a major determinant of yeast interphase nuclear organization. J. Cell Sci. 113, 1903–1912 42 Belmont, A.S. et al. (1999) Large-scale chromatin structure and function. Curr. Opin. Cell Biol. 11, 307–311 43 Kanda, T. et al. (1998) Histone–GFP fusion protein enables sensitive analysis of chromosome dynamics in living mammalian cells. Curr. Biol. 8, 377–385 44 Tumbar, T. et al. (1999) Large-scale chromatin unfolding and remodeling induced by VP16 acidic activation domain. J. Cell Biol. 145, 1341–1354 45 Tsukamoto, T. et al. (2000) Visualization of gene activity in living cells. Nat. Cell Biol. 2, 871–878 46 Biggins, S. et al. Genes involved in sister chromatid separation and segregation in the budding yeast Saccharomyces cerevisiae. Mol. Biol. Cell (in press) 47 Toth, A. et al. (2000) Functional genomics identifies monopolin: a kinetochore protein required for segregation of homologs during meiosis I. Cell 103, 1155–1168
GFP applications series This Review by Andrew Belmont is part of a series of articles being featured in Trends in Cell Biology on the uses of GFP in a variety of applications and technologies. See last month’s issue for a review from Fred Wouters, Peter Verveer and Philippe Bastiaens, ‘Imaging biochemistry inside cells’. Next month’s issue will contain a review by Derek Toomre and Dietmar Manstein, ‘Lighting up the cell surface with evanescent wave microscopy’.
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