The Interphase Nucleus as a Dynamic Structure

The lnterphase Nucleus as a Dynamic Structure Umberto De Boni Department of Physiology, Faculty of Medicine, University of Toronto, Toronto. Ontario Canada M5S lA8

1. Introduction

The spatial organization of the contents of interphase nuclei is of increasing interest, largely because of an increasing body of evidence which shows that interphase nuclei in different tissues in general and in cells of the central nervous system (CNS) in particular exhibit distinct patterns of chromatin organization. The interphase nuclei of different CNS cells display distinct and nonrandom arrangements of specific DNA sequences (Manuelidis, 1984a,b; Manuelidis and Borden, 1988; Billia and De Boni, 1991; Holowacz and De Boni, 1991). This has been postulated to indicate that transcription may be, in part, controlled and regulated by nuclear compartmentalization (Manuelidis, 1985a,b). This concept is supported by the argument that the primary DNA sequence alone, which is common to the cells of a given organism, cannot fully account for the different tissue- or cell type-specific gene expressions which occur within interphase nuclei (Pienta et al., 1991). Further support comes from observations of an association between changes in chromatin organization and altered functional states in cells. In fact, as early as 1949 it was recognized that electrical stimulation of hypoglossal neurons in situ resulted in a significant repositioning of nucleolar satellite DNA, concurrent with intense RNA synthesis (Barr and Bertram, 1949, 1951). More recently, specific chromatin domains have also been shown to reorganize and move significantly during neuronal differentiation (Manuelidis, 1985) as well as in neurons that exhibit altered functional states such as those which occur within epileptic foci in the human cerebral cortex (Borden and Manuelidis, 1988). In addition, observations of cells in vitro have shown that chromatin moves in a quantifiable manner within the spatial confines of interphase nuclei, a motion traditionally Internarional Reuien of Cyrolog)~,Vol. I S 0

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termed nuclear rotation (Pomerat, 1953; Pomerat et al., 1967; De Boni and Mintz, 1986). The rate of such motion in neurons in uitro is altered by nerve growth factor or neurotransmitters (Fung and De Boni, 1988), agents which also alter neuronal gene expression (Greenberg et al., 1985, 1986). Together, these observations correlate changes in dynamic positions of gene sequences within the three-dimensional (3-D) space of neuronal interphase nuclei with changes in gene expression, and suggest causal links. The existence of such links is further supported by the emerging concept that the 3-D organization of the genome may play an important role in the control of gene expression (Pienta et al., 1991). The work presented here is intended to summarize some of the evidence describing the 3-D organization of interphase nuclei and to correlate this evidence with results that show that the organization of interphase nuclei is dynamic and under the control of physiological stimuli.

II. Nuclear Rotation: Chromatin Motion in lnterphase Nuclei in Vitro

Motion of chromatin domains within interphase nuclei has been described for several cell types maintained in v i m . It is particularly pronounced in neurons, a cell type which is permanently arrested in interphase. When assayed by quantification of nucleolar displacement over time, this motion is termed “nuclear rotation.” In much of the work described here the quantification of nuclear rotation was derived from time-lapse photomicrographs and was expressed as the planar angle, subtended at the center of nuclei of circular geometry, through which nucleoli move over time. The term “nuclear rotation” itself is derived from the seminal work carried out by Pornerat (1953) and by Paddock and Albrecht-Buehler ( 1986a,b) and is based on the observation that intranuclear motion is most readily discernible and quantifiable in cells in which nucleoli are located near the nuclear periphery. As indicated later, the motion of chromatin structures is not restricted to circular trajectories, but occurs in three dimensions within the spatial confines of the nucleus. Therefore, it might be considered prudent to replace the traditional term “nuclear rotation” with the term “chromatin motion.” For this reason “nuclear rotation” will be retained where appropriate for the citations employed but will be replaced by the term “chromatin motion” where this latter terminology more appropriately describes the results.

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The first observations of nuclear rotation, which was seen as the motion of nucleoli in neuronal nuclei, were made by Murnaghan (1941) and by Nakai (1956) and were quantitatively analyzed by Pomerat and colleagues (1967) and Lodin and colleagues (1970). Nuclear rotation in non-neuronal cells was shown to occur in epithelial cell types (Pomerat, 1953; Leone et a!., 1955; Hinschke, 1956; Capers, 1960; Bourgeois et al., 1981). In some of these cell types, nuclear rotation during interphase is frequently either very slow or absent altogether, unless these cells are treated with agents which alter membrane permeability, such as monensin or phytohemagglutinin. In contrast, several neuronal types in uitro, at least while actively differentiating, invariably exhibit nuclear rotation. This observation speaks against the argument that nuclear rotation is related to events initiating mitosis (Bard et al., 1985), events which do not occur in differentiated neurons. Nuclei of many cell types and especially those of neurons maintained in uitro have a spherical geometry. It is unlikely, then, that the motion of nucleoli in these cells would be restricted to a particular nuclear plane; rather, it is likely that nucleolar motion extends throughout the karyoplasm. Indeed, dot-product vector analyses of consecutive XYZ positions of chromatin domains showed that nuclear rotation represents the motion of chromatin domains along curvilinear trajectories, occurring in three dimensions throughout the karyoplasm (De Boni and Mintz, 1986). Using living neurons in uitro whose nuclei were labeled with the DNA-specific dye 4,6-diarnidino-Zphenylindole (DAPI), this work also showed that displacement of chromatin domains is not restricted to motion of nucleoli but that it includes additional chromatin domains. Nuclear rotation occurs independently of the concurrent motion of juxtanuclear, cytoplasmic structures (Albrecht-Buehler, 1984; De Boni and Mintz, 1986; Hay and De Boni, 1991); this observation supports the hypothesis that the motion described as nuclear rotation is driven from within the nucleus (see the following discussion). We had proposed that nuclear rotation may function in the transposition of specific chromatin domains to transcriptionally active compartments (De Boni and Mintz, 1986; De Boni, 1988a; Fung and De Boni, 1988). Nuclear rotation would thus be expected to be intermittent rather than continuous since changes in gene expression are clearly synchronized in time with induced altered neuronal activity (Greenberg et al., 1986; Cole ef al., 1989). Subsequent work clearly showed that nuclear rotation is indeed saltatory and includes reversal in direction in all three spatial dimensions (De Boni and Mintz, 1986; Hay and De Boni, 1991). These saltatory aspects of motion are obscured when quantifying mean rates of nuclear rotation

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as the cumulative distance traversed by chromatin domains, such as phase dark nucleoli and major domains stained supravitally with DNA-specific, fluorescent dyes (Pomerat et af., 1967; De Boni and Mintz, 1986; Fung and De Boni, 1988; De Boni, 1988a; Schiffmann and De Boni, 1991). This paradigm provides information on mean rates of chromatin motion. However, it smoothes the actually saltatory nature of nuclear rotation, which is clearly evident in plots of the instantaneous position of chromatin domains versus time (Figs. 1,2a,b). The saltatory behavior of nuclear rotation is also evident in frequency power spectra (Fig. 2c) derived from fast fourier transform analyses of chromatin motion over time (Park and De Boni, 1991). This latter work with dorsal root ganglion neurons and additional work with PC12 cells in uitro (Figs. 1,2) clearly established nuclear rotation as a periodic motion of chromatin domains, within spatially restricted nuclear compartments (see the following discussion). In dorsal root neurons, such domains move at mean rates of 2.2 f 0.04 deg/min, with dominant power bands at frequencies ranging from 0.47 c/hr to 2.91 c/hr. Resonance in these power spectra, indicated by observations of significant power fractions at frequencies corresponding to multiples of the fundamental wavelength, suggests that nuclear rotation consists of forced harmonic motion, a motion under multiple levels of control. While such mechanisms of control remain enigmatic, it has been demonstrated that changes in intracellular calcium concentrations alter nuclear rotation rates (Fung and De Boni, 1988). It may thus be speculated that one level of control may be related to the cycling, free calcium levels reported to occur in nuclei of excitable cells (Przywara et af., 1991). Given that a large amount of the nuclear content of neurons in uitro exhibits saltatory motion, it is important to predict the location of the interface between moving karyoplasm and the relatively stationary cytoplasm. The presence of an extranuclear, cytoplasmic motor would result in motion of the nucleus in toto, including its envelope, as previously proposed. It has been argued (Albrecht-Buehler, 1984) that an interface for nuclear rotation between the outer nuclear membrane and the cytoplasm is most likely, because the evidence for stable connections between the endoplasmic reticulum and the outer nuclear membrane is uncertain. Rates of nuclear rotation increase in the presence of Monensin, whose action on the Golgi apparatus seems consistent with the production of breaks between the nuclear envelope and endoplasmic reticulum. This led to the conclusion that the ties between the nucleus and cytoplasm are dynamic (Albrecht-Buehler, 1984). Such a dynamic association between the nuclear envelope and the cytoskeleton is, however, not supported by results which show that the

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FIG. 1 A representative series of time-lapse, phase-contrast photomicrographs (10-min intervals between frames) of the nucleus of a PC12 cell in uitro, showing the phenomenon of nuclear rotation (chromatin motion). Note the pronounced changes in the intranuclear, spatial position of the nucleoli and the reversal of direction (g,h). Also note the changes in rate of motion per interval, indicating the saltatory nature of motion. The black dot marks a juxtanuclear, cytoplasmic structure, which remains relatively stationary while the nuclear content shows significant displacement. Magnification: 3000 X .

outer membrane of the nuclear envelope is intimately linked with intermediate filaments of the cytoskeleton (Georgatos and Blobel, 1987) and that the pattern of the juxtanuclear cytoskeleton is not dramatically different in cells with stationary nuclei and those that have rotating nuclei (Paddock and Albrecht-Buehler, 1986a,b).

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It is conceptually difficult to place the locus of the sliding interface for nuclear rotation between the outer nuclear membrane and the cytoplasm unless the filamentous structures between the nuclear envelope and cytoplasm are totally dynamic. Time-lapse video records show that nuclear rotation in dorsal root neurons in vitro and in PC12 cells (Fig. 1) does not include the motion of cytoplasmic organelles, such as cysternae of endoplasmic reticulum, mitochondria, and Nissl clusters in the juxtanuclear zone (Pomerat, 1953; Albrecht-Buehler, 1984; De Boni and Mintz, 1986). Moreover, juxtanuclear cytoplasmic structures located within fractions of a micrometer of the outer nuclear membrane remain stationary when chromatin is clearly seen moving past such structures, even when the intermediate filament network is disrupted by acrylamide (Hay and De Boni, 1991). This is not surprising in view of ultrastructral evidence which shows that the outer nuclear membrane of dorsal root sensory neurons is extensively continuous with membranes of the endoplasmic reticulum (De Boni, 1988a,b). The absence of resolvable changes in the morphology of the immediate juxtanuclear area during periods of significant motion of nucleoli supports the hypothesis that an interface between the outer nuclear membrane and the cytoplasm is unlikely. A relative motion between the membranes forming the nuclear envelope is also unlikely, as previously pointed out (Albrecht-Buehler, 1984; Paddock and Albrecht-Buehler, 1986a,b; De Boni, 1988a). A large number of nuclear pores connect inner and outer nuclear membranes, at densities of up to 18 pores per square micrometer of membrane in cerebral cortex cells (Lodin et al., 1978). There exists no evidence that pore complexes, organelles with an estimated, aggregate molecular weight of 10 x lo7 daltons (Davis and Blobel, 1986), undergo rapidly reversing cycles of disassembly and assembly, at rates compatible with observed chromatin motion. Additional constraints on motion between inner and outer nuclear membranes may be associated with their fusion to the spokes of nuclear pore complexes (Maul, 1977; Unwin and Milligan, 1982; Burke, 1990).

FIG. 2 The dynamics of motion of two nucleoli (solid and dashed lines, respectively) in two representative, binucleolate PC12 cells in uitro, (A) in the absence of nerve growth factor and (B) following differentiation by nerve growth factor (7 days, 100 ngfml). The angle between vectors is derived from dot-product vector analyses of the spatial positions of nucleoli within spherical nuclei (see the text and Park and De Boni, 1991). Saltatory motion is evident as changes in the rates of angular displacement over time (A,B). In the absence of nerve growth factor, the motion of both nucleoli is synchronous. This synchrony is also reflected in the similarity of frequency power spectra of motion (C) of the two nucleoli (solid and hatched bars, respectively) shown in A. In cells differentiated with nerve growth factor (B), such synchrony of motion between nucleoli is lost and the mean rate of motion is significantly decreased (note the different scale of the y-axis).

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An interface for motion between the lamina and the inner nuclear membrane is similarly unlikely, given the presence of specific, integral membrane proteins (Gerace er al., 1974, 1978) and lamin B, which anchor the lamina-pore complex to the inner nuclear membrane by a specific glycoprotein (Lebel and Raymond, 1984). Relative motion of the lamina along the inner nuclear membrane would require connections between these elements to be dynamic. Given the dimensions of nuclei of dorsal root neurons, together with measured rates of chromatin motion of 0.3 pm/min and more at the nuclear periphery, lamina sliding along the inner membrane would have to move through at least 3 pore diameters per minute. Ultrastructural evidence shows that the lamina is invariably interrupted in the regions underlying nuclear pores. Motion of the lamina relative to the inner nuclear membrane would therefore result in the presence of lamina underlying nuclear pores; this is never observed. As indicated above, disruption of the intermediate filament network by acrylamide results in a 4-fold increase in the mean rate of chromatin motion, within an apparently stationary envelope. This increase in the rate of chromatin motion occurs concurrently with significant changes in thickness of the nuclear lamina, as shown by ultrastructural examination of neurons exposed to acrylamide. Lamina thickness decreases significantly ( P < 0.0001) from 20.86 5.10 nm (n = 159) in controls to 16.05 2 4.37 nm ( n = 103) in nuclei of neurons treated with 4 mM acrylamide (Hay and De Boni, 1991). While acrylamide has actions on cell function other than disruption of intermediate filament bundles, including changes in phosphorylation of neurofilament protein (Howland and Alli, 1986), intermediate filaments and nuclear lamins share extensive alpha-core epitopes with intermediate filaments (McKeon er al., 1986). The reduced lamina thickness after acrylamide exposure indicates that such lamina epitopes may also be disrupted by acrylamide and that the associated increase in rates of nuclear rotation may be the result of removal of a damping constraint on chromatin motion, which is normally provided by an intact lamina. Together, the above arguments support the hypothesis that nuclear rotation represents the motion of chromatin domains within nuclei and not a motion of the nucleus in rum, including its envelope. In view of the controversy regarding the subcellular site of the interface of the observed motion, work was undertaken to determine whether nuclear rotation is associated with the motion of chromatin domains relative to each other (Park and De Boni, 1991).This work, using direct, quantitative analyses of the dynamics of nucleolar fusion during redifferentiation of living neurons in uitru, clearly showed that intranuclear domains may indeed move independently and relative to each other, although within spatially restricted nuclear territories. Moreover, the results showed that nucleolar fusion is invariably preceded by a significant burst in the rate of nuclear rotation.

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This work also provided additional evidence which shows that nuclear rotation is not restricted to motion of nucleoli. As nucleoli undergo fusion as a function of in uitro age and their number per nucleus decreases, the number of kinetochore signals, detected by immunocytochemistry , also decreases (De Boni et al. 1992; Park and De Boni, 1992). This decrease in the number of kinetochore signals is associated with a concomitant increase in the volume of the remaining signals detected. This observation is indicative of additional clustering and thus of motion of chromatin domains relative to each other. Together, the results indicate a clear correlation between the periodic and pulsatile nature of nuclear rotation and nucleolar fusion in neurons in uitro; they indicate that nucleolar fusion, clustering of specific chromatin domains, nuclear rotation, and differentiation are linked (Park and De Boni, 1991,1992). Moreover, the results support the hypothesis that nuclear rotation may be driven by an intranuclear motor, a hypothesis supported by additional evidence discussed later. We conclude, therefore, that the interface of chromatin motion must lie between the inner nuclear membrane on the outside and a component of chromatin on the karyoplasmic side. We have proposed that nuclear rotation may function in the transposition to transcriptionally competent compartments of those chromatin domains actively transcribed. This hypothesis is supported by the association of actively transcribed genes with specific intranuclear loci that are associated with “nuclear channels” (Hutchison and Weintraub, 1985), and by theoretical considerations of a phenotypically defined, albeit dynamic, three-dimensional structure of the cellular genome (Blobel, 1985). Observations of nuclear rotation in general, and of the pronounced motion of supravitally stained large chromatin masses which move in tandem with nucleoli (De Boni and Mintz, 1986) in particular, have been restricted to cells in uitro. Under these conditions nuclear rotation is especially pronounced during early stages of redifferentiation and may slow down or cease altogether in mature, fully differentiated neurons (Pomerat et al., 1967; Park and De Boni, 1991). It is not known whether pronounced motion of the nuclear contents occurs in neurons in situ, although an increasing body of indirect evidence indicates that such motion occurs in viuo during differentiation and adaptation to altered functional states, as decribed later.

111. Chromosome Topology in lnterphase Nuclei

During interphase, most of the chromatin within a nucleus is less condensed than it is during stages of the cell cycle associated with mitosis.

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As a result, individual chromosomes or components thereof, with the exception of nucleoli, cannot readily be discerned. Early work by Rabl (1885) indicated, however, that such interphase chromatin may retain a telophase-like arrangement of chromosomes, with centromeres and telomeres remaining associated with opposite poles of the nucleus. Evidence for this was confirmed in the specialized, polytene nuclei in salivary gland cells of Drosophila larvae (Agard and Sedat, 1983; Hochstrasser er al., 1986; Hochstrasser and Sedat, 1987a,b).In contrast, mammalian cell types frequently display a non-‘‘Rabl-type” chromosome topography (Moroi et al., 1981; Brinkley el al., 1986; Manuelidis, 1984b, 1985a; Haaf and Schmid, 1989, 1991; Haaf et al., 1990). With the advent of in situ hybridization, it has become possible to show, however, that different DNA sequences are not randomly positioned within interphase nuclei, but rather, that they are organized into a 3-D pattern which is cell type specific (Manuelidis, 1984a,b, 1985a,b; Manuelidis and Borden, 1988; Arnoldus et al., 1989; Billia and De Boni, 1991). Highly repetitive, nontranscribed chromosome domains such as centromeric satellite DNA (sDNA) (Jones, 1970; Pardue and Gall, 1970; Rae and Franke, 1972; Manuelidis, 1982, 1984b, 1985a,b; Joseph et al., 1989; Masumoto et al., 1989a,b) and telomeric DNA sequences (Mathog et al., 1984; Hochstrasser et al., 1986; Katsumoto and Lo, 1988; Rawlins and Shaw, 1990a,b; Billia and De Boni, 1991) become nonrandomly and reproducibly fixed to cell type-specific, intranuclear sites (Fig. 3). This rearrangement of patterns probably occurs during postmitotic, final differentiation (Manuelidis, 1984b). Similar results were obtained by immunocytochemistry used to detect kinetochore proteins associated with sDNA (Moroi et al., 1981; Earnshaw et al., 1984; Hadlaczky et al., 1986; Chaly and Brown, 1988; Holowacz and De Boni, 1991). These observations, together with the fact that such reproducible patterns are not observed in tumors of neurectodermal origin (Manuelidis, 1984a; Cremer et af., 1982) indicate that the observed patterns are related to the functional state of cells and possibly to cell type-specific gene expression. In fact, domains remote from centromeres, such as locus 1~36.3,for instance, may be dynamically and reversibly positioned into transcriptionally active regions in the nuclear interior without cytotypic orientation (Manuelidis and Borden, 1988). In this manner, the relatively fixed, repetitive centromeric and telomeric regions are envisioned to act as general organizing centers, still permitting other chromosome domains to occupy dynamic loci under control of transcriptional demands and possibly permitting efficient processing of both housekeeping and cell specific transcripts. The intranuclear position of those chromatin domains which are apparently fixed, as indicated by their cell type-specific, spatial position can,

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FIG. 3 Localization by in situ hybridization of satellite DNA in a neuron in the hippocampal CA3 region and in a neuron of cerebral cortical layer 111. (See the text and Billia et al., 1992). A,B. Phase-contrast micrographs of sections showing respective position (arrows) of the neuronal nuclei shown in a’ and b’. Two consecutive, optical sections (2-pm steps) show differences in the cell type-specific number and spatial positions of satellite DNA detected by irnmunofluorescence (a’””, CA3 neuron; b’,b”, cortical neuron). This is also evident in corresponding, computer-assisted 3-D reconstructions (a’ ”, b’ ”, sDNA is stippled, nucleoli are black) of the nuclei shown in a’ and b’. Magnification: A,B; bar= 100 wm. a”,a” ,b’ ,b” ; bar = 5 b m .

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however, be altered to some extent. Neurons within epileptic foci in human cerebral cortex show changes in the intranuclear, spatial positions of specific gene sequences, changes not detected in the same neuronal type in adjacent tissue, outside the epileptic focus (Borden and Manuelidis, 1988). This important finding provides a potential link between positioning of specific gene sequences and neuronal function. The nuclear content thus has been postulated to represent a dynamic system controlled physiological stimuli. A unifying theory for the function of differential chromosome patterns in different cell types has yet to be proposed. Moreover, it remains unclear whether the changes observed in the spatial distribution patterns of specific chromatin domains in interphase nuclei of cells which exhibit altered function are causal in altered function or are merely associative. Nevertheless, the emerging recognition that those physiological stimuli which result in activation of second messenger systems are also associated with rapid changes in gene expression may well provide a functional link between stimuli, altered gene expression, and the associated changes in the intranuclear, spatial position of specific DNA sequences.

IV. The Functional State of Cells and lnterphase Chromosome Patterns

As outlined earlier, a considerable body of evidence indicates that specific chromatin domains in interphase nuclei of cells in uitro may move within the global confines of interphase nuclei. In addition, chromosome patterns in cells in viuo are dynamic, either during differentiation when cells exhibit stage-specific, characteristic patterns of centromeric satellite DNA (Manuelidis, 1985a) or under conditions of altered functional states (Borden and Manuelidis, 1988). Together, these results indicate that rearrangement of the spatial positions of specific chromatin domains may depend upon transcriptional demands of the cell and may be taken to indicate the presence of a potential link between the positioning of specific gene sequences and cell function. Difficulties in interpretation arise when the rate of motion observed as nuclear rotation, which exceeds 0.4 pm/min under certain experimental conditions (Pomerat, 1953; De Boni and Mintz, 1986; Fung and De Boni, 1988; Hay and De Boni, 1991), is contrasted with the reproducible spatial positions of satellite and telomeric DNA sequences in neurons (Manuelidis, 1984a,b; Manuelidis and Borden, 1988; Billia and De Boni, 1991). The possibility must thus be considered that the pronounced saltatory motion of nuclear contents measured as nuclear rotation in cells in uitro

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in general and in neurons in uitro in particular may correlate with the redifferentiation of these cells which occurs following their transfer to in uitro conditions. In contrast, the more selective repositioning of chromatin domains in neurons in situ (Borden and Manuelidis, 1988) may reflect comparatively minor changes in chromatin organization, possibly associated with altered demands in gene expression. These considerations led to the hypothesis that changes in altered cell function in general and in gene expression in particular should be associated with changes in rates of chromatin motion, which is in turn associated with altered interphase chromatin patterns. The available evidence clearly shows that activation of membrane receptors, by neurotransmitters or their agonists, results in changes in gene expression within minutes, both in cells in uitro (Greenberg et al., 1986) and in functional, organotypic cell assemblies where neurotransmitters alter expression of specific neuronal “competence” genes of the proto-oncogene family (Douglas et al., 1988; Cole et al., 1989; Herrera and Robertson, 1990). Associations between activation of receptors by neurotransmitters and altered chromatin patterns have indeed been demonstrated. In dorsal root ganglion neurons in uitro, agents which alter neuronal gene expression, including neurotransmitters, also alter rates of chromatin motion. In these cells, chromatin motion is altered in a dose-dependent, reproducible manner by a-amino butyric acid (GABA), a neurotransmitter to which dorsal root ganglion neurons express receptors (Holz et al., 1986), and by nerve growth factor, with the changes in rates of motion occurring at the same time as the changes in gene expression (Fung and De Boni, 1988; De Boni, 1988a). While it has not been established which mechanism mediates the induced changes in chromatin motion, GABA can activate at least two receptor subtypes on dorsal root ganglion neurons. Of these, receptors of the GABA-A type directly control a chloride channel (Robertson, 1989) while GABA-B responses are mediated by a linking G-protein (Holz et al., 1986; Goh and Pennefather, 1989). It is thus possible that receptor activation and chromatin patterns are linked through a second messenger system-a hypothesis supported by results presented later. Analyses of the three-dimensional distribution of kinetochores by immunocytochemistry, and of centromeric satellite DNA sequences by in situ hybridization in nuclei of murine dorsal root ganglion neurons, showed that the changes in rates of chromatin motion observed in response to receptor activation by neurotrophic ligands were indeed associated with rearrangements of specific chromosome domains (Holowacz and De Boni, 1991). Moreover, in situ hybridization using a biotinylated mouse satellite DNA probe added further evidence for distinct, nonrandom patterns of interphase chromosomes in dorsal root sensory neurons.

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In these cells, kinetochores (Holowacz and De Boni, 1991) as well as satellite DNA sequences (Billia and De Boni, 1991) occur in clusters associated with nucleoli and throughout the nucleoplasm, with kinetochores colocalizing with the corresponding satellite DNA in a distinct orientation with respect to the loci of the satellite DNA. Specifically, nucleolus-associated kinetochores appose the corresponding satellite DNA at the interface between the satellite DNA and the nucleolar border. Nucleoplasmic kinetochores consistently occupied a 5-pm distance from the nuclear center which represented 70% of the spherical nuclear radius. The corresponding satellite DNA loci was frequently associated with the nuclear membrane, with kinetochores located on the surface of the satellite DNA that faced the nuclear interior. Such specific kinetochore-satellite DNA associations suggest a role for kinetochores in satellite DNA organization within the interphase nucleus. However, GABA induced a significant reorganization in the spatial distribution of kinetochores. Following exposure to GABA, the total number of kinetochore signals decreased, compared with controls. This significant change was associated with a decrease in the number of nucleoplasmic signals, that is, a decrease of those signals situated in the spatial domain between the nucleolus and the nuclear membrane. The effect occurred in a dose dependent manner, upon a 1-hr exposure to 0 . 5 , 5 , or 10 mM GABA, and was interpreted to indicate that GABA either induces additional clustering of kinetochores within the nucleoplasmic compartment or, alternatively, that some kinetochores within the nucleoplasmic compartment are relocated to those clusters already associated with the nucleolar periphery. Such a reorganization has been previously described as occurring in mouse Sertoli cells after nucleolar activation (Haaf el al., 1990), and in human lymphocytes where exposure to phytohemagglutinin results in the recruitment of ribosomal DNA sequences into actively transcribing nucleolar organizer regions (Wachtler e l al., 1986). Together the results indicate that reorganization of chromatin at the supramolecular level may occur in response to changes in the physiological state of cells. Alternatively, such reorganization may occur in association with changes in the state of cellular differentiation which are needed to accommodate a new transcriptional state. A body of evidence thus indicates that rearrangement of chromosome patterns may occur in cells in uitro as well as in cells in uivo under pathological conditions (Borden and Manuelidis, 1988). To ultimately assign a functional role to these phenomena, however, it would be necessary to show in intact cell assemblies that changes in the organization of chromatin domains within neuronal interphase nuclei may occur in response to physiological stimuli.

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The hippocampal slice preparation is considered an excellent model for studying of cellular interactions at levels which range from electrophysiology to detailed studies of molecular aspects associated with intercellular signaling. In this preparation, brief electrical stimulation of the perforant path with high-frequency pulses produces a long-lasting increase in the amplitude of the population spike, a phenomenon referred to as long-term potentiation (LTP) (Bliss and Lomo, 1973; Bliss and Gardner-Medvin, 1973). The long duration of LTP, together with the proposed role of the hippocampus in memory (Lynch and Baudry, 1984; Thompson, 1985; Teyler and DiScenna, 1985,1987), has led to the use of LTP as a model in studying the cellular changes associated with learning and memory. Some of these changes include the expression of “immediately early” genes (Cole et al., 1989; Dragunow et al., 1989a,b), RNA synthesis (Goelet et af., 1986), and de n o w protein synthesis (Abraham and Otani, 1989; Frey et af., 1988; Krug et al., 1984; Montarolo et al., 1985, 1986). In fact, LTP is most sensitive to inhibition of protein synthesis during or immediately following stimulation (Stanton and Sarvey , 1984). Inhibition of protein synthesis also prevents LTP-associated changes in synaptic morphology (Petit, 1988), a finding which supports a role for de nouo protein synthesis in LTP. In a test of the hypothesis that a change in compartmentalization of specific chromatin domains is associated with an altered functional state and is not restricted to isolated neurons in v i m , the topology of sDNA sequences was examined by in siru hybridization in conjunction with three-dimensional reconstruction in nuclei of hippocampal CA1 pyramidal neurons exhibiting LTP, and in neurons exposed to bath-applied N-methylD-aspartate (NMDA). This work showed that LTP is associated with a rearrangment of interphase chromatin, indicated by enhanced clustering of satellite DNA sequences (Billia er al., 1992), a clustering similar to that induced in dorsal root ganglion neurons exposed to GABA in uitro (Holowacz and De Boni, 1991). Such clustering was clearly restricted to that neuronal population which exhibited LTP and was absent in dentate neurons within the same slice-neurons which did not exhibit LTP with the experimental paradigm used. Clustering of sDNA sequences occurred regardless of whether LTP was induced by electrical stimulation or hippocampal CAI neurons were activated by NMDA. Moreover, the observation that the effect of NMDA was clearly abolished when the protein kinase inhibitor H-7 was coapplied with NMDA (Billia et al., 1992) suggests a link, through a second messenger system, between agonist-mediated receptor activation and chromatin pat terns.

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The cellular response to extracellular stimulation requires signal transduction to the nucleus via second and third messenger systems, and a recognition mechanism by which a subset of genes can be activated or repressed. Evidence exists which shows that calcium may be part of such a system. Changes in intracellular calcium concentrations alter rates of chromatin motion (Fung and De Boni, 1988) and it has been reported that a stimulated rise in intracellular calcium is greater in the nucleus than in the cytoplasm (Przywa e? al., 1991). These results add to a body of evidence which shows that the activation of the NMDA receptor leads to an increase in intracellular calcium (Kuba and Kumamoto, 1990; Lynch e? at., 1983; MacDermott e? at., 1986). This increase triggers events which lead to consolidation of LTP through protein kinase C, which in turn activates phosphorylation of nuclear transcription factors (Schonthal, 1990). Together, these observations of selective chromatin rearrangments in cells within complex neural networks, such as the hippocampal slice preparation, suggest that chromatin motion is not an event restricted to isolated cells in vitro. They also show that changes in interphase chromosome patterns are induced by physiological stimuli and are possibly mediated by second messenger systems, as postulated earlier.

V. An lntranuclear Motor: Contractile Proteins in lnterphase Nuclei

The mechanisms for generating the motive force involved in chromatin motion and rearrangement are unknown. It had previously been suggested that chromatin motion in cells may be driven by an extranuclear, cytoplasmic, actin-based motor (Leone et al., 1955; Paddock and AlbrechtBuehler, 1986a). This was based upon the demonstration that cytoplasmic actin filaments in 3T3 cells undergo reorganization at the same time as changes in the rates of chromatin motion. This hypothesis is, however, contradicted by the repeatedly reported observation that chromatin motion may occur independently of concurrent motion of juxtanuclear, cytoplasmic structures (Albrecht-Buehler, 1984; De Boni and Mintz, 1986; Hay and De Boni, 1991) and the demonstration that specific chromatin domains such as nucleoli may move relative to each other (see earlier discussion and Park and De Boni, 1991). Moreover, dorsal root sensory neurons in uitro, which exhibit significant chromatin motion, have no, or only trace amounts of, filamentous actin within the somata (Hay and De Boni, 1991). While antiactin antibody detected actin

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throughout somata and neuronal processes, fluorescein-labeled phalloidin, which is selective for f-actin (Estes et al., 1981), failed to decorate actin within somata. It did, however, react with f-actin in neuronal processes and in the cytoplasm of underlying non-neuronal cells in the same cultures (Hay and De Boni, 1991). The absence of f-actin within neuronal somata thus makes it unlikely that cytoplasmic actin acts on chromatin motion. This leads to the conclusion that there is adistinct possibility that the motor postulated to drive chromatin motion may be located in the subcellular compartment enclosed by the inner nuclear membrane-the karyoplasm. Biochemical analyses clearly show that actin and myosin are constituents of the nonhistone protein fraction of interphase nuclei and of isolated nuclear matrices (Ohnishi et al., 1963; 1964; Jockusch et al., 1974; Hauser et al., 1975; LeStourgeon et al., 1975; Clark and Merriam, 1977; Fukui, 1978; Bremer et al., 1981; Capco et al., 1982; Nakayasu and Ueda, 1983, 1984, 1985, 1986; Brasch, 1990), including a chemically distinct ic.,o form of actin found in isolated nuclei (Bremer et al., 1981; Kumar et al., 1984). A significant fraction of actin in interphase chromatin has been shown to exist in the f-actin form (Clark and Rosenbaum, 1979). This is supported by ultrastructural studies employing phalloidin-gold complexes, which have shown that the nucleoplasm of Physarum polycephalum cells is the most heavily labeled of all cellular compartments (Lachapelle and Aldrich, 1988). Both actin and myosin have been shown to be a constituent of the nuclear matrix (Douvas et al., 1975; Wunderlich and Herlan, 1977; LeStourgeon, 1978; Fukui, 1978). Actin has been shown to be associated with the nucleolus (Hauser et al., 1975), and a specific association of myosin I has been demonstrated with the inner nuclear membrane (Rimm and Pollard, 1989). While the function of these proteins is open to speculation, it was suggested that they might function in changes of nuclear shape (Horowitz et al., 1986), in chromatin condensation (Rungger et al., 1979), in nucleocytoplasmic transport (Schindler and Jiang, 1986; Ueyama et al., 1987), in gene expression (LeStourgeon et al., 1975), and as a transcription initiation factor (Egly et al., 1984; Scheer et al., 1984; Ankenbauer et al., 1989). Criticisms that nuclear actin is in equilibrium with cytoplasmic actin and that it is without function (Goldstein et al., 1977) were challenged by the finding of a form of actin specific to nuclei (Bremer et al., 1981; Kumar et al., 1984). In this work, analyses of actin derived from nuclear fractions indicated that nuclear actin, while similar to cytoplasmic actin, is chemically distinct. These same authors also found that nuclear and skeletal muscle actins are similar and have suggested that nuclear actin has the

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capacity to interact with myosin and thus has a potential function in force generation. Evidence for a second contractile protein within nuclei, myosin, which is associated with the nuclear envelope (Berrios and Fisher, 1986), was confirmed by the purification and characterization of a nuclear actinbinding protein in Acanthamoeba (Rimm and Pollard, 1989). This nuclear actin-binding protein is antigenically related to myosin I, although it does not exhibit most of the characteristics that define myosins, such as actinactivated ATPase and ATP-sensitive actin binding. Nuclear actin-binding protein does, however, exhibit binding to DNA, while actin, myosin I, and alpha-actinin do not. This nuclear actin-binding protein represents the first actin-binding protein to be localized exclusively to the nucleus, although its function is not yet known. Rimm and Pollard (1989) propose that this protein may attach DNA to the actin-containing nuclear matrix, as previously postulated (Armbruster et al., 1983; Nakayasu and Ueda, 1983,1985,1986;Crowley and Brasch, 1987). Despite this accumulated body of biochemical evidence which localized actin to interphase nuclei at relatively high concentrations, its precise intranuclear localization remained obscure. Recent and continuing work using laser confocal immunocytochemistry and immunogold techniques at the ultrastructural level, has shown, however, that actin and myosin occur as aggregates in nuclei of intact neurons (Milankov el al., 1991; Milankov and De Boni, 1993). Specifically, actin-like and myosin-like antigens were demonstrated to be present as aggregates in nuclei of fixed but intact neurons maintained in v i m , as well as in nuclei of neurons within intact sensory ganglia. Threedimensional reconstruction from confocal images as well as ultrastrucural localization place some of these aggregates at the nucleolar periphery, from where actin also extends into the nucleoplasm along chromatin fibers. Labeling with fluorescent-palloidin shows that only those actin aggregates associated with the nucleolar periphery contain sufficient f-actin to be detectable by phalloidin. It is of interest to note that use of antibodies against subclasses of alphaactin isoforms indicated that nuclear actin is more closely related to an alpha-sarcomeric isoform than to smooth muscle actin. This was indicated by decoration of nuclear actin aggregates by an antibody to alphasarcomeric actin, including a highly specific monoclonal antibody, but not by a highly specific antibody against a synthetic decapeptide unique to smooth-muscle actin. In contrast, actin in neuronal cytoplasm was labeled with antibody to smooth-muscle actin. Given the evidence for the occurrence of actin as well as myosin aggregates within interphase nuclei, it remained to be demonstrated that the

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intranuclear f-actin present has the capability to functionally interact with myosin. The existence of such an interaction is indicated by the ultrastructural localization of arrowhead patterns in nuclei of mildly fixed and perme1991), abilized neurons labeled with heavy meromyosin (Milankov et d., or by heavy meromyosin conjugated to colloidal gold (Amankwah and De Boni, 1994). Such labeling resulted in the decoration of putative intranuclear microfilaments typical of actin decorated by heavy meromyosin (Tshikawa et al., 1969; Schloss et al., 1977). In summary, the demonstrated presence of actin and myosin aggregates in interphase nuclei, together with the observation that nuclear actin may interact in a typical manner with a fragment of myosin, supports the hypothesis that these macromolecules may be the motor responsible for chromatin motion and the formation of chromatin patterns.

VI. Conclusions

The combined results of the work decribed in this chapter show that interphase nuclei exhibit cell type-specific, reproducible compartmentalization of specific DNA sequences such as sDNA, transfer DNA, and others, as well as compartmentalization of nucleoli and kinetochores. The results also indicate that such compartmentalization is dynamic, both during differentiation and during changes in functional state induced by physiological stimuli. The hypothesis that the spatial, intranuclear sites of pre-mRNA processing and transcription may differ between cell types and that the functions of small nuclear ribonucleoproteins (snRNPs) and actin within interphase nuclei may be related is supported by recent work by Sahlas and co-workers. In fact, rat pheochromocytoma cells, when differentiated into neuron-like cells with nerve growth factor, exhibit a snRNP topology which is clearly distinct from that which is observed in the same cell type before exposure to nerve growth factor. Moreover, based on quantitative evidence of changes in the extent of the association between snRNP and actin aggregates, it may be concluded that the functions of snRNPs and intranuclear actin may indeed be dynamically related (Sahlas et al., 1993). While the nature of the intranuclear motor which translocates specific chromatin domains during interphase remains enigmatic, the presence of intranuclear aggregates of actin and myosin, together with the demonstration that nuclear actin may functionally interact with myosin, suggests that actomyosin complexes may provide the motive force required for chromatin motion and thus patterning of chromatin in interphase nuclei. The demonstration of multiple intranuclear aggregates of such actomyosin

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complexes could also account for the forced harmonic motion described earlier, which is associated with the postulated multiple levels of control (see Section 11). The mechanisms that control chromatin motion remain enigmatic. Changes in intracellular calcium concentrations do, however, alter rates of nuclear rotation (Fung and De Boni, 1988). It may thus be speculated that one level of control may be related to the cycling, free calcium levels reported to occur in nuclei of excitable cells (Przywara et al., 1991). A vast amount of DNA within a nucleus must be dynamically organized and requires a structural system to coordinate the expression of genes which make life possible (Pienta et al., 1991). The definition of exact functional associations between the supramolecular organization of chromatin and cell function in general and gene expression in particular requires much additional effort but may ultimately lead to a critical test of the hypothesis that there is a causal link between chromatin motion and its organization into a pattern which permits gene expression.

References Abraham, W. C., and Otani, C. (1989). Soc. Neurosci. Abstr. 15, 85. Agard, D. A., and Sedat, J. W. (1983). Nature (London) 302, 676-681. Albrecht-Buehler, G.(1984). In “Cancer cells. The Transformed Phenotype” (A. J. Levine, G. F. Vande Woude, W. C. Topp and J. D. Watson, eds.), Vol. I , pp. 87-96. Cold Spring Harbor, NY. Amankwah, K. S . , and De Boni, U. (1994). Exp. Cell. Res. 210, (in press). Ankenbauer, T., Kleinschmidt, J. A., Walsh, M. J., Weiner, 0. H., and Francke, W. W. (1989). Nature (London) 342, 822-825. Armbruster, B. L., Wunderli, H., Turner, B. M., Raska, I., and Kellenberger, E. (1983). J . Histochem. Cytochem. 31, 1385-1393. Amoldus, E. P. J., Peters, A. C. B., Bots, G. T. A. M., Raap, A. K., and van der Ploeg, M. (1989). Human Genet. 83, 231-234. Bard, F., Bourgeois, C. A., Costagliola, D., and Bouteille, M. (1985). Biol. Cell54, 135-142. Barr, M. L., and Bertram, E. G. (1949). Nature (London) 163, 676-677. Barr, M. L., and Bertram, E. G. (1951). J . Anat. 85, 171-181. Benios, M., and Fisher, P. A. (1986). J. Cell Biol. 103, 711-724. Billia, F., and De Boni, U. (1991). J. Cell Sci. 100, 219-227. Billia, F., Baskys, A., Carlen, P., and De Boni, U. (1992). Mol. Brain Res. 14, 101-108. Bliss, T. V. P., and Gardner-Medwin, A. K. (1973). J. Physiol. 232, 357-374. Bliss, T. V. P., and Lomo, T. (1973). J. Physiol. 232, 331-356. Blobel, G. (1985). Proc. Natl. Acad. Sci. U.S.A. 82, 9527-9529. Borden, .I. and , Manuelidis, L. (1988). Science (Washington, D.C.) 242, 1687-1691. Bourgeois, C. A., Hemon, D., Beaure D’Augeres, C., Robineaux, R., and Bouteille, M. (1981). B i d . Cell 40, 229-232. Brasch, K. (1990). Biochem. Cell Biol. 68,408-426. Bremer, J. W., Busch, H., and Yeoman, L. C. (1981). Biochem. 20, 2013-1017. Brinkley, B. R., Brenner, S. L., Halk, J. M., Tousson, A., Balczon, R. D., and Valdivia, M. (1986). Chromosoma 94, 309-317.

THE INTERPHASE NUCLEUS AS A DYNAMIC STRUCTURE

169

Rurke, B. (1990). Current Opin. Cell Biol. 2, 514-520. Capco, D. G., Wan, K. M., and Penman, S. (1982). Cell (Cambridge, Mass.) 29, 847-858. Capers, C. R. (1960). J. Biophys. Biochem. Cytol. 7 , 559-566. Chaly, N., and Brown, D. L. (1988). J . Cell Sci. 91, 325-335. Clark, T. G., and Memam, R. W. (1977). Cell (Cambridge, Mass.) 12, 883-891. Clark, T. G., and Rosenbaum, J . L. (1979). Cell (Cambridge, Mass.) 18, 1101-1108. Cole, A. J., Saffen, D. W., Baraban, J. W., and Worley, P. F. (1989). Nature (London) 340,474-476.

Cremer, T., Cremer, C., Schneider, T., Baumann, H., Hens, L., and Kirsch-Volders, M. (1982). Human Genet. 62,201-209. Crowley, K. S., and Brasch, K. (1987). Cell Biol. Intl. Rep. 11, 537-546. Davis, L. I., and Blobel, G. (1986). Cell (Cambridge, Mass.) 45, 699-709. De Boni, U. (1988a). Anticancer Res. 8, 885-898. De Boni, U. (1988b). Biol. Cell 63, 1-8. De Boni, U., and Mintz, A. H. (1986). Science (Washington, D.C.) 234, 863-866. De Boni, U., Milankov, K., Amankwah, K. S.,and Park, P. (1992). Proc. 19th Ann. Meeting, EMSA pp. 562-563. Douglas, R. M., Dragunow, M., and Robertson, H. S. (1988). Mol. Brain Res. 4, 259-262. Douvas, A. S., Hamngton, C. A,, and Bonner, J. (1975). Proc.Nat. Acad. Sci. U.S.A. 72, 3902-3906.

Dragunow, M., Abraham, W. C., Goulding, M., Mason, S. E., Robertson, H. A., and Faull, R. L. M. (1989a). Neurosci. Lett. 101, 274-280. Dragunow, M., Cunie, R. W., Faull, R. L. M., Robertson, H. A., and Jansen, K. (1989b). Biobehau. Rev. W, 301-313. Earnshaw, W. C., Halligan, N., Cooke, C. A., and Rothfield, N. (1984). J. Cell Biol. 98, 352-357.

Egly, J. M., Miyamoto, N. G., Moncollin, V., and Chambon, P. (1984). EMBO J . 3, 2363-2371.

Estes, J. E., Seldon, L. A., and Gershman, L. C. (1981). Biochem. 20, 708-712. Frey, U., Krug, M., Reymannn, K. G., and Matthies, H. (1988). Brain Res. 452, 57-65. Fukui, Y. (1978). J. Cell Biol. 76, 146-157. Fung, L. C., and De Boni, U. (1988). Cell Motil. Cytoskel. 10, 363-373. Georgatos, S. D., and Blobel, G. (1987). J . Cell Biol. 105, 117-125. Gerace, L., Comeau, C., and Benson M. (1974). J . Cell Sci. (Suppl. 1) 137-160. Gerace, L., Blurn, A., and Blobel, G. (1978). J. Cell Biol. 79, 546-566. Goelet, P., Castellucci, V. F., Schacher, S., and Kandel, E. R. (1986). Nature (London) 322,419-422.

Goh, J. W., and Pennefather, P. S. (1989). Science (Washington, D.C.)244,980-983. Goldstein, L., Rubin, R., and KO, C. (1977). Cell (Cambridge, Mass.) 12, 601-608. Greenberg, M.E., Greene, L. A., and Ziff, E. B. (1985). J . B i d . Chem. 260, 14101-14110. Greenberg, M. E., Ziff, E. B., and Greene, L. A. (1986). Science (Washington, D.C.) 234, 80-83.

Haaf, T., and Schrnid, M. (1989). Human Genet. 81, 137-143. Haaf, T., and Schmid, M. (1991). Exp. Cell Res. 192, 325-332. Haaf, T., Grunenberg, H., and Schmid, M. (1990). Exp. Cell Res. 187, 157-161. Hadlaczky, Gy., Went, M.,and Ringertz, N. R. (1986). Exp. Cell Res. 167, 1-15. Hauser, M., Beinbrech, G., Groeschel-Stewart, U., and Jockusch, B. M. (1975). Exp. Cell Res. 95, 127-135.

Hay, M., and De Boni, U. (1991). Cell Motil. Cytoskel. 18, 63-75. Herrera, D. G., and Robertson, H. A. (1990). Neuroscience 35, 273-281. Hinschke, E. (1956). 2.Zellfonsch 43, 526-542. Hochstrasser, M.,and Sedat, J. W. (1987a). J. Cell Biol. 104, 1455-1470.

170

UMBERTO DE BONl

Hochstrasser, M., and Sedat, J. W. (1987b). J. Cell Biol. 104, 1471-1483. Hochstrasser, M., Mathog, D., Gruenbaum, Y., Saumweber, H., and Sedat, J. (1986). J . Cell. Biol. 102, 112-123. Holowacz, T., and De Boni, U. (1991). Exp. Cell Res. 197, 36-42 Holz, G. G., Rane, S. G., and Dunlap, K . (1986). Nature (London) 319, 670-672. Horowitz, S. B., Tluczek, L. J. B., and Paine, B. L. (1986). J . Cell Biol. Abstr. 103, 46a. Howland, R. D., and Alli, P. (1986). Brain Res. 363, 333-339. Hutchison, N., and Weintraub, H. (1985). Cell (Cambridge, Mass.) 43, 171. Ishikawa, H., Bischoff, R., and Holtzer, H. (1969). J . Cell Biol. 43, 312-328. Jockusch, B. M., Becker, M., Hindennach, I., and Jockusch, H. (1974). Exp. Cell. Res. 89,241-246. Jones, K.W. (1970). Nature (London) 225, 912-915. Joseph, A., Mitchell, A. R., and Miller, 0. J. (1989). Exp. Cell Res. 183, 494-500. Katsumoto, M., and Lo, C. W. (1988). J. Cell Sci. 90, 193-199. Krug, M., Lossner, B., and Ott, T. (1984). Brain Res. Bull. 13, 39-42. Kuba, K., and Kumamoto, E . (1990). Prog. Neurobiol. 34, 197-269. Kumar, A., Razinddin, Finlay, T. H., Thomas, J. O., and Szer, W. (1984). Biochem. 23, 6753-6757. Lachapelle, M., and Aldrich, H. C. (1988). J . Histochem. Cytochern. 36, 1197-1202. Lebel, S., and Raymond, Y. (1984). J . Biol. Chem. 259, 2693-2696. Leone, V., Hsu, T . C., and Pomerat, C. M. (1955). Z. Zellforsch 41, 481-492. LeStourgeon, W. M. (1978). In “The Cell Nucleus” Vol. VI, Part C pp. 305-326. Academic Press, New York. LeStourgeon, W. M., Forer, A , , Yang, Y. Z., Bertram, J. S., and Rusch, H. P. (1975). Biochem. Biophys. Acta. 379, 529-552. Lodin, Z., Booher, J., and Kasten, F. H. (1970). Exp. Cell Res. 60, 27-39. Lodin, Z., Blumajer, J., and Mares, V. (1978). Acta Histochem. 63, 74-79. Lynch, G., Larson, J., Kelson, S., Bamonuevo, G., and Schottler, F. (1983). Nature (London) 305, 719-721. Lynch, G., and Baudry, M. (1984). Science (Washington, D . C . ) 224, 1057-1063. MacDermott, A. B., Mayer, M. L., Westbrook, G. L., Smith, S. J., and Barker, J. L. (1986). Nature (London) 321, 519-522. Manuelidis, L. (1982). In “Genome Evolution” ( G . A. Dover and R. B. Flavell, eds.), pp. 263-285. Academic Press, London. Manuelidis, L. (1984a). J . Neuropath. Exp. Neurol. 43, 225-241. Manuelidis, L. (1984b). Proc. Natl. Acad. Sci. U.S.A. 81, 3123-3127. Manuelidis, L. (1985a). Ann. N. Y. Acad. Sci. 450, 205-221. Manuelidis, L. (1985b). Human Genet. 71, 288-293. Manuelidis, L., and Borden, J. (1988). Chromosoma 96, 397-410. Masumoto, H., Masukata, H., Muro, Y., Nozaki, N., and Okazaki, T. (1989a). 1. Cell Biol. 109, 1963-1973. Masumoto, H., Sugimoto, K., and Okazaki, T. (1989b). Exp. Cell Res. 181, 181-196. Mathog, D., Hochstrasser, M., Gruenbaum, Y., Saumweber, H., and Sedat, J. W. (1984). Nature (London) 308, 414-421. Maul, G. G. (1977). Int. Rev. Cytol. Suppl. 6, 76-186. McKeon, F. D., Kirschner, M. W., and Caput, D. (1986). Nature (London) 319, 463. Milankov, K., Amankwah, K. S., and De Boni, U. (1991). J . CellBiol. Abstr. 115, 95a. Milankov, K., and De Boni, U. (1993). Exp. Cell Res. 209, 189-199. Montarolo, P. G., Castellucci, V. F., Kandel, E. R., Goelet, P., and Schacher, S. (1985). SOC.Neurosci. Abstr. 15, 795. Montarolo, P. G., Goelet, P., Castellucci, V. F., Morgan, J., Kandel, E. R., and Schacher, S. (1986). Science (Washington, D.C.) 234, 1249-1254.

THE INTERPHASE NUCLEUS AS A DYNAMIC STRUCTURE

171

Moroi, Y., Hartman, A. L., Nakane, P. K., and Tan, E. M. (1981). J.CellBio1. 90,254-259. Murnaghan, D. P. (1941). Anat. Rec. 81, 183-203. Nakai, J. (1956). A m . J. Anat. 99, 81-89. Nakayasu, H., and Ueda, K. (1983). Exp. Cell. Res. 143, 55-62. Nakayasu, H., and Ueda, K. (1984). Cell Struct. Funct. 9, 317-325. Nakayasu, H., and Ueda, K. (1985). Cell Struct. Funct. 10, 305-309. Nakayasu, H., and Ueda, K. (1986). Exp. Cell Res. 163, 327-336. Ohnishi, T., Kawamura, H., and Yamamoto, T. (1963). J. Biochem. (Tokyo) 54, 298-300. Ohnishi, T., Kawamura, H., and Tanaka, Y. (1964). J. Biochem. (Tokyo) 56,6-15. Paddock, S. W., and Albrecht-Buehler, G. (1986a). Exp. Cell Res. 163, 525-538. Paddock, S. W., and Albrecht-Buehler, G. (1986b). Exp. Cell Res. 166, 113-126. Pardue, M. L., and Gall, J. G. (1970). Science (Washington, D.C.)168, 1356-1358. Park, P. C., and De Boni, U. (1991). Exp. Cell Res. 197, 213-221. Park, P. C., and De Boni, U. (1992). Exp. Cell Res. 203, 222-229. Petit, T. L. (1988). In “Neural Plasticity: A Lifespan Approach.” (T. L. Petit and G. 0. Ivy. eds.). pp. 201-234. Alan R. Liss., New York. Pienta, K. J., Getzenberg, R. H., and Coffey, D. S. (1991). Critical Reviews in Eukaryotic Gene Expression, 1, 355-385. Pomerat, C. M. (1953). Exp. Cell Res. 5 , 191-196. Pomerat, C. M., Hendelman, W. J., Raiborn, C. W., and Massey, J. F. (1967). In “The Neuron” (H. Hyden, ed.), pp. 119-178. Elsevier, New York. Przywara, D. A., Bhave, S. V., Bhave, A., Wakade, T. D., and Wakade, A. R. (1991). FASEB J . 5,217-222, Kabl, C. (1885). In “Morphologisches Jahrbuch” (C. Gegangaur, ed.), vol. 10, pp. 214-330. Rae, P. M. M., and Franke, W. W. (1972). Chromosoma 39, 443-456. Rawlins, D. J., and Shaw, P. J. (1990a). J. Micros. 157, 83-89. Kawlins, D. J., and Shaw, P. J. (1990b). J . Cell B i d . Absrr. 111, 504a. Robertson, B. (1989). J . Physiol. 411, 285-300. Rimm, D. L., and Pollard, T. D. (1989). J. Cell Biol. 109, 585-591. Kungger, D., Rungger-Brandle. E., Chaponnier, C., and Gabbiani, G. (1979). Nature (London) 282, 320-321. Sahlas, D. J., Milankov, K., Park, P. C., and De Boni, U. (1993). J. Cell Sci. 105,347-357. Scheer, U., Hinssen, H., Franke, W. W., and Jockusch, B. M. (1984). Cell (Cambridge, Mass.) 39, 111-122. Schiffmann, D., and De Boni, U. (1991). Mural. Res. 246, 113-122. Schindler, M., and Jiang, L. W. (1986). J . Cell Biol. 102, 859-862. Schloss, J. A . , Milsted A., and Goldman, R. D. (1977). J . Cell Biol. 74, 794-815. Schonthal, C. A. (1990). Cell Signal. 2, 215-225. Stanton, P. K . , and Sarvey, J. M. (1984). J. Neurosci. 4, 3080-3088. Teyler, T. J., and DiScenna, P.(1985). Neurosci. Biobehau. Res. 9, 377-389. Teyler, T. J., and DiScenna, P. (1987). Ann. Reu. Neurosci. 10, 131-161. Thompson, K. F. (1985). Science (Washington, D . C . ) 233, 941-947. UeYama, H., Nakayasu, H . , and Ueda, K. (1987). Cell Biof. In?. Rep. 11, 671-677. Unwin, P. N. T., and Milligan, R. (1982) J. Cell Biol. 93. 63-75. Wachtler, F., Hopman, A. H. N., Wiegant, J., and Schwarzacher, H. G. (1986). Exp. Cell Res. 167, 227-240. Wunderlich, F., and Herlan, G. (1977). J. Cell Biol. 73, 271-278.