Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, PA 15213, USA
* Author for correspondence (e-mail: csink{at}andrew.cmu.edu
Accepted 14 December 2004
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Nuclear organization, Chromatin dynamics, Differentiation, Heterochromatin
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
What influences this nuclear reorganization? One answer is that chromatin structure itself dictates the position of a chromosomal region: chromatin with intrinsic or induced heterochromatic properties is localized to a specific region of the nucleus while chromatin in a euchromatic state is localized elsewhere. An example of this was found in Drosophila where the brownDominant (bwD) allele contains a 1.6 Mb insertion of heterochromatin into the distal euchromatin of the right arm of chromosome 2 (2R) (Platero et al., 1998). Flies heterozygous for bwD exhibit a variegated eye phenotype, showing that the wild-type gene on the homologous chromosome is also inactivated. Fluorescent in situ hybridization (FISH) on central nervous system (CNS) cells dissected from third instar larvae heterozygous for bwD showed that the bw locus associates with centric heterochromatin of the second chromosome (2Rh) (Csink and Henikoff, 1996
; Dernburg et al., 1996
). It is thought that somatic pairing of the homologous interphase chromosomes (a phenomenon commonly found in almost all nuclei of Dipteran insects) allows the bwD allele to drag the wild-type homolog closer to heterochromatin, resulting in silencing of the wild-type gene.
Nuclei in higher organisms are known to undergo extensive changes in organization as they progress through both the cell cycle and development (for a review, see Francastel et al., 2000). Thus, it is not surprising that positioning of the bwD heterochromatic insertion varies over the cell cycle. The association between bwD and 2Rh is not apparent until at least 5 hours into G1. If the length of G1 in a specific cell type is sufficient for bwD-2Rh association to form, the association is broken at the beginning of the subsequent S phase. No association is formed during G2 (Csink and Henikoff, 1998
). The first 13 embryonic nuclear cycles contain only S and M phases. Consistent with this, the bwD locus is not associated with 2Rh in nuclei from cycle 13 embryos (Dernburg et al., 1996
). These studies suggest that silencing caused by interaction with heterochromatin or positioning in a heterochromatic compartment may not be possible until well into the G1 phase of the cell cycle. Studies in mammalian cells have also found that association of some silenced loci with pericentric heterochromatin occurs during G1 (Brown et al., 1999
).
The larval CNS consists of a heterogeneous population of cells that are rapidly cycling, differentiating or are terminally differentiated. The changes in nuclear organization seen in differentiated cells may either be regulated by a specific event during differentiation or could merely be a consequence of increased time spent in the G1/G0 phase of the cell cycle. Even though heterochromatic associations occur within a population of cycling cells, the possibility that the association between bwD and 2Rh is regulated by a specific event during differentiation was not ruled out (Csink and Henikoff, 1998). In this study, we have tested whether the change in nuclear organization of bwD is correlated with specific events during differentiation.
We found that the number of cells that show association between bwD and 2Rh is higher in differentiating cells as opposed to undifferentiated cells of the eye imaginal discs. However, we also observed that in larval CNS of developmentally arrested larvae, the number of nuclei where the bwD locus is close to 2Rh increased. These data are more consistent with the interpretation that the length of G1 is responsible for increased association, rather than specific signal during differentiation. Of course, as terminal differentiation is often accompanied by cell cycle exit, increased association will be an indirect consequence of developmental progression. Interestingly, when we observed chromatin dynamics using time-lapse fluorescence microscopy we found that chromatin was more constrained in differentiated cells. These findings suggest that change in nuclear organization, although not established by a differentiation specific event, may be maintained by a differentiation controlled change in overall chromatin dynamics.
![]() |
Materials and Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Yeast strains
A diploid yeast strain heterozygous for erg2 was obtained from the Saccharomyces genome deletion project (SGDP), strain record number 20788, MATA/
his3
1/his3
1 leu2
/leu2
met15
/MET15 lys2
/LYS2 ura3
/ura3
erg2
:: KANMX4/ERG2. The haploid erg2
delete was obtained by random sporulation. PCR was done using primers as suggested by SGDP to confirm the deletion. Yeast used as an ERG2 (WT) control was a haploid strain, MAT
his3
200ura3-52 leu2 lys2. Both the erg2
and the ERG2 control yeasts were in a S288C background.
Subcloning of mRFP-LacI-NLS fusion protein expressed under the ubiquitin promoter
The ubiquitin promoter was obtained as a 1980bp BamHI/KpnI fragment from plasmid pCa4UbnlsGFPh27 (Davis et al., 1995) kindly donated by P. O'Farrell (University of California, San Francisco). LacI-NLS and 3'UTR were obtained as a BamHI/ClaI fragment of plasmid p3'SS (Belmont et al., 1999
) provided by A. Belmont (University of Illinois, Urbana, IL). Using mRFP1 in plasmid pRSETB (Campbell et al., 2002
) obtained from R. Tsien (University of California, San Diego) as a template, the sequence for mRFP was PCR amplified with the forward primer 5'-CCATCGATATGGCCTCCTCCG-3' and reverse primer 5'-TCTTAGGATCCGGCGCCGGT-3'. ClaI and BamHI restriction sites in the primers enabled subcloning with the ubiquitin promoter and lacI sequences in frame. All the above fragments except mRFP were first individually subcloned into pBluescript II KS(±) and were later put together in frame using appropriate combinations of restriction enzymes in pBluescript II KS(±). The construct was finally subcloned into the KpnI/SacII site of pCasPer4 to obtain pCas4{Ubq-mRFP-LacI-NLS}. Expression and nuclear localization of the fusion protein were tested by transfecting SL2 cells before P-element transformation. The construct along with pP25.7wc
2-3 were micro-injected into w1118 embryos to obtain germline transformants as described previously (Spradling, 1986
).
Microscopy
All microscopy on nuclei was carried out using a Deltavision (Applied Precision) system. The images were gathered with a cooled CCD camera using either a 60x/1.4 na UPlanApo (FISH and BrdU experiments) or 100x/1.35na PlanApo (live imaging experiments) objectives (Olympus). In the FISH experiments, images were gathered as 3D stacks, deconvolved using the Softworx software package (Applied Precision) and projected into 2D for measurements.
Fluorescence in situ hybridization on eye imaginal discs
Fluorescence in situ hybridization was done on eye imaginal discs from late third instar female larvae. Larvae heterozygous for bwD were derived from a cross between P{w+mW.hs=GawB}elav[C155], w; P{w+mC=UAS-syt.eGFP} males and w1118; bwD female flies and compared to P{w+mW.hs=GawB}elav[C155], w; P{w+mC=UAS-syt.eGFP}, bw+ larvae. The eye-antennal imaginal discs were dissected from larvae along with the mouth hooks in nuclear structure preservation buffer (NSPB) (80 mM KCl, 20 mM NaCl, 15 mM PIPES pH 7.0, 2 mM EDTA, 0.5 mM EGTA, 0.5 mM spermidine, 0.15 mM spermine, 1 mM DTT) supplemented with 0.1% deoxy-cholate and 0.1% triton-X-100. The discs were teased with a sharp tungsten needle to remove the peripodial membrane, subjected to hypotonic treatment in 1.0% sodium citrate for 5 minutes and dissected at the morphogenetic furrow in NSPB buffer under an epifluorescence dissecting microscope. The expression pattern of GFP was used to demarcate the furrow. After separating the posterior end of the eye discs, the anterior end was dissected away from the antennal disc and mouth-hooks. The separated anterior and posterior ends of the eye imaginal discs were incubated in 0.3 mg/ml solution of collagenase (Sigma) for 5 minutes at room temperature. The 2 mg/ml stock solution of collagenase was prepared in divalent cation free buffer (Ashburner, 1989). The discs were fixed in methanol(11): acetic acid(11): water(1) (MAW) for 30 seconds, and transferred to a drop of 45% acetic acid on a slide for a minute as described previously (Csink and Henikoff, 1996
), except that the slide was pre-coated with 5-10 µl of 1 mg/ml stock solution of poly-lysine (Sigma) prepared in 50 mM sodium borate buffer pH 8.5. The tissue was further teased apart using a tungsten wire and squashed under a sialized coverslip. FISH was as described in (Csink and Henikoff, 1998
). Nuclei were counterstained with DAPI (Molecular Probes). The two dimensional distance between the signals for the two probes was measured and corrected for by the nuclear radius (as determined by the nuclear area assuming a perfect circle). To prevent bias the images were coded and scored blind.
Developmental arrest of larvae and FISH on CNS
w1118; bwD and w1118 larvae were used. In all, 70-80 females were allowed to lay eggs on a grape agar plate for 2 hours. The embryos were collected, washed in sterile dH2O, 70% ethanol and dechorionated by washing in 2.5% NaOCl (bleach) in 50% ethanol for 3 minutes and rinsed in Chang and Gehrings's solution (Ashburner, 1989). Bleach and ethanol prevent wild-type yeast contamination. Embryos were transferred to a sterile conical flask containing 5 ml of 0.5% dextrose in 2% water-agar gel and plugged with sterile rayon.
The agar was overlaid with either erg2 yeast or WT yeast. The yeast strains were grown as overnight cultures of 500 ml; care was taken to wash off the media in sterile distilled water. The pellet was suspended in twice the volume in mls of the wet weight of the pellet in dH2O. For each overlay approximately 0.5 ml of the yeast paste was usually added as adapted from (Bos et al., 1976
). More yeast was added 48 hours, 72 hours and 120 hours AED. The embryos were reared in these flasks at 25°C. They reached the third instar larval stage at around 96-100 hours AED. CNS from female larvae fed WT yeast as well as erg2
yeast were dissected at 110-115 hours AED. A second round of dissections was done for developmentally arrested larvae that had been fed erg2
yeast at 130-135 hours AED. By this timepoint all larvae reared on WT yeast had pupated. The protocol followed for dissection and FISH on CNS was as described previously (Csink and Henikoff, 1998
). The images were acquired and analyzed as described above.
Measuring mitotic index
w1118 larvae were reared on either erg2 yeast or WT yeast and dissected at 110-115 hours AED (day 1) and 130-135 hours AED (day 2) (developmentally arrested larvae only). CNS were dissected and prepared as described previously (Ashburner, 1989
). To prevent bias the slides were coded before imaging and each field was randomly chosen with the nuclei slightly out of focus before imaging, so that perception of mitotic figures would not influence field selection.
Determination of cell cycle phase by quantitating DAPI intensity
w1118; bwD, larvae were reared on erg2 yeast or WT yeast as described above. Larvae were used at 113 hours AED (reared on erg2
yeast or WT yeast) as well as 137 hours AED (reared on erg2
yeast). They were fed bromodeoxy-uridine (BrdU) laced food for 3 hours, prepared in the following manner: 0.3 g of dry agar was placed in a 35 mmx10 mm style cell culture dish (Corning). The agar was soaked with 1.6 ml of 1 mg/ml solution of BrdU stock solution (20 mg/ml in 40% ethanol). The stock solution of BrdU was diluted to 1 mg/ml in 0.5% dextrose and bromophenol blue tracker at 0.3 mg/ml. The larvae obtained from the conical flasks were fed the BrdU laced blue food for 3 hours at 25°C in a humid chamber, after which only those with blue food in the gut were selected. The larvae were washed and CNS dissected. The tissue and slides were processed as described (Csink, 2004
).
Fields for imaging were chosen while looking only at the DAPI channel and were acquired at a single plane of focus. Quantitation of DAPI intensity was performed using Softworx software (Applied Precision). The total DAPI intensity per unit area was calculated for each nucleus and the background intensity was subtracted. The median nuclear intensity for each field was used as the normalizing factor. On the basis of the distribution of the ratios in a histogram, and the corresponding BrdU label, the ratios obtained were designated into three classes: Aneuploids (A) with ratio <0.5 and >2.6, G1 if the ratio fell between 0.5-1.20 and S/G2 if the ratio was between 1.2-2.6. Statistical significance of the distribution of cell cycle for calculating the P-value was computed using the G-test (Sokal and Rohlf, 1981).
Measurement and imaging of chromatin movement
To study the dynamics of chromosomal movement w, y P{wmc, lacO}; P{wmc, lacO} Dcp-1/CyO females were crossed with w1118, P{wmc, Ubq-mRFP-lacI-NLS} males to obtain y w P{wmc, lacO}/w1118, P{wmc, Ubq-mRFP-lacI-NLS}; P{wmc, lacO} Dcp-1/+ for imaging. Eye-antennal discs were dissected from third-instar female larvae obtained from the cross described above. The larvae were washed in 1x Ringers and the dissections were carried out in MM3 media (Ashburner, 1989) supplemented with 2.5% FBS. The discs were transferred to a poly-lysine-coated 40 mm coverslip and imaging was done in an FCS2 chamber (Bioptech) at 21° to 24°C, allowing us to maintain the primary culture during microscopy (Li and Meinertzhagen, 1995
). Images were taken every 5 seconds at a single plane of focus. The length of the movies varied from of 140-500 seconds. For image analysis, polygons were drawn around the nucleus as well as the dots representing the lacO repeats on the X chromosome and near the bw locus on the 2nd chromosome, using the Softworx software. The distance between the center of mass of the two dots and the nuclear radius was computed using a Java program developed in lab. The mean step size (Table 2) was computed for
t=10 seconds. The mean square change in distance <
d2> for all possible values of
t was computed using a macro on Excel and statistical calculations were performed on StatView (version 4.5). For increasing values of
t the number of data points used to calculate <
d2> decreases. This adds to noise in the data observed at later time points, confirming earlier findings that the approximation <
d2> is reliable only for first 25-50% of the collection period for each nucleus (Heun et al., 2001
; Vazquez et al., 2001
). Hence to evaluate chromatin motion we first computed <
d2> values for individual nuclei using the entire range of data. Since the duration of movies varied from 140 to 500 seconds, values corresponding to the first 50% of the length of movie (70-250 seconds) for each individual nucleus were used to evaluate chromatin motion as the average <
d2>. The values obtained until
t=120 seconds are represented here. The diffusion coefficient (D) can be computed from the slope of the graph, based on the relation <
d2>=4D <
t> (Qian et al., 1991
; Vazquez et al., 2001
). Since chromatin motion is constrained, the change in distance over time reaches a standard value. The slope was hence calculated by taking only the linear portion of the graph into account at
t=5-10 seconds for both differentiated as well as undifferentiated cells. For undifferentiated cells at
t=5 seconds data was representative of 814 data points, from 19 nuclei in 7 eye discs. The value of average <
d2> at
t=120 was calculated from 18 nuclei. In the case of differentiated cells, at
t=5 seconds the data was obtained from 668 data points, 21 nuclei from 9 eye discs. The number of nuclei used to compute average <
d2> at
t=120 was 18.
|
|
|
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
To aid in the separation of the two cell types, we used a fly-line that expresses GFP only in the differentiated cells posterior to the morphogenetic furrow. We dissected eye imaginal discs from crawling third instar larvae at the morphogenetic furrow, separating the differentiating cells from undifferentiated cells (Fig. 1A). The dissected tissues were subject to FISH on separate slides. No significant difference in the distance between bw-2Rh (corrected by the radius) in differentiated and undifferentiated cells from bw+ larvae was found (Fig. 1B). However, in bwD heterozygotes the distance between bw-2Rh significantly decreased in differentiated cells (Fig. 1B,C) when compared with undifferentiated cells. Interestingly, we also found a statistically significant decrease in the bw-2Rh distance in the undifferentiated cells from bwD heterozygotes when compared with bw+. Association between bw-2Rh in undifferentiated bwD/bw+ nuclei shows that differentiation is not strictly required for association. This observation suggests that increased association in differentiated cells may simply be a stochastic event and the increased association in the differentiated cells is due to their spending more time in G1. The experiments described below were performed to further test this possibility.
Heterochromatic associations in developmentally arrested larvae
To test if the repositioning of the bwD heterochromatic block is specific to differentiation, we measured the association between bwD and 2Rh in developmentally arrested larvae. If differentiation promotes association between bwD and 2Rh, then developmental arrest, which prevents differentiation, should decrease association. Developmental arrest was accomplished by starving the larvae of the hormone ecdysone. Ecdysone is essential for morphogenesis in insects and regulates many aspects of cellular differentiation (Thummel, 1995). In Drosophila melanogaster, a small amount of ecdysone precursor is provided maternally, but later morphogenesis requires that the precursor be provided in the diet. This precursor, in turn, is the product of the erg2 pathway in Saccharomyces cerevisiae, the primary food of D. melanogaster. When larvae are restricted to erg2 yeast, their development stalls in the third instar stage and they fail to pupate (Fig. 2A) (Bos et al., 1976
). Larvae reared on the mutant yeast developed at the same rate as those fed wild-type yeast until mid third instar stage, 110-115 hours after egg deposition (AED) (referred to as `day 1'). At around 118-120 hours AED larvae fed wild-type yeast started to pupate, whereas those fed mutant yeast remained in the third instar stage (Fig. 2A). Even at 144 hours AED, larvae fed mutant yeast failed to pupate, did not grow in size and would remain in the third instar stage till they begin to die about 5 days later. Larvae fed wild-type yeast stopped eating during the late third instar stage. By contrast, the developmentally arrested larvae continued to eat at least until 140 hours AED (referred to as `day 2'). By feeding them wild-type yeast, we could rescue the developmentally arrested larvae at 120 or 135 hours AED; however, the frequency of rescue was slightly lower when rescued at 135 hours AED. All the larvae that successfully pupated, eclosed as fertile flies that appeared normal.
|
We compared heterochromatic association in the CNS dissected from bw+ and bwD larvae 110-115 hours AED (day 1) that had been fed erg2 yeast and in CNS from similarly aged bw+ and bwD larvae fed wild-type yeast. We also compared CNS derived from developmentally arrested larvae reared on erg2
yeast that had stalled at this stage for about 24 hours, at 135-140 hours AED (day 2). At this time all larvae that had been fed wild-type yeast had pupated, thus no corresponding control of larvae fed wild-type yeast was possible. We chose to use CNS rather than eye imaginal discs, because CNS dissection and preparation was easier and the GFP expression in the fly line used in our earlier analysis varied greatly among larvae that had been fed the mutant yeast.
Contrary to our expectations, we found that in bwD nuclei from larvae 130-140 hours AED (day 2) that had been fed erg2 yeast, there was a significant decrease in the distance between bw-2Rh (Fig. 2B,C). On day 1 the apparent association was not significantly different in nuclei that were fed either the wild-type yeast or the erg2
yeast. In the bw+ nuclei, there was a slight difference in the distribution of the distance between bw-2Rh in larvae reared on erg2
or wild-type yeast at day 1, which seemed to be lost in the day 2 larvae. Taken together, these data, along with our observation in eye imaginal discs, argue against a causative role for a differentiation specific signal in regulating the changes in nuclear organization with respect to bwD.
Cell cycle profile of developmentally arrested larvae
To follow up on our data from the developmentally arrested larvae, we examined the cell cycle profile of the larvae reared on erg2 yeast. We first compared the mitotic index in the CNS of larvae from the three different classes in Table 1. The difference in mitotic index was not statistically significant, indicating that feeding the larva erg2
yeast does not cause a cell cycle arrest during mitosis.
|
We next determined the proportion of cells in the G1 and S/G2 phase of the cell cycle by quantifying the DNA content in cells based on relative DAPI intensity. Larvae reared on the wild-type yeast were fed BrdU at 113 hours AED (day 1); those reared on erg2 yeast were fed BrdU at 113 hours (day 1) or at 137 hours (day 2) AED. In each case BrdU-laced food was fed for 3 hours, following which the CNS was dissected. Using the DAPI intensity and the presence or absence of BrdU labeling, we were able to determine the proportion of nuclei in G1 or S/G2 phases of the cell cycle (Csink and Henikoff, 1998
). In addition, we were also able to determine if there was a change in the proportion of cells entering the S phase in larvae reared on erg2
yeast. In larvae fed erg2
yeast the proportion of cells in G1 was higher than that observed in larvae reared on wild-type yeast (P<0.01) (Table 1). In larvae reared on erg2
yeast, the proportion of cells in G1 was similar for day 1 as well as day 2. However, there was a significant decrease in the percentage of cells that were labeled with BrdU in the developmentally arrested larvae on day 2 as compared with day 1 (P<0.001). These results are consistent with an overall retardation of the cell cycle, a slight lengthening of G1 and/or a partial block of the G1-S transition.
Collectively, these results suggest that increased association between bwD and 2Rh in the developmentally arrested larvae at 135 hours AED is because of an increase in the length of G1. In the day 1, erg2 fed larvae, although the proportion of cells in G1 was higher than that observed in larvae undergoing normal development, we did not observe a similar increase in association between bwD and 2Rh. Since in these larvae the proportion of cells entering the S phase was only slightly different from those reared on wild-type yeast, we hypothesize that on day 1 more of the cells are in the earlier part of G1 where we do not expect to see association. On day 2 the cells would have been in G1 for a long enough period of time for a larger number of associations to form.
Dynamics of chromosomal movements in primary cultures of eye imaginal discs
On the basis of our studies in fixed tissue, we were interested in learning if the difference in nuclear organization of bwD arose due to a disparity in the dynamics of chromosomal movement in differentiated versus undifferentiated cells. Changes in chromatin dynamics during differentiation could play a major role in the changing patterns of gene expression. With advances in microscopy and the development of the lac operator (lacO) and LacI-GFP fusion technique, the dynamics of movement of chromosomal regions can now be studied in live tissue (Belmont et al., 1999). Crosses were performed to produce flies that contained two copies of the lacO repeats, one marking the X chromosome and the other the bw locus on the second chromosome (Fig. 3A) and expressed a mRFP-LacI fusion protein. Nuclei from these larvae contained two bright dots. The unbound, but nuclear localized, mRFP-LacI was less bright than the spots and was used to demarcate the interphase nucleus (Fig. 3B, Figs S1, S2 and S3 in Supplementary material).
We compared the dynamics of chromosomal movement in differentiated cells posterior to the morphogenetic furrow to undifferentiated cells anterior to the furrow in primary cultures of eye imaginal discs obtained from third instar larvae. Differentiated cells were easy to recognize as they were part of the ommatidial clusters in the eye disc (Fig. 3B). Images were taken at a time interval of 5 seconds at a single plane of focus (for movies of nuclei from anterior and posterior cells, see Figs S1 and S2 in Supplementary material). To deduce the dynamics of movement, the distance between the two dots (d) was computed over time. A measurement of distance between the center of mass of two loci eliminates the need for correction for apparent motion that may arise due to nuclear rotation and translation (Vazquez et al., 2001). To determine the characteristics of chromatin movement, the mean square change in distance <
d2> which is the average of
d2 values over all possible combinations of timepoints separated by
t (Qian et al., 1991
; Vazquez et al., 2001
) was computed for each nucleus (Fig. 3C).
The diffusion coefficient (D) for undifferentiated cells was calculated at t=5-10 to be 3.3x104 µm2/seconds and was comparable to that for differentiated cells 2.1x104 µm2/seconds (Fig. 3C and Table 2). An unpaired t-test comparing the average <
d2> values in differentiated versus undifferentiated cells for
t values less than 25 seconds found no significant difference. However, for
t values greater than 25 seconds the t-test indicated that the values were significantly different. The average value of step size (
d) for both differentiated and undifferentiated cells at a time interval of 10 seconds was found to be equivalent (Table 2). Although the rate of chromatin diffusion was found to be similar in both cell types, the plot for differentiated cells reached a plateau as early as
t=50 seconds. The plateau height on the <
d2> graph is referred to as the radius of confinement and is the size of the region through which the locus is free to diffuse. A similar plateau was not seen in undifferentiated nuclei during the time period studied (Fig. 3C), suggesting that chromatin motion is much more constrained in differentiated cells. This is exemplified by the smaller radius of confinement for the tagged chromosomes in differentiated cells, calculated from the graph in Fig. 3C and shown in Table 2. A comparison of the nuclear radius of undifferentiated and differentiated cells shown in Table 2 suggests that the decrease in the radius of confinement in differentiated cells may be due to a general nuclear contraction and/or chromatin compaction that accompanies differentiation.
Dynamics of chromosomal movement in developmentally arrested larvae
Our observations suggest that the association between bwD and 2Rh is a factor of time spent in G1/G0 phases of the cell cycle since the previous mitosis. This, in turn, implies that the movement of bw towards 2Rh could either be a slow directed movement or a random walk event. In both these cases the probability of association would increase if more time is available for the event. If this is indeed the case, then the establishment of association between bwD and 2Rh observed in the developmentally arrested larvae should not arise due to a change in the rate of chromatin movement. We therefore compared chromatin movement in primary cultures of the CNS derived from larvae fed wild-type yeast to those reared on erg2 yeast (for movies of nuclei from CNS cells, see Fig. S3 in Supplementary material). We followed the movement of the bw locus with respect to the X chromosome, as was done in eye imaginal discs (Fig. 4). As theorized, the average <
d2> among larvae reared on wild-type yeast when compared with those reared on erg2
yeast on either day 1 or day 2 was not statistically significant at any
t (computed by unpaired t-test). The diffusion coefficients were also similar (Table 2). As a control we also imaged fixed nuclei and found that the plot for average <
d2> did not change with increasing time intervals. The diffusion coefficient was found to be 1.0x107 µm2/seconds, confirming that the change in average <
d2> over time, observed in live tissue, was not an artifact of drift or jitter that may arise during microscopy. The average step size of chromosomal movements at a time interval of 10 seconds was also found to be similar irrespective of the yeast fed or the time of development (Table 2).
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
One may imagine that the diffusion coefficient would be influenced by several factors, including the degree of compaction of the chromatin fiber, the anchoring of chromatin to nuclear structures and the rigidity or density of materials that may contribute to a nuclear matrix. Studies of Sedat and colleagues have found that the diffusion coefficient is different in various cell types in Drosophila. In spermatocytes derived from third instar larvae, the rate of chromatin diffusion in nuclei in mid G2 was found to be 1x103 µm2/seconds, whereas for late G2 the value is 5x105 µm2/seconds (Vazquez et al., 2001). The diffusion coefficient of chromatin in nuclei from early Drosophila embryos has been reported as 2x103 µm2/seconds (Marshall et al., 1997
). The calculated diffusion coefficients in our studies fall between these two extremes; they ranged from 1.9-3.7x104 µm2/seconds (Table 2) and are close to the value of 1.2x104 µm2/seconds found for chromatin in mammalian tissue culture cells (Chubb et al., 2002
). However, it is interesting to note that their seems to be little to no change in the diffusion coefficient over development in the system we have examined.
We have also shown that a specific rearrangement of interphase nuclear organization, the association of two blocks of constitutive heterochromatin, is correlated with differentiation. However, our results suggest that the heterochromatic association between bwD and 2Rh is not a result of a discrete event during differentiation. Rather, association is a consequence of time spent, since the last mitosis, in either G1 or G0 phase of the cell cycle. The increased association between bwD-2Rh is seen not only in cells undergoing differentiation but also in the developmentally arrested larvae with a lengthened G1 phase, as well as some undifferentiated cells of the eye imaginal disc. Association is also observed in cells that will re-enter S phase (Csink and Henikoff, 1998). These findings suggest that the association could be regulated by a factor that accumulates or is lost slowly from the nucleus. This gradual change would promote heterochromatin cohesion after mitosis, independent of differentiation. Indeed, some of the changes in nuclear organization seen during differentiation may be simply due to the fact that differentiation is accompanied by an extended G1 or exit from the cell cycle. We did see an increase in the confinement of a locus on differentiation. Therefore, it is possible that once a specific nuclear organization is established, its maintenance is assisted by a differentiation specific event that confines chromosomal movement.
What aspects of chromatin and nuclear structure could gradually change to promote association? Heterochromatin is known to be inherently sticky, and although this interaction is sequence independent (Sage and Csink, 2003), it is probably mediated by proteins that recognize the repetitive nature of heterochromatin (Dorer and Henikoff, 1997
). The delayed association between bwD and 2Rh and the fact that centromeric regions do not associate to form a chromocenter in most diploid tissues of Drosophila show that heterochromatin does not always self associate. These findings suggest that heterochromatic cohesion is regulated. Indeed, there are proteins that are known to associate with heterochromatin during mitosis, but gradually dissociate at a slow rate usually over a period of a few hours during interphase (Platero et al., 1998
). Other proteins, like HP1 are removed from chromatin just before mitosis and redeposited as the cell exits mitosis (Kellum et al., 1995
; Murzina et al., 1999
). Moreover, the binding of HP1 to stable heterochromatic domains has been found to be highly dynamic (Cheutin et al., 2003
). It has been speculated that this transient binding allows for competition among proteins to bind to heterochromatin and regulate its chromatin structure and cohesion.
There are several possible factors that participate in the alterations in nuclear structure during development and may contribute to the changes that increase chromatin confinement and promote bwD association. The composition of nuclear lamina is known to change as a cell undergoes differentiation and progresses through the cell cycle. Lamins have been shown to directly and indirectly interact with chromatin or chromosomal binding proteins (reviewed by Goldman et al., 2002). Therefore, the increased confinement to a smaller nuclear space during differentiation that we see in the eye discs may be due to specific changes in the nuclear lamina or its associated proteins that in turn regulate their interaction with chromatin. HP1, a protein known to promote bwD-2Rh associations (Csink and Henikoff, 1996
), interacts with the nuclear envelope protein lamin B Receptor (LBR) (Ye and Worman, 1996
). Changes in the phosphorylation profile of HP1 have been correlated with its cell cycle dependent assembly on chromatin, protein-protein interactions and heterochromatinization (reviewed by Kellum, 2003
). Modification of HP1, during differentiation, may alter its interaction with the nuclear lamina as well as with chromatin. This in turn might help maintain the nuclear organization necessary to obtain the transcription profile accompanying differentiation.
Because the bw locus is present on the tip of the right arm of the second chromosome, its association with 2Rh in nuclei heterozygous for bwD requires large-scale chromosomal movements. If the movement of bwD towards 2Rh is a fast directional event we would expect to see triggered, saltatory chromatin reorganization in a short time window. So far, we have detected no such chromosomal movements. Instead, it appears that the movement of bwD towards 2Rh is a slow directed movement or a random walk event where the probability of association would increase if more time was available for the event. This aspect of differentiation could then be viewed as being due to an extended G1, which would provide more time for establishment of final nuclear organization. Previous studies in Drosophila spermatocytes (Vazquez et al., 2001) and in S. cerevisiae (Heun et al., 2001
) show that loci undergo both rapid, local saltatory movements as well as occasional long range movements (Gasser, 2002
). The direction of these movements seems to be quite random. We postulate that bwD randomly moves about its area of confinement within the nucleus in early G1. If it encounters 2Rh during early G1 it should be able to move away, but if it encounters 2Rh during late G1, due to the increase in the stickiness of heterochromatin, it can associate with 2Rh. Such association, and other later changes in nuclear organization, would then be stabilized by an overall increase in the confinement of chromatin.
![]() |
Acknowledgments |
---|
![]() |
Footnotes |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Ashburner, M. (1989). Drosophila: A Laboratory Manual. Cold Spring Harbor: Cold Spring Harbor Laboratory Press.
Belmont, A. S., Li, G., Sudlow, G. and Robinett, C. (1999). Visualization of large-scale chromatin structure and dynamics using the lac operator/lac repressor reporter system. Methods Cell Biol. 58, 203-222.[Medline]
Bos, M., Burnet, B., Farrow, R. and Woods, R. A. (1976). Development of Drosophila on sterol mutants of the yeast Saccharomyces cerevisiae. Genet. Res. 28, 163-176.[Medline]
Brown, K. E., Guest, S. S., Smale, S. T., Hahm, K., Merkenschlager, M. and Fisher, A. G. (1997). Association of transcriptionally silent genes with Ikaros complexes at centromeric heterochromatin. Cell 91, 845-854.[Medline]
Brown, K. E., Baxter, J., Graf, D., Merkenschlager, M. and Fisher, A. G. (1999). Dynamic repositioning of genes in the nucleus of lymphocytes preparing for cell division. Mol. Cell 3, 207-217.[Medline]
Campbell, R., Tour, O., Palmer, A., Steinbach, P., Baird, G., Zacharias, D. and Tsien, R. (2002). A monomeric red fluorescent protein. Proc. Natl. Acad. Sci. USA 99, 7877-7882.
Cheutin, T., McNairn, A. J., Jenuwein, T., Gilbert, D. M., Singh, P. B. and Misteli, T. (2003). Maintenance of stable heterochromatin domains by dynamic HP1 binding. Science 299, 721-725.
Chubb, J. R., Boyle, S., Perry, P. and Bickmore, W. A. (2002). Chromatin motion is constrained by association with nuclear compartments in human cells. Curr. Biol. 12, 439-445.[CrossRef][Medline]
Csink, A. K. (2004). Analysis of chromosomes of the larval CNS by FISH and BrdU labeling. Methods Mol. Biol. 247, 343-352.[Medline]
Csink, A. K. and Henikoff, S. (1996). Genetic modification of heterochromatic association and nuclear organization in Drosophila. Nature 381, 529-531.[CrossRef][Medline]
Csink, A. K. and Henikoff, S. (1998). Large-scale chromosomal movements during interphase progression in Drosophila. J. Cell Biol. 143, 13-22.
Csink, A. K., Bounoutas, A., Griffith, M. L., Sabl, J. F. and Sage, B. T. (2002). Differential Gene Silencing by trans-heterochromatin in Drosophila melanogaster. Genetics 160, 257-269.
Davis, I., Girdham, C. H. and O'Farrell, P. H. (1995). A nuclear GFP that marks nuclei in living Drosophila embryos; maternal supply overcomes a delay in the appearance of zygotic fluorescence. Dev. Biol. 170, 726-729.[CrossRef][Medline]
Dernburg, A. F., Broman, K. W., Fung, J. C., Marshall, W. F., Philips, J., Agard, D. A. and Sedat, J. W. (1996). Perturbation of nuclear architecture by long-distance chromosome interactions. Cell 85, 745-759.[Medline]
Dorer, D. R. and Henikoff, S. (1997). Transgene repeat arrays interact with distant heterochromatin and cause silencing in cis and trans. Genetics 147, 1181-1190.
Francastel, C., Schubeler, D., Martin, D. I. and Groudine, M. (2000). Nuclear compartmentalization and gene activity. Nat. Rev. Mol. Cell Biol. 1, 137-143.[CrossRef][Medline]
Gasser, S. M. (2002). Visualizing chromatin dynamics in interphase nuclei. Science 296, 1412-1416.
Goldman, R. D., Gruenbaum, Y., Moir, R. D., Shumaker, D. K. and Spann, T. P. (2002). Nuclear lamins: building blocks of nuclear architecture. Genes Dev. 16, 533-547.
Heun, P., Laroche, T., Shimada, K., Furrer, P. and Gasser, S. M. (2001). Chromosome dynamics in the yeast interphase nucleus. Science 294, 2181-2186.
Kellum, R. (2003). HP1 complexes and heterochromatin assembly. Curr. Top. Microbiol. Immunol. 274, 53-77.[Medline]
Kellum, R., Raff, J. W. and Alberts, B. M. (1995). Heterochromatin protein 1 distribution during development and during the cell cycle in Drosophila embryos. J. Cell Sci. 108, 1407-1418.
Kosak, S., Skok, J., Medina, K., Riblet, R., Le Beau, M., Fisher, A. and Singh, H. (2002). Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296, 158-162.
Li, C. and Meinertzhagen, I. A. (1995). Conditions for the primary culture of eye imaginal discs from Drosophila melanogaster. J. Neurobiol. 28, 363-380.[Medline]
Manuelidis, L. (1985). Individual interphase chromosome domains revealed by in situ hybridization. Hum. Genet. 71, 288-293.[Medline]
Marshall, W. F., Straight, A., Marko, J. F., Swedlow, J., Dernburg, A., Belmont, A., Murray, A. W., Agard, D. A. and Sedat, J. W. (1997). Interphase chromosomes undergo constrained diffusional motion in living cells. Curr. Biol. 7, 930-939.[Medline]
Murzina, N., Verreault, A., Laue, E. and Stillman, B. (1999). Heterochromatin dynamics in mouse cells: interaction between chromatin assembly factor 1 and HP1 proteins [In Process Citation]. Mol. Cell 4, 529-540.[Medline]
Platero, J., Csink, A., Quintanilla, A. and Henikoff, S. (1998). Changes in chromosomal localization of heterochromatin binding proteins during the cell cycle in Drosophila. J. Cell Biol. 140, 1297-1306.
Qian, H., Sheetz, M. P. and Elson, E. L. (1991). Single particle tracking. Analysis of diffusion and flow in two-dimensional systems. Biophys. J. 60, 910-921.[Abstract]
Sage, B. T. and Csink, A. K. (2003). Heterochromatic self-association, a determinant of nuclear organization, does not require sequence homology in Drosophila. Genetics 165, 1183-1193.
Sokal, R. R. and Rohlf, F. J. (1981). Biometry. New York: W. H. Freeman.
Spector, D. L. (2003). The dynamics of chromosome organization and gene regulation. Annu. Rev. Biochem. 72, 573-608.[CrossRef][Medline]
Spradling, A. C. (1986). P element-mediated transformation. In Drosophila a practical approach (ed. D. B. Roberts), pp. 175-198. Washington, D.C.: IRL Press.
Thummel, C. S. (1995). From embryogenesis to metamorphosis: the regulation and function of Drosophila nuclear receptor superfamily members. Cell 83, 871-877.[Medline]
Trask, B., van den Engh, G., Pinkel, D., Mullikin, J., Waldman, F., van Dekken, H. and Gray, J. (1988). Fluorescence in situ hybridization to interphase cell nuclei in suspension allows flow cytometric analysis of chromosome content and microscopic analysis of nuclear organization. Hum. Genet. 78, 251-259.[CrossRef][Medline]
Vazquez, J., Belmont, A. S. and Sedat, J. W. (2001). Multiple regimes of constrained chromosome motion are regulated in the interphase Drosophila nucleus. Curr. Biol. 11, 1227-1239.[CrossRef][Medline]
Verschure, P. J., van Der Kraan, I., Manders, E. M. and van Driel, R. (1999). Spatial relationship between transcription sites and chromosome territories. J. Cell Biol. 147, 13-24.
Ye, Q. and Worman, H. J. (1996). Interaction between an integral protein of the nuclear envelope inner membrane and human chromodomain proteins homologous to Drosophila HP1. J. Biol. Chem. 271, 14653-14656.