Quantitative Analysis of Chromosome Condensation in Fission Yeast
Boryana Petrova,a,b Sascha Dehler,a Tom Kruitwagen,a* Jean-Karim Hériché,a Kota Miura,c Christian H. Haeringa,d
Cell Biology & Biophysics Unit,a International PhD Programme,b Centre for Molecular and Cellular Imaging,c and Structural & Computational Biology Unit,d European
Molecular Biology Laboratory, Heidelberg, Germany
Chromosomes undergo extensive conformational rearrangements in preparation for their segregation during cell divisions. Insights into the molecular mechanisms behind this still poorly understood condensation process require the development of new approaches to quantitatively assess chromosome formation in vivo. In this study, we present a live-cell microscopy-based chromosome condensation assay in the fission yeast Schizosaccharomyces pombe. By automatically
tracking the three-dimensional distance changes between fluorescently marked chromosome loci at high temporal and
spatial resolution, we analyze chromosome condensation during mitosis and meiosis and deduct defined parameters to
describe condensation dynamics. We demonstrate that this method can determine the contributions of condensin, topoisomerase II, and Aurora kinase to mitotic chromosome condensation. We furthermore show that the assay can identify
proteins required for mitotic chromosome formation de novo by isolating mutants in condensin, DNA polymerase ␧, and
F-box DNA helicase I that are specifically defective in pro-/metaphase condensation. Thus, the chromosome condensation
assay provides a direct and sensitive system for the discovery and characterization of components of the chromosome condensation machinery in a genetically tractable eukaryote.
T
o ensure their proper partitioning during mitotic and meiotic
cell divisions, chromosomes must undergo substantial conformational rearrangements from what is thought to be a loose
assembly of interphase chromatin fibers into compact rod-shaped
structures. The molecular mechanisms behind this large-scale
chromosome reorganization process are still poorly understood
(1, 2).
Biochemical analysis identified topoisomerase II (Topo II) and
multisubunit protein complexes named condensins I and II as
abundant nonhistone components of mitotic chromosomes (3–
9). Depletion or inactivation of condensins or Topo II in different
model systems frequently leads to the formation of structurally
unstable mitotic or meiotic chromosomes and inevitably causes
failures in the resolution and segregation of sister chromatids during anaphase (10, 11). Consistent with a role in mitotic chromosome formation, condensins and Topo II localize to the longitudinal axes of metazoan metaphase chromosomes (12–14). The
chromosomal association and activity of condensins are controlled by mitotic kinases, including Aurora B (reviewed in reference 15).
Unexpectedly, chromosomes can still compact to considerable
levels in a number of cultured cell lines after gene knockout or
RNA interference (RNAi) depletion of condensin subunits (16–
20), Topo II (21–23), or both (24). Similarly, inhibition or depletion of Aurora B appears to have little effect on the extent of
prophase chromosome condensation in Caenorhabditis elegans or
human cells (25–27). These findings suggest that there exist additional molecular players that induce the compaction of mitotic
chromosomes, including a hypothetical regulator of chromosome
architecture (RCA) that promotes chromosome condensation
when cyclin-dependent kinase (CDK) activity is high (28). Despite significant efforts, including mapping the proteome of
mitotic chromosomes (29) and profiling all genes required for
mitotic cell divisions (30), the identities of chromosome condensation factors like RCA remain unknown.
Why does the identification of the proteins that drive chromosome condensation prove to be so difficult? Condensation most
likely requires the cooperative action of multiple proteins or protein complexes. Inactivation of just a single of these proteins may
cause only subtle defects in the compaction levels of mitotic chromosomes, which might be difficult to detect by merely qualitative
methods. Measuring chromosome condensation in living cells
quantitatively is, on the other hand, challenging due to its dynamic and transient nature, and the quantitative methods used to
study mitotic chromosomes in vitro probably suffer from artifacts
that are frequently introduced by isolation or fixation procedures
(2, 31). Progress has recently been made by approaches that measure either fluorescence resonance energy transfer (FRET), projection intensity, or the volume occupied by histones tagged with
green fluorescent protein (GFP) in live mammalian cell lines or C.
elegans embryos (26, 32, 33). Although these studies provide excellent descriptions of the degrees and kinetics of mitotic chromosome condensation, they require complex microscopy and analysis methods that may hinder their application for the search for
novel condensation factors in high throughput. Incomplete depletion of proteins or off-target effects by RNA interference in
these systems may, furthermore, hamper genome-wide screens
for such factors. A complete understanding of the chromosome
condensation machinery will therefore require alternative approaches that quantitatively assess chromosome condensation in
a systematic manner.
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Molecular and Cellular Biology
Received 17 October 2012 Returned for modification 5 November 2012
Accepted 18 December 2012
Published ahead of print 21 December 2012
Address correspondence to Christian H. Haering, [email protected].
* Present address: Tom Kruitwagen, Department of Biochemistry, ETH Zurich,
Zurich, Switzerland.
Supplemental material for this article may be found at http://dx.doi.org/10.1128
/MCB.01400-12.
Copyright © 2013, American Society for Microbiology. All Rights Reserved.
doi:10.1128/MCB.01400-12
March 2013 Volume 33 Number 5
Quantitative Analysis of Chromosome Condensation
The full repertoire of genetic tools available makes yeasts excellent model systems to screen for novel condensation factors.
However, the small size of yeast chromosomes and, hence, the
inability to visualize individual chromosomes is still a major bottleneck, even when taking the most recent developments in microscopy technologies into account. Previous studies of chromosome condensation in the budding yeast Saccharomyces cerevisiae
relied on the visualization of individual chromosome regions in
fixed cells by in situ hybridization with fluorescently labeled
probes (fluorescent in situ hybridization [FISH]) (34). These measurements suggested that most of the genome, with the exception
of the ribosomal DNA (rDNA) cluster, undergoes only very minor
degrees of compaction during mitosis. Studies that evaluated the
association between fluorescently labeled chromosome loci in live
cells found that the distances between certain locus combinations
decreased during mitosis, while the distances between other loci
remained unchanged (35, 36), indicating tha only certain regions
of Saccharomyces cerevisiae chromosomes condense to appreciable
levels.
In comparison to the 16 budding yeast chromosomes, the 3
chromosomes of the fission yeast Schizosaccharomyces pombe are
considerably longer and exhibit higher levels of complexity, including heterochromatin formation and centromere structures
closer to those of mammalian cells (37). Taking advantage of the
longer and more complex chromosomes that make S. pombe ideal
to study the chromosome condensation machinery in a genetically
tractable organism, we established a chromosome condensation
assay that measures three-dimensional (3D) distances between
fluorescently labeled loci in live fission yeast cells. We show that
this assay is capable of providing quantitative readouts for the
compaction of chromosomes during mitotic and meiotic cell
divisions, allowing us to define quantitative parameters for the
extent and dynamics of chromosome condensation. We demonstrate that these parameters can be used to describe the relative contributions of condensin, Topo II, and Aurora kinase to
chromosome condensation. We furthermore show that the
condensation assay can be used in medium to high throughput
to determine genes required for chromosome condensation in
genetic screens by identifying from a pool of random yeast
mutants eight new conditional condensin alleles, as well as mutants in DNA polymerase ε (pol ε) and F-box DNA helicase I,
which had not been previously implicated in mitotic chromosome condensation.
MATERIALS AND METHODS
Growth and synchronization of yeast strains. Strains were grown in either complete medium with supplements (YE5S) or minimal medium
(EMM2) supplemented with the appropriate additives (Leu, Lys, His,
Ade, Ura). For live-cell imaging, cells were grown either in YE5S or in a
mix of YE5S and EMM2 at a ratio of 1:8 (MiMe), where indicated. Yeast
strains expressing slp1⫹ from the Pnmt41 promoter were grown in EMM2,
and thiamine (vitamin B1 [VB1]) was added to 15 ␮M to repress slp1⫹
expression. For microtubule depolymerization, methyl 2-benzimidazolecarbamate (BCM; Sigma-Aldrich) was added to the medium to a
final concentration of 50 ␮g/ml. For histone deacetylase (HDAC) inhibition, trichostatin A (TSA; Sigma-Aldrich) was added to a final
concentration of 12.5 to 25 ␮g/ml. To inhibit the Aurora B kinase
mutant ark1-as3, 4-amino-1-tert-butyl-3-(1=-naphthylmethyl)pyrazolo[3,4-d]pyrimidine (1NM-PP1; Toronto Research Chemicals) was
added to a final concentration of 5 ␮M.
To accumulate cells in G2 phase for imaging, 50 ml of a culture at an
March 2013 Volume 33 Number 5
optical density at 600 nm (OD600) of 0.5 to 0.8 was centrifuged for 2 min
at 2,200 ⫻ g. The pellet was resuspended in 750 ␮l H2O, layered on top of
a 7 to 30% lactose gradient, and centrifuged for 8 min in a swing-out rotor
at 210 ⫻ g with minimum acceleration and deceleration. A fraction of 300
␮l from the top layer was immediately diluted in 1 ml medium, centrifuged for 2 min at 960 ⫻ g, and resuspended in 200 ␮l medium. Sporulation of diploid cells was induced at 25°C by shifting them to EMM2 lowglutamate medium for 13 to 15 h before the onset of imaging. Genotypes
are listed in Table S2 in the supplemental material.
Yeast strain construction. To generate S. pombe strains with fluorescent reporter arrays at centromere and arm regions of chromosome I
(cen-arm), a gene encoding the tetracycline repressor (TetR) fused to tandem Tomato fluorescent protein (tdTomato) (38) was integrated into the
Z locus of strain C1280 containing a lactose operator (LacO) array at lys1⫹
and expressing the lactose repressor (LacI) fused to GFP (39). Tandem
arrays of the tetracycline operator (TetO) sequence were subcloned from
pRS306-TetO2⫻112 (40) into pFA6a-hphMX4 (41) via SacI/SapI. Five
intergenic chromosomal regions on chromosome I and one on chromosome II were PCR amplified and cloned into the NotI site of pFA6ahphMX4-TetO2⫻112 (see Table S3 in the supplemental material for
chromosomal locations, PCR primers, unique restriction sites for integration, and TetO and LacO repeat numbers). Correct integration of the
linearized plasmids was confirmed by fluorescence microscopy, colony
PCR, and Southern blot analysis.
To generate strains with fluorescent reporter arrays at different arm
regions of chromosome I (arm-arm), a tandem array containing 256
lactose operators (LacO) was cloned from pAFS52 (42) into pFA6anatMX6 (41) via SacI/SapI. Three intergenic chromosomal regions
(see Table S3 in the supplemental material) were cloned into the NotI
site of pFA6a-natMX6-LacO⫻256. The Tnmt1 terminator region from
pREP1 (43) was inserted into the XmaI/SacI sites of pJK148 (44). The
SalI/SmaI fragment from pNATZAtetR-tdTomato (38) encoding tdTomato was inserted to yield pJK-tdTom-Tnmt1. The Pura4 promoter
region was PCR amplified from chromosomal DNA and cloned via
SalI/BamHI into the resulting plasmid. Finally, the TetR was PCR
amplified from pNATZA and inserted into the BamHI site, yielding
pJK-Pura4-tdTom-Tnmt1. This plasmid was integrated along with plasmids pFA6a-natMX6-LacO⫻256 and pFA6a-hphMX4-TetO2⫻112
into strain C2616 expressing LacI-GFP. Correct integration was confirmed by colony PCR and Southern blotting.
To generate a strain with fluorescently labeled tubulin, the atb2⫹ gene,
including the promoter and terminator regions, was PCR amplified and
cloned into pJK148 via XmaI/SacI. The coding sequence for monomeric
Cherry fluorescent protein (mCherry) or enhanced cyan fluorescent protein (eCFP) was PCR amplified and cloned via BssHII or MluI/BssHII,
respectively, into an MluI site that had been introduced after the start
codon of atb2⫹ to yield pJK-Patb2-mCherry-atb2 and pJK-Patb2-eCFPatb2, respectively. The Patb2-mCherry-atb2 gene was then subcloned into
pFA6a-kanMX (41) via XmaI/SacI, and a PCR-amplified chromosomal
region for integration targeting was inserted into the NotI site of the
resulting vector.
Time-lapse microscopy. Cells were placed onto microscopy dishes
(MatTek; iBiDi) that had been coated with 1 mg/ml Bandeiraea simplicifolia lectin (Sigma) for 10 min and were left to settle for a minimum of 10
min. Dishes were preequilibrated on a DeltaVision microscope (Applied
Precision) equipped with a controlled temperature chamber and a Coolsnap-HQ ICX285 camera for 10 min. Images were recorded using an
Olympus UPlanApo objective at ⫻100 magnification (numerical aperture
[NA], 1.35; oil) at up to four preprogrammed positions with 8 z stacks at
a step size of 0.4 ␮m every 40 s for a period of 30 min to 1 h (mitosis) or
with 12 z stacks at a step size of 0.4 ␮m every 2.5 min for 3 h (meiosis),
using a dual-band filter set of GFP-dsRed for GFP or mCherry and tdTomato fluorescence controlled by softWoRx software. Exposure times varied from 60 to 100 ms to account for intensity variations of the Olympus
U-LH100HG fluorescence lamp.
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High-throughput imaging of the mutant library was performed in a
controlled-environment chamber on a Leica AF7000 wide-field microscope equipped with a Leica DFC3860FX camera. Images of 12 z stacks
with a step size of 0.41 ␮m at up to 12 positions were recorded for a period
of 45 min with 60-s intervals, using exposure times of 80 ms per GFP or
mCherry/tdTomato channel (YR2 495/595 dichroic filters for excitation/
emission) and a Leica HCX PlanApo objective at ⫻63 magnification (NA,
1.30; glycerol).
The ImageJ macro 3D Distance Tool (http://rsbweb.nih.gov/ij/macros
/tools/) was used to determine the distance between marker arrays. In the
few cases where one of the marker arrays split, the distance between the
nearest markers was measured. Time-lapse sequences of strains expressing mCherry-tubulin were deconvolved using Huygens image analysis
software (http://www.svi.nl/HuygensSoftware).
Automated image analysis. Preprocessing, segmentation, and distance measurement steps were implemented as a plug-in in ImageJ opensource software (45). The plug-in is available for free download from http:
//cmci.embl.de. Noise was first removed by band-pass filtering (lower, 2
pixels; upper, 10 pixels). Since photobleaching of fluorescence signals
caused undersegmentation, especially in the later part of the image sequences, images were corrected for bleaching by a histogram-matching
algorithm (46). The pixel intensity histogram of the three-dimensional
stack of the first frame was used as a template, and the histograms of stacks
in successive frames were matched by extending a code written by Burger
and Burge (46). These two steps stabilized and improved both signal-tonoise ratios and baseline intensities through the time course.
After preprocessing, locus signals were binarized by thresholding for
each channel individually. Threshold levels were adjusted automatically
in each three-dimensional stack so that the volume of segmented loci was
within a specified range, which was determined from empirical studies.
The three-dimensional boundary for each locus was detected using the
functionality of the 3D object counter plug-in. Centroid coordinates of
the signals were then extracted for each locus using a center-of-mass algorithm, and the distance between two loci at each time point was calculated. Locus splitting was seen along with nuclear division, and for time
frames with split loci, pairs were determined according to the nearestneighbor principle.
Isolation of ts mutants and mutant analysis. To generate an S. pombe
temperature-sensitive (ts) mutant library, random mutations were introduced into cells of strain C2926 using a UV light mutagenesis protocol
(47). Plates were incubated at 23°C for about 4 days until colonies appeared, which were then replica plated onto YE5S plates containing 20
mg/liter phloxine B (Merck) and incubated overnight at 34°C. Colonies
that appeared dark pink or red at 34°C were identified, and the corresponding colonies from the 23°C plate were picked as potential candidates, rechecked for the temperature-sensitive phenotype, categorized
microscopically, and frozen as glycerol stocks.
Time-lapse images were recorded for ⬃300 mutant strains that
showed similar morphological phenotypes as condensin mutants at 36°C.
For one-third of the recordings, no distance graphs could be calculated
due to insufficient numbers of dividing cells or high intracellular background fluorescence. Defects in prophase chromosome condensation
were detected in 24 of the ⬃200 average-distance graphs. Twenty-two of
the 24 mutants could be backcrossed three times to wild-type strain
C3041. After backcrossing, the mutants were reimaged on the DeltaVision
microscope, and new average-distance graphs were generated and analyzed by curve fitting. At least one condensation parameter differed significantly (⬎2 standard deviations) from that for the wild type for half of
the reimaged mutants (see Table 3).
Mapping of ts mutations by massive parallel sequencing. All colonies
from 3 to 6 tetrads of the third backcross were inoculated in 5 ml YE5S
medium and grown at 25°C. Equal numbers of cells from all ts cultures or
of all non-ts cultures were combined to a total of 70 to 360 OD units (2 ⫻
109 to 1 ⫻ 1010 cells/ml). Cell pools could be frozen at this stage after
addition of glycerol to 15%. About 20 ␮g genomic DNA was prepared
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from the ts pool and the non-ts pool using a Genomic-tip 100/G column
(Qiagen) following the manufacturer’s protocol.
DNA samples were bar coded, sequenced with a coverage varying between 4- and 100-fold, and mapped to the S. pombe reference genome
using the ELAND (version 2) alignment tool (Illumina Inc.). Alignment
files were redundancy filtered using the Picard library, and single-nucleotide variation (SNV) calling was performed with the SAMtools mpileup
tool (48). SNVs were intersected with information on gene-coding regions using the ANNOVAR framework (49). SNVs were classified as exonic, intronic, or intergenic. All exonic SNVs were further classified as
synonymous or nonsynonymous. Using custom scripts, all SNVs were
annotated with the nucleotide distribution at that position in the corresponding sample to determine the alternative allele frequency. Only
SNVs that occurred with an allele frequency of 1 in the ts sample pools
but were absent in the corresponding non-ts sample pools and in the
parental strains were considered a hit. For most mutants, only a single
hit was identified using this approach (see Table 3). A second mutation
in the tlh2⫹ gene in the pol2-am7 mutant could be excluded by a
further round of backcrossing and Sanger sequencing. One mutant
(cnd2-S1) was identified through genetic crosses and Sanger sequencing of the cnd2 gene.
Immunofluorescence detection. Immunofluorescence staining was
performed as described previously (50) after digesting the cell wall with 50
␮g/ml zymolyase T-100 (Seikagaku) for 30 min at 37°C. DNA was stained
with Hoechst 33342, and tubulin was detected with TAT1 antibody (a
kind gift from Keith Gull) and anti-mouse IgG antibodies coupled to
Alexa Fluor 594 (Invitrogen). Antibodies for Western blot detection of
Slp1 were a kind gift from Tomohiro Matsumoto.
Mathematical data fit. In order to extract quantitative parameters
from the compaction curves for each experiment, the average and standard deviations of the distances (in pixels) at each time point were computed. Using the nonlinear least-squares (nls) function of the R programming language, the following function was fitted to the averaged curve for
the 20 time points before anaphase onset: d ⫽ [c/(1 ⫹ ea ⫻ t ⫹ b)] ⫹ dmax,
where d is the distance between the marker arrays in pixels and t is the time
in number of frames relative to anaphase onset. The parameters produced
by the curve fitting procedure are dmax, a, b, and c, where dmax is the
maximum distance defined as the upper asymptote, a is the decay rate, b is
proportional to the inflection point, and c is the difference between the
asymptotes. The following parameters were derived from the fitted curve:
compaction ratio (r), which is equal to dmax/dmin, where dmin is the minimum distance; duration (tdur), which is equal to t95% ⫺ t5%, where t95%
and t5% are the times required for 95% and 5% compaction, respectively;
and timing to anaphase (tana), which is equal to t0 ⫺ t50%, where t0 and
t50% are time of anaphase onset and the time from anaphase onset to 50%
compaction, respectively.
RESULTS
A live-cell imaging assay for chromosome condensation in fission yeast. Our chromosome condensation assay is based on the
presumption that the physical distance between any two loci located on the same chromosome arm should decrease as chromosomes condense between the entry into cell division and sister
chromatid splitting at anaphase onset (Fig. 1A). To track chromosomal loci by live-cell imaging, we integrated tandem arrays of
prokaryotic lactose operator (LacO) or tetracycline operator
(TetO) sequences into two different positions of chromosome I,
the longest fission yeast chromosome, and labeled them by the
simultaneous expression of lactose repressor fused to green fluorescent protein (LacI-GFP) and tetracycline repressor fused to
tandem Tomato fluorescent protein (TetR-tdTomato) (Fig. 1B).
These labeled loci can be identified as distinct diffraction-limited
dots by fluorescence microscopy (38, 39) (Fig. 1C) (see Movie S1
in the supplemental material).
Molecular and Cellular Biology
Quantitative Analysis of Chromosome Condensation
FIG 1 Live imaging of chromosome condensation in fission yeast. (A) The three-dimensional distance between two fluorescent marker arrays located on the
same chromosome arm is expected to decrease during mitotic chromosome condensation; (B) different chromosome loci were fluorescently labeled by
integration of TetO or LacO arrays and expression of TetR-tdTomato (TetR-tdTom) or LacI-GFP; (C) images of mitosis in a cell in which chromosome I was
labeled with the 1.2-Mb cen-arm array (strain C2926) (see Movie S1 in the supplemental material).
For the first set of experiments, we generated fission yeast
strains with the LacO array positioned close to the centromere of
chromosome I and the TetO array positioned at increasing distances from the centromere along the chromosome arm (cen-arm
marker combinations). After enrichment of cells in G2 phase by
lactose gradient centrifugation, we recorded ⬃80 cells simultaneously by dual-channel wide-field fluorescence microscopy at 40-s
intervals for the course of 1 h. We selected the cells that had entered mitosis and initiated anaphase during this time period and
measured for each cell the distance between the three-dimensional positions of the GFP and tdTomato markers at every time
point (Fig. 2A). We then aligned the measurements from individual cells to anaphase onset, which we define by the splitting of the
centromere-proximal markers on the sister chromatids, and plotted mean distances over time (Fig. 2B).
As expected, the distances measured in G2-phase cells for
markers that were integrated closer to each other were lower than
the distances measured for markers that had been integrated further apart, ranging from 0.5 to 2.0 ␮m (Fig. 2C). Mean distances
remained constant until ⬃10 min before anaphase onset, at which
point they started to steadily decrease to reach approximately half
the G2-phase distances by the end of metaphase (Fig. 2B and Table
1). After anaphase onset, distances continued to decrease further.
The continued decrease in marker distances during anaphase is in
line with the continued shortening of chromosome arms described in mammalian cells (32, 33).
In order to correlate the observed changes in marker distances
with mitotic progression, we monitored mitotic spindle formation, extension, and disassembly by expression of ␣-tubulin fused
March 2013 Volume 33 Number 5
to the monomeric Cherry fluorescent protein (mCherry-Atb2) in
cells that contained the 1.2-Mb cen-arm marker array (see Fig. S1
and Movie S2 in the supplemental material). The distance between the marker dots started to decrease shortly after the appearance of mitotic spindle asters, consistent with the onset of chromosome condensation and mitotic spindle formation at the
beginning of prophase (51). The distance between the marker dots
continued to shorten during anaphase spindle elongation and
reached a minimum shortly before spindle disassembly during
telophase. The distances between the marker dots then started to
increase again, in agreement with chromosome decondensation at
the exit from mitosis (see Fig. S1 in the supplemental material). A
reason why distances did not increase to the same values as those
during G2 phase might be the confinement of chromosomes into
the smaller volume of G1/S-phase nuclei (52). Confinement
within a nucleus of ⬃3 ␮m in diameter presumably also limits the
maximum distance that two marker arrays can separate, which
explains why also during G2 phase we could not observe mean
distances of more than ⬃2.3 ␮m, even when we increased the
spacing between the arrays to 2.2 Mb (Fig. 2B and C).
To test if the decrease in distances between two chromosomal
loci is indeed a measure of global chromosome condensation and
not merely of local changes in chromatin structure at, for example,
centromeres, we generated a second set of strains that contained
both marker arrays on the left arm of chromosome I (Fig. 2D). For
all three arm-arm marker combinations tested, we measured a
decrease in distances within a 10-min period before anaphase onset similar to the distance changes that we had observed for the
cen-arm marker combinations. In order to further validate that
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Molecular and Cellular Biology
Quantitative Analysis of Chromosome Condensation
TABLE 1 Condensation parameters for differently spaced marker arrays
Markers (marker
spacing [Mb])
dmax (␮m)
dmin (␮m)
r
arm-cen (0.1)
arm-cen (0.7)
arm-cen (1.2)
arm-cen (1.7)
arm-cen (2.2)
arm-arm (0.5)
arm-arm (1.0)
arm-arm (1.5)
0.50
1.52
1.72
1.88
2.03
1.00
1.27
1.26
0.27
0.78
0.99
1.16
1.10
0.66
0.73
0.82
1.84
1.93
1.74
1.62
1.85
1.53
1.74
1.54
these changes in marker distances are caused by chromosome
condensation and not merely by clustering of chromosomes, we
measured distances for marker arrays located on the arms of two
different chromosomes (Fig. 2E). Even though the distances between the two marker arrays sharply dropped at anaphase onset,
presumably due to the concerted movement of the three sister
chromatids toward the same spindle pole, we could not detect the
pronounced decrease in distances during pro- and metaphase that
we had observed when the markers dots were located on the same
chromosome arm.
Condensation is maintained during metaphase and is independent of mitotic spindle formation or histone acetylation.
Our experiments so far do not exclude the possibility that chromosome attachment to spindle microtubules and/or biorientation and not chromosome condensation per se is responsible for
the observed changes in marker distances prior to anaphase. We
therefore sought to compare marker distances in cells arrested in
metaphase before and after spindle depolymerization. In order to
arrest fission yeast cells in metaphase with intact spindles, we
placed the gene encoding the anaphase-promoting complex activator Cdc20 (slp1⫹ in S. pombe) under the control of the thiamine
(VB1)-repressible Pnmt41 promoter (53). Even though steady-state
Slp1 protein concentrations were notably higher when expressed
from this promoter in the absence of VB1 (see Fig. S2A in the
supplemental material), they dropped to levels undetectable by
immunoblotting within 4 h of vitamin addition (see Fig. S2B in
the supplemental material). Notably, more than 90% of Pnmt41Slp1 cells arrested with a stable metaphase spindle within 3 h of
VB1 addition (Fig. 3A). Addition of VB1 had, in contrast, no effect
on the cell cycle progression of cells expressing Slp1 from its endogenous promoter (Fig. 3A). Slp1 repression therefore enables
synchronization of fission yeast cells in metaphase in a manner
analogous to that for Cdc20 depletion in budding yeast (54).
We next followed the distances between the 1.2-Mb cen-arm
marker arrays as cells entered the Slp1 arrest and aligned the measurements from individual cells to the time point that a stable
mitotic spindle had formed (Fig. 3B; see Movie S3 in the supplemental material). In agreement with the measurements of unperturbed mitoses, marker distances decreased approximately 2-fold
(from ⬃1.5 ␮m to ⬃0.7 ␮m) and remained stable at this distance
for at least 90 min of metaphase arrest. We next depolymerized
spindle microtubules in metaphase-arrested cells by addition of
BCM (see Fig. S2C in the supplemental material) and asked
whether marker distances would change in response to spindle
depolymerization. In contrast to the increase in chromosome
marker distances after mitotic spindle depolymerization observed
in budding yeast (35), we measured a further decrease in the cenarm marker distances (to ⬃0.6 ␮m) after BCM addition (Fig. 3C).
This is consistent with the report that fission yeast chromosomes
appear condensed after spindle inactivation using a cold-sensitive
tubulin mutant (55) and the hypercondensation observed for
chromosomes isolated from nocodazole-arrested mammalian
cells (1). We conclude that the decrease in distance between the
marker arrays is stably maintained in cells arrested in metaphase
and is independent of the presence of a mitotic spindle. It therefore presumably represents a direct measure of mitotic chromosome condensation.
Reports that histone acetylation of mammalian chromosomes
decreases during mitosis (56, 57) and that inhibition of HDACs
affects chromosome condensation in Xenopus oocytes (58)
prompted us to test whether HDAC inhibition had any effect on
the mitotic condensation of fission yeast chromosomes measured
by our assay. Since addition of the HDAC inhibitor TSA caused a
notable delay in mitotic entry of a considerable fraction of cells
when added during G2 phase (59), we added TSA to cells arrested
in metaphase by Slp1 depletion and assayed whether the distances
between the 1.2-Mb cen-arm arrays increased 1 h after addition of
the inhibitor (Fig. 3D). Even though TSA addition caused a slight
increase in marker distances, the increase was not statistically significant. In addition, chromosome condensation dynamics were
similar to those of wild-type cells in the small fraction (⬃20%) of
cells that entered mitosis with normal timing after TSA addition
during G2 phase (data not shown). These findings suggest that
histone deacetylation plays no major role in mitotic chromosome
condensation in fission yeast cells.
Quantitative and automated analysis of chromosome condensation. A considerable advantage of our assay is that it provides a time-resolved and quantitative measure of chromosome
condensation, which should enable us to describe the decrease in
distances between the marker arrays in mathematical terms. We
tested multiple alternatives to fit the data points for the period of
pro-/metaphase condensation (for details, see Materials and
Methods) and noted that the compaction behavior of wild-type
cells could be best described with a sigmoid curve. From this fit, we
extracted parameters for the maximal and minimal distances be-
FIG 2 Quantitative analysis of condensation dynamics. (A) Three-dimensional distances (d) between GFP- and tdTomato-labeled arrays were measured in the
image stacks. (B) Distances between a GFP-labeled LacO array at the centromere (cen) and tdTomato-labeled TetO arrays positioned on the left arm (arm) of
chromosome I (chrI) with increasing spacing (s) from cen were recorded by time-lapse microscopy at 36°C (strains C2572, C2574, C2570, C2568, and C2566).
The measurements from the indicated number (n) of individual cells were aligned to anaphase onset (time zero), and mean distances ⫾ standard deviations (blue
bars) were plotted. (C) Maximum cen-arm distances during G2 phase (dmax) were plotted against the spacing between the marker arrays (Table 1). The gray line
indicates a logarithmic fit (R2 ⫽ 0.998). (D) As in panel B, plotting of distances between a tdTomato-labeled TetO array 1.5 Mb from the left telomere and
GFP-labeled LacO arrays positioned with various spacings on the left arm of chromosome I (strains C2774, C2779, and C2724). (E) Distances between a
GFP-labeled LacO array 1.95 Mb from the left telomere of chromosome I (armI) and a tdTomato-labeled TetO array on the right arm of chromosome II (armII)
were plotted as in panel B (closed circles; strain C3245). Distances from the 1.2-Mb arm-cen combination with a similar G2 phase distance are plotted for
comparison (open circles; strain C2570).
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FIG 3 Chromosome condensation in metaphase-arrested cells. (A) Thiamine (VB1) was added to exponentially growing cultures of yeast cells expressing slp1⫹
from either its endogenous promoter or the Pnmt41 promoter (strains C1277 and C2489). The fraction of cells in interphase, metaphase, or anaphase judged by
immunofluorescence staining of tubulin and Hoechst staining of DNA was scored for 100 cells at the indicated time points after VB1 addition. (B) Cells
containing the 1.2-Mb cen-arm arrays and expressing slp1⫹ from Pnmt41 and mCherry-labeled ␣-tubulin (strain C3365) were enriched in G2 phase and incubated
for 1 h in the presence of VB1 before imaging at 32°C. Individual time traces were aligned to the first time that a stable metaphase spindle had formed (see Movie
S3 in the supplemental material), and mean distances ⫾ standard deviations (blue bars) were plotted. (C) Asynchronous cultures of cells expressing slp1⫹ from
Pnmt41 and eCFP-labeled ␣-tubulin (strain C2852) were grown at 30°C for 3 h in the presence of VB1 to arrest cells in metaphase. Then, BCM was added to half
of the culture for an additional 1 h, and microtubule depolymerization was confirmed by fluorescence microscopy (see Fig. S2C in the supplemental material).
Distances between 1.2-Mb cen and arm arrays in 100 cells from asynchronous or arrested populations were measured, and mean values ⫾ standard deviations
were plotted; P values were calculated from Student’s t tests. (D) As in panel C, but the HDAC inhibitor TSA was added instead of BCM.
tween the marker arrays (dmax and dmin) and calculated from these
values the compaction ratio (r) (see Fig. S3A in the supplemental
material). We also determined the duration of compaction, which
we defined to be the time required from 5% to 95% compaction,
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and the relative timing to anaphase, which we defined to be the
time from 50% compaction to anaphase onset.
In order to analyze large numbers of time-lapse recordings, we
established an automated image analysis pipeline for marker de-
Molecular and Cellular Biology
Quantitative Analysis of Chromosome Condensation
FIG 4 Chromosome condensation in condensin, Topo II, and Aurora kinase mutants. (A) Temperature-sensitive single or double mutant strains of condensin’s
Cut14 subunit (cut14-208) or Topo II (top2-191) containing the 1.2-Mb cen-arm array (strains C3005, C3061, and C3136) were imaged at 36°C. Time traces were
aligned to anaphase onset, and mean distances ⫾ standard deviations (blue bars) of the indicated number of cells (n) were plotted. (Insets) Sigmoid fits for each
plot (red) compared to the average fit of 4 independent plots of wild-type cells (green) (see Fig. S3D in the supplemental material). (B) Wild-type or analogsensitive ark1-as3 Aurora kinase mutant cells containing the 1.2-Mb cen-arm array (strains C2926 and C3130) were imaged after addition of 1NM-PP1 or
dimethyl sulfoxide (DMSO) solvent at 32°C. Distance-time traces of the indicated number of cells were aligned to anaphase onset, and average distances ⫾
standard deviations (blue bars) were plotted for each time point. (Insets) Sigmoid-fit curves (red) compared to the average-fit curve of 4 independent graphs
obtained for wild-type cells in the absence of the ATP analogue at 32°C (green) (see Fig. S3E in the supplemental material). No sigmoid curve fit could be obtained
for ark1-as3 in the presence of 1NM-PP1.
tection and distance calculation (see Fig. S3B in the supplemental
material and Materials and Methods). In short, image stacks were
first preprocessed to remove noise and to correct for photobleaching, segmented, and converted to binary images using an automatic threshold. Finally, the centroids of the marker array signals
were detected, and the three-dimensional distance between them
was calculated. A comparison between distance curves calculated
manually or automatically showed only a very subtle variation
between the two detection methods (see Fig. S3C and Table S1 in
the supplemental material), confirming that the automatic pipeline was able to reliably reproduce distance measurements.
The contributions of condensin, topoisomerase II, and Aurora kinase to mitotic chromosome condensation. To test
whether the automated assay can quantitatively assess differences
in mitotic chromosome condensation dynamics between wildtype cells and condensin or Topo II mutants, we generated yeast
strains that contained the 1.2-Mb cen-arm marker arrays and tem-
March 2013 Volume 33 Number 5
perature-sensitive alleles of the condensin subunit Cut14 (cut14208) (60, 61), Topo II (top2-191) (62), or both (cut14-208 top2191). We then imaged cells at 36°C to inactivate the mutant alleles.
While the centromere-proximal sister arrays segregated correctly
toward opposite cell poles at anaphase onset in both single mutants and the double mutant, the arm marker arrays frequently
failed to separate (see Movies S4 and S5 in the supplemental material), which is consistent with the segregation failures originally
described for these mutants (63). Segregation of centromere but
not arm markers resulted in a sudden increase in intermarker
distances after anaphase onset, which could be clearly identified in
the distance graphs (Fig. 4A).
In order to quantify pro-/metaphase condensation dynamics
in the mutants, we fit the last 20 data points before entry into
anaphase with sigmoid curves (Fig. 4A, insets), extracted fit parameters, and compared them to the values generated from several
independent measurements of wild-type cells (Table 2). As ex-
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TABLE 2 Condensation parameters for condensin, Topo II, and Aurora kinase mutants
Parameter and genotype (solvent)
dmax (␮m)
Avg for wild type at 36°C
cut14-208
top2-191
cut14-208 top2-191
Avg for wild type at 32°C
ark1⫹ (1NM-PP1)
ark1-as3 (DMSO)
ark1-as3 (1NM-PP1)
a
b
dmin (␮m)
r
Duration (min)
Timing (min)
1.36 ⫾ 0.07
1.34
1.18
1.40
0.66 ⫾ 0.02
1.11
0.84
1.27
2.06 ⫾ 0.13
1.21
1.40
1.10
5.89 ⫾ 2.18
5.84
3.44
6.69
4.35 ⫾ 0.46
5.46
5.00
4.47
1.28 ⫾ 0.05
1.45
1.16
NDb
0.54 ⫾ 0.03
0.72
0.77
ND
2.38 ⫾ 0.13
2.02
1.51
ND
9.82 ⫾ 2.49
7.75
3.51
ND
4.80 ⫾ 0.82
3.72
5.81
ND
a
Data represent standard deviations.
ND, parameters were not determined when no sigmoid fit could be generated.
pected, the change in marker distances was considerably reduced
in the cut14 condensin mutant. This resulted in a compaction
ratio of only ⬃1.2, which is significantly less than the ⬃2-fold
compaction observed in wild-type cells. Likewise, the compaction
ratio in the top2 mutant was reduced to 1.4. These measurements
are in agreement with the condensation defects described for
cut14 and top2 mutants in fixed cells by FISH or DAPI (4=,6diamidino-2-phenylindole) staining (61), suggesting that our assay is capable of measuring condensation defects quantitatively in
live cells. Interestingly, combination of the cut14 and top2 mutant
alleles had an additive effect, resulting in a further reduction of the
compaction ratio to 1.1 (Table 2; note that a compaction ratio of
1.0 equals no change in distances). Condensin and Topo II therefore have to at least some extent nonoverlapping functions in the
formation of mitotic chromosomes in fission yeast.
To confirm that the condensation defects that we measured
were not restricted to marker combinations containing centromeric regions, which have high levels of condensin bound (36,
64), we introduced cut3-477 and cut14-208 mutant alleles into
yeast strains containing the 1.0-Mb arm-arm marker arrays. While
in wild-type cells the average distance of this marker combination
decreased from ⬃1.0 ␮m during G2 phase to ⬃0.6 ␮m during
pro- and metaphase, the decrease was considerably reduced in the
cut14 and cut3 mutants (see Fig. S4 in the supplemental material).
We also noticed a temporary increase in the arm-arm marker distances shortly after anaphase onset, even though this increase was
markedly lower than the distance increase that we observed for the
cen-arm marker combinations during anaphase (Fig. 4A).
Segregation defects had been observed after inactivation of
Ark1, the single Aurora kinase present in fission yeast, which plays
a role in loading condensin onto chromosomes (65–68). We
therefore tested whether inactivation of a version of Ark1 (ark1as3) that can be selectively inhibited by the addition of the ATP
analogue 1NM-PP1 (69) would result in pro-/metaphase condensation defects. Addition of 1NM-PP1 had only subtle effects on
the changes in 1.2-Mb cen-arm marker distances in wild-type cells
(Fig. 4B and Table 2). In ark1-as3 cells in the absence of the ATP
analogue, marker distances were slightly reduced during G2 phase
and temporarily increased during the first minutes of anaphase,
suggesting that Aurora kinase function might be partially compromised by the ark1-as3 mutation. Nevertheless, during pro- and
metaphase, the marker distances decreased to the same values as in
wild-type cells. In contrast, pro-/metaphase condensation was
greatly reduced in ark1-as3 cells after addition of 1NM-PP1. Upon
entry into anaphase, we observed stretching of chromatin (Fig. 4B;
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see Movie S6 in the supplemental material) similar to that in condensin and Topo II mutants (Fig. 4A). Our findings support the
notion that Aurora kinase activity contributes to chromosome
condensation in fission yeast cells.
Chromosome condensation during meiotic divisions. We
next tested whether our live-cell imaging approach can also be
used to assay chromosome segregation during meiotic cell divisions. We generated diploid yeast strains containing the 1.2-Mb
cen-arm marker arrays on one homolog of chromosome I
(heterozygous dots). We induced cells to enter meiosis, enriched
for cells at the horsetail stage of meiotic prophase, and then followed them by live microscopy as they passed through the first and
the second meiotic divisions. Since sister arrays cosegregate to the
same pole during the first meiotic division, we used the directional
movement of the centromere-proximal marker array toward one
pole to define the time point of anaphase I onset. Splitting of
centromere-proximal markers then defined anaphase II onset
(Fig. 5A). Since the time period between anaphase I and anaphase
II varied considerably between individual cells, we generated separate distance graphs for each meiotic division by aligning the
measurements from individual cells to either anaphase (Fig. 5B).
During the horsetail stage of premeiotic prophase, the cen-arm
marker arrays were, on average, ⬃1.4 ␮m apart, which is similar to
the distance that these markers were separated during mitotic G2
phase. Starting at about 30 min before anaphase I onset, the distance between the marker arrays started to steadily decrease to a
value of ⬃0.8 ␮m just before anaphase onset (Fig. 5B and C; see
Movie S7 in the supplemental material) and then continued to
decrease during segregation of dyad chromosomes during anaphase I, similarly to what we had observed during mitotic anaphases. Interestingly, we measured only a slight increase in distances during the transition from the first to the second meiotic
division, suggesting that chromosomes remain largely condensed
between divisions. At approximately 15 min before anaphase II
onset, the distances between the marker arrays started to decrease
further to reach a value ⬃0.5 ␮m before the onset of the second
anaphase. Distances continued to decrease during segregation of
chromatids during anaphase II to reach a minimum of ⬃0.3 ␮m
and remained at this value while spores started to form. We conclude that our condensation assay is able to capture the dynamics
of chromosome condensation not only during mitotic divisions
but also over the complete course of meiosis.
Identification of components of the chromosome condensation machinery. The availability of a highly sensitive and quantitative method to measure chromosome condensation dynamics in
Molecular and Cellular Biology
Quantitative Analysis of Chromosome Condensation
FIG 5 Chromosome condensation during S. pombe meiosis. (A) Meiosis in a strain with heterozygous dots. Anaphase I was defined by the directed movement
of the centromere-proximal marker arrays toward one cell pole (left); homologous recombination between the cen and arm markers could result in arm marker
splitting during anaphase I (right). Anaphase II was defined by sister marker splitting. (B) Sporulation of diploid cells in which one of the two homologs of
chromosome I was labeled with the 1.2-Mb cen-arm array (strain C3064) was induced at 25°C. Distance-time traces were aligned to the first time point at which
cen arrays moved toward one cell pole (anaphase I onset) or to splitting of the cen arrays (anaphase II onset), and separate mean distance ⫾ standard deviation
(blue bars) plots were generated. (C) Images of a cell passing through both meiotic divisions (see Movie S7 in the supplemental material).
vivo presents a unique opportunity to de novo identify cellular
components that play a role in mitotic chromosome condensation. Since we expected that any protein with a key role in condensation—like condensin, Topo II, or Aurora kinase—would most
likely be indispensable for cell proliferation, we decided to screen
yeast cells containing conditional mutations in essential genes for
mutants with condensation defects. We introduced random mutations into a yeast strain containing the 1.2-Mb cen-arm marker
combination using a UV light radiation protocol (47) and selected
for cells that showed normal growth at 25°C but no or severely
impaired growth at 34°C. This protocol resulted in a pool of
⬃1,100 temperature-sensitive mutants. To limit the number of
strains to be analyzed, we first grouped all mutants according to
the morphologies that the mutant cells displayed at 34°C. Since we
had noticed previously that at the restrictive temperature condensin and Topo II mutants accumulated mixed populations of small
nondividing cells and large cells with no or abnormal septa, we
March 2013 Volume 33 Number 5
decided to initially focus on the mutants from the pool that displayed similar morphologies.
We recorded time-lapse movies of ⬃300 mutant strains starting 1
h after shifting cells to the restrictive temperature and then used the
automated image analysis pipeline to generate average-distance
graphs. We identified a set of mutants that displayed obvious anaphase segregation defects (see Fig. S5A in the supplemental material)
similar to those of the condensin or Topo II mutants that we had
characterized previously (Fig. 4A). In addition, we identified a second
set of mutants that displayed no or only very mild anaphase defects
but whose condensation parameters differed significantly from those
of wild-type cells at 36°C (Table 3; see Fig. S5B in the supplemental
material). After genetic cleanup, we used a massive parallel sequencing-based approach to identify mutations in the two sets of mutants
(for details, see Materials and Methods). For each mutant, this approach identified a single nonsynonymous mutation that cosegregated with the temperature-sensitive phenotype. Strikingly, all five
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TABLE 3 Condensation parameters for condensin, pol2, and fhb1 mutants
Parameter or allele
Avg for wild type at 36°C
cut3-m26
cut3-l23
cut14-r8
cut14-aa14
cnd1-ae7
cnd2-s1
cnd2-ae9
cnd3-j29
pol2-am7
fbh1-c18
fbh1⌬
a
b
Mutation
dmax (␮m)
dmin (␮m)
r
Duration
(min)
Timing
(min)
L289F
S1116P
G10D
T558L
D368N
S85F
L103P
R205Q
S400F
D734N
Deletion
1.36 ⫾ 0.07a
NDb
1.39
1.26
1.37
1.38
1.76
ND
1.57
1.23
1.20
1.21
0.66 ⫾ 0.02
ND
0.85
0.77
1.06
1.06
0.95
ND
0.75
0.81
0.63
0.79
2.06 ⫾ 0.13
ND
1.64
1.64
1.29
1.30
1.85
ND
2.09
1.52
1.89
1.52
5.89 ⫾ 2.18
ND
1.48
6.60
0.73
4.02
19.97
ND
15.21
2.80
6.77
4.96
4.35 ⫾ 0.46
ND
3.65
2.21
5.78
3.93
9.66
ND
5.30
5.37
3.85
5.17
Data represent standard deviations.
ND, parameters were not determined when no sigmoid fit could be generated.
mutants of the first set and three of the five mutants of the second set
contained an amino acid substitution in one of the five condensin
subunits (Fig. 6 and Table 3).
In the second set of mutants, we also identified missense mutations in the pol2⫹ (also named cdc20⫹) gene encoding the catalytic subunit of DNA polymerase ε (pol ε) and in the fbh1⫹ gene
encoding F-box DNA helicase I. Since fbh1⌬ cells are viable at
25°C (70), we used the deletion mutant to measure average-distance graphs at 36°C (see Fig. S6 in the supplemental material).
The compaction ratio was notably reduced in the pol2-am7 and in
the fbh1⌬ mutant (Table 3), suggesting that pol ε and F-box DNA
helicase I are both required for mitotic chromosome condensation. In further support of this conclusion, we found that combinations of pol2-am7 or fbh1-c18 with the cnd1-ae7, cnd2-ae9, or
cnd3-j29 condensin mutants further reduced growth at the restrictive temperature (data not shown). The fact that our microscopybased screening using the live-cell condensation assay identified
new mutants (including three new condensin alleles) that are im-
FIG 6 Novel condensin mutants identified by the condensation assay. Screening of a library of conditional S. pombe mutants identified eight novel temperature-sensitive alleles in the five condensin subunits (white stars). Previously
characterized condensin mutants are indicated as a reference (gray stars).
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paired in pro-/metaphase condensation but show no obvious
chromosome segregation defects (see Fig. S5B and S6 in the supplemental material) demonstrates that—in contrast to previous
approaches—it can reveal genes involved in mitotic chromosome
condensation directly on the basis of changes in chromosome
compaction and independently of secondary effects.
DISCUSSION
A direct and quantitative method to assay chromosome condensation in fission yeast. In this study, we describe the development
of a live-cell imaging assay to record chromosome condensation
parameters during mitotic and meiotic divisions of Schizosaccharomyces pombe cells. We demonstrate that the three-dimensional
distances between two marker arrays located on the same chromosome decrease to approximately half their G2 interphase distance during pro-/metaphase, independent of the marker positions (Fig. 2). Distances then continue to shorten during
chromosome movements toward the cell poles. Strikingly, the
timing and extent of the changes in marker distances are similar, if
not identical, to the degree of chromosome condensation measured by alternative means in mammalian cells (32, 33), suggesting that the genetically tractable fission yeast system provides an
excellent model to analyze molecular mechanisms of mitotic
chromosome formation that are likely to be conserved in mammals.
Previous studies had found that in the budding yeast Saccharomyces cerevisiae, the distances between marker arrays integrated
on chromosome IV (the longest chromosome) or on chromosome XII (which contains the rDNA cluster) were shorter in mitotic cells than in interphase cells (35, 36). Surprisingly, several
other marker combinations did not display this behavior (35, 71),
implying that only certain chromosome regions (for example, the
rDNA repeats) display considerable condensation during budding yeast mitosis. In contrast, we find that in fission yeast, the
distances between all marker combinations tested decrease with a
similar timing and to a similar degree (roughly 2-fold) (Fig. 2B
and D). These findings are therefore consistent with the conclusion that our assay measures large-scale changes in chromosome
folding, which are independent of the specific loci where the
markers had been integrated. Moreover, we find that fission yeast
cells arrested in metaphase maintain chromosome compaction
for extended periods of time (Fig. 3B), which is not the case in
Molecular and Cellular Biolog
Quantitative Analysis of Chromosome Condensation
nocodazole-arrested budding yeast cells (35). Mitotic spindle depolymerization during the arrest instead causes a further decrease
in marker distances (Fig. 3C), which is consistent with the observed hypercondensation of chromosomes following spindle depolymerization in fission yeast (55) or mammalian (1, 17) cells.
Hypercondensation may be a consequence of the loss of tension
between sister chromatids upon microtubule depolymerization,
since we cannot detect an increase in condensation in metaphasearrested cells with an intact spindle. The release of bipolar tension
at anaphase onset might also explain why condensation proceeds
further during anaphase and telophase (Fig. 2; see Fig. S1 in the
supplemental material).
A major advantage of the live-cell condensation assay is the fact
that it provides a direct, time-resolved, and quantitative readout
for the compaction of yeast chromosomes, which should allow the
detection of even subtle alterations in the global structure of mitotic chromosomes (see below). Despite its sensitivity, the assay
does not detect significant changes in the distances between
marker arrays after inhibition of HDAC activity (Fig. 3D). The
decrease in global histone acetylation that has been observed for
mammalian mitotic chromosomes (56, 57) can therefore probably not account for most of the condensation that we measure in
fission yeast cells.
Condensin, Topo II, and Aurora kinase regulate pro-/metaphase condensation in S. pombe. To test the sensitivity with
which the assay system can measure defects in pro-/metaphase
condensation following inactivation of proteins with known
roles in the formation of mitotic chromosomes, we measured
condensation in fission yeast strains expressing temperaturesensitive versions of Topo II or condensin’s Cut14 subunit.
Consistent with the extended chromosome shapes that had
been observed in dividing cells of either mutant (61, 72), we
found that the prophase compaction ratios between cen-arm
marker arrays are reduced from ⬃2.1 to ⬃1.4 or ⬃1.2 in temperature-sensitive mutants of Topo II and Cut14, respectively
(Fig. 4A). Similarly, compaction of arm-arm arrays is greatly
decreased in mutants of either condensin structural maintenance of chromosomes (SMC) subunit (see Fig. S4 in the supplemental material). Simultaneous inactivation of condensin
and Topo II reduces the compaction ratio even further, which
implies that condensin and Topo II may perform at least to
some extent independent functions in condensation. It is notable that our assay is capable of detecting the additive effect of
condensin and Topo II inactivation, which previous methods
that assessed mitotic chromosome morphology qualitatively
might not have been able to discern (24). Nevertheless, marker
distances still decrease during pro-/metaphase even in the double mutant. This is consistent with the findings that mitotic
chromosomes can still form to appreciable degrees in mammalian cells depleted of condensin complexes (16, 17, 19) or Topo
II (23). While the condensation observed in these cells may be
the consequence of only partial inactivation of protein function in the temperature-sensitive yeast mutants or incomplete
depletion by RNAi in mammalian cells, we favor the possibility
that there exist other yet unknown factors that contribute to
mitotic chromosome condensation (see below).
Similar to condensin or Topo II mutants, fission yeast cells
depleted of the Aurora kinase Ark1 were found to display severe
chromosome segregation defects during anaphase, resulting in cut
phenotypes (73, 74). Consistent with a role of Aurora kinase in
March 2013 Volume 33 Number 5
chromosome condensation (35, 75), we found that upon inhibition of an analog-sensitive version of Ark1, the change in marker
distances during pro-/metaphase is greatly reduced and chromosomes frequently fail to segregate (Fig. 4C). One role of
Ark1 during condensation is the recruitment of condensin to
chromosomes by phosphorylation of its Cnd2 subunit (65, 67,
68, 76). Interestingly, the condensation that we measured during meiotic prophase (Fig. 5) was almost completely abolished
in ark1-as3 cells (data not shown), supporting a requirement of
Aurora kinase in chromosome condensation also during meiotic cell divisions.
A novel tool to identify components of the chromosome condensation machinery. To prove that the condensation assay can
be successfully applied to identify proteins that are required for
mitotic chromosome condensation, we screened a pool of fission
yeast strains with conditional mutations in random essential genes
using an automated format of the condensation assay. Remarkably, this screen identified temperature-sensitive mutants in all
five subunits of the condensin complex (Fig. 6). Five of the novel
condensin mutants displayed obvious anaphase segregation defects (see Fig. S5A in the supplemental material), which are similar
to those in the originally isolated Cut3 and Cut14 mutants (Fig.
4A). While previous methods to identify condensation mutants
needed to rely on these segregation defects (61), which may result
from mitotic defects independent of chromosome condensation
failure, our assay assesses chromosome condensation directly.
Based merely on pro-/metaphase condensation parameters, our
screen picked up three novel condensin mutants that showed at
most only very subtle segregation defects during anaphase (cut3m26, cut3-l23, and cnd3-j29) (see Fig. S5B in the supplemental
material). Thus, the live-cell assay is capable of detecting mutants
with condensation defects with unprecedented specificity and
sensitivity.
In addition to mutations in condensin subunits, we also isolated mutations in the catalytic subunit of DNA polymerase ε and
in the F-box DNA helicase I. While prophase condensation was
notably reduced in strains expressing the mutant version of pol ε
or lacking Fbh1, anaphase segregation was not notably affected
(see Fig. S6 in the supplemental material), explaining why an involvement of the two proteins in chromosome condensation had
not been detected previously. At this point, the potential roles of
pol ε and Fbh1 in condensation are unclear. pol ε functions during
S phase as the leading-strand replicase (77–79). It is possible that
inactivation of pol ε in cells enriched for late S and G2 phase
prevents the completion of DNA replication. Entry into mitosis
with partially unreplicated chromosomes might subsequently interfere with condensin binding to chromosomes and/or condensation, consistent with the reduction in chromosomal condensin
levels in mitotic budding yeast cells that had not undergone DNA
replication (80) and the condensation defects observed in Drosophila cells expressing mutant versions of the DNA replication
origin recognition complex (81, 82). The finding that pol ε can be
coimmunoprecipitated with condensin I from human cells following induction of DNA single-strand breaks suggests an alternative link between the two protein complexes (83). Fbh1 is involved in the processing of Rad51-dependent intermediates
generated during DNA damage repair by homologous recombination (84), a function that is essential for chromosome segregation during meiosis (85). It is conceivable that the presence of
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these unresolved intermediates impedes chromosome condensation (86).
Several attempts have recently been undertaken to identify
candidates for chromosome condensation factors, yet the identities of condensation factors like RCA are still unknown. Here, we
focused our proof-of-principle screen on yeast mutants that displayed morphological phenotypes similar to those of already characterized condensin mutants and consequently identified mostly
novel condensin alleles. When expanded to genome-wide screens
(63, 70), we expect that the assay that we presented will provide a
powerful new method for the identification and quantitative characterization of yet unknown chromosome condensation factors in
the future.
15.
16.
17.
18.
19.
ACKNOWLEDGMENTS
We thank Jutta Metz and Carmen Aguirre Hernández for excellent
technical assistance, Damian Brunner, Tony Carr, Jan Ellenberg, Iain
Hagan, Michael Knop, Johannes Lechner, Edward Lemke, Sabine
Strahl, and all members of the C. H. Haering lab for advice and suggestions, and Kathy Gould, Keith Gull, Silke Hauf, Tomohiro Matsumoto, Yoshinori Watanabe, Mitsuhiro Yanagida, and the Yeast Genetic Resource Center Japan for reagents and strains. We are grateful
for the outstanding support from all members of the EMBL Advanced
Light Microscopy Facility and from Vladimir Benes and Tobias Rausch
of the EMBL Genomics Core Facility.
This work was supported by the German Research Foundation (DFG)
Priority Programme 1384 and EMBL.
20.
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Molecular and Cellular Biology
Quantitative Analysis of Chromosome Condensation in Fission Yeast SUPPLEMENTAL TABLES
TABLE S1. Condensation parameters using manual or automatic measurements
Genotype
Max. distance (μm)
Min. distance (μm)
Compaction ratio
Duration (min)
Timing (min)
Manual
1.69
0.83
2.04
7.62
7.03
Automatic
1.49
0.70
2.13
8.05
7.38
TABLE S2. Yeast strains
Strain Genotype
C1277
C1283
C1388
C2489
C2566
C2568
C2570
Source
h+, ade6-M216
Kohli
+
h , lys1-131, his7-366, leu1-32, ura4-D18, ade6-M210
Javerzat
+
h , nda3-KM311, leu1-32, ura4-D18, his2
+
Wang
+
h , kanMX-Pnmt41-slp1 , ade6-M210 or ade6-M216
this study
-
+
+
this study
-
+
+
this study
-
+
+
this study
-
+
+
h , LacO::lys1 , LacI-GFP::his7 , ChrI 1.5Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32, ura4-D18, ade6-M210
h , LacO::lys1 , LacI-GFP::his7 , ChrI 1.95Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32, ura4-D18, ade6-M210
h , LacO::lys1 , LacI-GFP::his7 , ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32, ura4-D18, ade6-M210
C2572
h , LacO::lys1 , LacI-GFP::his7 , ChrI 3.6Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32, ura4-D18, ade6-M210
this study
C2574
h-, LacO::lys1+, LacI-GFP::his7+, ChrI 3.0Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32, ura4-D18, ade6-M210
this study
C2616
h-, LacI-GFP::his7+, lys1-131, leu1-32, ura4-D18, ade6-M210
this study
C2724
C2774
-
+
+
h , ChrI 3.0Mb::LacO-natMX, LacI-GFP::his7 , ChrI 1.5Mb::TetO-hphMX, TetR-tdTomato::leu1 , lys1-131, ura4-D18, ade6-M210
-
+
+
h , ChrI 1.95Mb::LacO-natMX, LacI-GFP::his7 , ChrI 1.5Mb::TetO-hphMX, TetR-tdTomato::leu1 , lys1-131, ura4-D18, ade6-M210
-
+
+
C2779 h , ChrI 2.49Mb::LacO-natMX, LacI-GFP::his7 , ChrI 1.5Mb::TetO-hphMX, TetR-tdTomato::leu1 , lys1-131, ura4-D18, ade6-M210
+
+
+
this study
this study
this study
C2852 h?, kanMX-Pnmt41::slp1 , LacO::lys1 , LacI-GFP::his7 , ChrI 2.49Mb:: TetO-hphMX, Z locus::TetR-tdTomato-natMX
Patb2-eCFP-atb2::leu1+
this study
C2926 h-, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32, ura4-D18, ade6-M210
this study
C2929 h?, cut3-477, ChrI 2.49Mb::LacO-natMX, LacI-GFP::his7+,ChrI 1.5Mb::TetO-hphMX, TetR-tdTomato::leu1+, ade6-M210
this study
+
+
C2930 h?, cut14-208, ChrI 2.49Mb:: LacO-natMX, LacI-GFP::his7 , ChrI 1.5Mb::TetO-hphMX, TetR-tdTomato::leu1 , ade6-M210
-
this study
+
C3005 h , cut14-208, LacO::lys1 , LacI-GFP::his7 , ChrI 2.49Mb::TetO-hphMX,Z locus::TetR-tdTomato-natMX,
ura4-D18, ade6-M210
this study
C3041 h+, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX
leu1-32, ura4-D18, ade6-M210
this study
C3061 h-, top2-191, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX
this study
+
+
-
+
+
C3064 h /h , LacO::lys1 /lys1-131, LacI-GFP::his7 /his7, ChrI 2.49Mb::TetO-hphMX /ChrI 2.49Mb,
Z locus::TetR-tdTomato-natMX/Z-locus, leu1-32/leu1-32, ura4-D18/ura4-D18, ade6-M210/ade6-M216
this study
C3130 h-, ark1-as3::hphMX, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb:: TetO-hphMX, Z locus::TetR-tdTomato-natMX
leu1-32, ade6-M210
this study
C3136 h?, top2-191, cut14-208, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX
this study
C3164
h-, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX,
ChrI 0.52Mb::Patb2-mCherry-atb2-kanMX, leu1-32, ura4-D18, ade6
this study
C3245
h-, ChrI 1.95Mb::LacO-natMX, LacI-GFP::his7+, ChrII 3.6Mb::TetO-hphMX, TetR-tdTomato::leu1+,
lys1, ade6-M210
this study
C3358
h+/h-, ark1-as3::hphMX/ark1-as3::hphMX, TetR-tdTomato::leu1+/ leu1+, Z-locus::TetR-tdTomato-natMX/Z-locus,
LacO::lys1+/lys1+, LacI-GFP::his7+/ his7+, ChrI 2.49Mb::TetO-hphMX /ChrI, ade6-210/ade6-216
this study
1
TABLE S2 (continued). Yeast strains
Strain
Genotype
Source
C3365
h?, kanMX-Pnmt41::slp1+, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX,
Z locus::TetR-tdTomato-natMX, ChrI 0.52Mb::Patb2-atbp2-mCherry-kanMX
this study
C3510
h?, fbh1::kanMX, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3532
h?, cut14-r8, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3533
h?, cut14-aa14, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3534
h?, cnd1-ae7, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3535
h?, cnd2-ae9, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3536
h?, cnd2-s1, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3537
h?, cut3-m26, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3538
h?, cut3-l23, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3539
h?, cnd3-j29, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
C3640
h?, pol2-am7, LacO::lys1+, LacI-GFP::his7+, ChrI 2.49Mb::TetO-hphMX, Z locus::TetR-tdTomato-natMX, leu1-32,
ura4-D18, ade6-M210
this study
TABLE S3. Integration of TetO and LacO sequences
Strain
Spacing
Position
Primers
Site
Repeats
gtagtagtGCGGCCGCCTTTATGTCTTGTTCTAGCATGG
catcatcaGCGGCCGCCCCCACGTCTAGCGTCTGCC
gtagtGCGGCCGCCGCACTTGAAACTTTCATGTC
catcaGCGGCCGCCATTCTACTCCGAAGTATAGGG
gtagtagtGCGGCCGCAAATTGAGCACGATCAATAACTG
catcatcaGCGGCCGCGATCTGATTTAGGCAGCGCAGAGC
gtagtagtGCGGCCGCCCGAATGATGCGTCAAATTAGC
catcatcaGCGGCCGCGCTCATATAGTTGGCATCGCCAC
gtagtagtGCGGCCGCCTTGATTTTAGTCCTTTATGC
catcatcaGCGGCCGCCGTAGACTACACGATCTAGGGAAC
gtagtagtGCGGCCGCAAATTGAGCACGATCAATAACTG
catcatcaGCGGCCGCGATCTGATTTAGGCAGCGCAGAGC
EcoRV
1×
SwaI
1×
NheI
1×
BsaAI
1×
XcmI
1×
NheI
2×
ChrI 1.5 Mb
gtagtagtGCGGCCGCCTTGATTTTAGTCCTTTATGC
catcatcaGCGGCCGCCGTAGACTACACGATCTAGGGAAC
XcmI
ChrII 3.6 Mb
gtagtagtGCGGCCGCGTACTGTATCATAGCGTACTATAG
catcatcaGCGGCCGCGCTAACGGATTTGATGCTCAGC
MfeI
2×
3×
2×
1x
BsaAI
1×
NheI
1×
SwaI
1×
BsaAI
1×
TetO cen-arm
C2572
0.1 Mb
ChrI 3.6 Mb
C2574
0.7 Mb
ChrI 3.0 Mb
C2570
1.2 Mb
ChrI 2.49 Mb
C2568
1.7 Mb
ChrI 1.95 Mb
C2566
2.2 Mb
ChrI 1.5 Mb
C2926
1.2 Mb
ChrI 2.49 Mb
TetO arm-arm
C2774
C2779
C2724
C3245
1.0 Mb
LacO arm-arm
C2774
0.5 Mb
C2779
1.0 Mb
C2724
1.5 Mb
C3245
2
ChrI 1.95 Mb
gtagtagtGCGGCCGCCCGAATGATGCGTCAAATTAGC
catcatcaGCGGCCGCGCTCATATAGTTGGCATCGCCAC
ChrII 2.49 Mb gtagtagtGCGGCCGCAAATTGAGCACGATCAATAACTG
catcatcaGCGGCCGCGATCTGATTTAGGCAGCGCAGAGC
ChrI 3.0 Mb
gtagtGCGGCCGCCGCACTTGAAACTTTCATGTC
catcaGCGGCCGCCATTCTACTCCGAAGTATAGGG
ChrI 1.95 Mb gtagtagtGCGGCCGCCCGAATGATGCGTCAAATTAGC
catcatcaGCGGCCGCGCTCATATAGTTGGCATCGCCAC
SUPPLEMENTAL FIGURES
A
14
Distance (d) cen-arm (μm)
10
1.5
8
6
1.0
Spindle length (μm)
12
2.0
4
0.5
2
0
0
-15
0
15
30
45
Time to anaphase onset (min)
merge
B
LacI-GFP
TetR-tdTom
tubulin-mCh
5 µm
-13.3
-12.0
-10.7
-9.3
-8.0
-6.7
-5.3
-4.0
-2.7
-1.3
0
1.3
2.7
4.0
5.3
6.7
8.0
9.3
10.7
12.0
Time to anaphase onset (min)
FIG S1 Correlation of chromosome condensation with spindle dynamics. (A) Cells in which chromosome I was labeled with the
1.2 Mb cen-arm array and α-tubulin was labeled with mCherry (strain C3164) were imaged by time-lapse microscopy at 32°C.
Distances between cen and arm arrays (blue circles) were measured in parallel to spindle length (green diamonds) and aligned to
the time point of cen array splitting. Error bars represent standard deviations. (B) Images of a cell measured in (A) (see Movie S2).
3
A
B
6h 30°C
6h 18°C
0h
nda3-KM311
Pslp1-slp1+
0h
72 kDa
72 kDa
2h
4h
Pnmt41-slp1+
6h
0h
2h
4h
6h
α-Slp1
Time after VB1 addition
α-Slp1
55 kDa
55 kDa
α-Tubulin
55 kDa
α-Tubulin
55 kDa
Tubulin-eCFP
C
5µm
+VB 1
+VB 1 +BCM
Distance (d) arm-arm (μm)
FIG S2 slp1+ promoter shut-off. (A) To estimate metaphase protein levels of Slp1, nda3-KM311 β-tubulin coldsensitive mutants (strain C1388) were either shifted to 18°C or maintained at 30°C for 6 h and Slp1 was detected by
immunoblotting. Note that due to its unstable nature, Slp1 could only be robustly detected in cultures of cells
arrested by activation of the spindle assembly checkpoint in nda3-KM311 cells but not in asynchronous cultures. (B)
Thiamine (VB 1) was added to cells expressing Slp1 from its endogenous promoter (Pslp1; strain C1277) or the nmt41
promoter (Pnmt41; strain C2489) and protein levels were estimated by immunoblotting. Even though Slp1 is expressed
from Pnmt41 at higher levels than from Pslp1, Slp1 was undetectable by 4 h after VB 1 addition.
1.5
cut3
+
cut14
+
cut3-477
cut14 +
cut3 +
cut14-208
n = 40
n = 44
n = 42
1.0
0.5
0
-45
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
Time to anaphase onset (min)
FIG S4 Chromosome arm condensation in condensin mutants. Wild-type, cut3-477, or cut14-208 temperature-sensitive condensin mutant
strains in which chromosome I was labeled with the 1.0 Mb arm-arm array (C2779, C2929 and C2930) were imaged at 36°C. Distance time
traces of the indicated number (n) of cells were aligned to anaphase onset and average distances ± standard deviation (blue bars) plotted for
each time point.
4
45
A
B
d max
LacI-GFP
TetR-tdTom
Timing
(t 50%- t 0 )
d min
Select regions of interest
0
Duration
(t 95% - t 5% )
C
Manual
2.0
Distance (d) cen-arm (μm)
1.5
1.0
Substract background
Normalize intensity
0.5
0
Automatic
2.0
1.5
1.0
Auto-threshold
0.5
0
-30
-25
-20
-5
-10
36°C
2.0
1.5
1.0
0.5
-10
-5
0
5
10
15
E
Distance (d) cen-arm (μm)
Distance (d) cen-arm (μm)
D
-15
0
Time to anaphase onset (min)
20
32°C
2.0
1.5
Determine coordinates
Compute distances
1.0
User quality control
0.5
-10
-5
0
Time to anaphase onset (min)
Align to anaphase onset
Generate average distance ± SD plot
FIG S3 Automated imaging and image analysis pipeline. (A) Condensation parameters. A sigmoid curve fit to the 20 time points before
anaphase onset defines maximum and minimum distances (d max and dmin), the compaction ratio (r = dmax/dmin), the duration of condensation as
the time between 5% and 95% compaction, and the timing of condensation as the time between 50% compaction and anaphase onset (t = 0).
(B) Image analysis pipeline. Cells that undergo mitosis within 1 h of imaging were selected manually (step 1) and submitted for automated
image processing (step 2). After thresholding to binarize images (step 3), the centroids of both marker positions were determined in 3D
(yellow and red dots) and distances between them (blue lines) were calculated (step 4). The correct assignment of the center positions was
confirmed visually before using distance time traces from individual cells for alignment and generation of an average distance plot. (C) An
identical dataset from wild-type cells (strain C2779) was analyzed either manually or using the automated pipeline. Note that automated
measurements always returned slightly lower distance values than manual measurements. (D) Time traces from 4 independent datasets of
wild-type cells (strain C2926) imaged at 36°C were fit to calculate an average condensation curve (green) and average condensation parameters
± standard deviations. (E) as in (D) but imaged at 32°C.
5
A
wild-type
6.0
Distance arm-cen (μm)
5.0
n = 38
2.0
cut14-r8
n = 30
2.0
cut14-aa14
n = 14
2.0
1.5
1.5
1.5
1.0
1.0
1.0
0.5
0.5
0.5
4.0
3.0
-10
-5
0
-10
-5
-10
0
-5
0
2.0
1.0
0
-45
Distance arm-cen (μm)
6.0
5.0
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
45
cnd1-ae7
cnd2-ae9
n = 16
n = 29
2.0
1.5
1.0
2.0
-45
-30
-15
0
15
30
45
cnd2-s1
n = 20
2.0
1.5
1.5
1.0
1.0
0.5
0.5
4.0
0.5
3.0
-10
-5
0
-10
-5
-10
0
-5
0
2.0
1.0
0
-45
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
45
Time to anaphase onset (min)
B
cut3-m26
Distance arm-cen (μm)
6.0
n = 16
2.0
cut3-l23
n = 20
2.0
1.5
1.5
cnd3-j29
n = 10
2.0
1.5
5.0
1.0
1.0
4.0
0.5
0.5
3.0
1.0
-10
-5
0.5
-10
0
-5
0
-10
-5
0
2.0
1.0
0
-45
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
45
-45
-30
-15
0
15
30
Time to anaphase onset (min)
FIG S5 Identification of novel condensin mutants. The indicated numbers of cells (n) of mutants that displayed anaphase segregation defects
(A, strains C3532-C3536) or no obvious anaphase defects (B, strains C3537-C3539) were imaged at 36°C and distance time traces for each
single cell aligned to the time of anaphase onset. Average distances ± standard deviations (blue bars) were plotted. Insets show sigmoid fit
curves (red) compared to the average fit of plots of wild-type cells (green, see Fig. S3D). No sigmoid fits could be generated for mutants cnd2ae9 and cut3-m26.
6
45
pol2-am7
Distance (d) cen-arm (μm)
2.5
n = 17
2.0
2.0
1.5
1.5
1.0
1.0
0.5
0.5
-10
-5
0
-10
-5
0
0
fbh1Δ
Distance (d) cen-arm (μm)
2.5
2.0
n = 16
2.0
1.5
1.5
1.0
1.0
0.5
0.5
0
-45
-30
-15
0
15
30
45
Time to anaphase onset (min)
FIG S6 Condensation defects in pol2 and fbh1Δ mutants. The indicated numbers of cells (n) of the
pol2-am7 (strain C3640) or fbh1Δ (strain C3510) mutants were imaged at 36°C and distance time
traces for each single cell aligned to the time of anaphase onset. Average distances ± standard
deviations (blue bars) were plotted. Insets show sigmoid fit curves (red) compared to the average fit
of plots of wild-type cells (green, see Fig. S3D).
SUPPLEMENTAL MOVIE LEGENDS
MOVIE S1 Time-lapse epifluorescence microscopy of mitosis in a fission yeast cell (strain C2926) with the 1.2 Mb cen-arm marker array
labeled with LacI-GFP (cen, green) and TetR-tdTomato (arm, red) imaged at 36°C. Frames were taken every 40 sec for 60 min.
MOVIE S2 Time-lapse epifluorescence microscopy of mitosis in a fission yeast cell (strain C3164) with α-tubulin Atb2 fused to mCherry
(red) and the 1.2 Mb cen-arm marker array labeled with LacI-GFP (cen, green) and TetR-tdTomato (arm, red) imaged at 32°C. Frames were
taken every 40 sec for 60 min.
MOVIE S3 Time-lapse epifluorescence microscopy of mitosis in a fission yeast strain containing the 1.2 Mb cen-arm marker array labeled
with LacI-GFP (cen, green) and TetR-tdTomato (arm, red), α-tubulin Atb2 fused to mCherry (red), and slp1+ under control of the Pnmt41
promoter (strain C3365) imaged at 32°C in the presence of 15 μM VB1. Frames were taken every 80 sec for 120 min.
MOVIE S4 Time-lapse epifluorescence microscopy of mitosis in a cut14-208 mutant strain with the 1.2 Mb cen-arm marker array (strain
C3005) labeled with LacI-GFP (cen, green) and TetR-tdTomato (arm, red) imaged at 36°C. Frames were taken every 40 sec for 60 min.
MOVIE S5 Time-lapse epifluorescence microscopy of mitosis in a top2-191 mutant strain with the 1.2 Mb cen-arm marker array (strain
C3061) labeled with LacI-GFP (cen, green) and TetR-tdTomato (arm, red) imaged at 36°C. Frames were taken every 40 sec for 60 min.
MOVIE S6 Time-lapse epifluorescence microscopy of mitosis in a strain with the 1.2 Mb cen-arm marker array labeled with LacI-GFP (cen,
green) and TetR-tdTomato (arm, red) and the analogue-sensitive ark1-as3 allele (strain C3130) imaged at 32°C in the presence of 5 µM
1NM-PP1. Frames were taken every 40 sec for 60 min.
MOVIE S7 Time-lapse epifluorescence microscopy of meiosis in a diploid yeast cell (strain C3064) with a heterozygous 1.2 Mb cen-arm
marker array labeled with LacI-GFP (cen, green) and TetR-tdTomato (arm, red) imaged at 25°C. Frames were taken every 150 sec for 180
min.
7