A system for imaging the regulatory noncoding
Xist RNA in living mouse embryonic stem cells
Karen Ng*, Nathalie Daigle†, Aurélien Bancaud‡, §, Tatsuya Ohhata║, Peter Humphreys║,
Rachael Walker║, Jan Ellenberg† & Anton Wutz*,║
* Research Institute of Molecular Pathology, Dr. Bohr-Gasse 7, 1030 Vienna, Austria
† Cell Biology and Biophysics Unit, European Molecular Biology Laboratory (EMBL),
Meyerhofstr. 1, 69117 Heidelberg, Germany
‡ CNRS ; LAAS ; 7 avenue du colonel Roche, F-31077 Toulouse, France
§ Université de Toulouse ; UPS, INSA, INP, ISAE ; LAAS ; F-31077 Toulouse, France
║ Wellcome Trust Centre for Stem Cell Research, Tennis Court Road,
Cambridge CB2 1QR, UK
Correspondence should be addressed to Anton Wutz or Jan Ellenberg
Phone: +44-1223-760234 fax: +44-1223-760241 email: email@example.com
Phone: +49-6221-387328 fax: +49-6221-38798328 email: firstname.lastname@example.org
Running head: Imaging of Xist RNA in living cells
Wordcount: 9282 (Abstract 200 words)
2 Supplementary Figures
1 Supplementary Movie
In mammals, silencing of one of the two X chromosomes in female cells compensates for the
different number of X chromosomes between the sexes. The noncoding Xist RNA initiates X
chromosome inactivation. Xist spreads from its transcription site over the X chromosome territory
and triggers the formation of a repressive chromatin domain. To understand localization of Xist over
one X chromosome better we aimed to develop a system for investigating Xist in living cells. Here,
we report successful visualization of transgenically expressed MS2-tagged Xist in mouse embryonic
stem cells. Imaging of Xist during an entire cell cycle shows that Xist spreads from a single point to
a steady state when the chromosome is covered with a constant amount of Xist. Photobleaching
experiments of the established Xist cluster indicate that chromosome-bound Xist is dynamic and
turns over on the fully Xist covered chromosome. It appears that in interphase the loss of bound Xist
and newly produced Xist are in equilibrium. We also show that the turnover of bound Xist requires
transcription and Xist binding becomes stable when transcription is inhibited. Our data reveal a
strategy for visualizing Xist and indicate that spreading over the chromosome might involve
dynamic binding and displacement.
In mammals the dosage difference that arises from the different number of X chromosomes between
the sexes is compensated by X inactivation (Heard and Disteche, 2006; Payer and Lee, 2008). Early
in female development one of the two X chromosomes is selected for inactivation in a random
manner. This process involves counting the number of X chromosomes, choice of which
chromosome to inactivate and keeping the other X active in a reciprocal manner (Minks and Brown,
2009; Barakat et al., 2010). Chromosome-wide inactivation of one X chromosome requires the
noncoding Xist RNA (Borsani et al., 1991; Brockdorff et al., 1991; Brown et al., 1991a; Brown et
al., 1991b). Xist is expressed from the future inactive X chromosome (Xi) and spreads over the X
chromosome territory (Clemson et al., 1996). Thereby, Xist serves as a paradigm for studying
noncoding RNA function in regulating chromatin organisation.
The process of random X inactivation is recapitulated during the differentiation of female
mouse embryonic stem cells. ES cells have therefore been extensively used for studying the
mechanism of Xist function. Accumulation of Xist over the chromosome leads to the exclusion of
RNA polymerase II and factors associated with transcription and splicing from the X chromosome
nuclear domain (Okamoto et al., 2004; Chaumeil et al., 2006). Loss of activating histone
modifications and recruitment of Polycomb group complexes lead to the formation of a repressive
compartment (Chaumeil et al., 2002; Chaumeil et al., 2006). Subsequently, chromatin
modifications including tri-methylation of histone H3 lysine27 and ubiquitination of histone H2A
contribute to the establishment of facultative heterochromatin on the Xi (Heard and Bickmore,
2007). It has been shown that the formation of a repressive compartment can be separated from
gene silencing by specific mutations within Xist. The 5´-end of Xist that contains the repeat A
sequence motif is required for gene repression (Wutz et al., 2002). Expression of Xist with a
mutation of repeat A leads to the formation of a repressive compartment without initiation of gene
repression (Chaumeil et al., 2006; Pullirsch et al., 2010). Recruitment of genes to the repressive
compartment requires repeat A and involves additional pathways (Blewitt et al., 2008; Agrelo et al.,
2009). From these observations it is clear that localization of Xist over the X chromosome does not
require gene silencing. Consistent with this previous studies have observed that expression of Xist
in most differentiated cells does not lead to the initiation of gene silencing whereas Xist RNA
localization appears normal (Beard et al., 1995; Wutz and Jaenisch, 2000; Sado et al., 2004;
Savarese et al., 2006).
Xist localization is not dependent on an X chromosomal context. Using transgenes it has
been shown that Xist can spread in cis over autosomes and cause repression of at least certain
autosomal genes (Lee et al., 1996; Herzing et al., 1997; Heard et al., 1999; Wutz and Jaenisch,
2000; Hall et al., 2002; Chow et al., 2007; Tang et al., 2010). This suggests that X-chromosome
specific sequences are not crucial for attachment of Xist to the chromosome. However, analysis of
translocation chromosomes indicates that the X chromosome might be more permissive for Xist
spreading (Popova et al., 2006). In interphase Xist localizes to the core of the X chromosome
territory that is composed of mainly noncoding sequences and genomic repeats (Chaumeil et al.,
2006; Clemson et al., 2006). Particularly, long interspersed elements (LINE) have been implicated
in X inactivation (Lyon, 2003; Tang et al., 2010). Taken together, these findings suggest that Xist
might initially attach to genomic repeat sequences. Xist has also been shown to be retained in the
nuclear matrix when chromatin is extracted (Clemson et al., 1996). The ability of Xist to attach to
the nuclear matrix has led to the proposal that Xist performs a structural role associated with the
stability of the Xi territory. Consistent with this idea chromosomal binding of Xist has been shown
to be stable with a half-life between 4 to 8 hours when transcription is blocked (Panning et al.,
1997; Wutz and Jaenisch, 2000; Sun et al., 2006). Xist localization requires the scaffold attachment
factor A (SAF-A) gene (Hasegawa et al., 2010). SAF-A has been shown to be associated with the
Xi nuclear matrix and contributes to a stable nuclear structure of the Xi in somatic cells (Helbig and
Fackelmayer, 2003). The binding of SAF-A on the Xi has been measured under conditions that do
not block transcription and shown to be statically bound with a half life over hours (Fackelmayer,
2005). The apparent static binding of Xist and SAF-A to the Xi raises the question how Xist is
distributed from the Xic locus over the chromosome territory.
Here we describe a new tool for studying Xist dynamics and apply it to investigate the
mechanism of Xist spreading. By visualizing the non-coding RNA in living mouse embryonic stem
cells we present data indicating that Xist dynamically associates with the chromosome.
In vivo labelling of Xist RNA in embryonic stem cells
In order to visualize Xist RNA in living cells we chose a system for fluorescently labelling RNA in
vivo. For this we adopted a strategy of tagging Xist with an RNA motif from the MS2 phage which
can be bound by an MS2 RNA binding protein. This system has been used extensively in imaging
cellular RNAs in a variety of organisms (Bertrand et al., 1998; Fusco et al., 2003; Fusco et al.,
2004; Janicki et al., 2004). A U to C mutation in this RNA sequence (C-variant) has been identified
that interacts much stronger with the MS2 binding protein and forms a complex with picomolar
binding affinity (Lowary and Uhlenbeck, 1987). We introduced 24 C-variant MS2-binding sites
(5’-ACAUGAGGAUCACCCAUGU-3’) at the 3´-end of Xist (Xist-MS2). For regulating Xist-MS2
expression we used a tetracycline-inducible promoter. Similar transgenes have been shown
previously to recapitulate silencing on autosomes in male mouse ES cells (Wutz and Jaenisch,
2000; Tang et al., 2010). The Xist-MS2 transgene was introduced into J1:R26/N-nlsrtTA ES cells
(Wutz and Jaenisch, 2000), which contained a tetracycline inducible transactivator expressed from
the ubiquitous ROSA26 locus (Figure 1A). Subsequently a transgene for expression an the MS2
RNA-binding protein GFP fusion (MCP-GFP) (Bertrand et al., 1998; Fusco et al., 2003; Fusco et
al., 2004; Janicki et al., 2004) was introduced into these XM ES cells (Figure 1B). The resulting
XMG ES cells contained seven copies of the Xist-MS2 transgene integrated into a single site in the
distal third of chromosome 7 (Figures 1, C-E).
We verified Xist expression by RNA FISH analysis using probes specific for Xist and MS2
sequences. Xist-MS2 accumulated in the vicinity of its transcription site upon induction of
expression with doxycycline (Figure 1F). Importantly, the Xist domain could also be observed by
immunostaining with an antibody detecting the RNA binding MCP-GFP fusion protein in XMG ES
cells showing that the MS2 RNA tag on Xist-MS2 was recognized by the MS2 RNA binding
protein in living cells (Figure 1G).
In order to determine if the MS2 modified Xist retained its function we performed an
analysis of chromatin modifications. We observed tri-methylation of histone H3 lysine 27 and
mono-ubiquitination of histone H2A lysine 119 overlapping the Xist-MS2 domain demonstrating
that transgenic Xist-MS2 expression led to epigenetic marks which are associated with X
inactivation (Figure 1H). To further assess gene silencing in the autosomal context we analyzed the
expression of several imprinted genes on chromosome 7. Imprinted genes are expressed from one of
the parental chromosomes, and therefore repression of genes that are expressed from the Xist-MS2
transgenic chromosome can be measured. We observed repression of the paternally expressed
imprinted Peg3, Snrpn, and Ndn genes on chromosome 7 upon Xist induction (Figure 1I). In
contrast the maternally expressed imprinted H19 gene was not changed upon induction suggesting
that the Xist-MS2 transgene had integrated into the paternally inherited chromosome. Xist-MS2
induction caused strong repression of Snrpn and weaker repression of Peg3. Ndn was slightly
repressed. Our data show that Xist-MS2 represses autosomal genes to various degrees. This is
consistent with a recent study that comprehensively analyzed gene repression by autosomal Xist
transgenes (Tang et al., 2010). Taken together our observations show that Xist-MS2 retains several
aspects of Xist function in an autosomal context.
Analysis of Xist localization during the cell cycle
We next attempted to image Xist in living cells. A clearly defined green fluorescent territory was
observed in the nucleus of living XMG ES cells after induction with doxycycline (Figure 2A). The
fluorescence signal was low and required optimizing of the imaging conditions to avoid
photobleaching. Using a confocal microscope which had been previously optimized for GFP
imaging we were able to capture a z-stack of images at high resolution under low excitation levels.
3D reconstruction of confocal images revealed a high density of Xist in the center and small clusters
in the periphery of the Xist domain which might reflect a heterogenous distribution of Xist on the
chromosome (Figure 2B; Supplementary movie 1).
In order to analyze spreading of Xist RNA we used time-lapse confocal microscopy.
Adjusting imaging conditions for recording multiple z series without significant bleaching allowed
recording at a fairly high resolution. First, we studied Xist localization after initial induction with
doxycycline in interphase (Figure 3A). Xist first appeared as a single diffraction limited focus of
less than 0.5 µm2 which progressively expanded reaching an area of 4.3 ± 0.8 µm2 within ~1.5
hours (Figure 3C). This expansion seemed to result from the binding of additional Xist molecules to
a larger area, as the local concentration of Xist remained constant (Figure 3D). During the
expansion of the Xist covered area, the number of Xist molecules increased and reached a stable
level within ~1.5 hours as judged by the total fluorescence intensity of the domain (Figure 3D). Our
observations give a first indication of the timescale for the spreading of Xist and suggest that a
significant amount of time is required to accumulate Xist RNA over the chromosome territory.
Xist dissociated from the chromosome when the cells entered into mitosis (Figure 3B). After
cell division Xist reappeared as a spot under conditions where Xist-MS2 expression was
continuously induced with doxycycline. We measured the reappearance of Xist after mitosis (Figure
3B). Strikingly, the kinetics of Xist accumulation after initial induction in interphase and during
postmitotic reestablishment were indistinguishable under our conditions (Figure 3, C and D). This
observation suggests that chromatin modifications and Polycomb complex recruitment that were
established during the initial spreading of Xist did not alter the kinetics of Xist spreading after cell
division in a measurable manner.
We were able to follow one cell through an entire division cycle (Figure 4, A and B). In this
cell we noted an intriguing biphasic behavior of the Xist covered area. Measurement of the area of
the Xist territory showed that a plateau was reached 1.5 hours after the initial detection of Xist
(Figure 4B). The area of the Xist cluster then remained stable for the next 3 hours and increased
again over the following ~2.5 hours, when it reached a second plateau. We were able to measure
this larger Xist cluster for 3 hours before the cell entered mitosis and Xist was displaced. To test if
the increase in the Xist territory size between the two plateaus could be caused by the progression of
the cell through S-phase, we measured the area of the Xist cluster in image series of cells that had
progressed through mitosis. Analysis of 10 individual cells showed that the Xist cluster size
increased from 6.1 ± 1.1 µm2 to 12.3 ± 2.5 µm2 between G1 and G2 phase of the cell cycle,
respectively. To further investigate this observation we measured the fluorescence intensity of the
Xist cluster in fixed XMG ES cells. Nuclei were classified as either G1 or G2 based on the total
DNA content as measured by DNA staining using Hoechst 33342 fluorescent dye. On average the
Xist territory contained twice as much GFP fluorescence intensity in G2 than in G1 (Figure 4C). We
further isolated G1 and G2 fractions of XMG ES cells by flow sorting. After staining DNA in living
cells with Hoechst 33342 dye a clear G1 peak and a broader S-G2 area was observed from which a
G2/M area could be selected (Figure 4D). Analysis of Xist by quantitative real time PCR showed
that G2 cells contain 3 times the amount of Xist of G1 cells when normalized to total RNA (Figure
4D). This compares to a twofold increase in Gapdh mRNA. Taken together our data show that
between G1 and G2 phase cells the area of the Xist cluster and the amount of bound Xist increase
consistent with the idea that replication of chromosomal DNA increases the capacity of the
chromosome to bind Xist.
Dynamics of Xist binding within the chromosome territory
In order to investigate the dynamics of Xist after the cluster had reached a plateau we performed
fluorescence recovery after photobleaching (FRAP) experiments (Cole et al., 1996). First we
investigated cells under continuous induction with doxycycline. We bleached chromosome bound
Xist by exposing an area overlapping the Xist cluster to high intensity light. Under our conditions
between 60% and 90% of the total fluorescence of the cluster were lost after bleaching. Timelaps
imaging after photobleaching showed that within 20-30 minutes fluorescence recovered to about
60% to 80% of the initial intensity throughout the territory and then appeared to remain stable
We next investigated Xist when new synthesis of Xist is inhibited by blocking transcription
with Actinomycin D (ActD). After blocking transcription the Xist territory remained stable for more
than one hour (Figure 5C). When the Xist cluster was photobleached in the presence of ActD
fluorescence recovery of Xist was not observed (Figure 5B). This suggested that Xist dynamics
required transcription. It was important to rule out any effects of ActD treatment. For this we
stopped the production of Xist by washing out doxycycline from cells that had been induced
overnight. We then carried out FRAP experiments on cells showing a stable Xist cluster 30 minutes
after doxycycline depletion. Under these conditions no recovery of the Xist territory was observed,
whereas in control cells which were cultured in the presence of doxycycline fluorescence recovered
rapidly (Supplementary Figure 1, A and B). Taken together these observations indicate that
chromosome bound Xist displays greater dynamics when transcription is not blocked.
In order to visualize the displacement of chromosome bound Xist from the chromosome
territory we used inverse FRAP (iFRAP) experiments. We photobleached the unbound pool of
MCP-GFP excluding the Xist cluster and measured the fluorescence of the Xist cluster over time
(Figure 6A). We observed that the total fluorescence intensity of the Xist cluster decayed rapidly
after iFRAP and reached a constant level after 10 minutes (Figure 6B). The decay of fluorescence
intensity over time fitted well to a single exponential model indicating a first order kinetics (Figure
6, C and D). Taken together our observation suggest that Xist reversibly attaches to the chromosome
consistent with a dynamic turnover of Xist within the established cluster.
Here we describe a system for imaging Xist RNA in living cells. Albeit, clearly the system does not
reflect endogenous X inactivation we can show that gene repression and histone modifications are
recapitulated by transgenic Xist-MS2 in an autosomal context. We suggest that our system might be
useful for initial observations of Xist spreading given that there has been little progress with
attempts for imaging endogenous Xist. Several considerations make our transgenic system
preferably. First, analysis of endogenous X inactivation requires imaging in difficult conditions
such as the early mouse embryo or differentiating ES cell cultures whereby cell types and
physiological parameters are changing. We can image in mouse ES cells which are a highly
homogenous culture system. Secondly, using inducible expression of Xist provides enhanced
experimental control over Xist transcription which enabled us to perform crucial control
Using this system we have confirmed previous studies of Xist localization in living cells and
made several new observations. Consistent with earlier work we show that Xist spreads from a
single spot over the chromosome until it is lost at mitosis. We extend previous studies by showing
that spreading from a single site leads to saturation of the chromosome with Xist. Our study also
gives a first idea of the time required to cover a chromosome with Xist. We determined that Xist
accumulated within 1.5 hours which is a significant amount of time at the initiation of X
inactivation. We further find that the amount of Xist approximately doubles between G1 and G2
phase of the cell cycle indicating that the binding capacity is proportional to the amount of
chromosomal DNA. This is compatible with the idea that distinct DNA sequences such as LINE
elements or chromatin regions act as Xist attachment sites and their duplication during replication
leads to an increase in Xist binding. One surprising finding is that the spreading of Xist after initial
induction and reestablishment of the Xist cluster after mitosis follow similar kinetics. This
observation suggests that chromatin modifications do not have a measurable influence on Xist
spreading. One limitation of this result is that we performed our studies in mouse ES cells where X
inactivation is reversible. Therefore, our data do not rule out that changes in Xist spreading might be
induced after entry into differentiation.
Our live imaging system afforded us the opportunity to analyze the dynamics of chromosome bound
Xist. Measurements by FRAP and iFRAP indicate that Xist is dynamically bound and is replaced by
newly synthesized Xist within the established cluster. This analysis critically depends on the
stability of the Xist MS2 MCP-GFP interaction. Exchange of MCP-GFP on the Xist-MS2 RNA in
the time scale of our measurements could seriously undermine our analysis. In vitro measurements
have previously estimated that the complex of the C-variant MS2 RNA motif and the RNA binding
protein dissociates with a half life of 408 minutes or more than 6 hours, which would certainly be
long enough for measuring in the 30 minute interval of our experiment (Lowary and Uhlenbeck,
1987). However, this might be changed in the in vivo context of our experiment. A recent study has
used a similar system to measure the dynamics or gurken mRNA in Drosophila egg chambers with
half-lives in the order of minutes (Jaramillo et al., 2008). In our setup quasi-static binding of Xist
when transcription is blocked (Clemson et al., 1996; Wutz and Jaenisch, 2000; Seidl et al., 2006;
Sun et al., 2006) can serve as an additional control by separating the two components of
fluorescence recovery. When Xist transcription is blocked fluorescence recovery can only come
from the exchange of bleached for green MCP-GFP molecules on the bound Xist-MS2. We did not
observe measurable recovery of fluorescence within 30 minutes after FRAP. Results after washout
of doxycycline or Actinomycine D transcription inhibition confirm a long lived interaction between
the MCP-GFP protein and the MS2-tagged Xist-MS2 in our system. We therefore believe that our
observations on the dynamic binding of Xist might be mechanistically relevant.
To obtain an estimate for the half-live of bound Xist in our system we have used the loss of
fluorescence in our iFRAP experiments. These data fitted well to a single exponential model where
the steady state Xist concentration in the cluster results from the balance of transcription of new Xist
and dissociation of old bound Xist from the territory (Figure 5C). From this a rate of dissociation for
Xist of 0.006 ± 0.0007 s-1 was obtained. This corresponds to a residence time of Xist within the
chromosome territory in the order of 3 min (Figure 6, C and D). How this dynamics compares to the
endogenous Xist is unclear. It has been reported that Xist is continuously transcribed even in
differentiated cells (Clemson et al., 1996). Therefore, turnover of Xist is expected to be associated
with the Xi. Our measurements might provide an upper bound for Xist turnover. Taking into
account that a high rate of Xist production from the 7 copies of Xist in XMG ES cells almost
certainly leads to overexpression of Xist to some degree it is likely that Xist is less dynamic during
normal X inactivation. Furthermore, it is unknown if the addition of the GFP protein molecules to
Xist could have influenced its behavior. However, we note that in our transgenic system several
features of X inactivation are recapitulated under conditions of highly dynamic Xist turnover.
Our iFRAP measurements consistently give shorter recovery times than our FRAP
measurements. This might be explained by the need to bind MCP-GFP molecules to newly
transcribed Xist-MS2 whereas the iFRAP experiment measures the dissociation of already MCP-
GFP labeled Xist. This suggest that a significant amount of time is required for MCP-GFP binding
which could involve folding of the MS2 RNA motif and diffusion of MCP-GFP molecules within
the chromosome territory. Albeit, dynamics measurements by fluorescence correlation spectroscopy
(FCS) indicate that MCP-GFP diffusion is not limiting our kinetic measurements (Supplementary
Figure 2), local effects within the chromosome territory could lead to diffusion kinetics that differ
substantially from the overall measurements in the nucleus. The idea of local effects in MCP-GFP
diffusion and binding could eventually explain the fact that we did rarely observe more than 80%
recovery of the Xist cluster after FRAP. Local binding of bleached MCP-GFP to newly transcribed
Xist is a likely explanation for this observation. Formally the difference between the kinetics
measured in our FRAP and iFRAP experiments could also be the result of different amount of light
induced changes or damage of chromatin during photobleaching. However, we consider this
unlikely as the low fluorescence signal of the Xist cluster and the nuclear MCP-GFP were easily
photobleached with relative low light intensity. Under these conditions nuclear damage can be ruled
out making it unlikely that Xist dynamics was affected by photobleaching.
Less than complete recovery of fluorescence also opens up the possibility that a fraction of
up to 20% of Xist is stably bound. We do not favor this idea as a fraction of 20% of Xist would have
been easily detected in our iFRAP experiments, where we find no indication. Notably, over 90% of
Xist within the cluster becomes stably bound when transcription is arrested showing that the switch
to stable binding is not caused by different populations of Xist.
In our study we observe that the turnover of Xist within an established cluster is twice as fast as the
initial establishment of the Xist cluster after mitosis or induction of expression. This suggests that
once the chromosome is covered with Xist newly synthesized Xist can bind more rapidly than on a
chromosome that is not decorated with Xist. We think this might reflect changes in the organization
of chromatin that are induced by Xist binding. Chromatin modifications might play a lesser role for
Xist binding as we have observed similar kinetics of Xist accumulation after initial induction and
after mitosis when chromatin modifications are inherited from the previous cell cycle.
The kinetics of Xist turnover with and without transcription differ by nearly two orders of
magnitude with half lives of 3 minutes and 4-6 hours, respectively. The very long half life of Xist
after transcription block indicates a very stable attachment and a corresponding high binding
energy. An interesting question is then how this substantial energy barrier is overcome when newly
synthesized Xist displaces chromatin bound Xist. We speculate that the binding energy could be
overcome if Xist were displaced by a stepwise takeover of individual interactions along the RNA.
Alternatively, Xist binding could be regulated consistent with a report that Aurora B kinase has a
role in the displacement of Xist during mitosis (Hall et al., 2009).
Observations from visualization of Xist in living cells have uncovered a dynamic behavior that
might be important for understanding the mechanism of Xist function. Further studies will be
needed to confirm our results during normal X inactivation. Xist has exclusively evolved in
placental mammals (Duret et al., 2006; Elisaphenko et al., 2008) and presents a model for
noncoding RNA function in chromatin compartmentalization (Heard and Bickmore, 2007). In the
light of recent discoveries of large sets of long noncoding RNAs (Rinn et al., 2007; Guttman et al.,
2009; Khalil et al., 2009; Ponting et al., 2009) live imaging might also be a useful approach for
studying the role and dynamics of other noncoding transcripts in mammalian cells. We are
confident that dynamics will prove a vital aspect of noncoding RNA function and our live imaging
system might introduce a new tool.
Materials and Methods
ptetOP-MS2 was generated by directional cloning of the BamHI(blunt)-SphI fragment of 24x MS2
(Fusco et al., 2003) into the EcoNI (blunt)-SphI site of ptetOP H/X vector (Wutz and Jaenisch,
2000). The ClaI-PvuI fragment was then inserted into ptetOP-Xist-PA vector (Wutz and Jaenisch,
2000) digested with ClaI-PvuI, giving ptetOP-Xist-MS2-PA. For pCAG-MCP-GFPnls the ClaI
digested nlsMCP(dIFG)-GFP fragment from plasmid hsp83-MCP(dIFG)-GFP (a gift from P.
Becker) was inserted into the EcoRI digested pCAG vector (Niwa et al., 1991).
Cell culture and generation of cell lines
The XMG cell line was generated by a two step random integration strategy into J1 ES cells, which
contained nlsrtTA targeted in the ROSA26 locus (Wutz and Jaenisch, 2000). Firstly, 100 µg
ptetOP-Xist-MS2-PA and 10 µg pPGKpuro plasmid, both linearized, were co-electroporated into J1
ES cells to create the XM cell line. Puromycin (2 µg/ml)-resistant colonies were individually
selected and screened for expression by Xist/MS2 RNA FISH after 24h of induction on ROBOZ
slides (CellPoint). Secondly, 100 µg pCAG-MCP-GFP-nls with 10 µg pPGKhygro, both linearised,
were co-integrated into the XM cell line. Stable expressing clones were identified after selection
with Hygromycin B (130 µg/ml) for 8 days. Positive clones were verified under the fluorescence
microscope after 24h induction on ROBOZ slides (CellPoint), and confirmed by flow cytometry
(FACsort; BD Biosciences). After further subcloning a cell line was isolated that allowed imaging
of Xist in the majority of cells.
RNA and DNA FISH analysis
ES cells were attached to poly-l-lysine coated slides (Sigma) by using a Cytospin 3 centrifuge
(Thermo Shandon). RNA FISH was performed as described (Wutz and Jaenisch, 2000). Briefly,
cells were permeabilized with CSK buffer/0.5 % Triton X-100 for two min, fixed with 4% PFA in
PBS for 10 min and dehydrated progressively in ethanol. Hybridizations were performed overnight
at 37ºC in a dark and humid chamber, followed by washing 3 times in 50% formamide/2xSSC, 3
times in 2xSSC for 5 min each at 39ºC, and once in 1xSSC for 10 min at room temperature. Slides
were counterstained with DAPI, mounted with coverslips and analyzed. The coordinates of the
cells on the slides were recorded on the microscope. Following RNase treatment (100 µg/ml) at
37ºC for 30 min, cells were denatured in 70% formamide/2xSSC at 80ºC for 10 min and rinsed in
2xSSC prior to overnight hybridization for sequential DNA FISH. For DNA FISH analysis cells
were fixed with methanol-acetic acid (3:1), dropped onto poly-l-lysine coated slides, denatured at
80ºC for 5 min, and subsequently hybridized.
Probes were labelled using random priming (PrimeIt, Stratagene) with cy5-dCTP or cy3-
dCTP (Amersham). Xist was excised from ptetOP-Xist-PA (Wutz and Jaenisch, 2000) and MS2
from 24x MS2 (Fusco et al., 2003).
Immunofluorescence and protein analysis
Immunofluorescence combined with RNA FISH was performed as described (Chaumeil et al.,
2006). For GFP immunofluorescence, cells were fixed for 10 min at room temperature in 4 % PFA
in PBS, permeabilized for 5 min at RT in 0.1 % Na-Citrate/ 0.1 % Triton X-100 and blocked for 60
min at RT in PBS containing 5 % (wt./vol.) BSA, 0.1 % Tween-20. GFP was detected using a
mixture of mouse monoclonal clone 7.1 and 13.1 anti-GFP antibody at 1:500 (#11814460001;
Roche) followed by Alexa A-11034 Fluor 488 goat anti-mouse IgG (H+L) at 1:500 (Molecular
Probes). For combined H2AK119ub1 / H3K27me3 immunofluorescence, cells were pre-extracted
in 100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 10 mM Pipes pH 6.8 and 0.5 % Triton for two
minutes at room temperature. Histone modification H3K27me3 was detected using α-H3K27me3
at 1:1000 (#6523; (Schoeftner et al., 2006)), followed by Alexa A-11034 Fluor 488 goat anti-rabbit
IgG (H+L) at 1:500 (Molecular Probes). H2AK119ub1 primary antibody α-ubiquityl-Histone H2A
at 1:50 (clone E6C5; #05-678; Upstate Biotechnology) followed by secondary antibody Alexa A-
11004 Fluor 568 goat anti-mouse IgG (H+L) at 1:500 (Molecular Probes). After
immunofluorescence, cells were fixed in 4 % PFA in PBS for 10 min at room temperature, de-
hydrated, hybridized and washed as described for RNA FISH.
For Western analysis ES cells were grown for two days without feeders. Nuclear extracts
were prepared and analyzed by SDS-PAGE as described (Schoeftner et al., 2006). GFP was
detected with a mixture of mouse monoclonal clone 7.1 and 13.1 anti-GFP antibody at 1:200
(#11814460001; Roche) and a HRP-conjugated AffiniPure goat anti-mouse IgG (H+L), 1:5000
(Jackson ImmunoResearch Laboratories. Inc.) secondary antibody. Chemiluminescence reagent
(ECL, Amersham) and imaging film (Biomax, Eastman-Kodak) was used for detection.
Live cell microscopy and data analysis
Confocal images were acquired on a Zeiss LSM510 microscope. For live cell observation, cells
were grown in LabTek chamber cover glasses (Nunc) overnight in the presence of feeders. 1 µg/ml
DOX was added to the cells 4 hours before imaging. Prior to imaging, the medium was changed to
pre-warmed CO2-independent medium without phenol red (Invitrogen) containing 1 µg/ml DOX,
unless otherwise stated, and was sealed with silicon grease. Where indicated, DNA was stained in
live cells by 0.2 µg/ml Hoechst 33342. For Actinomycin D (ActD) experiments, 20 µg/ml ActD
(Sigma) was added as a 2x solution to imaging medium for 10 min before the experiment. Single
confocal sections or three-dimensional stacks of live cells were captured automatically (Rabut and
Ellenberg, 2004) using a Zeiss LSM510 confocal microscope with a 63x oil immersion objective
(Gerlich et al., 2006). For FRAP experiments three iterations of photobleaching at 100-500-fold the
acquisition laser intensity with 100% transmission of 488nm laser on either the entire Xist cluster or
half the Xist cluster or the nucleoplasm were used. The first post-bleach frame was acquired
immediately after photobleaching. A time series of typically 6 confocal z sections (3 µm pinhole
with 2 µm z-intervals) was recorded with two minute intervals for a total of 30 min for untreated
cells, and 60 min for ActD treated cells.
Images were adjusted by the brightness and contrast, and filtered using a low pass filter
implemented in the LSM510 software, then assembled in ImageJ and cropped. For 3D
reconstruction Imaris (Bitplane) was used. Quantification was carried out using ImageJ.
Fluorescence recovery in photobleaching experiments was measured in fixed-size user-defined
cellular regions in 3D projection of the z-stacks at all time points. To measure the area of Xist
territories during cell cycle, images were thresholded with the mean background value plus two
times the standard deviation of the background. FRAP experiments, which show a signal-to-noise
ratio >2, and Xist cluster ~2 µm in diameter were quantified, were normalized to the initial
fluorescence-intensity distribution and to total fluorescence, thereby correcting for movements in z-
plane and for fluorescence decay due to postbleach acquisition (Rabut and Ellenberg, 2004). FRAP
and iFRAP experiments were normalized to the initial Xist cluster intensity, and the normalized
intensity value for the Xist cluster, Inorm, was measured for each time point:
where IXist and INu are the Xist cluster and total nuclear background corrected intensities,
respectively, and prebl is prebleached and postbl is postbleach.
Kinetic modeling: Xist is produced by transcription at a constant rate (k1), it then binds to
chromatin and its association is characterized by a dissociation constant (k2). At steady state,
production and dissociation are balanced and define the number of Xist bound to the chromosome,
Xistst=k1/k2. Xist dynamics can be described with a first order kinetics :
This equation was used to fit FRAP/iFRAP experiments, in which the MCP-GFP shuttling can be
For experiments in Supplementary Figure 1 involving washing out doxycycline for turning
Xist transcription off cells were grown on ibiTreat µ–Slide 8 well slides (ibidi GmbH) coated with
gelatine. Cells were cultured overnight in the presence of 1 µg/ml Doxycycline. Before imaging
doxycyline was washed out by 5 times replacing the medium with fresh prewarmed medium
without doxycycline. After 30 minutes Xist transcription became negligible. These ibidi slides
provided superior adhesion for ES cells allowing for repeated washes without cell displacement.
Best imaging conditions were found using a 63x oil immersion objective on a Leica SP5 confocal
microscope with inverted DMI6000 stands fitted with environmental cube housings for live cell
Fluorescence correlation spectroscopy (FCS)
FCS was performed at room temperature using Zeiss LSM FCS Confocor2 with a 40X water
immersion objective (NA=1.2) It records fluorescence intensity fluctuations, which were analyzed
using the auto-correlation function of the intensity signal that can be adjusted with the anomalous
diffusion model. This technique enabled us to measure the MCP-GFP nucleoplasmic diffusion
coefficient and concentration based on anomalous diffusion fitting (Haustein and Schwille, 2007):
with 〈N〉 the mean number of tracers in the confocal volume, τD the mean residence time in the
confocal volume, S2 the structure parameter, which is defined by the confocal volume equatorial to
axial dimension ratio, and β the anomaly parameter. According to Zeiss specifications, the confocal
volume axial and equatorial dimensions are typically 0.3 µm and 1.5 µm, respectively, so S was set
5 . 0
to 5. The fit using (S1) enables us to measure MCP-GFP nucleoplasmic residence time
(Supplementary Figure 2), and the nucleoplasmic concentration of MCP-GFP given the confocal
volume dimension. Combining FCS with confocal imaging, the onset of MCP-GFP concentration in
the Xist territory was subsequently extracted, providing an estimate for the number of MCP-GFP
bound to Xist. Assuming that 33 MCP-GFP bind to one Xist RNA (Fusco et al., 2003), the steady
number of Xist molecule could therefore be calculated.
Expression analysis of G1 and G2 phase cells
XMG ES cells were cultured overnight in the presence of 1 µg/ml doxycycline. Approximately 5 x
106 cells were harvested and incubated with Hoechst 33342 dye diluted 1:100 in medium containing
1 µg/ml doxycycline for 30 minutes at 37 ºC. Sorting of the cells was performed on a MoFlo cell
sorter (Dako). Total RNA was isolated from equal numbers of G1 and G2 phase cells using Trizol
(Invitrogen). RNA concentration was determined on a NanoDrop spectrophotometer.
Quantitative real-time PCR of gene expression was performed using the FastSYBR Green
Master Mix and the Step One Plus Real-Time PCR System (Applied Biosystems). Sequences for
primers for detection of Xist were
CAGGAGCACAAAACAGACTC. Gapdh was detected using the previously published primer set
Gapd F and Gapd R2 (Sado et al., 2005).
We thank L. Klein and M. Busslinger for critical discussion of the manuscript and P. Becker for
providing the hsp83-MCP(dIFG)-GFP plasmid. J.E. acknowledges support from EMBL and the
German National Research Council (DFG EL 246/2-2). A.B. was funded through a FEBS
postdoctoral fellowship. A.W. is supported by a Wellcome Trust Senior Research Fellowship (grant
reference 087530/Z/08/A). This work was supported by the IMP and by a grant from the Austrian
Science Fund (grant reference SFB17 FWF).
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