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Probing chromatin accessibility
Dr. Lars Hennig
Swedish University of Agricultural Sciences
Department of Plant Biology and Forest Genetics
PO-Box 7080
SE-75007 Uppsala, Sweden
Tel. +46 18 67 3326
Fax +46 18 67 3389
Breakthrough technology
Plant Physiology Preview. Published on July 8, 2013, as DOI:10.1104/pp.113.220400
Copyright 2013 by the American Society of Plant Biologists
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Measuring Arabidopsis chromatin accessibility using DNase I-PCR and DNase I-chip
assays
Huan Shu; Wilhelm Gruissem; Lars Hennig*
Department of Biology and Zurich-Basel Plant Science Center, ETH Zurich, CH-8092,
Zurich, Switzerland (H.S., W.G., L.H.)
Functional Genomics Center Zurich, University of Zürich/ETH Zürich, CH-8057 Zurich,
Switzerland (W.G.)
Department of Plant Biology and Forest Genetics, Uppsala BioCenter, Swedish University of
Agricultural Sciences and Linnean Center for Plant Biology, SE-75007 Uppsala, Sweden
(L.H.)
Science for Life Laboratory, SE-75007 Uppsala, Sweden (L.H.)
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1 This work was supported by the Sixth Framework Program of the European Commission
through the AGRON-OMICS Integrated Project (LSHG–CT–2006–037704), by the Swedish
Research Council, and by the Swiss National Science Foundation (31003A_ 130272 and
31003A_132971).
* Corresponding author; e-mail [email protected]
The author responsible for distribution of materials integral to the findings presented in this
article in accordance with the policy described in the Instructions for Authors
(www.plantphysiol.org) is: Lars Hennig ([email protected]).
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Abstract
DNA accessibility is an important layer of regulation of DNA-dependent processes. Methods
that measure DNA accessibility at local and genome-wide scales have facilitated a rapid
increase in knowledge of chromatin architecture in animal and yeast systems. In contrast,
much less is known about chromatin organization in plants. We developed a robust DNase I-
PCR protocol for the model plant Arabidopsis thaliana. DNA accessibility is probed by
digesting nuclei with a gradient of DNase I followed by locus-specific PCR. The reduction in
PCR product formation along the gradient of increasing DNase I concentrations is used to
determine the accessibility of the chromatin DNA. We explain a strategy to calculate the
decay constant of such signal reduction as a function of increasing DNase I concentration.
This allows describing DNA accessibility using a single variable: the decay constant. We also
used the protocol together with Agronomics1 DNA tiling microarrays to establish genome-
wide DNase I sensitivity landscapes.
One sentence summary
Differential DNA accessibility, which is established by local chromatin environments and
strongly affects gene expression, can be assayed by the presented protocol utilizing DNase I
digestion coupled to detection by PCR or on tiling arrays.
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INTRODUCTION
Chromatin has a major impact on genome organization and gene activity. Differential
accessibility of DNA is thought to be a major consequence of locally different chromatin
composition and structure (Li et al., 2007). Chromatin sensitivity to nucleases has proven to
be a powerful tool to probe DNA accessibility in chromatin. Frequently used nucleases
include deoxyribonuclease (DNase), micrococcal nuclease (MNase), and restriction enzymes.
The resolution of restriction enzymes is limited by their sequence specificity, and MNase is
more often used to determine nucleosome occupancy (Schones et al., 2008). Chromatin
sensitivity to DNase I has often been used to define the “openess” of chromatin relative to its
higher order structures. Its applicability has been manifested by detecting regulatory
elements, such as promoters, enhancers and insulators, as DNase I hypersensitive sites (Wang
et al., 2001, Crawford et al., 2004, Dorschner et al., 2004, Crawford et al., 2006, Sabo et al.,
2006, Boyle et al., 2008, Naughton et al., 2010, Pique-Regi et al., 2011). DNase I sensitivity
can also be used as a measure for the general accessibility of chromatin (Weil et al., 2004).
Initially, chromatin accessibility of local genomic regions to DNase I was probed by
Southern blotting (Mather et al., 1983, Bender et al., 2000, Wang et al., 2001, Bulger et al.,
2003). However, Southern blotting is tedious and lacks sensitivity, and the interpretation of
results can be challenging. Therefore, analysis methods based on PCR have been developed
(Pfeifer et al., 1991, Feng et al., 1992, McArthur et al., 2001, Dorschner et al., 2004, Martins
et al., 2007). In recent years, DNase I assays were coupled to high-throughput genome-wide
profiling strategies such as genome tiling arrays and next-generation sequencing (Crawford et
al., 2004, Sabo et al., 2004, Weil et al., 2004, Crawford et al., 2006, Sabo et al., 2006). While
much has been learned about accessibility of chromatin in animal and yeast systems, our
knowledge of chromatin accessibility in plants is limited. Most studies have focused on
selected genomic regions such as the GRF1 gene, the Adh1 and Adh2 genes in maize (Paul et
al., 1998a, Paul et al., 1998b), or the GRF gene, the Adh gene, and an 80 kb genomic region
harbouring 30 protein coding genes in Arabidopsis (Vega-Palas et al., 1995, Paul et al.,
1998a, Paul et al., 1998b, Kodama et al., 2007). The technique used in these reports was
exclusively DNase I treatment and analysis of accessibility using Southern blotting. More
recently, we have combined the DNase I sensitivity assay with whole genome tiling arrays in
Arabidopsis to generate a genome-wide chromatin accessibility profile (Shu et al., 2012).
Here we present a robust, optimized DNase I sensitivity assay protocol for
Arabidopsis tissues based on PCR. This protocol can be adapted to different samples or
experimental objectives; the strategies for optimizing each step are also discussed. Analysis
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of relatively large fragments by PCR has proven to be highly robust as a first step in probing
DNase I sensitivity in any region of the genome. We also introduce a new strategy for
presenting DNase I sensitivity of the tested regions using a decay constant calculated by
fitting PCR product intensity values from a gradient digestion. In this way, the sensitivity of
each region is characterized by a single value, facilitating comparisons between different
regions or samples. Finally, we describe how our protocol can be combined with genomic
techniques for genome-wide profiles of chromatin accessibility.
RESULTS AND DISCUSSION
DNase I PCR assays often monitor the digestion of nuclei over a time course (Feng et
al., 1992, Wang et al., 2001) or in a gradient of enzyme concentrations (Vega-Palas et al.,
1995, McArthur et al., 2001, Martins et al., 2007). Here we discuss the entire procedure in the
following sections: (1) Isolation of permeabilized nuclei, (2) DNase I digestion and DNA
recovery, (3) analysis of chromatin sensitivity by PCR and (4) data interpretation (Fig. 1).
Assays to probe chromatin accessibility should use freshly prepared nuclei. The time
for isolating nuclei and the delay before digestion should be minimized to reduce unspecific
damage to chromatin. Yield and reproducibility of the nuclei extraction can be pre-tested by
measuring the DNA content from identical or similar extractions. The key to reproducible
DNase I sensitivity assays is to keep an invariable DNase-to-chromatin ratio between
replicates. An optimal ratio has to be determined empirically for each tissue. The DNA
recovered from such digests is a mixture of long and short fragments depending on the
accessibility of the chromatin. Samples must be handled with much care to minimize
unspecific fragmentation. We recommend using large-pore pipette tips and gentle handling of
nuclei and chromatin suspensions. Rigorous mixing, e.g. by using Vortex mixers, must be
avoided. To control for possible differential sensitivity of the DNA template, a digestion of
isolated DNA can be included. Ideally, the DNA is extracted from identical material as the
nuclei to be used in the experiment.
Isolation of permeabilized nuclei
Plant material
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Because cells in expanded green tissues usually contain large vacuoles, the nuclei/cell
(w/w) ratio is much lower in older than in young tissues. In our experience, it is often more
efficient to isolate nuclei from plant cell cultures, inflorescences or young seedlings than
from mature green tissues. Fresh samples can be used for extracting nuclei. However, it is
often more practical to use frozen material. In this way, not only can plant material be stored
almost infinitely at -80˚C, but nuclei isolation can also be easily synchronized when multiple
samples are involved.
Isolation of nuclei
To minimize the time for nuclei isolation, we developed a one-incubation-step
protocol. The short isolation step also favors a higher yield of nuclei. The entire isolation
procedure should be carried out on ice or at 4˚C.
Frozen plant tissue is ground to a fine powder in liquid nitrogen to disrupt cell walls
and other fibrous structures. The disrupted plant tissue is then treated with a cell lysis buffer
to release the nuclei. Cells are lysed by a surfactant in the buffer for disrupting membrane
structures. A nonionic surfactant, e.g. Triton X-100 or Igepal CA 630, is normally used at
concentrations between 0.3 -1% (v/v) depending on the specific requirement of each
experiment (Bowler et al., 2004, Wagschal et al., 2007). Normally, a 15 min treatment with
cell lysis buffer containing 0.5% Triton X-100 is sufficient for most Arabidopsis green
tissues. It is advisable to include sucrose or hexylene glycol in the buffer (Bowler et al.,
2004) to maintain a physiological osmotic pressure. Such osmotic agent also increases
viscosity of the buffer system slightly and forms an additional protection for the nuclei. After
cell lysis, the lysate is filtered through multiple layers of Miracloth (Calbiochem, Germany)
or nylon mesh filters such as CellTrics (Partec, Germany) to remove cell debris. We
recommend the CellTrics as they guarantee a more reproducible nuclei yield. When
Miracloth is used, the Miracloth filter should be pre-saturated with cell lysis buffer before
filtering to minimize nuclei retention in the fabric. Furthermore, the Miracloth should not be
squeezed, as this can lead to considerable organelle (and organelle DNA) contamination and
therefore an unreliable nuclei yield estimation. After filtering, crude nuclei can be collected
by mild centrifugation (see the materials and methods section for details).
DNase I digestion
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DNase I time course or gradient digestion can be used, but the latter can be more
practical when taking advantage of multichannel pipettes and PCR cyclers for handling
multiple samples. In this way, adding enzyme or stop buffer to the reaction mixture can be
carried out in synchrony for all samples. PCR cyclers are robust instruments for providing
precise temperature control for the incubation of the digestion reaction (Fig. 2). The
procedure for the DNase I gradient digestion is detailed below and steps for time course
digestion need to be adjusted accordingly.
Before digestion, the crude nuclei are washed at least once with digestion buffer to
remove the detergent carried over from the cell lysis buffer. Afterwards, the nuclei are re-
suspended in digestion buffer. For easier handling, the nuclei suspension should be split into
equal aliquots of the planned gradient digestion steps (Fig. 2). Because isolated nuclei
sediment rapidly, equal aliquotting requires pipetting the nuclei suspension up-and-down
gently before taking each aliquot. In parallel, aliquots of a DNase I dilution series are also
transferred to PCR tubes. Once the reaction is ready to start, equal volumes of the enzyme
dilutions are added to the nuclei suspensions using a multichannel pipette (Fig. 2) and mixed
by pipetting up-and-down. The digestion mixtures should be immediately transferred to the
preheated PCR cycler for incubation (Fig. 2). After the desired reaction time (typically 3 to 5
min), digestions are stopped by adding stop buffer (EDTA and/or EGTA; see the materials
and methods section for details) to all the reaction mixtures using the multichannel pipette. If
the reactions are incubated at a sub-optimal temperature for DNase I activity, e.g. 30˚C
(Kodama et al., 2007, Song et al., 2010) instead of 37˚C, incubation times can be extended to
10 to 15min. Such an extended incubation time is advantageous because it reduces effects
caused by errors in handling time.
There may be no best number of DNase I dilutions fitting all experiments. A rule of
thumb is to include enough dilutions to detect smooth transitions of band intensities and
differences between insensitive and sensitive control regions for the particular experiment
(see below). In addition, a mock digested (no DNase I) or minimally-digested (highly dilute
DNase I concentration) step should be included as input chromatin control.
After the DNase I treatment is completed, DNA should be immediately purified to
avoid further degradation. We recommend against column-based DNA purification
techniques, because they usually retain large genomic DNA fragments and therefore
introduce a digestion bias. Instead, organic solvent-based extraction and ethanol precipitation
work best. Precipitation carriers such as glycogen can be added for more efficient recovery of
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the DNA. DNA pellets should be immediately reconstituted in 10 mM Tris-HCl buffer (pH
8.0).
After a limited DNase I digest, a large proportion of the inaccessible DNA remains in
very long DNA fragments. Precipitated very long DNA can be difficult to resuspend.
However, we strongly advise against high temperature incubation, fierce vortexing or
pipetting of the pellet to facilitate dissolving. Instead, the DNA pellet should not be over-
dried and reconstitution buffer should be added to the DNA pellet when it is still moist. After
adding the buffer, samples can be incubated on ice or in the refrigerator for 30 min to
overnight. The DNA samples are then gently pipetted or tapped to ensure homogenous
distribution in the buffer. Repeated freeze-thaw cycles should be avoided by storing the DNA
samples at 4°C. We found that recovered DNA samples can be stored at 4°C for at least one
month without detectable changes in the PCR results (data not shown).
Analysis of digested DNA by PCR
Chromatin sensitivity to DNase I results in reduced accumulation of PCR products in
treated samples in comparison to input controls (Fig. 3). There are two major factors affecting
the signal reduction in such a DNase I PCR assay: (1) the ratio of DNase I to chromatin in the
reaction, and (2) the length of the PCR amplicon: The shorter the PCR amplicon, the lower
the probability that cleavage events occur within the probed region. Therefore, longer
amplicons are better suited to probe the DNase I sensitivity. Because of qPCR relies usually
on very short amplicons, its power is limited in such DNase I-PCR assays. However, qPCR
based strategies can be used if long stretches of genomic regions are probed by tiling
amplicons (Dorschner et al., 2004) or if prior knowledge of DNase I hypersensitive sites
allows precise positioning of amplicons (McArthur et al., 2001, Martins et al., 2007). In
contrast, when prior knowledge is lacking we found conventional PCR most powerful to scan
genomic regions for differential sensitivity. Usually, 1.5~2kb long amplicons (covering ~ 7 to
10 nucleosomes) work best at this stage. Once a sensitive region is identified, higher
resolution probing by qPCR can be performed.
Here, we describe a strategy to probe the sensitivity of a genomic region by
conventional PCR using decay constants calculated from gradient digestions. As example, the
chromatin sensitivity was analyzed for two silent transposable element (TE) genes (TA2 and
Cinful-like) and two active genes (GAPDHα and ACTIN7) in Arabidopsis. We and others
have shown that transcription is generally associated with accessible chromatin (Weil et al.,
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Probing chromatin accessibility
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2004, Boyle et al., 2008, Bell et al., 2010, Shu et al., 2012, Zhang et al., 2012). Thus, we
predict that the two TE genes should be much less sensitive to DNase I than the two active
genes and therefore can be used as insensitive and sensitive controls in future assays.
Reduced DNA accessibility is now also considered a silencing mechanisms established by
Polycomb group (PcG) proteins in animals (Bell et al., 2010, Eskeland et al., 2010, Naughton
et al., 2010, Bantignies et al., 2011) and plants (Shu et al., 2012). Therefore, two Arabidopsis
PcG target genes (AG and FT) were included in the assays.
In the absence of particular hypersensitive sites the probability of cleavage within a
region is proportional to the length of this region. Thus, it is important to maintain similar
amplicon sizes for the different regions to be probed. Because inefficient PCR and saturated
PCR will strongly distort results, PCR conditions have to be carefully optimized for each
primer pair by establishing at least (1) an optimal annealing temperature and (2) an optimal
cycle number within the exponential phase. Cycle numbers might differ between primer pairs
but should lead to comparable PCR signals for the input sample.
It is important to include insensitive and sensitive controls in a DNase I sensitivity
experiment. Conventional choices for the controls are DNA for heterochromatin regions and
active genes. If PCR conditions were correctly optimized but differences between insensitive
and sensitive controls are not easily seen, the DNase I digestion conditions are not
appropriate. Typical problems are over-digestion, under-digestion or damaged chromatin in
input nuclei. A gradient digestion with isolated DNA can also be included to control for
possible differential sensitivity of the DNA template, although this does not seem to be a
common problem (Sabo et al., 2006).
DNase I digestion of native chromatin and purified DNA was carried out in triplicate
(Fig. 4). For the chromatin digestion, PCR signals of the two heterochromatic TE genes
remained nearly constant while PCR signals of the two active genes dropped rapidly,
confirming our hypothesis of differences between heterochromatin regions and active genes.
This is in agreement with genome-wide profiles (Shu et al., 2012) and establishes these
regions as insensitive and sensitive controls for future assays. Interestingly, the sensitivity of
the two PcG targeted genes AG and FT seemed to be in between the TE genes and the active
genes. In contrast, there were no systematic differences in DNase I sensitivity of isolated
DNA: for all tested fragments, PCR signals became reduced following similar patterns. This
control confirms that (1) DNA in chromatin is indeed much less accessible to nuclease
digestion than after purification and (2) the observed differences in DNase I sensitivity of
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chromatin reflect differences in chromatin properties and not differential intrinsic sensitivity
of the DNA template.
PCR product quantification and calculation
Gel images such as in Figure 4 can reveal strong differences but comparison of many
probed fragments is not trivial. Therefore, a quantitative description of the results is desirable.
PCR products can be quantified from gel images using image processing software such as
ImageJ (Schneider et al., 2012). This does not require specialized equipment and can be
implemented on any computer. However, quantification from gel images is error-prone. It is
therefore recommended to use a microfluidics-based electrophoresis system such as
Bioanalyzer (Agilent, USA) or Experion (Biorad, USA). Of note, some organic components
in the PCR buffer can interfere with the measurement using such systems (see manufacturer’s
instruction for more details). Because purification of PCR products can introduce biases, it is
best to consider using PCR buffers with only basal salts.
DNase I sensitivity can be quantified using the PCR signal as a function of increasing
DNase I concentration; the steeper the slope, the more accessible is the DNA fragment. The
PCR signal y can be expressed as an exponential decay function of DNase I concentration x
(U ml-1):
(1) y = Y0 e-λxlt
where Y0 is the input PCR product intensity, λ (U-1 ml kb-1 min-1) is the decay constant
characteristic for a given DNA fragment, l is the amplicon length (kb) and t is the digestion
time (min). Note that the decay constant of the curve is largely independent of primer
efficiency. Nevertheless, we recommend using primer pairs with similar high efficiency when
comparing different amplicons. Figure 5 shows the fitted curves for the quantified PCR
products shown in Figure 4. The fitted curves show that Ta2 and Cinful-like were most
resistant to DNase I than ACT7 and GAPDHα and that the DNase I sensitivity of AG and FT
was intermediate between the active and heterochromatic genes with a slightly higher
sensitivity of FT than AG.
Figure 6 displays the fitted λ values on a logarithmic "heat" axis. By summarizing the
results from the three replicates one can easily describe and visualize the DNase I sensitivity
of different genomic fragments. The sensitivity of the chromatin of the tested genomic
regions varied widely. The most sensitive region was ACT7 with an average decay constant
of 0.18, and the least sensitive region was Ta2 with an average decay constant of 0.02. The
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Probing chromatin accessibility
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PcG target genes AG and FT were between the insensitive and sensitive controls with FT
most similar to the sensitive regions. In contrast, isolated DNA was always highly sensitive,
and the differences between the decay constants for the various regions were minor, probably
reflecting experimental noise (Fig. 6).
Together, the decay constant λ can be used as a convenient quantitative descriptor of
chromosomal DNA accessibility but comparisons should be restricted to similar amplicon
sizes and digestion times.
Genome-wide DNA sensitivity profiles
DNase I sensitivity can be probed locally but also genome-wide, where DNase I
treatment can be coupled to DNA size fractionation. Depending on the profiling of hyper-
sensitivity or hypo-sensitivity, different size fractions of DNA are used. For profiling hypo-
sensitivity, the least cleaved DNA fragments should be enriched. In our hands, isolation of
fragments larger than 17kb proved to work well. This fraction was shown to be highly
enriched for DNase I hypo-sensitive DNA including transposable element genes such as Ta2
from pericentric heterochromatin over active genes such as ACT7 (Fig.7, right 3 columns). In
contrast, no enrichment was seen for the undigested control (Fig. 7, left 3 columns). Isolated
DNA can be labeled and hybridized to genomic DNA tiling arrays (Shu et al., 2012).
Alternatively, isolated DNA can be subjected to next generation sequencing. On the other
hand, the hyper-sensitive DNA population can be enriched by minimal DNase I treatment. In
this case, small-sized DNA should be enriched, and high enrichment of active genes versus
heterochromatin fragments is expected (Sabo et al., 2006). Alternatively, end-labeling of
DNA after minimal DNase I treatment can be used to enrich for hypersensitive sites
(Crawford et al., 2006, Follows et al., 2006, Sabo et al., 2006).
CONCLUSION
We have presented a robust and reproducible protocol for DNase I sensitivity assays
using Arabidopsis nuclei and have discussed points to consider when adapting the protocol
for specific experiments. In principle, this protocol should be easily adaptable to other plant
species. We have shown that two TE genes are highly insensitive to DNase I, and that two
active genes are highly sensitive. These genes can serve as insensitive and sensitive controls
in future chromatin DNase I sensitivity assays. We also have shown that two PcG targets are
less sensitive than the active genes, supporting the model that PcG proteins employ
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Probing chromatin accessibility
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compaction of the chromatin as a repression mechanism not only in animals but also in
plants. Subsequent to assays of DNase I sensitivity by PCR higher resolution can be achieved
by qPCR (Dorschner et al., 2004) or by high-throughput DNase-chip or DNase-sequencing
for genomic chromatin accessibility profiling (Crawford et al., 2006, Sabo et al., 2006, Song
et al., 2010, Shu et al., 2012).
Because altered DNA accessibility is a major effect of many histone modifications,
histone variants and non-histone proteins, testing DNA accessibility will become more
common in future studies on chromatin function.
MATERIALS AND METHODS
Plant material
Two hundred milligrams of 15-day old seedlings of Arabidopsis thaliana accession
Col-0 were harvested for each replicate. Seedlings were immediately frozen in liquid nitrogen
and ground to power with pre-cooled mortar and pestle. The powder was transferred to 15 ml
tubes stored at -80˚C or used for nuclei extraction immediately.
Nuclei isolation
Frozen powder of Arabidopsis plant tissue was suspended in 10 ml of ice cold Nuclei
Isolation Buffer (1 M hexylene glycol; 20 mM PIPES-KOH, pH 7.6; 10 mM MgCl2; 1 mM
EGTA; 15 mM NaCl; 0.5 mM spermidine; 0.15 mM spermine; 0.5% Triton X-100, v/v; 10
mM β-mercaptoethanol; 1× protease inhibitor cocktail (Roche, Switzerland)). β-
mercaptoethanol and protease inhibitor were added immediately before use. The mixture was
incubated for 15 min at 4˚C with gentle rotation. The suspension was filtered through 30 nm
CellTrics, and the eluate was centrifuged for 10 min at 1500×g at 4˚C. The supernatant was
discarded, and the pellet was collected as crude nuclei extract.
DNase I digestion and DNA recovery
Nuclei extracts were washed once with 1 ml of Digestion Buffer (40 mM Tris-HCl,
pH 7.9; 0.3 M sucrose; 10 mM MgSO4; 1 mM CaCl2; 1× protease inhibitor cocktail (Roche))
and gently resuspended in 670 μl of fresh Digestion Buffer by pipetting until no clumps were
visible. Aliquots of 80 µl were transferred to five PCR tubes and kept on ice.
A DNase I (RQ1 RNase-Free DNase, Promega USA) dilution series was prepared by step-
wise dilution using Digestion Buffer to establish the following concentrations: 0, 0.125, 1.25,
2.5, and 5 U/ml. Aliquots of 30 µl were transferred to five PCR tubes and kept on ice.
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Probing chromatin accessibility
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Nuclei suspensions were pre-heated on a PCR cycler for 2 min at 37˚C. Twenty microliters of
diluted DNase I were transferred to the corresponding nuclei suspensions using a
multichannel pipette and mixed briefly by pipetting. The final DNase I concentrations in the
digestion mixtures were 0, 0.025, 0.25, 0.5, and 1 U/ml. The digestion mixtures were
incubated at 30˚C for 15 min. Reactions were stopped by adding 10 μl of 100 mM EDTA.
Reaction mixtures were then returned to ice. DNA was recovered using phenol–chloroform-
isoamyl alcohol (PCI; 25:24:1, pH 8.0) extraction and ethanol precipitation.
For digestion of isolated DNA, genomic DNA was extracted from 240 μl of the nuclei
suspension using PCI extraction and ethanol precipitation. The DNA was re-dissolved in 240
μl of Digestion Buffer and 80 μl aliquots of the extracted DNA were digested with DNase I at
final concentrations of 0, 0.025, and 0.05 U/ml. Digested isolated DNA was recovered
similarly.
PCR analysis
The precipitated DNA was re-dissolved in 150 μl of nuclease-free water, and 5 μl
were used for each PCR reaction. PCR was performed using EuroTaq (Euroclone, Italy) in
25-μl-reactions according to the manufacturer’s protocol. The PCR fragments were designed
to overlap the start codons of AG, FT, ACT7 and GAPDHα, or to locate within the annotated
sequence of Cinful-like and Ta2. For PCR primer sequences see Table 1. The absolute
quantity of the PCR product was measured using DNA 7500 kit (Agilent) on an Agilent 2100
Bioanalyzer platform (Agilent). PCR measurements were averaged across the three digestion
replicates. qPCR was performed as described before (Shu et al., 2012) using the Universal
Probe Library system (Roche). For qPCR primer sequences see Table 2.
Analysis on tiling arrays
Alternatively, the precipitated DNA was re-dissolved in 15 µl of nuclease-free water
and resolved on 3 % agarose gels. DNA fragments of sizes above 17 kb were extracted from
the gel using freeze/thaw method. The extracted DNA was amplified and labeled with biotin
using the BioPrime® DNA Labeling System (Invitrogen, California). Labeled DNA was
hybridized to Affymetrix AGRONOMICS1 Arabidopsis tiling arrays (Affymetrix, Santa
Clara, CA) as recommended by the manufacturer and previously described (Rehrauer et al.,
2010; Shu et al., 2012). Data analysis using freely available R packages has been described
(Shu et al., 2012).
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ACKNOWLEDGMENTS
We thank the Functional Genomics Centre Zurich for providing the Agilent 2100
Bioanalyzer platform.
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Figure legends
Figure 1. Outline of the DNase-PCR procedure. This outline represents the DNase I-PCR
procedure as described in the text.
Figure 2. Schematic illustration of the digestion procedure. This illustration represents the
procedure of the digestion step as described in the text. For easier handling and better
synchronization, sample aliquots can be transferred using multichannel pipettes. Digestion
can be performed taking advantage of PCR tube strips and PCR cyclers.
Figure 3. Principle of using PCR to detect the accessibility of DNA fragments in
chromatin. After a limited DNase I digest of native chromatin, more accessible chromatin is
cleaved more frequently within a short distance resulting in fragmentation of the DNA and
reduced signals from PCR reactions (right). Inaccessible chromatin is cleaved less frequently
and therefore the DNA remains nearly intact, resulting in strong PCR signals (left). The green
arrows represent primer positions.
Figure 4. DNase I-PCR shows differential DNase I sensitivity of chromatin in selected
genomic regions. Equal amounts of chromatin and purified DNA were subjected to gradient
DNase I treatments. Extracted DNA was used as template for PCR for two pericentric TE
genes (CINFUL-like and Ta2), two PcG target genes (AG and FT), and two active genes
(ACTIN-7 and GAPDHα). The experiment was replicated three times (rep. 1-3)
Figure 5. Fitting curves of the DNase I-PCR results for the selected genomic regions.
PCR products as shown in Fig. 4 were quantified using an Agilent Bioanalyzer. The
measurements were normalized to the input values and fitted to an exponential decay model.
The fitted curves for all tested genomic regions are shown for the 3 replicates (rep. 1~3) for
both digested chromatin (top) and digested isolated DNA (bottom).
Figure 6. The distribution of the decay constants of the tested genomic regions on a
“heat” axis. The fitted decay constants were averaged over the three replicates for digested
chromatin (blue text) and digested isolated DNA (red text), respectively. The results were
plotted on a logarithmic axis. Error bars show the standard error of the decay constants. The
intercept table shows a summary of the calculated results (upper right corner).
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Figure 7. qPCR confirmation of enrichment of hypo-sensitive over hyper-sensitive DNA
regions in different size fractions after controlled DNase I digestion of nuclei. DNA
fractions of four size ranges (> 17 kb, 6 kb-17 kb, 3 kb-6 kb, <3 kb) were recovered
separately from undigested (left 3 columns) and digested (right 3 columns) nuclei.
Enrichment of hypo-sensitive (inaccessible) pericentric heterochromatin versus hyper-
sensitive (accessible) euchromatic active genes is represented by enrichment of signal
abundance for the TE gene Ta2 against the active ACTIN-7 gene as measured by qPCR. High
levels of enrichment are shown in blue and depletion in red.
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Table 1. Primers for DNase-PCR
Locus Primer Sequence Amplicon length (bp)
Ta2 TACAGTGGCTACCACATTGC
1680 ATGAAAGCGGTCCCATCA
CINFUL-like ACCGCAATATCCGATACGAC
1651 TTCAACGTCGAACGAGTCCTAG
AG GTGAAACAAATTTTCCTGCAGAATGTC
1631 TCCTAGCTCCGATTGGTACG
FT TCTTTTAGAACGTTTTCGCTTTCG
1695 GGTTGCTAGGACTTGGAACATC
GAPDHα CTCCCTTGGAAGGAGCTAGG
1648 AATTGTGAATAACTAGCTTTAGCATTGGA
ACTIN-7 GAACTGCTCTTGGCTGTCT
1646 AGCATTGTCTCTCCCAGATTTT
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Table 2. Primers and Universal Probes for q-PCR
Locus Primer Sequences Probe No.
Ta2 ATGAAAGCGGTCCCATCA
#119 CGACTGCTATTCCCTTGTCC
ACTIN-7 GGAAACATCGTTCTCAGTGGT
#31 CTTGATCTTCATGCTGCTAGGT
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