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INTRODUCTION |
The assembly of eukaryotic DNA into folded nucleosomal arrays has
drastic consequences for many nuclear processes that require access to
the DNA sequence, including RNA transcription, DNA replication, recombination, and repair. The nucleosome, which consists of 147 bp1 of DNA wrapped nearly
twice around an octamer of histones H2A, H2B, H3, and H4, can occlude
DNA sequences both in vivo and in vitro. The
nucleosome is not a static structure but appears to be a dynamic and
flexible assembly. For instance, moderate concentrations of NaCl can
lead to several distinct changes in nucleosome conformation (1-4). In
addition, nucleosomes isolated from transcriptionally active chromatin
appear to be depleted of histone H2A-H2B dimers (discussed in Ref. 5)
and contain histone octamers the interiors of which are more accessible
to enzymatic and chemical modifications (6-8). Nucleosomes from
transcriptionally active chromatin can also be visualized
microscopically as extended, largely unfolded structures (9, 10). These
and other studies have led to the view that regulatory factors might
antagonize the repressive effects of chromatin by disrupting the
structure or conformation of the histone octamer (discussed in Ref.
11).
Two types of highly conserved chromatin remodeling enzymes have been
implicated as regulators of the repressive nature of chromatin
structure (12, 13). Several members of the SWI2/SNF2 family of
DNA-stimulated ATPases use the energy of ATP hydrolysis to disrupt
nucleosome structure, which can lead to an enhanced mobility of
nucleosomes (14-16, 64). The second type consists of the nuclear
histone acetyltransferases that covalently modify lysine residues
within the flexible N-terminal domains of the histone proteins. The
Saccharomyces cerevisiae SWI-SNF complex is the prototype
ATP-dependent chromatin remodeling complex. This widely
conserved 2-MDa multisubunit assembly is required for the inducible
expression of a number of diversely regulated yeast genes and for the
full functioning of many transcriptional activators (reviewed in Refs.
17 and 18). SWI-SNF can be recruited to target genes via direct
interactions with gene-specific activators (19-21), and in several
cases, SWI-SNF facilitates the binding of activators to nucleosomal
sites in vivo (22, 23). In vitro, the purified
SWI-SNF complex is a DNA-stimulated ATPase that can use the energy of
ATP hydrolysis to disrupt histone-DNA interactions. Although the
mechanism by which SWI-SNF disrupts nucleosome structure is not known,
this "remodeling" reaction leads to an enhanced accessibility of
nucleosomal DNA to DNase I (24-26), restriction enzymes (27, 28), and
sequence-specific DNA-binding proteins (24, 25, 29). More recently,
SWI-SNF has been shown to increase DNA accessibility of nucleosomal
arrays in a catalytic manner that is dependent on the presence of
histone N-terminal domains (27, 28).
Four different models have been proposed to explain the mechanism by
which ATP-dependent remodeling by the SWI-SNF complex increases nucleosomal DNA accessibility. 1) Several studies have suggested that SWI-SNF might remove or rearrange the H2A-H2B dimers (24, 30-34). 2) SWI-SNF remodeling may induce a novel conformation of
the histone octamer (Refs. 35 and 36; discussed in Ref. 11) that might
involve conformational changes in the (H3-H4)2 tetramer
analogous to the transcription-associated transitions described above.
3) SWI-SNF may use the energy of ATP hydrolysis to translocate along
DNA and destabilize histone-DNA interactions (discussed in Ref. 37).
This model is similar to the octamer spooling mechanism described by
Studitsky et al. (38) for passage of polymerases through
nucleosomes. 4) Finally, SWI-SNF might bind directly to nucleosomes and
use the energy of ATP hydrolysis to change the path of nucleosomal DNA
(25) or to peel DNA off the surface of the histone octamer without
changing octamer structure (39).
In this study, we have directly tested whether
ATP-dependent chromatin remodeling by the SWI-SNF complex
alters the composition or conformation of the histone octamer. We use a
nucleosomal array remodeling assay (27, 28) to quantify SWI-SNF
activity on arrays of histone (H3-H4)2 tetramers and on
nucleosomal arrays reconstituted with histone octamers containing
internally cross-linked tetramers. In order to monitor more subtle or
transient changes in octamer structure, we also measured the effects of
SWI-SNF remodeling on the steady state fluorescence of nucleosomal and tetramer arrays harboring a histone H3 derivative site specifically modified at residue 110 with the fluorescent probe
acetylethylenediamine-(1,5)-naphthol sulfonate (AEDANS) (1, 2, 40).
Taken together, our data are consistent with a model in which substrate
recognition by SWI-SNF requires an intact histone octamer, and
subsequent ATP-dependent remodeling disrupts histone-DNA
contacts without a concomitant loss of histone proteins or perturbation
of the histone octamer.
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EXPERIMENTAL PROCEDURES |
Reagent Preparation--
Array DNA template was isolated by
digestion of plasmid pCL7c with NotI, HindIII,
and HhaI (New England Biolabs) followed by FPLC purification
on Sephacryl-500 (Amersham Pharmacia Biotech) essentially as described
(27, 41). Array DNA template was end-labeled as described (27, 41).
Chicken erythrocyte histone octamers were purified from chicken whole
blood (Pel-Freez) as described previously (42). Tetramers were provided
as a kind gift from Jeff Hansen and were purified by stepwise elution
from hydroxylapatite columns as described (42). Tetramers were dialyzed
against Buffer T (1 M NaCl, 10 mM Tris-HCl, pH
8.0, 0.25 mM EDTA, 0.1 mM dithiothreitol) prior to array reconstitution. Disulfide-linked histone octamers were generated by first diluting histone octamers 2-fold with Buffer D1 (10 M urea, 2 M NaCl, 20 mM Tris-HCl,
pH 8.0) followed by dialysis against Buffer D2 (5 M urea, 2 M NaCl, 10 mM Tris-HCl, pH 8.0) at 4 °C with
constant agitation for 4 days. Histone octamers were then reconstituted
by dialysis against Buffer D3 (2 M NaCl, 10 mM
Tris-HCl (pH 8.0), 0.25 mM EDTA). Histone H3 oxidation
efficiency was analyzed by SDS-polyacrylamide gel electrophoresis in
the absence of reducing agent (see Fig. 1B, lane 3).
AEDANS-modified histone H3 was generated by first diluting histone
octamers 2-fold with Buffer D1, followed by addition of
iodoacetylethylenediamine-(1,5)-naphthol sulfonate (1,5-IAEDANS)
(Molecular Probes, Inc.) at a molar ratio of 20:1 (1,5-IAEDANS:H3) and
incubation for 2 h at room temperature on a nutator (Adams).
Reactions were quenched with an excess of 
-mercaptoethanol (Sigma)
and dialyzed against Buffer D2 to remove unreacted 1,5-IAEDANS. Samples
were then dialyzed against Buffer D3 to reconstitute histone octamers.
Labeling specificity was verified by visualization of the corresponding
fluorescent histone band upon illumination with long wave ultraviolet light.
SWI-SNF complex was purified from yeast strain CY396 or CY743
(sin3
) as described (41). The concentration of complex
was determined to be approximately 300 nM by comparative
Western blot and by ATPase assays (27, 41).
Reconstitution and Analysis of Nucleosomal Arrays--
Arrays
were reconstituted onto the 208-11 S DNA template (Fig. 1A)
in a Slide-a-lyzer dialysis cassette (Pierce) using the salt dialysis
protocol of Hansen and Lohr (43). Octamer concentrations were
determined by A230 (44). Array saturation and
nucleosome/tetramer positioning was determined by EcoRI or
MspI digestions using approximately 20 nM array
in remodeling buffer (5 mM MgCl2, 50 mM NaCl, 10 mM Tris-HCl (pH 8.0), 1 mM dithiothreitol) as described previously (27, 28, 41,
42). Arrays were digested for 30 min at 37 °C, and the reactions
were electrophoresed on 4% native polyacrylamide gels (see Fig.
1C). The gel was briefly soaked in 2 µg/ml ethidium bromide and photographed under ultraviolet illumination.
Assay Conditions--
Coupled array remodeling-restriction
reactions were performed in a final concentration of 5 mM
MgCl2, 50 mM NaCl, 10 mM Tris-HCl (pH 8.0), 1 mM dithiothreitol, 0.1 mg/ml bovine serum
albumin, 1 mM ATP, and 500 units/ml HincII (New
England Biolabs) as described previously (27, 28, 41).
Fluorescence spectroscopic studies were carried out using a PTI
QM1/S.E.901/S.E.910Q fluorescence spectrophotometer. Samples were
excited at 344 nm (slit width, 2.5 nm), and emission intensities were
recorded from 450 to 500 nm (slit width, 4 nm). Samples (80 µl in
quartz microcuvettes) contained 4 nM AEDANS-labeled octamer or tetramer array in remodeling buffer (described above) at room temperature. In experiments assessing effects of monovalent salt, emission intensities were recorded initially 0.5 min after adjustment to 50, 75, 150, and 300 mM NaCl and 0.5, 60, 120, 240, 480, and 960 min after subsequent adjustment to 600 mM NaCl. In
experiments assessing effects of SWI-SNF, emission intensities were
recorded initially at 5 min after addition of a final concentration of 2, 10, or 20 nM purified SWI-SNF and 5, 10, and 60 min
after subsequent addition of a 1 mM final concentration of
ATP. For samples in which SWI-SNF, ATP, or salt was added to the
arrays, data were corrected for a buffer addition control. All data
were corrected for dilution.
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RESULTS |
SWI-SNF Remodels Arrays of Histone (H3-H4)2
Tetramers--
It has been proposed that SWI-SNF increases the
accessibility of nucleosomal DNA by depletion or rearrangement of the
histone H2A-H2B dimers (Refs. 24, 30, 31, 32, and 34; discussed in Ref.
33). In order to investigate whether loss of the dimers is equivalent
to the SWI-SNF remodeled state, we used purified chicken erythrocyte
histone octamers or (H3-H4)2 tetramers (Fig. 1B, lanes 1 and 4)
to reconstitute nucleosomal or tetramer arrays. The DNA template used
for these reconstitutions is composed of 11 head-to-tail repeats of a
nucleosome positioning sequence from the Lytechinus
variegatus 5 S rRNA gene, each of which is flanked by
EcoRI restriction sites. Nucleosomes that are reconstituted onto each of these 5 S repeats assume a major translational position that is present in at least 50% of the population (45, 46). Minor
translational positions also exist and differ from the major frame by
multiples of 10 bp. The central repeat of our array template also bears
a unique SalI/HincII restriction site close to
the dyad axis of symmetry of a nucleosome positioned at the major frame
(27, 47). Array reconstitutions were analyzed for extent of DNA repeat
saturation and for correct positioning by restriction enzyme cleavage
(Fig. 1C; see also under "Experimental Procedures") (27,
41, 42). EcoRI digestion of nucleosomal or tetramer arrays
releases primarily mononucleosome-sized particles and few of the high
molecular weight partial digestion products that would be indicative of
alternative positioning or oversaturation (Fig. 1C).
Digestion of nucleosomal or tetramer arrays with MspI (the site of which is located ~30 bp from the predicted dyad axis of a
nucleosome positioned at the major frame) demonstrates that these sites
are fully protected by nucleosomal arrays but accessible in tetramer
arrays (Fig. 1C). These data are consistent with the fact
that an (H3-H4)2 tetramer assumes the same translational positions as an intact octamer (48, 49) but that the tetramer assembles
less DNA (34).
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Fig. 1.
Array reconstitutions. A,
schematic representation of the 208-11 S template DNA used to
reconstitute nucleosomal and tetramer arrays. It consists of 11 head-to-tail repeats of a 5 S rRNA nucleosome positioning sequence, the
central repeat bearing an unique SalI/HincII
restriction enzyme site. Each repeat is flanked by EcoRI
restriction enzyme sites. In addition, an MspI site is
located ~30 bp from the predicted dyad axis of symmetry of each
positioned nucleosome. B, analysis of histone proteins used
for array reconstitutions by 18% SDS-polyacrylamide gel
electrophoresis and Coomassie staining. Lane 1, chicken
histone octamers; lane 2, chicken histone octamers contained
1,5-IAEDANS conjugated to histone H3; lane 3, oxidized
chicken histone octamers electrophoresed under nonreducing conditions;
lane 4, chicken (H3-H4)2 tetramers.
C, analysis of saturation and positioning of reconstituted
arrays by restriction enzyme digestion and native polyacrylamide gel
electrophoresis. Digestion of arrays with EcoRI releases
primarily mononucleosome- or monotetramer-sized particles ( ), as
well as some free 208-bp 5 S repeats ( ). The ratio of free 5 S
repeats to mononucleosome/monotetramer particles provides a qualitative
measurement of array saturation (60-90% saturation for the arrays
shown here). Higher molecular weight species represent partial
EcoRI digestion products. Nucleosome assembly on the 5 S
repeats results in occlusion of the MspI sites, whereas
these sites are accessible for the tetramer arrays and yield a
monotetramer-sized particle (), indicating that the
(H3-H4)2 tetramers protect less DNA as expected.
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We then exploited a sensitive nucleosomal array remodeling assay in
which SWI-SNF remodeling activity is coupled to restriction enzyme
activity (27, 28). Previously we used this assay to determine the
kinetic parameters of ATP-dependent nucleosomal array
remodeling by the SWI-SNF and RSC (remodels the
structure of chromatin) complexes (27, 28, 41).
To quantify the remodeling capacity of SWI-SNF complex, 3 nM of nucleosomal or tetramer array was exposed to 500 units/ml HincII, either in the presence or absence of 3 nM SWI-SNF complex and 1 mM ATP (Fig.
2A). As described previously,
HincII digestion of nucleosomal arrays is biphasic (27); the
first, rapid phase of the reaction represents digestion of arrays
harboring HincII sites positioned between nucleosomes, and
the second, slow phase represents digestion of the nucleosomal HincII sites. By limiting our analysis to the second phase
of HincII digestion, the first order rate of
HincII cleavage yields a quantitative measurement of
nucleosomal DNA accessibility (27, 28, 47).
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Fig. 2.
Tetramer arrays are not optimal substrates
for SWI-SNF. A, representative time course for
HincII digestion of 3 nM nucleosomal (left
panel) or (H3-H4)2 tetramer (right panel)
arrays in the presence ( ) or absence () of 3 nM
SWI-SNF. Time point 0 reflects a 20-min preincubation with
HincII in the absence of SWI-SNF. The high percentage of
tetramer array cleaved in the first phase of the digestion is due in
part to slight undersaturation of these arrays. We note that similar
levels of undersaturation of nucleosomal arrays have no effect on
SWI-SNF remodeling rates (data not shown; see also Ref. 41).
B, quantification of the data shown in A. Data
are presented as the concentration of nucleosomal () and tetramer
() arrays cleaved by HincII in the presence of SWI-SNF
during the initial 10 min of the reaction. Similar results were
obtained in at least three different experiments using both different
SWI-SNF preparations and independent nucleosomal and tetramer array
reconstitutions.
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In the absence of SWI-SNF, the first order rate of HincII
cleavage was 1.3 × 10
5 for tetramer arrays and
2.4 × 106 for nucleosomal arrays (Fig. 2; see also
Table I). The 5-fold higher rate for the
tetramer arrays is comparable to the rate observed previously for
nucleosomal arrays reconstituted with histones that lack their
trypsin-sensitive N-terminal domains (28) and is consistent with
nucleosomal DNA being more accessible in the absence of the histone
H2A-H2B dimers (31, 48, 50, 51). In the presence of SWI-SNF, the first
order rate of HincII cleavage was 6.4 × 105 for tetramer arrays and 9.2 × 105
for nucleosomal arrays (Fig. 2; Table I). The fact that the rate of
cleavage of nucleosomal arrays in the presence of SWI-SNF significantly
exceeded the rate of cleavage of tetramer arrays in the absence of
SWI-SNF (see Table I) indicates that remodeling is not equivalent
simply to loss of histone H2A-H2B dimers. The fact that the rate of
cleavage of tetramer arrays in the presence of SWI-SNF is 5-fold higher
than the rate in the absence of SWI-SNF indicates further that SWI-SNF
can remodel tetramer arrays. However, SWI-SNF remodeling of tetramer
arrays appears to be quantitatively less efficient than SWI-SNF
remodeling of nucleosomal arrays, as the rate of cleavage of tetramer
arrays in the presence of SWI-SNF was 30% slower than that of
nucleosomal arrays in the presence of SWI-SNF (Fig. 2C; see
also Table I).
SWI-SNF Activity Is Not Catalytic on Arrays of (H3-H4)2
Tetramers--
Previously, we showed that SWI-SNF is able to
catalytically remodel multiple nucleosomal arrays and that histone
N-terminal domains are required for this reaction (27, 28). As
remodeling of tetramer arrays appears to be quantitatively less
effective than remodeling of nucleosomal arrays (see above), we
investigated whether SWI-SNF was able to function catalytically, or
only stoichiometrically, on tetramer arrays. Remodeling assays were
carried out in which there was a 10-fold molar excess of array to
remodeling complex (12 nM array to 1.2 nM
SWI-SNF). Consistent with previous data, SWI-SNF was able to stimulate
HincII cleavage of nucleosomal arrays throughout the 150-min
time course (Fig. 3A, left
panel; see also Refs. 27 and 28). However, there was only minor
stimulation of HincII cleavage by SWI-SNF on tetramer arrays
(Fig. 3A, right panel). In fact, the rate of
HincII cleavage on tetramer arrays in the presence of
SWI-SNF approximated the rate determined in the absence of remodeling
complex (Fig. 2A, right panel; and Table I). Fig.
3B shows the quantification of the amount of
SWI-SNF-dependent array cleavage during the 150-min time
course. As expected, SWI-SNF was able to perform approximately 2.5 rounds (3 nM of array cleaved by 1.2 nM
SWI-SNF) of nucleosomal array remodeling in 150 min (~50 min per
round; see also Ref. 27). In contrast, SWI-SNF was unable to complete
even one round of remodeling on tetramer arrays (0.3 nM
array cleaved due to 1.2 nM SWI-SNF) during the time
course. Therefore, SWI-SNF is unable to catalytically remodel multiple
arrays of (H3-H4)2 tetramers.
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Fig. 3.
SWI-SNF activity is not catalytic on arrays
of (H3-H4)2 tetramers. A, representative
time course for HincII digestion of 12 nM
nucleosomal (left panel) or (H3-H4)2 tetramer
(right panel) arrays in the presence () or absence ( )
of 1.2 nM SWI-SNF complex. Time point 0 reflects
a 30-min preincubation with HincII in the absence of
SWI-SNF. B, quantification of the data shown in
A. Data are presented as the remodeler-dependent
HincII cleavage of either nucleosomal () or tetramer
() arrays versus time. Remodeler-dependent
HincII cleavage events were obtained by subtracting the
fraction of cleaved arrays in the absence of SWI-SNF (A,
) from the fraction of cleaved arrays in the presence of SWI-SNF
(A, ). Data are presented as nanomolar remodeled
nucleosomal arrays versus time.
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One possible explanation for the lack of catalytic remodeling of
tetramer arrays may be that SWI-SNF has a higher affinity for tetramer
arrays and is thus defective for product release (analogous to the
explanation for the lack of catalytic remodeling of trypsinized
nucleosomal arrays (28)). In order to investigate this possibility,
SWI-SNF remodeling reactions were assembled that contained labeled
nucleosomal array and a 3- or 12-fold molar excess of free DNA,
nucleosomal, trypsinized nucleosomal, or tetramer competitor array.
Under these conditions, tetramer arrays competed for SWI-SNF activity
to the same extent as nucleosomal arrays or free DNA. In contrast,
trypsinized nucleosomal arrays were markedly more potent in competing
for SWI-SNF activity (data not shown; see also Ref. 28). This suggests
that the defect in catalytic remodeling of tetramer arrays, unlike the
defect in catalytic remodeling of trypsinized nucleosomes, is not due
to a higher affinity of SWI-SNF for these arrays.
SWI-SNF Remodeling Does Not Require Disruption of the
(H3-H4)2 Tetramer--
Chicken histone octamers contain
only two cysteine residues (Cys-110 of each of the two copies of
histone H3), and these cysteine residues are in close apposition within
the interior of the histone octamer near the dyad axis of symmetry (34,
52). The cysteine residues normally are inaccessible to chemical
modification, although denatured histone H3 can be disulfide-linked or
chemically modified in vitro with minimal perturbation to
the subsequently reconstituted histone octamer (52-54). In contrast,
these cysteine residues appear to be more exposed to solvent in
nucleosomes isolated from transcriptionally active chromatin (6-8) or
when nucleosomes are exposed to higher salt concentrations (1, 2).
To investigate whether SWI-SNF action might require a structural
transition of the (H3-H4)2 tetramer, we reconstituted
nucleosomal arrays with histone octamers that contain disulfide-linked
histone H3 (Fig. 1B, lane 3; see under "Experimental
Procedures" for details). As described above, 3 nM of
nucleosomal or disulfide-linked nucleosomal array was incubated in the
presence or absence of SWI-SNF and 1 mM ATP. In the absence
of SWI-SNF, the first order rate of cleavage by HincII was
3.8 × 106 for the disulfide-linked nucleosomal
array compared with 2.4 × 106 for nucleosomal
arrays (Fig. 4A; Table I).
This is consistent with previous observations that disulfide-linked
histone octamers can be reconstituted into nucleosomes that are not
grossly different from canonical nucleosomes (Refs. 52-54; see also
Fig. 1C). We then quantified the ability of SWI-SNF to
increase the accessibility of nucleosomal DNA to restriction enzyme
cleavage on the disulfide-linked and nucleosomal arrays. In the
presence of SWI-SNF complex, HincII cleavage was stimulated
25-fold on the disulfide-linked arrays and 37-fold on the control
nucleosomal arrays (Fig. 4A; Table I). Furthermore, the
first order rate of HincII cleavage in the presence of
SWI-SNF was nearly identical for the disulfide-linked and control
nucleosomal arrays (Fig. 4B; Table I). These data strongly
suggest that structural perturbation of the histone
(H3-H4)2 tetramer is not an obligatory intermediate or
product of ATP-dependent nucleosome remodeling.
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Fig. 4.
SWI-SNF can remodel nucleosomal arrays
reconstituted with disulfide-linked histone octamers.
A, representative time course for HincII
digestion of 3 nM nucleosomal (left panel) or
disulfide-linked nucleosomal (right panel) array in the
presence () or absence () of 3 nM SWI-SNF. Time
point 0 reflects a 20-min preincubation with HincII in
the absence of SWI-SNF. B, quantification of data shown in
A. Results are presented as the concentration of nucleosomal
and tetramer arrays cleaved by HincII in the presence of
SWI-SNF during the initial 10 min of the reaction. Similar results were
obtained in at least three different experiments using independent
nucleosomal () and disulfide-linked () nucleosomal array
reconstitutions and SWI-SNF preparations.
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SWI-SNF Action Does Not Alter the Steady State Fluorescence of
AEDANS-modified Arrays--
The cysteine residue of histone H3 can
also be modified with sulfhydryl-specific fluorescent groups (1, 2, 40,
55-57). For instance, reconstitution of nucleosomes with
AEDANS-labeled H3 does not cause a significant perturbation of
nucleosome structure (1, 2, 54), and steady state AEDANS fluorescence
has been used to detect changes in octamer conformation as a function
of monovalent cation concentration (1, 2). We used this method in an
attempt to detect more subtle or transient changes in histone octamer
integrity and conformation as a result of ATP-dependent SWI-SNF activity.
Nucleosomal and tetramer arrays were reconstituted after modification
of histone H3 with the sulfhydryl-specific fluorescent probe AEDANS
(1-2; see under "Experimental Procedures" for details). To confirm
that the modification did not disrupt the structure of the arrays or
affect the activity of SWI-SNF, we quantified the kinetics of
HincII cleavage of the AEDANS-modified nucleosomal arrays in
the presence or absence of SWI-SNF activity. In the absence of SWI-SNF,
the first order rate of cleavage by HincII was 6.6 × 106 for the AEDANS-modified nucleosomal arrays and
2.4 × 106 for the nucleosomal arrays (Fig.
5A; Table I). The higher rate of cleavage observed for the AEDANS nucleosomal arrays indicates that
this histone H3 modification may cause a slight perturbation in the
structure or stability of the arrays. In the presence of SWI-SNF
complex, however, the rate of HincII cleavage was nearly identical for the AEDANS-modified and nucleosomal arrays (Fig. 5A; Table I). Thus, the data indicate that SWI-SNF is fully
functional on arrays reconstituted with AEDANS-H3.
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Fig. 5.
Effects of SWI-SNF on steady state
fluorescence emission intensity of nucleosomal and tetramer arrays
reconstituted with AEDANS-H3. A, SWI-SNF can remodel
AEDANS-labeled nucleosomal arrays (see also Table I). Representative
time course for HincII digestion of 3 nM
nucleosomal (left panel) or AEDANS-modified nucleosomal
(right panel) arrays in the presence () or absence ()
of 3 nM SWI-SNF. Time point 0 reflects a 20-min
preincubation with HincII in the absence of SWI-SNF. Similar
results were obtained in at least three different experiments using
independent nucleosomal and AEDANS-modified nucleosomal array
reconstitutions. B, fluorescence emission spectra of
AEDANS-modified nucleosomal and tetramer arrays. The upper
and lower left panels illustrate the effect of increasing
concentrations of NaCl on the steady state fluorescence of the modified
nucleosomal arrays and tetramer arrays, respectively. Fluorescence
emission intensities are shown in arbitrary units for AEDANS-modified
arrays at equilibrium in 50 mM NaCl, 30 s after
adjustment to 75, 150, 300, and 600 mM NaCl and 1, 2, 4, 8, and 16 h after adjustment to 600 mM NaCl. The
upper and lower right panels illustrate the
effect of SWI-SNF and ATP on the fluorescent-labeled nucleosomal and
tetramer arrays, respectively. Fluorescence emission intensities are
shown in arbitrary units for AEDANS-modified arrays at equilibrium in 5 mM MgCl2, 50 mM NaCl, 10 mM Tris-HCl (pH 8.0), and 1 mM dithiothreitol
(solid line); 5 min after addition of SWI-SNF to 10 nM (long dashes); and 5, 10, and 60 min after
subsequent addition of ATP to 1 mM (medium dashes,
short dashes, and dots, respectively). The experiment
yielded identical results in the presence of 2 or 20 nM
SWI-SNF (data not shown).
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AEDANS-modified nucleosomal and tetramer arrays were titrated with
increasing NaCl concentrations in the absence of SWI-SNF, and
fluorescence emission intensities were recorded 30 s after each
adjustment of NaCl concentration. Consistent with previous studies (1,
2), a dramatic decrease in fluorescence emission intensity was detected
with increasing NaCl concentration, suggesting increased solvent
accessibility of the internal structure of the histone octamer (Fig.
5B, left panels). After adjustment of NaCl concentration to
600 mM, fluorescence emission intensities of nucleosomal
and tetramer arrays were monitored as a function of time (Fig.
5B, left panels). Consistent with previous results (1, 2), a
biphasic decrease in fluorescence emission intensity was observed, with
a fast component on the second time scale (complete within 30 s)
and a slow component on the hour time scale (complete within 4 h).
These salt-dependent decreases in fluorescence emission intensity were observed for both nucleosomal and tetramer arrays, indicating that the observed decreases are not due to disruption of the
interface between the (H3-H4)2 tetramer and the H2A-H2B dimers. However, we note that AEDANS-modified tetramer arrays exhibit a
~3-fold lower fluorescence emission intensity than AEDANS-modified nucleosomal arrays (see Fig. 5B, left panels), suggesting
that the presence of the H2A- H2B dimers decreases solvent
accessibility of the (H3-H4)2 tetramer.
We next measured the effect of SWI-SNF remodeling activity on the
steady state fluorescence of AEDANS-modified nucleosomal or tetramer
arrays. SWI-SNF (2, 10, 20 nM) was added to arrays (4 nM) equilibrated in remodeling buffer (see under
"Experimental Procedures" for details) in the absence of ATP, and
emission intensities were recorded (Fig. 5B, right panel,
and data not shown). Subsequently, ATP was added to a final
concentration of 1 mM, and emission intensities were
recorded 5, 10, and 60 min after its addition (Fig. 5B; right panel). In striking contrast to NaCl titration (Fig. 5B,
left panels), neither the binding of SWI-SNF (which occurs in the
absence of ATP) nor ATP-dependent remodeling altered the
fluorescence emission intensities of arrays (Fig. 5B, right
panels). Addition of HincII directly to the reaction
cuvettes confirmed that SWI-SNF and ATP stimulated HincII
cleavage of the arrays and thus was active under these conditions (data
not shown). Thus, these data indicate that SWI-SNF remodeling is not
accompanied by the loss of the H2A-H2B dimers or by disruption or
changes in the internal accessibility of the (H3-H4)2 tetramer.
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DISCUSSION |
The S. cerevisiae SWI-SNF complex provides a paradigm
for a family of eukaryotic protein assemblies that function in an
ATP-dependent manner to alter chromatin structure. Although
it is evident that chromatin remodeling by SWI-SNF and other related
complexes results in enhanced accessibility of nucleosomal DNA and an
increased mobility of nucleosomes, the mechanism by which this reaction occurs remains controversial. This study was designed to directly test
simple predictions for several models that have been proposed for
ATP-dependent chromatin remodeling.
The "Dimer Disruption" Model--
In the extreme case, the
dimer disruption model predicts that ATP-dependent
remodeling by SWI-SNF will generate a (H3-H4)2 tetramer.
Although assembly of a (H3-H4)2 tetramer is sufficient to
position at least 90 bp of DNA (34), several studies have shown that
DNA-tetramer particles are more accessible to DNA-binding proteins (31,
50, 51), and removal of the histone H2A-H2B dimers facilitates
transcription in vitro (58). A simple prediction of such a
model is that the accessibility of DNA within (H3-H4)2 tetramer arrays should be equivalent to that of nucleosomal DNA after
SWI-SNF remodeling. However, we find that DNA wrapped around a
(H3-H4)2 tetramer is only about 5-fold more accessible to
HincII digestion compared with DNA assembled onto a complete
histone octamer. SWI-SNF action, on the other hand, enhances
nucleosomal DNA accessibility by ~35-fold (see Table I). Thus, the
remodeled state of a nucleosome is not equivalent to a
(H3-H4)2 tetramer. The dimer disruption model also predicts
that SWI-SNF will not be able to enhance the accessibility of
(H3-H4)2 tetramers. We found, however, that SWI-SNF can
remodel arrays of (H3-H4)2 tetramers, which leads to an
additional 5-fold enhancement of HincII cleavage rates.
Finally, we show that fluorescence emission intensity of AEDANS-modified tetramer arrays is significantly lower than that of
AEDANS-modified nucleosomal arrays (presumably due to increased solvent
accessibility of the internal structure of the tetramer) (Fig.
5B). The fact that SWI-SNF remodeling of AEDANS-modified nucleosomal arrays does not decrease fluorescence emission intensity from the level characteristic of nucleosomal arrays to the level of
tetramer arrays (Fig. 5B, right panels) also indicates that remodeling does not involve dimer disruption. Thus, SWI-SNF action does
not convert a nucleosome into a tetramer.
The "Nucleosome Spooling" Model--
The nucleosome spooling
model for SWI-SNF action is based on the mechanism for passage of some
RNA polymerases through a nucleosome (38). This model proposes that the
energy of ATP hydrolysis might be used to translocate SWI-SNF along DNA
and around a nucleosome in a "wave-like" fashion (discussed in Ref.
37). Such ATP-driven translocation of SWI-SNF along the DNA would
disrupt histone-DNA contacts and may also lead to movement of the
histone octamer. Such a reaction mechanism might also result in
transfer of intact histone octamers onto an acceptor DNA, which has
been observed during nucleosome remodeling by the yeast RSC complex
(59). The nucleosome spooling model predicts that SWI-SNF will not
discriminate between a tetramer and nucleosomal substrate. In fact, one
might predict that the absence of H2A-H2B dimers might facilitate the ability of SWI-SNF to translocate through the residual histone-DNA interactions of the (H3-H4)2 tetramer. Although SWI-SNF
does remodel the (H3-H4)2 tetramer arrays, the apparent
rate of remodeling was approximately 30% slower than the rate for
nucleosomal array remodeling. Furthermore, SWI-SNF was inactive on the
(H3-H4)2 tetramers in remodeling reactions in which the
concentration of tetramer array was in excess over SWI-SNF. These
results demonstrate that arrays of (H3-H4)2 tetramers are
poor substrates for ATP-dependent remodeling by SWI-SNF, a
result that is not predicted by the nucleosome spooling model.
Furthermore, SWI-SNF does not show ATP-dependent tracking
activity in a DNA supercoiling assay (60), nor do other SWI2/SNF2
family members (e.g. Mot1p) demonstrate DNA tracking activity (61). Together, these data suggest that
ATP-dependent remodeling by SWI-SNF does not involve DNA
tracking, nor is it equivalent to the loss of the H2A-H2B dimers. Our
data do indicate that efficient remodeling activity requires a
canonical histone octamer that contains both an (H3-H4)2
tetramer and one or more H2A-H2B dimers.
SWI-SNF Action Does Not Perturb Octamer Structure--
Several
groups have recently suggested the alternative possibility that SWI-SNF
activity might induce a novel conformation of the nucleosome that may
involve rearrangement of the histone octamer without loss of histone
proteins (32, 35, 36). This model is consistent with recent electron
microscopy studies that indicated that ATP-dependent
remodeling by SWI-SNF does not change the protein mass of a nucleosome
and that remodeling is relatively insensitive to addition of an
external cross-linking reagent, dimethyl suberimidate to
mononucleosomes or nucleosomal arrays (39). What is this alternate
conformation? Lee et al. (32) proposed that human SWI-SNF
might use the energy of ATP hydrolysis to rearrange one or both H2A-H2B
dimers such that only the flexible N-terminal domain contacts DNA close
to the nucleosomal dyad. This novel octamer conformation might then
have a propensity to form the dinucleosome-like particle that was
previously observed (35, 36). Alternatively, SWI-SNF action might lead
to a conformational change in the (H3-H4)2 tetramer that
might mimic the "split nucleosome" or "lexosome" structure that
that has been proposed for the structure of transcriptionally active
chromatin (6, 9, 10).
To test these possibilities, we took advantage of the single cysteine
residue found within each copy of chicken histone H3. These two
cysteines are buried within the histone octamer and are located very
close to each other at the nucleosomal dyad axis (34, 52). To test for
gross changes in the structure of the (H3-H4)2 tetramer, we
monitored the apparent rates of remodeling of nucleosomal arrays that
contain disulfide-linked (H3-H4)2 tetramers. We found that
SWI-SNF remodeled these substrates with rates equivalent to nucleosomal
arrays. These data indicate that SWI-SNF action does not require a
significant rearrangement of the tetramer.
To probe for more subtle changes in the structure of the histone
octamer, we also monitored the effects of ATP-dependent
remodeling on the steady state fluorescence of AEDANS-labeled
nucleosomal and tetramer arrays. Steady state fluorescence of AEDANS-H3
has been used to detect at least three distinct conformational states of the nucleosome as a function of salt concentration (1, 2). Furthermore, the solvent accessibility of the AEDANS group, and thus
its fluorescence emission intensity, is predicted from previous fluorescence studies to be highly dependent on the presence of one or
both histone H2A-H2B dimers (Ref. 57; see also Fig. 5B, left
panels).
We found that in contrast to increased salt concentrations, which have
large effects on fluorescence emission intensities of nucleosomal or
tetramer arrays (Refs. 1 and 2; see also Fig. 5B, left
panels), addition of SWI-SNF (one SWI-SNF per two nucleosomes),
with or without ATP, had no measurable effect on fluorescence emission
intensities (Fig. 5B, right panels). These data suggest that
SWI-SNF does not change the overall structure or the solvent
accessibility of the histone octamer or tetramer.
What Is ATP-dependent Chromatin Remodeling?--
Our
data are consistent with previous suggestions that
ATP-dependent nucleosome remodeling by SWI-SNF disrupts
histone-DNA contacts without a structural change in the histone octamer
(Refs. 25 and 39; discussed in Ref. 62). How does SWI-SNF accomplish this feat? The remodeling reaction randomizes the rotational setting of
both wraps of nucleosomal DNA without removing most of the DNA from the
octamer surface (25). Furthermore, SWI-SNF action does not enhance the
reactivity of nucleosomal DNA to potassium permanganate, indicating
that remodeling does not involve an unwinding of the DNA double helix
(25). We favor models in which the energy of ATP hydrolysis is used to
rotate the DNA helix along its long axis relative to the histone
octamer. In one such model, SWI-SNF would remain at a fixed position
relative to the histone octamer, and both SWI-SNF and the octamer would
remain in a fixed translational position relative to DNA. In this
model, SWI-SNF would rotate the DNA helix back and forth with each
round of ATP hydrolysis, changing rotational phasing and disrupting
histone-DNA contacts throughout both wraps of nucleosomal DNA. In
another, similar model, SWI-SNF would remain in a fixed position
relative to the octamer, but SWI-SNF and the octamer would not remain
in a fixed translational position relative to the DNA. In this model
ATP-driven DNA helix rotation would "screw" the octamer along the
DNA helix, changing the translational position of the octamer
(analogous to a model proposed in Ref. 63). Either of these models
would be consistent with previous data (see above and Ref. 25), as well
as with recent studies indicating that several members of the SWI-SNF
family of chromatin remodeling complexes can enhance nucleosome
mobility (14-16, 64). Studies designed to test and distinguish between
these models are in progress.
,
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ABSTRACT
INTRODUCTION
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