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J. Biol. Chem., Vol. 275, Issue 25, 18864-18870, June 23, 2000
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From the
Received for publication, April 3, 2000, and in revised form, April 17, 2000
ATP-dependent chromatin remodeling
enzymes antagonize the inhibitory effects of chromatin. We compare six
different remodeling complexes: ySWI/SNF, yRSC, hSWI/SNF, xMi-2,
dCHRAC, and dNURF. We find that each complex uses similar amounts of
ATP to remodel nucleosomal arrays at nearly identical rates. We also
perform assays with arrays reconstituted with hyperacetylated or
trypsinized histones and isolated histone (H3/H4)2
tetramers. The results define three groups of the
ATP-dependent family of remodeling enzymes. In addition we
investigate the ability of an acidic activator to recruit remodeling
complexes to nucleosomal arrays. We propose that
ATP-dependent chromatin remodeling enzymes share a common reaction mechanism and that a key distinction between complexes is in
their mode of regulation or recruitment.
The assembly of eukaryotic DNA into folded nucleosomal arrays is
likely to have a major impact on the efficiency or regulation of
nuclear processes that require access to the DNA sequence, including
RNA transcription, DNA replication, recombination, and repair. In fact,
it is now generally recognized that disruption or remodeling of
chromatin structure is a rate-determining step for most of these
nuclear DNA transactions (1-3). Two classes of highly conserved
chromatin remodeling enzymes have been implicated as regulators of the
repressive nature of chromatin structure, the first class includes
enzymes that covalently modify the nucleosomal histones
(e.g. acetylation, phosphorylation, methylation,
ADP-ribosylation; reviewed in Ref. 4), and the second class is composed
of multisubunit complexes that use the energy of ATP hydrolysis to
disrupt histone-DNA interactions (reviewed in Refs. 5 and 6).
Each member of the ATP-dependent family of chromatin
remodeling enzymes contains an ATPase subunit that is related to the SWI2/SNF2 subfamily of the DEAD/H superfamily of nucleic
acid-stimulated ATPases (7). Seventeen members of the SWI2/SNF2 family
have been identified in the yeast genome (6), and to date, four of
these ATPases have been purified as subunits of distinct chromatin remodeling complexes ySWI/SNF (8, 9), yRSC (10), ISW1 and ISW2 (11).
Additional ATP-dependent remodeling complexes that harbor
SWI2/SNF2 family members have been identified in Drosophila (dACF (12), dNURF (13), dCHRAC (14), Brahma (15, 16)), human (hSWI/SNF
(17), hNURD (18-20), hRSF (21)), and frog (xMi-2 (22)). Although these
complexes have a variable number of subunits (i.e. 3-15),
and many different types of assays have been used to monitor the
activity of individual complexes, each enzyme can apparently use the
energy of ATP hydrolysis to alter chromatin structure and enhance the
binding of proteins to nucleosomal DNA-binding sites (3, 5).
Furthermore, in the case of the ySWI/SNF, Drosophila Brahma,
and hSWI/SNF complexes, remodeling is required for transcriptional
regulation of target genes in vivo (Refs. 23 and 24, for
review, see Ref. 5).
ATP-dependent chromatin remodeling complexes have been
further divided into three groups based on whether the sequence of the
ATPase subunit is more related to yeast SWI2 (ySWI/SNF, yRSC, Brahma,
and hSWI/SNF), Drosophila ISWI (ISW1, ISW2, dNURF, dCHRAC, dACF, and hRSF), or human Mi-2 (hNURD, xMi-2) (reviewed in Ref. 3).
Although each of these ATPases share a SWI2/SNF2-like ATPase domain,
they harbor additional, unique sequence motifs adjacent to the ATPase
domain that are characteristic of each group, the SWI2 group contains a
bromodomain (25), the ISWI group contains a SANT domain (26), and the
Mi-2 group contains a chromodomain (27). Differences among some groups
are also apparent in the nucleic acid cofactor required for stimulation
of ATPase activity. For enzymes that contain a SWI2-like ATPase
(ySWI/SNF, yRSC, and hSWI/SNF), ATPase activity is stimulated equally
well by "free" DNA or nucleosomes (8, 10, 28). In contrast, the
ATPase activity of enzymes that contain an ISWI-like or Mi-2-like
ATPase is optimally stimulated by nucleosomes (18-20, 22, 29). In the
case of ISWI-like ATPases, this requirement for nucleosomes may reflect
obligatory interactions with the trypsin-sensitive, histone N-terminal
domains (30).
Here we report the first direct comparison of the biochemical
properties of six different chromatin remodeling enzymes (ySWI/SNF, yRSC, dCHRAC, dNURF, hSWI/SNF, and xMi-2) which encompass all three
previously suggested groups. Surprisingly, each complex shows similar
ATPase activity on nucleosomal array substrates, and they are each able
to facilitate nucleosome mobilization within an array at nearly
equivalent rates. We have also investigated the nucleosome substrate
requirements for each enzyme by using arrays reconstituted with
hyperacetylated or trypsinized histone octamers, as well as histone
(H3/H4)2 tetramers. ATPase and remodeling assays with these
different substrates identify new common features, as well as new
distinctions among enzymes. In addition, we test the ability of the
GAL4-VP16 chimeric transcriptional activator to recruit these
remodeling complexes to a nucleosomal array substrate. We report that
ySWI/SNF is uniquely potent for recruitment by GAL4-VP16 in this assay.
Our data are consistent with the differential regulation of
ATP-dependent enzymes that each share a similar mechanism
of nucleosome remodeling.
Reagent Preparation--
The array DNA template contains 11 tandem, head-to-tail repeats of a 208-base pair sea urchin 5 S rRNA
gene (31, 32). Template was isolated by digestion of plasmid pCL7c
(208-11S) or pCL8b (208-11S-Gal4) with NotI,
HindIII, and HhaI (New England Biolabs) followed
by fast protein liquid chromatography purification on Sephacryl-500
(Amersham Pharmacia Biotech) essentially as described (31, 32). Array
DNA template was end-labeled by Klenow fill-in reaction with
[
Chicken erythrocyte histone octamers were purified from chicken whole
blood (Pel-Freez Biologicals) as described previously (33).
Hyperacetylated histone octamers were purified from butyrate-treated HeLa cells as described (34). Trypsinized histone octamers and (H3/H4)2 tetramers were purified as described (33, 35).
(H3/H4)2 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.
ySWI/SNF complex was purified from yeast strains CY396 or
CY743(sin3 Reconstitution and Analysis of Substrate Arrays--
Histone
proteins used for array reconstitutions were analyzed by 18%
SDS-polyacrylamide gel electrophoresis and Coomassie staining. Octamer
concentrations were determined by A230 (37). Histone octamers were reconstituted onto the 208-11S DNA templates (or
208-11S-Gal4 for recruitment assay) in a slide-a-lyzer dialysis cassette (Pierce) by salt gradient dialysis as described previously (38). Each repeat of the 208-11S template (or 208-11S-Gal4 for recruitment assay) is flanked by EcoRI restriction enzyme
sites. In addition, a unique MspI site is located 30 base
pairs from the predicted dyad axis of symmetry of each positioned
nucleosome. Array quality, saturation, and positioning was determined
by EcoRI or MspI digestion 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 (31,
32, 38, 39). Arrays were digested for 30 min at 37 °C and
electrophoresed on a 4% native polyacrylamide gel. The gel was briefly
soaked in 2 µg/ml ethidium bromide and photographed under ultraviolet
illumination. Saturation of arrays was analyzed by digestion with
EcoRI and comparison of the ratio of nucleosome bound repeat
to uncomplexed 208-base pair 5 S repeat. Positioning was analyzed by
digestion with MspI (32, 39). Whereas, nucleosomal,
trypsinized, and hyperacetylated arrays were inaccessible to digestion
with MspI, (H3/H4)2 tetramer arrays digested
with MspI released a mononucleosome size fragment indicating
that the (H3/H4)2 tetramers protect less DNA as expected
(40).
Assay Conditions--
ATPase reactions were performed with
respect to the optimal temperature for remodeling complex activity:
27 °C for xMi-2, dCHRAC, and dNURF, 30 °C for yRSC and ySWI/SNF,
and 37 °C for hSWI/SNF using 100 µM ATP and 0.2 µCi
of [
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 (31, 32, 41). Assays were performed
with respect to the optimal temperature for remodeling complex activity
(see above). HincII cleavage was quantified by
PhosphorImager analysis, and first-order rates were determined by curve
fitting. In multiple independent experiments, the first-order rates of
restriction enzyme cleavage for each particular combination of array
and remodeler varied by less than 20%.
ATPase and Remodeling Activities of Chromatin Remodeling
Enzymes--
In order to quantify the nucleosome remodeling activity
of ATP-dependent remodeling enzymes, we have developed a
biochemical assay where nucleosome remodeling activity is coupled to
restriction enzyme activity such that remodeling is revealed as an
enhancement of restriction enzyme cleavage rates (31, 32). This assay uses a nucleosomal array substrate in which the central nucleosome of
an 11-mer nucleosomal array contains a unique
SalI/HincII site located at the predicted dyad
axis of symmetry (31, 32). Restriction enzyme kinetics are biphasic in
this system; the first phase is rapid and reflects the fraction of
SalI/HincII restriction sites that are not
occluded by a nucleosome (due primarily in our assays to nucleosomes
that occupy minor translational positions; see Refs. 32, 42, and 43).
The second phase is slow and reflects a dynamic equilibrium between the
occluded and "open" nucleosomal DNA states (44, 45). In previous
studies, addition of yeast SWI/SNF and ATP stimulated the second phase
of SalI/HincII digestion 20-30-fold (32, 41).
Recently we have found that SWI/SNF remodeling leads to a rapid
redistribution of nucleosome positions within these arrays and that the
apparent rate of remodeling determined in this assay provides an
estimate of the rate of nucleosome mobilization (46).
Purified preparations of ySWI/SNF, yRSC, hSWI/SNF, dCHRAC, dNURF, and
xMi-2 were analyzed in parallel for nucleosome-stimulated ATPase
activity (see "Experimental Procedures"). Each complex was titrated
in an ATPase reaction which contained 100 µM ATP and 12 nM of a reconstituted, 11-mer nucleosomal array.
Surprisingly, the approximate concentration of each remodeling complex
that was required to achieve equivalent velocities of ATP hydrolysis were similar; for ySWI/SNF (2 nM), yRSC (2 nM),
hSWI/SNF (5 nM), dCHRAC (2 nM), and dNURF(4
nM), each complex catalyzed the hydrolysis of 450-600 nmol
of ATP/min (Fig. 1A, see also
Ref. 41). xMi-2 was slightly less active in this assay as ~15
nM was required to achieve this level of ATPase activity
(Fig. 1A). Given that our estimates of active enzyme
concentrations are only approximate (see "Experimental
Procedures"), the data shown in Fig. 1A indicate that each
of these enzymes have nucleosome-stimulated ATPase activities that are
similar within an order of magnitude. The similar levels of ATPase
activity among complexes was unexpected given that each complex has
different associated subunits which, at least in the case of the
hSWI/SNF complex, can have a large impact on the ATPase activity of the
catalytic subunit (i.e. BRG1; Ref. 28).
To assess the capacity of the six different complexes to remodel an
11-mer nucleosomal array, each remodeling enzyme (1-5 nM
ySWI/SNF, yRSC, hSWI/SNF, dCHRAC, dNURF, or 15 nM xMi-2)
was added to 1.5 nM nucleosomal array and the initial rates
of HincII digestion were measured in parallel reaction time
courses in the presence of ATP. We found that all six complexes
enhanced the rate of HincII digestion essentially
equivalently (Fig. 1B, see also Ref. 41 for a detailed
comparison of ySWI/SNF and yRSC). The dCHRAC complex reproducibly
yielded an approximately 2-fold lower rate of HincII
digestion than all other complexes which probably reflects the fact
that a significant amount of the ATPase activity of dCHRAC appears to
be contributed by topoisomerase II (see below). Since the initial rate
of HincII digestion provides an indirect measurement of the
rate of remodeling, these data indicate that all six enzymes use
similar amounts of ATP to remodel nucleosomal arrays at similar rates.
Furthermore, since it appears that this coupled restriction
enzyme-remodeling assay monitors the rate of nucleosome mobilization
(46), all six enzymes can apparently redistribute nucleosomes within an
array at comparable rates.
A hallmark of our nucleosomal array assay is that the
SWI/SNF-dependent enhancement of restriction enzyme
accessibility requires continuous ATP hydrolysis (32, 46). This
requirement reflects a state of constant redistribution of nucleosome
positions in the presence of ATP, and the subsequent inactivation of
SWI/SNF "freezes" a random positioning of nucleosomes which is
characterized by a general occlusion of restriction enzyme sites (46).
We carried out similar remodeling/"reversal" assays with hSWI/SNF, dCHRAC, dNURF, or xMi-2 and in all cases the enhanced rates of HincII digestion were lost after ATP was enzymatically
removed with apyrase (data not shown; for analysis of yRSC, see Ref.
41). Thus, these results indicate that all six complexes use the energy of ATP hydrolysis to create a dynamic, reversible state of nucleosome mobilization. Our results are consistent with previous demonstrations of mononucleosome mobilization catalyzed by ySWI/SNF (46, 49), dCHRAC
(47) or dNURF (48).
Nucleosome Moiety Requirements of the Chromatin Remodeling
Complexes--
Previous studies have demonstrated that optimal ATPase
activity of dNURF (29), dCHRAC (14), and xMi-2/NURD (18-20, 22) complexes requires nucleosomes, whereas the ATPase activities of
hSWI/SNF (17), ySWI/SNF (8), and yRSC (10, 41) complexes are stimulated
equally well by free DNA. Furthermore, in the case of the dNURF
complex, the nucleosome stimulation of ATPase activity requires one or
more trypsin-sensitive histone N-terminal domain(s) (30). To further
define the nucleosome moiety requirements for all six complexes, we
reconstituted nucleosomal arrays with hyperacetylated or trypsinized
histone octamers, as well as with histone (H3/H4)2 tetramers. To ensure that each type of array reconstitution was of
similar quality, all reconstitutions were analyzed for extent of DNA
repeat saturation and correct positioning by multiple restriction enzyme mapping and native polyacrylamide gel electrophoresis (see "Experimental Procedures"). We then measured the ability of these arrays to stimulate the ATPase activity of each complex (Fig. 2A). As expected, ySWI/SNF and
yRSC complex hydrolyzed ATP with similar kinetics on all substrates,
including free DNA (Fig. 2A; see also, Ref. 41). Likewise,
the ATPase activity of the hSWI/SNF complex was stimulated by all
substrates, with the exception that activity was consistently 40-50%
less in the presence of arrays reconstituted with histone
(H3/H4)2 tetramers (Fig. 2A).
In agreement with previous studies, we also found that the ATPase
activity of the dNURF complex was maximally stimulated only by
nucleosomal arrays (for analysis of ATPase activity with DNA or
hyperacetylated substrates, see Ref. 30); little ATPase activity was
detected with arrays reconstituted with trypsinized histone octamers or
histone (H3/H4)2 tetramers. Given that the ATPase activity
of dNURF requires one or more histone N-terminal domain(s) (Fig.
2A; see also, Ref. 30), the lack of ATPase activity in the
presence of the histone (H3-H4)2 tetramer arrays suggested that the N-terminal domains of the histone H2A/H2B dimers might play a
key role. However, nucleosomal arrays reconstituted with hybrid histone
octamers composed of intact histone (H3-H4)2 tetramers and
tail-less histone H2A-H2B dimers yielded maximal stimulation of dNURF
ATPase activity.2 Thus, the
inability of (H3-H4)2 tetramer arrays to stimulate the
ATPase activity of dNURF does not reflect a key role for the N-terminal
domains of histones H2A/H2B dimers.
The dCHRAC complex, like dNURF, contains ISWI, which is a
nucleosome-stimulated ATPase. In addition, dCHRAC also contains topoisomerase II which is a DNA-stimulated ATPase (14). Thus, the
ATPase activity associated with dCHRAC is a composite of ISWI and
topoisomerase II which complicates the analysis of the substrate preferences of this complex (Fig. 2A). The ATPase activity
of dCHRAC was stimulated by all substrates, although ATPase activity is
reproducibly higher in the presence of nucleosomal or hyperacetylated arrays. Since the ATPase activity of dNURF is only stimulated by a
nucleosomal or hyperacetylated substrate (Fig. 2A, see also Ref. 30), our data suggest that only 30-40% of the overall ATPase activity of CHRAC is due to the ISWI subunit, and the remaining DNA-stimulated ATPase activity is due to topoisomerase II.
The ATPase activity of the xMi-2 complex was distinct from both the
SWI/SNF (ySWI/SNF, yRSC, hSWI/SNF) and ISWI groups (dNURF, dCHRAC) of
ATPases (Fig. 2A). Like the ISWI group, the ATPase activity
of xMi-2 was maximally stimulated by nucleosomal arrays, although free
DNA did stimulate a significant amount of ATPase activity (27% of the
nucleosomal level). In contrast to the ISWI group, arrays reconstituted
with trypsinized histones were still able to stimulate the ATPase
activity of xMi-2 to nearly 70% the level of intact nucleosomal
arrays. Likewise, arrays reconstituted with hyperacetylated histones or
histone (H3-H4)2 tetramers were more similar to the
nucleosomal arrays. Thus the observed preference for nucleosomal arrays
does not reflect an obligatory interaction with the histone N-terminal
domains. Thus, based on a preference for a nucleosomal substrate and a
lack of histone tail dependence, xMi-2 appears to define a third group
of the ATP-dependent chromatin remodeling family.
We also performed coupled restriction enzyme-remodeling assays for most
of the different array substrates and each remodeling complex. As shown
in Fig. 2B, remodeling of the different substrate arrays
paralleled the ATPase activity of the complexes except in three cases.
First, although arrays reconstituted with histone (H3-H4)2
tetramers were able to stimulate the ATPase activity of ySWI/SNF and
hSWI/SNF, the apparent rate of remodeling of these tetramer arrays was
reduced 10-50-fold compared with remodeling of nucleosomal arrays
(Fig. 2B; see also Ref. 39 for an extensive discussion).
Second, although dCHRAC showed high levels of ATPase activity with all
substrates, it was not able to remodel arrays reconstituted with either
trypsinized histones or with the histone (H3/H4)2 tetramers
(Fig. 2B). These results suggest that the ATPase activity
that is presumably contributed by the topoisomerase II subunit of
dCHRAC is not sufficient to enhance restriction enzyme accessibility in
these array assays. Furthermore, the results from this remodeling
analysis indicate that dCHRAC and dNURF, which each contain the ISWI
ATPase, have indistinguishable histone moiety requirements. And
finally, although the ATPase activity of xMi-2 was stimulated well by
arrays reconstituted with (H3-H4)2 tetramers (Fig.
2A), these arrays were only poorly remodeled by xMi-2 (Fig.
2B).
Targeting of the Chromatin Remodeling Complexes by Transcriptional
Activators--
Recently we have shown that the remodeling activity of
ySWI/SNF can be targeted to reconstituted nucleosomal arrays by GAL4 derivatives that contain an acidic transcriptional activation domain
(50). For these targeting assays we used a modified array DNA template
which contains five high affinity GAL4-binding sites adjacent to the 5 S repeat that harbors the HincII/SalI site
(208-11S-GAL4; see Ref. 50). Reconstitution of nucleosomal arrays with
this DNA template positions the GAL4-binding sites in the linker region between two positioned nucleosomes (50). Targeting of remodeling activity is then assayed in HincII reactions which contain a
32P-labeled 208-11S-GAL4 array and 15-fold molar excess of
an unlabeled 208-11S array (which lacks GAL4 sites). In the absence of
targeting, the remodeling enzyme is sequestered by the excess
unlabeled, competitor array and there is little stimulation of
HincII digestion kinetics. Targeting of remodeling activity
is scored by any stimulation of HincII digestion kinetics
due to a functional GAL4 derivative (50). Note in this assay that a
GAL4 derivative does not affect HincII digestion kinetics in
the absence of remodeling complex or when the labeled and unlabeled
arrays lack GAL4-binding sites. Previously, using this assay we were
able to detect targeting of ySWI/SNF remodeling activity by GAL4-VP16
and GAL4-AH acidic activators (50).
We wished to investigate whether other members of the
ATP-dependent family of remodeling enzymes could also be
recruited by acidic activators in our reconstituted nucleosomal array
system. Each of the six remodeling complexes were added in parallel to HincII targeting assays which contained 0.2 nM
32P-labeled 208-11S-GAL4 nucleosomal array, 3 nM unlabeled 208-11S nucleosomal array, and 10 nM of a GAL4 derivative (Fig.
3). Under these reaction conditions,
little remodeling of the labeled 208-11S-GAL4 array was observed in the
absence of activator-dependent targeting. Likewise,
addition of the isolated GAL4 DNA-binding domain did not enhance
HincII digestion kinetics in the presence or absence of
remodeling enzyme (data not shown). Furthermore, similar to our
previous studies (50), the remodeling activity of ySWI/SNF was
effectively targeted to the 208-11S-GAL4 array by both the GAL4-VP16
and GAL4-AH acidic activators, as visualized by an activator- and
ySWI/SNF-dependent stimulation of HincII
digestion kinetics (Fig. 3 and data not shown; see also Table
I). In contrast, the remodeling activity
of yRSC, hSWI/SNF, dNURF, dCHRAC, or xMi-2 was not significantly
targeted by either GAL4-VP16 or GAL4-AH activators (Fig. 3A
and data not shown; see also Table I). In fact, we reproducibly
observed some activator-dependent inhibition of remodeling
by dCHRAC and yRSC (Fig. 3). Similar results were obtained in several
independent experiments and with a range of remodeler concentrations.
In the case of hSWI/SNF we also failed to observe targeting in this
assay using an immunoaffinity purified form of this enzyme
(51).3 Thus, for the six
purified remodeling complexes tested here, only the remodeling activity
of ySWI/SNF is effectively targeted by prototype acidic activators.
Our results indicate that six different members of the
ATP-dependent family of chromatin remodeling enzymes use
similar levels of ATP hydrolysis to rapidly establish a dynamic state
of enhanced nucleosome mobilization. This "fluid" chromatin state
is characterized by an enhanced accessibility of restriction enzymes
and DNA binding transcription factors. Furthermore, the nearly
identical rates of nucleosomal array remodeling (Fig. 1B)
and the common requirement for histone H2A/H2B dimers (Fig. 2) are
consistent with a similar remodeling mechanism for all members of this
ATP-dependent family. Although the mechanistic details of
"remodeling" are not clear, all of these enzymes can apparently
transduce the energy of ATP hydrolysis into an enhanced mobilization of
nucleosomes within linear arrays (as suggested in Ref. 52).
In contrast to our studies with nucleosomal arrays, some differences
between remodeling complexes have been observed with mononucleosome
substrates. For example, ySWI/SNF, NURF, and recombinant ISWI have been
shown to move a histone octamer from a central position to an end
position (46-48), whereas dCHRAC and ySWI/SNF can also move histone
octamers in the opposite direction (47, 49). dCHRAC, however, also
contains the ISWI ATPase, and thus these differences are not intrinsic
to the catalytic subunit or to the basic mechanism of remodeling.
Alternatively, the differences in the direction of histone octamer
movement may reflect the propensity of some complexes (such as dCHRAC)
to bind to DNA ends. In this scenario, protection of the DNA ends may
block end-directed movements and favor movements from the ends to more
central locations. In contrast, on nucleosomal arrays, where free DNA
ends do not flank individual nucleosomes, we propose that the direction
of histone octamer movement is random for all remodeling complexes.
This situation is consistent with our observation that the rates of nucleosome remodeling in the coupled array assay are similar for all complexes.
We were surprised to discover that arrays reconstituted with histone
(H3-H4)2 tetramers are not efficiently remodeled by any of
the complexes tested. In the absence of remodeling enzyme, arrays of
(H3-H4)2 tetramers are digested at rates only 3-5-fold faster than nucleosomal arrays (data not shown; see also Ref. 39),
whereas ATP-dependent remodeling of nucleosomal arrays typically yields 20-30-fold increases in restriction enzyme rates. Thus, arrays of (H3-H4)2 tetramers still provide a potent
barrier to factor access and, furthermore, the inability to score
remodeling of tetramer arrays is not due to a high level of restriction
enzyme cleavage in the absence of remodeling enzyme. Interestingly this requirement for the histone H2A-H2B dimers also does not reflect an
obligatory need for the N-terminal domains of these two histones, since
xMi-2, ySWI/SNF, and hSWI/SNF are insensitive to removal of all the
N-terminal domains (Fig. 2A) (41, 53). Instead, we favor a
model in which all of these enzymes require a canonical nucleosome
structure either for substrate recognition or for the mechanism of
remodeling. For instance, these enzymes may need to interact with two
adjacent gyres of DNA in order to induce nucleosome mobilization (see
also, Ref. 39).
Our data also suggest the delineation of three groups within the
ATP-dependent family: 1) a SWI/SNF group (ySWI/SNF, yRSC, and hSWI/SNF) whose ATPase activity does not require an intact nucleosome and whose remodeling function is independent of the histone
tails; 2) an ISWI group (dCHRAC, dNURF) whose ATPase activity requires
an intact nucleosome and whose remodeling function is histone tail
dependent; and 3) a Mi-2 group (xMi-2) whose optimal ATPase activity
requires an intact nucleosome and whose remodeling function is histone
tail independent. In fact, we found that the remodeling activity of
xMi-2 was actually enhanced by removal of the histone N-terminal
domains (Fig. 2B). Similar results have been obtained
previously using a subset of the complexes tested here as well as
recombinant BRG1 (ATPase subunit of hSWI/SNF) and ISWI (8, 14, 17, 28,
30, 41, 53). We note, however, that recombinant ISWI also shows
significant stimulation of ATPase activity by free DNA (54).
Furthermore, like dNURF and dCHRAC, the histone N-terminal domains
promote efficient remodeling by ySWI/SNF and yRSC complexes under
different reaction conditions where these enzymes must be catalytic
(41). Thus, although the different nucleosome moiety requirements are
important for defining distinctions among enzymes, these distinctions
are likely to reflect subtle differences in nucleosome recognition or
in regulation of the remodeling cycle (41, 56), rather than key
differences in the basic remodeling mechanism.
Molecular phylogenetic analysis has been used to organize the SWI2/SNF2
family of DNA-stimulated ATPases into multiple subfamilies (7). These
studies included sequence comparisons among different SWI2/SNF2 ATPase
domains as well as among sequences N-terminal or C-terminal to the
ATPase domain. Interestingly, ATPases from three of the subfamilies
defined by phylogenetic analysis are the catalytic subunits associated
with the three groups of ATP-dependent remodeling enzymes
delineated by our biochemical analyses (e.g. SWI2, ISWI,
Mi-2). This correspondence between such completely different
experimental approaches was not expected, since the homology among the
ATPase domains of SWI2, ISWI, and Mi-2/CHD proteins is very high (7).
One possibility is that a small number of amino acid changes can lead
to large differences in nucleic acid substrate requirements
(i.e. nucleosomes versus free DNA). Consistent
with this view, previous studies have found that ATPase domain swaps
between two members of the same subfamily (i.e. brahma and
SWI2/SNF2) yield a SWI2 protein that retains function in
vivo in yeast, whereas swaps between members of different families
(i.e. ISWI and SWI2/SNF2) are not functional (15). Alternatively, sequence elements that are unique to each subfamily that
lie outside of the ATPase domain (i.e. bromodomains, SANT domains, chromodomains) might also contribute to interactions with the
histone N-terminal domains or other nucleosomal components.
Although our comparative analysis delineates three groups of
ATP-dependent remodeling enzymes, our data also suggests
that individual enzymes within a single group are likely to be subject to differential modes of regulation. For instance, we found that ySWI/SNF was recruited by an acidic activator in our nucleosomal array
system, whereas other members of the SWI/SNF group (e.g. yRSC, hSWI/SNF) were not. We anticipate that yRSC and hSWI/SNF can be
recruited by other types of activators in this assay. Likewise, members
of the ISWI or Mi-2 groups are likely to be recruited by nonacidic
activators or by transcriptional repressors. These ideas are consistent
with several previous studies. First, acidic activators are unable to
recruit yRSC complex to an immobilized DNA template from a yeast
nuclear transcription extract (50). Second, xMi-2 complexes are
believed to function in transcriptional repression (36, 57) and thus it
is not surprising that an acidic activator is unable to recruit xMi-2.
And finally, hSWI/SNF has recently been demonstrated to be targeted
in vivo by the glucocorticoid receptor (58), an isoform of
C/EBP- Clearly, acidic activators are likely to recruit
ATP-dependent remodeling complexes in Drosophila
and mammalian cells. One possibility is that there exists additional,
uncharacterized members of the ATP-dependent remodeling
family that can be recruited by acidic activators and which might play
a key role in acidic activator function. It is also possible that
regulatory subunits, which might facilitate interactions with acidic
activators, have been lost during purification of one or more of the
remodeling complexes that we have tested here. Alternatively, several
of the more abundant complexes (e.g. dCHRAC and yRSC) may
establish more global domains of fluid chromatin, and thus they may not
rely on gene-specific targeting proteins.
We thank members of the Peterson laboratory
for helpful discussions throughout the course of this work. We are
especially grateful to Bradley Cairns (University of Utah) for the
generous gift of yRSC complex.
*
This work was supported by National Institues of Health
Grants GM49650 (to C. L. P.) and GM56244 (to A. N. I.) and by a fellowship from the Human Frontiers Science Program
Organization (to C. L.).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
§
Contributed equally to the results of this work.
¶
Current address: Dept. of Molecular Biology, 6525 ED Nijmegen,
The Netherlands.
¶¶
To whom correspondence should be addressed: 373 Plantation St., University of Massachusetts Medical School, Biotech 2, Suite 301, Worcester, MA 01605. Tel.: 508-856-5858; Fax: 508-856-4289; E-mail: craig.peterson@ummed.edu.
Published, JBC Papers in Press, April 21, 2000, DOI 10.1074/jbc.M002810200
1
D. Hill and A. N. Imbalzano, unpublished observations.
2
P. Horn and C. L. Peterson, unpublished observation.
3
P. Horn, R. E. Kingston, and C. L. Peterson, unpublished observations.
Functional Delineation of Three Groups of the
ATP-dependent Family of Chromatin Remodeling Enzymes*
§,§¶,
,,,¶¶
Program in Molecular Medicine and Department
of Biochemistry and Molecular Biology, University of Massachusetts
Medical School, Worcester, Massachusetts 01605, the ** Laboratory of
Molecular Embryology, NICHD, National Institutes of Health, Bethesda,
Maryland 20892, the Adolf Butenandt-Institut, Molekularbiologie,
Schillerstr. 44, 80336 Munchen, Germany, the
§§ Department of Cell Biology, University of
Massachusetts Medical School, Worcester, Massachusetts 01655, and the
Laboratory of Molecular Cell Biology, NCI,
National Institutes of Health, Bethesda, Maryland 20892
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-32P]dATP as described (31, 32).
) as described in Logie and Peterson (31). The
concentration of complex was determined to be approximately 300 nM by comparative Western blot and ATPase assays (31, 32).
yRSC (10), xMi-2 (36), dCHRAC (14), dNURF (29), and hSWI/SNF "A"
(17) were purified as described previously. Approximate concentrations
were estimated from total protein concentration in the purified
fractions and complexes were assumed to be 100% active. Thus, our
concentration estimates are likely to be an overestimate. Most
complexes had a high degree of purity, but in the case of hSWI/SNF A,
purity was estimated to be ~10%. We confirmed that the activity
monitored was in fact due to hSWI/SNF by antibody inhibition, addition
of antisera directed to the BRG1 subunit of hSWI/SNF eliminated
remodeling activity, whereas addition of preimmune sera had no
effect.1 For all the studies
described here, each assay was performed with two independent
preparations of each remodeling complex, with the exception of yRSC.
-32P]ATP (Amersham Pharmacia Biotech) in 0.1%
Tween, 20 mM Tris, pH 8, 5% glycerol, 0.2 mM
dithiothreitol, 5 mM MgCl2, 1 mM
phenylmethylsulfonyl fluoride, and 0.1 mg/ml bovine serum albumin as
described (31, 32, 41). Released phosphate was monitored with time by
resolution of Pi from ATP on plastic plates coated with PEI
cellulose (EM Science) with 0.75 M KPO4 (pH
3.5) as solvent and quantified by PhosphorImager analysis.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (18K):
[in a new window]
Fig. 1.
Comparison of ATPase and remodeling
activities of ATP-dependent chromatin remodeling
complexes. A, ATPase assays. The indicated remodeling
complexes (~1-5 nM ySWI/SNF, hSWI/SNF, dNURF, dCHRAC, or
~15 nM xMi-2) were analyzed in ATPase reactions that
contained (closed symbols) or lacked (open
symbols) 12 nM nucleosomal array, and ATP hydrolysis
was monitored with time. Velocities of ATP hydrolysis were calculated
from at least three reaction time points. B, nucleosomal
array remodeling assays. HincII digestion of nucleosomal
arrays incubated in the presence (closed symbols) or absence
(open symbols) of the indicated remodeling complexes.
HincII digestion rates were calculated from the slopes of
plots of the natural logarithm of the fraction of uncut array
versus time. These results are representative of multiple,
independent experiments. Similar results were also obtained with at
least two independent enzyme preparations for each complex except
yRSC.
View larger version (45K):
[in a new window]
Fig. 2.
Nucleosome moiety requirements for
ATP-dependent chromatin remodeling enzymes.
A, ATPase assays. The indicated remodeling enzymes were
added to ATPase assays that contained either 208-11S DNA template (DNA)
or 208-11S arrays reconstituted with histone octamers, hyperacetylated
histone octamers, trypsinized histone octamers, or isolated
(H3-H4)2 tetramers. Each reaction represented an ATPase
time course, and ATP hydrolysis velocities were calculated for each
substrate. Data is presented as a percentage of the ATPase velocity
exhibited with the nucleosomal array substrate. Data shown for yRSC is
the result of a single experiment which essentially repeated our prior
study (41). *, denotes that ATPase assays with DNA and hyperacetylated
array substrates were not performed with dNURF (see Ref. 30 for
detailed analysis). B, nucleosomal array remodeling assays.
HincII digestion rates were determined for each enzyme on
each array substrate. For each substrate, rates were calculated from
the slopes of plots of the natural logarithm of the fraction of uncut
array versus time. Data is presented as a percentage of the
remodeler-dependent HincII digestion rate of the
nucleosomal array. With the exception of yRSC, the results shown
include experiments with at least two independent enzyme preparations
for each complex. *, denotes that remodeling of tetramer arrays was not
performed with yRSC. In addition, remodeling data shown for yRSC with
hyperacetylated and trypsinized nucleosomal arrays is from Logie
et al. (41) and is shown for comparison purposes only.
View larger version (30K):
[in a new window]
Fig. 3.
Recruitment of ATP-dependent
enzymes by GAL4-VP16. HincII digestion kinetics of
reactions containing 0.2 nM labeled 208-11S-GAL4 array, 3 nM unlabeled 208-11S competitor array in the presence or
absence of 10 nM GAL4-VP16, and in the presence or absence
of the indicated chromatin remodeling complexes. y axis
shows percent uncut nucleosomal array. Note that addition of GAL4-VP16
had no effect on HincII digestion kinetics in the absence of
remodeling complex. Experiment shown is representative of multiple,
independent experiments.
Recruitment of chromatin remodeling complexes by GAL4-VP16
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
(59) and erythroid kruppel-like factor (60).
ACKNOWLEDGEMENTS
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DISCUSSION
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