2222

Biophysical Journal

Volume 104

May 2013

2222–2234

Phosphorylation-Coupled Intramolecular Dynamics of Unstructured
Regions in Chromatin Remodeler FACT
Manami Hashimoto,†††6 Noriyuki Kodera,{6 Yasuo Tsunaka,‡‡‡6 Masayuki Oda,** Mitsuru Tanimoto,kToshio Ando,{k Kosuke Morikawa,‡§* and Shin-ichi Tate†††*†Department of Mathematical and Life Sciences, Hiroshima University, Hiroshima, Japan; ‡Institute for Integrated Cell-Material Sciences(iCeMS), Kyoto University, Kyoto, Japan; §International Institute for Advances Studies, Kyoto, Japan; {Bio-AFM Frontier Research Center,College of Science and Engineering, Kanazawa University, Kanazawa, Japan; kSchool of Mathematics and Physics, College of Science andEngineering, Kanazawa University, Kanazawa, Japan; **Graduate School of Life and Environmental Sciences, Kyoto Prefectural University,
Kyoto, Japan; ††SENTAN, Japan Science and Technology Agency, Tokyo, Japan; and ‡‡PRESTO, Japan Science and Technology Agency,
Tokyo, Japan

ABSTRACT The intrinsically disordered region (IDR) of a protein is an important topic in molecular biology. The functional significance of IDRs typically involves gene-regulation processes and is closely related to posttranslational modifications such as
phosphorylation. We previously reported that the Drosophila facilitates chromatin transcription (FACT) protein involved in chromatin remodeling contains an acidic ID fragment (AID) whose phosphorylation modulates FACT binding to nucleosomes. Here,
we performed dynamic atomic force microscopy and NMR analyses to clarify how the densely phosphorylated AID masks the
DNA binding interface of the high-mobility-group domain (HMG). Dynamic atomic force microscopy of the nearly intact FACT
revealed that a small globule temporally appears but quickly vanishes within each mobile tail-like image, corresponding to
the HMG-containing IDR. The lifespan of the globule increases upon phosphorylation. NMR analysis indicated that phosphorylation induces no ordered structure but increases the number of binding sites in AID to HMG with an adjacent basic segment,
thereby retaining the robust electrostatic intramolecular interaction within FACT even in the presence of DNA. These data lead tothe conclusion that the inhibitory effect of nucleosome binding is ascribed to the increase in the probability of encounter between
HMG and the phosphorylated IDR.

INTRODUCTION
In recent years data derived from bioinformatics, protein
physical chemistry, and tertiary structure analyses, suchas x-ray crystallography and NMR, have led to the conclusion that numerous functional proteins contain regions or
domains that do not form well-defined orderly threedimensional structures but rather exist as dynamic ensembles of interconverting flexible conformers (1–3). These
intrinsically disordered regions (IDRs) are ubiquitously
found in eukaryotic proteins that play crucial roles ingene regulation within nuclei (4–8). It is known that these
IDRs are involved in functionally important molecular
recognition, where they fold into ordered conformations
through binding to rigid ordered protein subunits or domains, a process that is generally termed coupled folding(9,10). Conversely, IDRs in a number of proteins do notnecessarily undergo global disorder-to-order transitions
upon binding. Moreover, several important regulatory interactions involve dynamic complexes in which main
chains of IDRs continue to fluctuate without forming
defined architectures (11,12). These binding events appear
to be modulated by critical posttranslational modifications

Submitted October 10, 2012, and accepted for publication April 4, 2013.
6

Manami Hashimoto, Noriyuki Kodera and Yasuo Tsunaka contributed
equally to this work.
*Correspondence: tate@hiroshima-u.ac.jp or kmorikawa@icems.kyoto-u.
ac.jp

including phosphorylation, acetylation, methylation, andubiquitination (8,11,13).
Our previous work reported that high-speed atomic force
microscopy (HS-AFM) can simultaneously visualize eachbehavior of IDRs and rigid domains of Drosophila chromatin transcription-facilitating protein (dFACT) on a substrate surface in solution (14). This method is suitable forstudying proteins involved in gene expression, such as transcriptional regulators and chromatin-remodeling factors.
For example, FACT, classified as a chromatin remodeler,
is a heterodimer complex that consists of SSRP1 and
SPT16 subunits (Fig. 1). Both subunits contain several rigid
domains and IDRs that play crucial roles in chromatin
remodeling and transcriptional elongation (15–19). Thus,
FACT is a suitable target for investigating the mechanistic
and functional aspects of IDRs in solution through HS-AFM
and NMR analyses.
In a previous study on interactions between dFACT andnucleosomes, we found that dFACT initially binds nucleosomes and/or nucleosomal DNA via a high-mobility-groupdomain (HMG) and an HMG-flanking basic ID segment(BID) of the dSSRP1 subunit, which jointly forms a
DNA-binding element (Fig. 1) (20). The acidic ID segment(AID), adjacent to BID, forms intramolecular interactions
with both HMG and BID (Fig. 1). Extensive phosphorylation of AID strikingly increases the number of negative
charges, thereby strengthening the intramolecular interactions. As a result, the binding of dFACT to nucleosomal

Editor: Daniel Muller.
Ó 2013 by the Biophysical Society
0006-3495/13/05/2222/13 $2.00

http://dx.doi.org/10.1016/j.bpj.2013.04.007

Phosphorylated ID region dynamics

2223
phorylation of the AID and AB-HMG fragments by casein kinase II (CK2)
are described in the Supporting Material.
For the dynamic AFM experiments, the His-tagged Drosophila SPT16(401–887) (dSPT16) and Drosophila SSRP1 (dSSRP1) were cloned intoa pFastBacDual plasmid. To obtain the Ser/Thr-to-Ala mutants at the
10 phosphorylation sites (10SA; S443A, S472A, S476A, T477A, S488A,
S496A, S500A, S502A, S506A, and S515A (Fig.S1 in the Supporting
Material)), site-directed mutagenesis of the dFACT proteins was performedby the QuikChange method (Stratagene, La Jolla, CA). The nearly intact
dFACT heterodimers (His-dSPT16 (401–887) þ dSSRP1) were coexpressed as the fully phosphorylated form (wild–type (WT)) and a nonphosphorylated form (10SA) in Sf9 insect cells and purified as described
previously (20).

FIGURE 1 A schematic drawing of the domain organization of dSPT16
and dSSRP1, the two subunits in dFACT. The fragments used in this work
are listed: AID; SB-HMG (residues 543–554), which contains a part of the
BID region colored in purple (residues 519–554); and AB-HMG. These fragments jointly constitute the regulatory region of dSSRP1 binding to DNA.
The heterodimeric complex, consisting of dSPT16 (residues 401–887) andthe full-length dSSRP1, was used for HS-AFM measurements.

DNA is blocked, preventing the formation of rigid structures(20). The physiological significance of this control mechanism is highlighted by rapid chromatin transactions during
early embryogenesis through dephosphorylation in thematernally transmitted dSSRP1 after fertilization (20). In
humans, the dephosphorylated SSRP1, including theHMG, is essential for FACT to exhibit higher binding affinity with nucleosomes (19). In contrast, the yeast FACTHMG functions as an isolated Nhp6a/b protein (15,21,22).
Thus, the phosphorylation-dependent regulation is likely
to be conserved only in higher eukaryotes. Despite theimportance of AID phosphorylation, the detailed blocking
mechanism remains unresolved at the atomic level.
To clarify the mechanism of intramolecular interaction
between the AID segment and the DNA-binding elements,
we investigated the detailed effects of phosphorylation by
combining NMR spectroscopy and HS-AFM. Specifically,
we prepared large amounts of various fragments, whichwere labeled with stable isotopes for NMR in both the phosphorylated and nonphosphorylated states, whereas FACT
molecules nearly intact in size were observed by HS-AFM.
Thus, this study allowed us to conclude that dense phosphorylation causes an electrostatic reinforcement of the interactions between the acidic ID region and the DNA-binding
elements, and that the essence of the interactions is dynamic
and transient, thus lacking specific and stable binding sites
in domains or fragments.
MATERIALS AND METHODS
Protein preparation
The dFACT used was the same as that described in detail in our previous
study (20). The fragments used in this work are schematically drawn in
Fig. 1: AID, the short basic IDR plus HMG (SB-HMG), and the acidic
and basic IDRs plus HMG (AB-HMG). These fragments were expressedin Escherichia coli. Details of the purification of each fragment and phos-

Dynamic AFM analysis of the dFACT heterodimer
Dynamic AFM imaging experiments of the nearly intact dFACT heterodimers (Fig. 1) were carried out as described previously (14) using a laboratory-built HS-AFM (23,24). Briefly, we diluted the dFACT-WT ordFACT-10SA samples to ~2 nM with buffer A (20 mM Tris-HCl,
pH7.5, 50 mM KCl, 10 mM MgCl2, and 0.5% glycerol (vol/vol)). The
diluted samples were used within 3 h. A droplet (2 mL) of a diluted sample
was deposited on a freshly cleaved mica surface (~1 mm in diameterand <0.05 mm in thickness), which had been glued onto a glass stage(2 mm in diameter and 2 mm in height). After incubation for 3 min, molecules that were not attached to the mica surface were removed by rinsing
with ~20 mL of buffer A. The sample surface was not allowed to dry. Subsequently, the sample stage was immersed in a liquid cell filled with~60 mL of buffer A in which a small cantilever had been fixed. Imaging
was carried out by HS-AFM in tapping mode. Details of the HS-AFM
imaging experiments were summarized in a recent work (25). For image
analysis, we applied three filters in the following order. First, a low-passfilter to remove spike noise; second, a flattening filter to make the xy-planeflat; third, a line-to-line base collection filter to minimize the base height
difference between lines.

NMR spectroscopy
The backbone 1H/13C/15N resonance assignments of AID and phosphorylated AID fragments were performed with a standard set of triple resonanceexperiments on a DMX600 spectrometer (Bruker, Billerica, MA). (BMRB
entry 11511) Details are available in the Supporting Material.

NMR titration experiments
The interactions between the isolated AID and SB-HMG fragments were
performed in two ways: one experiment used nonlabeled AID titrated
into 15N-labeled SB-HMG, and the other experiment involved titrating nonlabeled SB-HMG into 15N-labeled AID. To explore how phosphorylation of
AID influenced its interaction with binding partners, a phosphorylated AID
fragment was also used in the titration experiments. The chemical-shiftchanges observed in a series of two-dimensional (2D) 1H-15N heteronuclear
single-quantum coherence (HSQC) spectra during the titration wereanalyzed by numerical curve fitting to elucidate the number of effective
binding sites, n, and the dissociation constants, KD, according to a previously published procedure (26). In the calculations, we applied a global
fitting procedure that incorporates all the chemical-shift-change profiles
for the residues under study, with n and KD as global adjustables (27).
Monte Carlo simulation was applied to estimate the statistical errors forn and KD, assuming an uncertainty for each peak position on a 1H-15N
HSQC spectrum of 0.002 ppm (1H) and 0.02 ppm (15N), respectively,
and using 64 synthetic data sets with Gaussian noise (28). Experimental details in the titration experiments between the isolated AID and SB-HMG
fragments are described in the Supporting Material.
Biophysical Journal 104(10) 2222–2234

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NMR analysis on the interaction between AB-HMG
and double-stranded DNA
We carried out NMR spectral analysis for the 15N-labeled nonphosphorylated
and phosphorylated AB-HMG fragments in the absence and presence of DNA(Fig. 1). The limited solubility and the severe spectral overlap in the spectraof AB-HMG did not permit complete assignment of the resonances arisingfrom the backbone nuclei. Only a limited number of resonances were assigned for AB-HMG. We used double-stranded DNA (dsDNA) with thesequence 50 -d(CGCGATATCGCG)2-30 . The reaction mixture including thedsDNA was passed through a NAP-10 column to exchange the buffer to
50 mM Tris-HCl, pH 6.6, the buffer used for the NMR experiments.

RESULTS
Molecular features of phosphorylated andnonphosphorylated dFACT heterodimers
visualized by HS-AFM
The wild-type dFACT (dFACT-WT) is spontaneously phosphorylated in Sf9 insect cells and is expressed as the fully
phosphorylated form at 10 phosphorylation sites. In
contrast, a dFACT mutant containing Ser/Thr-Ala substitutions at each of these sites, dFACT-10SA, is expressed as
the nonphosphorylated form, as described previously (20).
The molecular features of dFACT-WT (the phosphorylatedform) and dFACT-10SA (the nonphosphorylated form)
attached to mica surfaces in the buffer solution were directly
visualized by HS-AFM (Fig. 2, A and B, respectively; also
see Movies S1 and S2). At first glance, it appears thatboth molecules consist of a large globular domain and along tail region. The large globular domain appears to bewell adsorbed onto the mica surface, and its position does
not change significantly over time. In contrast, the longtail region exhibits rapid fluctuations because of thermal
agitations, indicating that this region has less affinity to
the mica surface. These molecular features are consistent
with the results of our previous study (14). Considering
our previous data (14), we can assign the large globular
domain (termed GD1, see Fig. 2 C) to the structured domain
consisting of dSPT16 (401887) and dSSRP1 (1404)(Fig. 1). The long tail region can be assigned to the IDR
connected to the HMG (Fig. 1).
With careful observations, we noticed that the IDR has
two small globular domains, which were connected with
IDR1–2 and IDR2–3 (Fig. 2 C). One of the small globular
domains (GD3 (see Fig. 2 C)) was consistently observed at
the end of the IDR and its appearance is not substantiallydifferent between the two constructs. In our previous study,
this small globular domain was not observed, although thesample was prepared using the E. coli expression system.
Furthermore, the amino acid sequence indicates no tertiary
structure at the end of the IDR. Taken together, we postulatethat unknown posttranslational modifications would induce
a globular domain at the end of the IDR.
On the other hand, the other small globular domain (GD2
(Fig. 2 C)) appears temporally around the middle of the IDR
Biophysical Journal 104(10) 2222–2234

Hashimoto et al.

but shortly vanishes. In other words, we observed that theheight of GD2 changes over time. In addition, the figure of
GD2 appears to be retained for a longer period in dFACTWT than in dFACT-10SA, whereas the length of the IDR
appears to be slightly shorter in dFACT-WT than indFACT-10SA. Notably, the distance between GD1 and
GD2 appears to be much shorter in dFACT-WT; however,the distance between GD2 and GD3 is similar in both constructs. Similar tendencies were observed for most of theother protein molecules examined in each construct.
Quantitative analysis of distinct HS-AFM images
between phosphorylated and nonphosphorylateddFACTTo quantitatively evaluate minute variations of the abovementioned molecular features, we performed image analysis. Because the IDRs are highly flexible and can adopt
similar molecular features at an arbitrary time, successive
AFM images must be analyzed without skipping images
to maintain arbitrariness of data sampling. To identify
IDRs that contribute to changes in molecular features, astraightforward approach is required for contour length
analysis of IDRs. However, considering the spatiotemporal
resolution of the HS-AFM, it is currently not possible toapply contour-length analysis to successive AFM images
without skipping images. As an alternative, we created asimple schematic representation of the molecular featuresof dFACT observed by HS-AFM (Fig. 2 C). In this schematic, the heights of the three globular domains (H1–H3)
and the end-to-end distances between the two globular
domains (D1–2, D2–3, and D1–3) were obtained from a single
AFM image. This approach allowed us to analyze almost all
AFM images, even when the AFM images had poor signal/
noise (S/N) ratios. The end-to-end distance can be used toadequately evaluate the length of a biological polymer ona surface, as described previously (29), and our data therefore support our interpretation. The analysis was performedon three typical molecules for the phosphorylated (dFACTWT) and nonphosphorylated (dFACT-10SA) constructs, andthe results are summarized in Table 1. We believe that thethree molecules selected lie within the population as a wholeof each construct, because most of the molecules observed
displayed similar molecular features. For example, we
can see four molecules of dFACT-10SA simultaneously(Movie S3), all of which appear to have similar molecularfeatures. Note that the parameters described herein are not
likely to have reached the most probable values, because
we analyzed 4586–5770 images of only three molecules
for each construct. However, we believe that these parameters will be convincing for our conclusions because they
represent well the molecular features of each construct.
The details are described below.
The distributions of H1 appeared to be similar for bothconstructs, and the mean height was 3.9 nm (Fig. 3, A

Phosphorylated ID region dynamics

2225

FIGURE 2 Typical HS-AFM images showing the molecular features of dFACT-WT (A) and dFACT-10SA (B). These HS-AFM images were clipped fromthe movie files (see Movie S1 for dFACT-WT and Movie S2 for dFACT-10SA). Every image was taken at 67.08 ms/frame (~15 frames/s). The time from the
beginning of the clip is indicated at the upper left of each image. Scanning area, 100  100 nm2 with 80  80 pixel; Z-scale, 4.0 nm. The observed molecularfeatures of dFACT are schematized according to the definitions in C. These schematics were drawn freehand by tracing the AFM images by visual estimation.
(C) Top-view (upper) and side-view (lower) schematics simply represent the characteristics of dFACT observed by HS-AFM. Gray-colored ellipses andblack-colored thick solid lines represent the GDs and the IDRs, respectively. The symbols used for image analysis are also depicted. For every image,
we selected three points (P1–P3) (green dots) representing the peak heights (Hi) in their respective GDs. Using these points, a distance from the highest pointof one GD to that of another was determined. For example, the distance from P1 of GD1 to P2 of GD2 was expressed as D1–2. In some images, the appearanceof GD2 or GD3 was unclear; there was no point with distinct heights around the area usually seen. In this case, P2 and P3 with the highest height around the
middle and the end of the IDR were selected.

and D, and Table 1), indicating no significant difference in
the GD1 for the two constructs. This height is the same asthat obtained in our previous study (14). Although the distributions of H3 appeared to differ, the mean height was
1.6 nm, with similar standard deviations for both constructs(Fig. 3, C and F, and Table 1), indicating that the GD3
domain may not differ between the two constructs. In
contrast, distributions for H2 were notably higher for
dFACT-WT than for dFACT-10SA (Fig. 3, B and E). Twopeaks could be seen in the H2 distribution of dFACT10SA around 1.1 nm and 1.7 nm (Fig. 3 E). The peak around
1.1 nm was higher and wider than that around 1.7 nm, andthe ratio of the areas of the two peaks was ~2:1 (low/high)(Fig. 3 E). Because we observed at least two physical states

in GD2 (i.e., a lower-height and a higher-height state), thisresult indicates that the GD2 of dFACT-10SA tends towarda lower-height state than a higher one. On the other hand,the H2 distribution of dFACT-WT appears to be a combination of a large peak around 1.6 nm and a small peak around
1.1 nm because a small shoulder around 1.1 nm could beseen in the distribution (Fig. 3 B). The area of the large
peak was more than twice as wide as that of the small
peak, indicating that the GD2 of dFACT-WT tends towarda higher-height state than a lower one. Consequently, theseresults suggest that the GD2 of dFACT-WT resides in thehigher-height state much longer than does that of dFACT10SA. Considering the two height values seen in the H2
distributions (i.e., 1.1 nm for the lower-height state andBiophysical Journal 104(10) 2222–2234

2226
TABLE 1

Hashimoto et al.
Summary of the AFM image analysis

Parameter

dFACT-WTdFACT-10SA

H1
H3
D1–2
D2–3
D1–3
D1–2/(D1–2 þ D2–3)
Number of molecules analyzed
Number of frames analyzed

3.9 5 0.3 nm
1.6 5 0.3 nm
15 5 5 nm
10 5 3 nm
21 5 5 nm
0.59 5 0.11
35770

3.9 5 0.3 nm
1.6 5 0.4 nm
17 5 5 nm
11 5 4 nm
23 5 7 nm
0.62 5 0.11
34586

Heights and distances are shown by mean 5 SD. Units of areas are arbitrary. The accuracy of the HS-AFM measurement for the z-direction (i.e.,background noise) was 0.15 nm. The standard deviations of the Gaussianfits were therefore restricted to >0.15 nm. Note that we did not apply
Gaussian fittings to the H2 distributions because the results obtained had
no statistical relevance.

1.6–1.7 nm for the higher-height state), the height of the
HMG on one side corresponded to ~1 nm (30) and the heightof the IDR was determined to be 0.4–0.6 nm. These datacollectively suggest that the lower-height state (H2 of~1.1 nm) and higher-height state (H2 of 1.6–1.7 nm) represent the HMG alone and the HMG associated with the IDR,
respectively.
The distributions of D2–3 were similar between dFACTWT and dFACT-10SA (Fig. 3, H and K), and the mean
length of D2–3 showed almost the same values (Table 1).
The length of IDR2–3 did not change in response to phosphorylation. Meanwhile, the mean length of D1–2 ofdFACT-WT (15 5 5 nm) was shorter by ~2 nm than thatof dFACT-10SA (17 5 5 nm) (Fig. 3, G and J, and Table 1).
Therefore, the shorter state of IDR1–2 within dFACT-WT(the phosphorylated form) is generated by the interaction
between HMG and IDR1–2, which contains the phosphorylated AID, but not that between HMG and IDR2–3. The
mean length of D1–3 of dFACT-WT (21 5 5 nm) was also
shorter by ~2 nm than that of dFACT-10SA (23 5 7 nm)(Fig. 3, I and L, Table1). This difference can be ascribedto the difference in the D1-2 for the two constructs because
we observed no difference greater than 2 nm in D2-3.
To estimate the mean position of the GD2 in the long taildomain, the ratios of D1–2/(D1–2 þ D2–3) for dFACT-WT(0.59 5 0.11) and dFACT-10SA (0.62 5 0.11) were determined (Fig. 4), indicating that phosphorylation of IDR1–2
shifts the mean position of GD2 to GD1 but not to GD3.
Combined with the results in Fig. 3, these results demonstrate that the height change for GD2 and the length changes
for D1–2 in dFACT-WT result from the interaction between
the HMG and IDR1–2, but not between the HMG and
IDR2–3. This observation is in good agreement with our previous model (20).
To summarize the HS-AFM data, GD2 temporally appears and then vanishes in both constructs. However, it isimportant that the lifetime of the higher-height state indFACT-WT is much longer than that in dFACT-10SA. In
addition, dFACT-WT seems to have a shorter IDR thanBiophysical Journal 104(10) 2222–2234

dFACT-10SA. These data indicate that the higher-heightstate of GD2 originates from the interaction between the
HMG and IDR1–2, which contains the phosphorylated AID(Fig. 1). Analysis by HS-AFM is highly suitable for directly
observing the dynamic behavior of large protein complexeson the surface of a substrate in solution. The findings
described here should be analyzed by NMR, which canprovide detailed local and atomic information on proteinprotein interactions.
Structural characterization of the phosphorylated
AID fragment
The fragments used in the NMR analysis are schematicallydrawn in Fig. 1. These fragments were expressed in E. coli
and were phosphorylated by CK2 in vitro (Fig. S2). The
limited signal dispersion of the isolated AID fragment in
the 2D 1H-15N HSQC spectrum is typical of a disordered
protein (Fig. 5 A). The 15N{1H} heteronuclear nuclear Overhauser effect (NOE) values for most residues of the AID
were negative, indicating that the fragment is essentiallyin a disordered state. Residues around 453 showed positive
heteronuclear NOE values. This region is rich in hydrophobic amino acids and is therefore likely to have restricted
local backbone motions; although the motional properties
do not seem functionally relevant (Fig. 5 C). CK2 successfully phosphorylated all nine canonical phosphoacceptor
serines within the AID, as revealed from high-field NMR
chemical shift changes in both the 1H and 15N dimensions(Fig. 5 B); T477 was not phosphorylated in this sample
preparation, presumably due to the lower activity of CK2
toward Thr residues (31). The phosphorylation of AID did
not induce any structural changes, because the increase in
the resonance dispersion in the 2D 1H-15N HSQC spectrum(data not shown) was negligible. The 15N{1H} heteronuclear NOE profile for the phosphorylated AID (pAID) also
indicated that it is in a disordered state (Fig. 5 C). Nonetheless, the heteronuclear NOE values of pAID did show an
overall small but significant increase in the size of theNOE values when compared with the data for the nonphosphorylated form (Fig. 5 C). Presumably, electrostatic repulsion among phosphate groups may induce some fragment
stiffness.
Interactions between the isolated AID
and SB-HMG fragments
To elucidate the binding modes between isolated fragments,
both the AID and pAID were titrated to the 15N-labeled
SB-HMG (Fig. 6, A and B), which contains a short basic
IDR and the HMG (Fig. 1). The normalized chemical-shiftchanges observed in the 2D 1H-15N HSQC spectra of15
N-labeled SB-HMG in the titration (Fig. 6, A and B) areplotted versus the residue numbers (Fig. 6, C and D). The
comparison between the two profiles showed that AID andPhosphorylated ID region dynamics

2227

FIGURE 3 Height distributions of three GDs and distributions of distance between two GDs of dFACT. Height distributions (H1–H3) are shown fordFACT-WT (A–C) and dFACT-10SA (D–F). Distance distributions (D1–2, D2–3 and D1–3) are shown for dFACT-WT (G–I) and dFACT-10SA (J–L). Greenlines represent single-Gaussian fitting. Note that we did not apply Gaussian fittings to the H2 distributions because the results obtained had no statisticalrelevance. A summary of the analysis is presented in Table 1. These results were obtained from analysis applied to three molecules for each construct.

pAID share similar binding sites on SB-HMG, irrespectiveof phosphorylation. Although they do have contact with
the DNA binding surface of the HMG, both fragments primarily bind to the BID, because the residues in the BID
showed greater chemical shifts than those in the HMG(Fig. 6, C and D). Furthermore, we found in our previous
work that the BID is essential for intramolecular interaction
with the AID (20). The magnitudes of the chemical shiftchanges by the addition of pAID are overall greater thanthose observed for AID binding, indicating that pAID is
likely to have higher affinity to SB-HMG. That is, theincrease in the population of the bound form of SB-HMG
with pAID causes greater spectral changes relative to thefree-state spectrum (27).
The numerical fittings to the chemical shift changes
against the molar ratios of AID or pAID to SB-HMG

showed that phosphorylation alters their binding modes to
SB-HMG (Fig. 6, E and F) (26,32). To determine the KD
and n values from the experiments by using 15N-labeled
SB-HMG, we included in our calculation 11 residues for
AID and 30 residues for pAID, showing that the significant
changes deviated more than the mean þ 1 SD (Fig. 6, C andD, red bars). The number of SB-HMG-binding sites on AID
was estimated at n ¼ 2.0 5 0.1 with a dissociation constantof KD ¼ 9.7 5 0.3 mM, assuming that all the binding sites
on AID have equivalent affinities for SB-HMG. In the caseof SB-HMG binding to pAID, the number of binding sites
was estimated at n ¼ 3.9 5 0.1 with KD ¼ 1.01 5
0.02 mM. This indicates that phosphorylation of AID doubles its number of SB-HMG-binding sites in addition toenhancing its affinity for SB-HMG. In agreement with thesedata from NMR, isothermal calorimetry (ITC) experiments
Biophysical Journal 104(10) 2222–2234

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FIGURE 4 Plots showing the ratio of D1–2/(D1–2 þ D2–3), which indicate
the mean positions of GD2 in the tail regions of dFACT-WT (A) and dFACT10SA (B). Green lines represent single-Gaussian fitting. A summary of theanalysis is presented in Table 1. These results were obtained from analysis
applied to three molecules for each construct.

using the SB-HMG and AID or pAID showed that the number of binding sites on AID approximately doubles, depending upon phosphorylation (data not shown).
The SB-HMG titration to 15N-labeled AID or pAID was
also examined. The spectral changes in the 2D 1H-15N
HSQC spectra for AID and pAID are shown in Fig. 7, A
and B, respectively. The 20 residues in AID showed significant spectral changes (chemical-shift changes deviating by
more than the mean þ 1 SD of the entire data) upon binding
to SB-HMG. It is thus likely that these residues are involved
in binding to SB-HMG (Fig. 7 E). The residues are classified
into two groups according to their affinity (nine high-affinity
residues (Fig. 7 C, left, and Fig. 7 E, blue bars) and 11 lowaffinity residues (Fig. 7 C, right, and Fig. 7 E, red bars)).
The global fitting calculation, using the chemical shift profiles for the nine high-affinity residues gave a KD value of
3.7 5 0.1 mM, whereas the KD value for the 11 low-affinity
residues was 43 5 1 mM. It should be noted that one-to-one
binding was assumed in these calculations. The residues
showed linear spectral changes according to SB-HMG concentration (Fig. 7 A), indicating that the high- and low-affinity residues do not form individual binding sites. If residues
with different affinities form individual binding sites, thechemical-shift-change profiles should become nonunidirectional (33). Therefore, the binding surfaces on AID for
SB-HMG may contain both high- and low-affinity residues.
In fact, the residues of 15N-labeled SB-HMG (Fig. 6 C, redbars) showed a slightly lower affinity for AID (KD ¼ 9.7 5
0.3 mM) than did the high-affinity residues in AID (KD ¼
3.7 5 0.1 mM). These results imply that the high- andBiophysical Journal 104(10) 2222–2234

Hashimoto et al.

low-affinity residues of AID do not interact with SB-HMG
in an individual manner but form multiple complexes in a
dynamic equilibrium (i.e., a dynamic complex) (34).
The titration experiments using 15N-labled pAID showedthat pAID could accommodate up to four SB-HMG molecules (Fig. 7, B and D), which is consistent with the result
from the analysis using 15N-labeled SB-HMG (n ¼ 3.9 5
0.1). Some residues in pAID showed nonunidirectional
spectral changes in the titration of SB-HMG (Fig. 7 B).
These residues are localized in the C-terminal half, which
is rich in phosphoserine and acidic residues (Fig. 7 F, greenbars). Similar to the nonlinear spectral changes in multiple
ligands binding to the protein (27,35), it is likely thatmultiple SB-HMGs bind to the phosphor-Ser-rich regionof pAID. The nonunidirectional spectral changes may therefore be caused by the expansion of the phosphor-Ser-rich region in response to the multiple bindings of SB-HMG.
Taken together, these data allowed us to conclude that nonphosphorylated AID contains binding surfaces that canaccommodate up to two SB-HMG molecules, whereas phosphorylation of AID expands the surface by a factor of 2. The
binding surfaces expanded by phosphorylation should increase the intramolecular encounter probability between
the AID and BID-HMG segments in the dSSRP1 subunit.
Changes of intramolecular interactions between
AID and BID-HMG in response to phosphorylationand dsDNA binding
To explore how AID forms intramolecular interactions with
BID-HMG, we compared the 2D 1H-15N HSQC spectra of
an AB-HMG fragment and the isolated AID (Fig. S3).
The limited solubility (<0.1 mM) and the longer IDR of
AB-HMG hampered assignment of the resonances of the
backbone nuclei. Only a limited number of signals for rather
isolated Ser residues were assigned based on the close proximity (i.e., similar chemical shifts) of the resonances for the
AB-HMG construct to those assigned in the spectra of isolated AID or SB-HMG fragments (Fig. S3).
The significant spectral differences between AB-HMG
and the isolated AID demonstrated that part of the AID
segment binds to the other part of the AB-HMG intramolecularly (Fig. S3 A). The spectral difference between the two
fragments remained in the presence of double-stranded
DNA (dsDNA) (Fig. S3 B). This observation shows that
the AID segment in the AB-HMG fragment still contacts
other parts of the fragment without being released to behave
as an isolated fragment, which would give NMR signals thatmatch more closely to those for the isolated AID fragment.
The spectral difference between the pAB-HMG and theisolated pAID was apparent, indicating that the pAID
segment in the pAB-HMG fragment has intramolecular contacts with other parts of the fragment (Fig. S3 C). The presence of the dsDNA did not completely reverse the spectral
difference (Fig. S3 D), indicating that it does not blockPhosphorylated ID region dynamics

2229

FIGURE 5 NMR spectral characterization of AID fragments. (A) Backbone resonance assignments of 15N-labeled AID in the 2D 1H-15N HSQC spectrum.
(B) Spectral comparison in a region of the 2D 1H-15N HSQC spectra for nonphosphorylated (red) and phosphorylated AID (blue). The backbone amide signals of the phosphorylated Ser shows an upfield shift in both the 1H and 15N dimensions. (C) Comparison of the 15N{1H} heteronuclear NOE profiles between
nonphosphorylated (red) and phosphorylated (blue) AID fragments.
Biophysical Journal 104(10) 2222–2234

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Hashimoto et al.

FIGURE 6 NMR titration experiments using the15
N-labeled SB-HMG fragment. (A) Chemicalshift changes in the titration of nonphosphorylated
AID fragment (AID) to 15N-labeled SB-HMG;
AID/SB-HMG molar ratios were 0.0 (black),
0.3 (red), 0.7 (blue), 1.0 (orange), 1.5 (pink),
2.0 (green), and 3.0 (yellow). (B) The corresponding spectral changes observed for phosphorylated
AID (pAID); pAID/SB-HMG ratios were0.00 (black), 0.07 (red), 0.21 (blue), 0.49 (orange),
0.70 (pink), and 1.04 (green). (C and D)Histograms of the chemical-shift differences in
SB-HMG upon binding with AID (C) and pAID
(D). Chemical-shift differences are plotted against
residue numbers of SB-HMG. Red bars indicatethat the chemical-shift differences are over the
average plus one standard deviation. Short bars incyan and purple along the x axis represent prolines
and the residues for which assignment information
was missing. Chemical-shift changes are shown for
the representative residues in SB-HMG in the titrations with AID (E) and pAID (F). The numericallydetermined dissociation constant, KD, and numberof binding sites, n, for the AID and pAID titration
experiments were KD ¼ 9.7 5 0.3 mM, n ¼ 2.0 5
0.1 and KD ¼ 1.01 5 0.02 mM, n ¼ 3.9 5 0.1,
respectively. Values for KD and n were determinedby global fitting using the residues marked by redbars in C and D (see Supporting Material).

the intramolecular contacts of the pAID segment with partsof the pAB-HMG fragment.
The absence of a complete set of resonance assignments
for the AB-HMG and pAB-HMG fragments allows only alimited analysis of the intramolecular interactions between
the AID and BID-HMG segments. Despite the experimental
limits of this study, the spectral comparison shows that intramolecular interactions between the segments happen irrespective of phosphorylation, and that these interactions
remain even in the presence of dsDNA; however, the modesof binding seem to change.
The intramolecular interactions were monitored using alimited number of assigned HMG signals (Fig. S4). In addition to the limited solubility of the AB-HMG fragment,
intense signals from the long unstructured part in the ABHMG prohibited observation of the signals from the HMG.
Despite the limited residues monitored, these NMR datacollectively indicate that, irrespective of the phosphorylation
Biophysical Journal 104(10) 2222–2234

state, the residues in HMG and BID retain contact with
the AID segment, as suggested by the spectral differencein the HMGs in the AB-HMG and pAB-HMG fragments
relative to the isolated HMG (Fig. S4). In the presence of
dsDNA, the intramolecular contacts of HMG with the AID
segment appear to change according to the phosphorylation
state of the AID segment, as evinced by the spectral comparisons for residues A606, K621, and G553 (Fig. S4).
The phosphorylation-dependent spectral changes for the
HMG induced by the addition of dsDNA suggest that phosphorylation of the AID segment alters its interaction with
the HMG box and thus changes the HMG-mediated binding
to dsDNA.
DISCUSSION AND CONCLUSIONSThe combination of HS-AFM and NMR analyses has shownthat phosphorylation of multiple serine residues in the AID

Phosphorylated ID region dynamics

2231

FIGURE 7 Spectral changes on 15N-labeled AID
and pAID in the titration with the unlabeled
SB-HMG. (A) Spectral changes induced by thetitration of AID; SB-HMG/AID ratios were0.0 (black), 0.1 (green), 0.3 (red), 0.7 (cyan),
1.0 (yellow), 1.5 (purple), 2.0 (blue), and
3.0 (pink). (B) Spectral changes by pAID;
SB-HMG/pAID ratios were 0.0 (black), 0.3 (green),
1.0 (red), 2.0 (cyan), 3.0 (yellow), 5.0 (purple),
7.0 (blue), and 9.0 (pink). (C) Chemical-shiftchange profiles for representative residues in AID,
showing the high (left) and low (right) affinities to
SB-HMG. (D) The 15N chemical-shift changes
observed for representative residues in pAID
according to the titration. Spectral changes determined that the maximal binding number of
SB-HMG to pAID is 4. (E and F) Histograms ofthe chemical-shift differences in AID (E) andpAID (F) upon binding with SB-HMG. Chemicalshift differences are plotted against residue
numbers for AID and pAID. Residues with amean þ 1 SD are colored. Residues in AID withblue bars have higher affinity (KD ¼ 3.7 5
0.1 mM) than those with red bars (KD ¼ 43 5
1 mM). The residues in pAID with green bars
showed kinked titration traces, as in B. Short
down bars in cyan and purple along the x axis indicate prolines and unassigned residues, respectively.
Short down bars in red indicate phosphorylated Ser
residues. Short down orange bars are residues
whose signals were incompletely traced due tosevere signal overlap during the titration.

modulates the intramolecular interactions between the AID
and the DNA-binding element, which consists of BID and
HMG. Our data collectively provide mechanistic insights
into dynamic interactions between AID and the DNA-binding elements in FACT, as follows.
1. AID makes intramolecular contact with the DNA-binding elements. Phosphorylation of AID expands the binding epitope to the DNA-binding element by a factor of 2
when compared with the nonphosphorylated region.
2. The intramolecular contact of the nonphosphorylated
AID with the DNA-binding element should be relatively

weak, as demonstrated by the limited spectral changes inresonances from the isolated AID (Fig. S3). This isconsistent with our HS-AFM observation that the nonphosphorylated FACT retains mostly extended IDRs
(Fig. 3). Thus, the intramolecular contact between thenonphosphorylated AID and the DNA-binding element
remains dynamic and transient.
3. Phosphorylation of AID reinforces its intramolecularinteraction with the DNA-binding element, as shownby the larger spectral changes for the resonances representing the phosphorylated serines in the AID whencompared with the same resonances in the isolated
Biophysical Journal 104(10) 2222–2234

2232

Hashimoto et al.
FIGURE 8 Summary of the mechanism underlying phosphorylation-dependent inhibition withrespect to nucleosomal DNA. The intramolecular
contact of nonphosphorylated AID with BIDHMG is nonspecific and dynamic. This situation
allows BID-HMG to transiently bind with the nucleosomal DNA. In contrast, dense phosphorylation
expands the AID region by elevating repulsive
forces. This results in an increase of the probabilityof encounter between BID-HMG and pAID, thereby
forming a more robust complex between BID-HMG
and pAID. AID and BID are symbolically represented by the red and purple strings, respectively.
Red and purple spheres indicate negative and positive net charges, respectively. The HMG is denotedby the cyan L-shaped structure. Red open circles
labeled P indicate phosphorylation sites.

pAID (Fig. S3). The phosphorylated AID retained intramolecular contacts with the DNA-binding elements inthe presence of DNA (Fig. S3), thus indicating that this
AID element impairs FACT binding to DNA. In the presence of DNA, several resonances representing residues
within the HMG showed chemical-shift changes, despite
the fact that these residues retain contact with phosphorylated AID (Fig. S4). Our HS-AFM observation revealed
dynamic behavior between folding and unfolding of theglobular domain, which is putatively formed by pAID
and BID-HMG (Fig. 3). Taken together, FACT involves
dynamic and transient intramolecular interactions, evenin the phosphorylated state.
The structure of the dSSRP1-HMG was similar to theyeast Nhp6a protein structure isolated from SSRP1 (30).
The DNA-binding surface on the dSSRP1-HMG structure
was similar to the DNA-binding surface in Nhp6A. The
NMR structure of the Nhp6A-DNA complex revealed acharacteristic L-shaped HMG fold, which contacts theminor groove of DNA, whereas an extended N-terminal
BID region interacts with the adjacent major groove (36).
The DNA-binding interface contains numerous conserved
lysine and arginine residues that participate in electrostaticinteractions with the phosphate backbone. Although somehydrophobic stacking/wedge interactions are formed
between a number of Nhp6A residues and DNA bases, theinteractions of Nhp6A with DNA are predominantly electrostatic. Our NMR data indicate that DNA and AID occupy
almost the same electrostatic interaction interfaces on the
HMG and BID regions regardless of the phosphorylation
state of AID.
An apparently similar mechanism was proposed for the
autoinhibition of DNA binding by phosphorylation of a transcription factor, Ets-1 (37,38). The affinity of Ets-1 to DNA
is allosterically regulated by the flexibility of the unstructured serine-rich region and the inhibitory module, whichis adjacent to the ETS domain responsible for DNA binding
Biophysical Journal 104(10) 2222–2234(38). Previous NMR experiments demonstrated that phosphorylation at several sites within the serine-rich region
gradually shifts the equilibrium more to the rigid-inactive
form, which is stabilized by stronger intramolecular interactions with both the inhibitory module and the DNA-binding
domain (39). However, this inhibitory effect of Ets-1 isconsiderably different from our previous results, whichshowed no additive reduction of DNA-binding inhibition
in response to the number of phosphorylated Ser/Thr mutations of dFACT (20). In the FACT mutant containing the
SSRC motif, AID, BID, and HMG (Fig. 1), Ser/Thr-toAla mutations at two phosphorylation sites showed negligible inhibition of DNA binding, whereas mutations atfour and six sites dramatically decreased the inhibitory
effect (as shown in Fig. 3 C of Tsunaka et al. (20)). In otherwords, the inhibitory effect appears to be ultrasensitive, but
not linear.
Our HS-AFM data provide, to our knowledge, a novel
structural view, where the dynamic behavior of the FACTIDR is drastically altered in response to phosphorylation.
The NMR data showed that dense phosphorylation of the
AID region increases binding sites with the DNA-binding
elements containing the HMG. Presumably, the phosphorylation expands the AID region by elevating the repulsive
forces, thus inducing a stronger interaction without protein
folding. Moreover, the increase in binding sites caused by
the phosphorylation of residues can be ascribed to a dynamic equilibrium among multiple binding states but not a
simple two-state equilibrium, thereby blocking DNA binding (Fig. 8). This notion relates to the concept of encounter
probability and differs to the mechanism observed for Ets-1.
This dynamic binding mode is rather similar to that in
the electrostatic model proposed for the interaction between Cdc4 and Sic1 (34,40); the SCF ubiquitin ligase
subunit Cdc4 interacts with the cyclin-dependent kinase inhibitor Sic1 in a dense phosphorylation-dependent manner(34). Multiple phosphorylation in the N-terminal IDRsof Sic1 leads to equilibrium engagement by anPhosphorylated ID region dynamics

interchange between phosphorylation sites (34). The degree of phosphorylation fine-tunes the complex formationof Sic1 with Cdc4 via long-range electrostatic interactions,ensuring ultrasensitivity of the Sic1-Cdc4 interaction
caused by a net charge reversal (40). In agreement withthis ultrasensitive change, our previous data showed thatsimultaneous mutations at four or six phosphorylation sites
drastically enhance the binding ability of FACT to nucleosomal DNA (as shown in Fig. 3 C of Tsunaka et al. (20)).
It is thus likely that the inhibitory mechanism of thephosphorylated FACT is essentially based on the electrostatic model.
Furthermore, a notable feature of this regulation mechanism is that the cooperative action of the tandemly linked
AID and BID regions directs nucleosomal DNA binding
through their interaction with the HMG in FACT. Thesetandemly linked IDRs may enhance the probability ofencounter between the DNA-binding elements and the phosphorylated IDR in the inhibitory mechanism (Fig. 8). In fact,
our HS-AFM analyses indicate that the lifetime of the globular domain in phosphorylated dFACT is much longer thanthat of the nonphosphorylated form.
Many atomic structures of protein complexes provide ageneral view that domain and/or subunit contacts take place
on well-ordered surfaces that complement each other. However, functionally important complexes frequently involve
dynamic, or transient, properties, which are derived from
unstructured IDRs even upon complex formation. It isvery likely that such flexibilities of protein complexes
should change in response to posttranslational modifications, thereby regulating physiological functions. Thus, thecombination of NMR and HS-AFM analysis used in thisstudy potentially can be applied to many other protein complexes in which IDRs play important roles.
SUPPORTING MATERIAL
Four figures, three movies, supporting materials, and methods are available at http://www.biophysj.org/biophysj/supplemental/S0006-3495(13)00433-5.
This work was supported by Grants-in-Aid for Scientific Research on Innovative Areas (Research in a Proposed Research Area) (Ministry of Education, Culture, Sports, Science and Technology KAKENHI Grant No.
21113002, 21121006, and 23107724); a Grant-in-Aid for Japanese Society
for the Promotion of Science Fellows (JSPS KAKENHI Grant No.
07J01196); the Mitsubishi Foundation; Japan Science and Technology(JST) Precursory Research for Embryonic Science and Technology; andJST Development of Systems and Technology for Advanced Measurement
and Analysis.

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