1. Structural Biology and Molecular Biophysics

Crystal structure of the full Swi2/Snf2 remodeler Mot1 in the resting state

  1. Agata Butryn
  2. Stephan Woike  Is a corresponding author
  3. Savera J Shetty
  4. David T Auble
  5. Karl-Peter Hopfner  Is a corresponding author
  1. Ludwig-Maximilians-Universität München, Germany
  2. University of Virginia Health System, United States
  3. Center for Integrated Protein Sciences Munich, Germany
https://doi.org/10.7554/eLife.37774
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Cite this article
  1. Agata Butryn
  2. Stephan Woike
  3. Savera J Shetty
  4. David T Auble
  5. Karl-Peter Hopfner
(2018)
Crystal structure of the full Swi2/Snf2 remodeler Mot1 in the resting state
eLife 7:e37774.
https://doi.org/10.7554/eLife.37774
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Abstract

Swi2/Snf2 ATPases remodel protein:DNA complexes in all of the fundamental chromosome-associated processes. The single-subunit remodeler Mot1 dissociates TATA box-binding protein (TBP):DNA complexes and provides a simple model for obtaining structural insights into the action of Swi2/Snf2 ATPases. Previously we reported how the N-terminal domain of Mot1 binds TBP, NC2 and DNA, but the location of the C-terminal ATPase domain remained unclear (Butryn et al., 2015). Here, we report the crystal structure of the near full-length Mot1 from Chaetomium thermophilum. Our data show that Mot1 adopts a ring like structure with a catalytically inactive resting state of the ATPase. Biochemical analysis suggests that TBP binding switches Mot1 into an ATP hydrolysis-competent conformation. Combined with our previous results, these data significantly improve the structural model for the complete Mot1:TBP:DNA complex and suggest a general mechanism for Mot1 action.

https://doi.org/10.7554/eLife.37774.001

Introduction

Swi2/Snf2 ATPases are members of the NTP-dependent helicase/translocase superfamily 2 (SF2) and are well known as the principal ATP hydrolyzing ‘engines’ of chromatin remodelers that govern processes such as transcription, replication, and DNA repair (Flaus et al., 2006; Narlikar et al., 2013; Hopfner et al., 2012; Becker and Workman, 2013). It is generally assumed, that the Swi2/Snf2 ATPase motor translocates on the minor groove of double-stranded DNA and that this universal core activity generates the force for the large diversity of remodeling reactions catalyzed by Swi2/Snf2 proteins (Saha et al., 2002; Whitehouse et al., 2003; Zofall et al., 2006; Dürr et al., 2005). However, very little is known about how groove tracking activity is converted into the diverse chemo-mechanical remodeling reactions (Hauk and Bowman, 2011; Narlikar et al., 2013; Blossey and Schiessel, 2018). In the absence of substrates, remodelers have been observed in catalytically inactive resting states (Hauk et al., 2010; Xia et al., 2016; Yan et al., 2016), but it is unclear how universal auto-inhibited resting states are. Recent work provides some insight into how Swi2/Snf2 chromatin remodelers interact with and reconfigure nucleosomal substrates (Liu et al., 2017; Farnung et al., 2017; Ayala et al., 2018; Eustermann et al., 2018; Sundaramoorthy et al., 2018). However, the architecture and chemo-mechanical mechanisms of the diverse types of remodeling reactions are not well understood for the great majority of enzymes in this class.

The single subunit remodeler Mot1 (Modifier of transcription 1) is a Swi2/Snf2 enzyme that either activates or represses transcription in a context-dependent manner by dissociating TATA box-binding protein (TBP) and Negative Cofactor 2 (NC2) from promoter DNA (Dasgupta et al., 2002; Zentner and Henikoff, 2013). Mot1 is an essential Swi2/Snf2 enzyme in yeast and the Mot1-TBP-NC2 regulatory axis is highly conserved in eukaryotes. The 140 – 210 kDa Mot1 protein has two functional domains. The N-terminal domain (Mot1NTD) recognizes TBP while the C-terminal domain (Mot1CTD) contains the catalytic Swi2/Snf2 ATPase that binds the DNA upstream from the TATA box (Moyle-Heyrman et al., 2012). The structure of the Mot1NTD has been determined by X-ray crystallography in complex with TBP (Wollmann et al., 2011) and in complex with TBP:NC2:DNA (Butryn et al., 2015). Low-resolution negative stain electron microscopy and chemical crosslinking coupled to mass spectrometry indicated the approximate location of the Mot1CTD near the opening of the Mot1NTD horseshoe (Butryn et al., 2015). However, the orientation of the Swi2/Snf2 domain and consequently the path of DNA remained elusive. As a result, the mechanism of Mot1-mediated dissociation of TBP complexes is still not well understood.

Here, we report the crystal structure of the near full-length Mot1 protein from Chaetomium thermophilium. Our structure reveals the location and orientation of the Swi2/Snf2 domain and, supported by mutagenesis studies, suggests a new type of resting state. Our data allow us also to derive a model for the Mot1 remodeler in complex with TBP and DNA.

Results and discussion

Architecture of CtMot1

We crystallized the near full-length Mot1 protein from Chaetomium thermophilum. The construct covers the entire Mot1NTD and Mot1CTD domains but lacks 50 amino acids from the C-terminus (Figure 1A). We determined the structure of this construct (residues 1–1836, Mot1∆C) harboring a point mutation in the Walker B motif (E1434Q) by Se-SAD to 3.2 Å (Table 1).

Figure 1 with 1 supplement see all
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Structure of the Chaetomium thermophilum Mot1.

(A) Domain organization of CtMot1. The HEAT repeats are in yellow. The latch is in pink, other insertions of CtMot1NTD and the linker are in brown. RecA-like subdomains of CtMot1CTD are in orange (1A) and green (2A). Swi2/Snf2-specific insertions 1B and 2B are in dark blue. Brace and bridge elements are in light blue and red, respectively. The boundary of the crystallization construct (residue 1836) is marked with the dotted line. (B) Cartoon representation of the structure. N- and C-termini are labelled N and C, respectively. HEAT repeats 1, 2, and 16 are labelled HR1, HR2, and HR16, respectively. Missing residues of the latch are represented by the dotted line. (C) Surface representation of CtMot1CTD lobe 1 (orange) and 2 (green). Regions where helicase motifs are located on each lobe are colored in red. (D) Side-by-side comparison of CtMot1CTD (top panel) and SsoRad54 (Dürr et al., 2005) (bottom panel). CtMot1NTD is represented as yellow surface. If not stated otherwise, all panels have color coding as in A.

https://doi.org/10.7554/eLife.37774.002
Table 1
Data collection and refinement statistics for the CtMot1 structure.
https://doi.org/10.7554/eLife.37774.004
Data collection
Space groupP21
Unit cell
a, b, c (Å)93.2, 96.9, 129.7
α, β, γ (°)90.0, 97.6, 90.0
Resolution (Å)48.7 (3.3–3.2)*
Total reflections239071 (10913)
Unique reflections36422 (1888)
Rmeas [%]14.4 (88.7)
I/σI11.8 (2.6)
CC1/20.99 (0.79)
Completeness (%)97.1 (68.8)
Redundancy6.6 (5.8)
Refinement
Resolution (Å)48.7 (3.3–3.2)
No. reflections36410 (2930)
Rwork0.19 (0.42)
Rfree0.24 (0.42)
No. atoms12390
Protein12390
Ligand/ion0
Water0
B factors (Å2)
Protein75
Ligand/ion
Water
R.M.S deviations
Bond lengths (Å)0.002
Bond angles (°)0.463
Ramachandran plot
Favored [%]97
Allowed [%]3
Outliers [%]0

  1. * Values in parentheses are for highest-resolution shell.

The CtMot1 enzyme is a ring-shaped protein (Figure 1B). The CtMot1NTD consists of 16 HEAT repeats (HR) with insertions at four sites and is similar to the much smaller Encephalitozoon cuniculi orthologue (EcMot1NTD) with some notable differences. The helices forming the HEAT repeats are not extended in number but in length and the insertion elements into the HEAT repeats are longer. Thus, genome compaction in E. cuniculi did not alter the overall architecture of Mot1, which appears to be highly conserved in evolution, consistent with its critical function.

The structure reported here is the first to visualize the position and orientation of the Swi2/Snf2 ATPase domain in Mot1. The ATPase domain contains two characteristic lobes connected by a short hinge helix. Each lobe consists of a RecA-like subdomain (1A or 2A) that harbors the SF2-specific sequence motifs responsible for ATP and DNA binding as well as Swi2/Snf2-specific helical subdomains 1B, 2B, and ‘brace’ that emanate from 1A, 2A and the C-terminus of 2A, respectively. Lobe 1 of the CtMot1CTD contacts the C-terminus of CtMot1NTD via HR16, a small insertion within HR12, and a ~ 45 amino acids linker. This highly hydrophobic surface has a total area of 2500 Å2. The tip of subdomain 1B interacts with HR1, thus lobe 1 effectively closes the ring structure of CtMot1. The architectural constraints imposed by a ring explain the conservation of the number of HEAT elements among Mot1 proteins. Lobe 2 binds the cleft between lobe 1 and HR1/2. The ~1900 Å2 interface between lobe 2 and the remainder of CtMot1 is dominated by hydrogen bonds and salt bridges.

In some remodelers, the brace is directly followed by a ‘bridge’ element (Hauk et al., 2010), also referred to as NegC (Clapier and Cairns, 2012) or SnAC (Sen et al., 2011; Xia et al., 2016). While EcMot1 does not possess this element, in CtMot1 it is 64 amino acids long (residues 1822 – 1886). The bridge can act as a positive or negative auto-regulatory element via mechanisms that are not understood (Wang et al., 2014; Xia et al., 2016; Yan et al., 2016; Clapier and Cairns, 2012; Carroll et al., 2014; Sen et al., 2011). The bridge was almost entirely omitted from our crystallization construct and the only included residues (1822 –1836) together with the C-terminal expression tag are not visible in the electron density maps.

In summary, the structure reveals the architecture of the CtMot1 protein. It forms a ring-like structure in which the substrate-interacting HEAT repeat ‘arch’ binds both lobes of the ATPase domain from opposing sites.

Apo CtMot1 adopts an auto-inhibited resting state

In all species tested (H. sapiens, S. cerevisiae, C. thermophilum, E. cuniculi), Mot1’s ATPase is robustly activated by TBP:DNA complexes, but very little if at all by DNA alone (Auble et al., 1997; Adamkewicz et al., 2000; Wollmann et al., 2011; Chicca et al., 1998). Interestingly, some Mot1 species are activated by TBP alone and do not require DNA, although a more robust activation is generally observed in the presence of both DNA and TBP. This suggests that the conformation of the Mot1CTD is structurally coupled to TBP binding to the Mot1NTD and that Mot1 alone is in an inactive state (Adamkewicz et al., 2000; Moyle-Heyrman et al., 2012). Indeed, comparison of CtMot1CTD to other SF2 enzymes shows that lobe 2 is flipped ~180° from an ‘active’ conformation in which the ATPase and DNA-binding motifs would be properly aligned, that is lobe 1’s motifs I-III are properly situated in the ATP-binding cleft, while lobe 2’s motifs IV-VI are situated on the outside and are fully solvent-exposed (Figure 1C). As more Swi2/Snf2 domain structures have become available, it has become evident that many show an auto-inhibited conformation with misaligned lobes 1 and 2 (Figure 1—figure supplement 1) (Dürr et al., 2005; Hauk et al., 2010; Xia et al., 2016; Yan et al., 2016). For example, the DNA binding site of the Saccharomyces cerevisiae Chd1 Swi2/Snf2 domain is directly occluded by the chromodomain, providing a means of specific activation of the enzyme by interaction with a nucleosomal substrate (Hauk et al., 2010; Farnung et al., 2017) (Figure 1—figure supplement 1A). Intriguingly, the ‘resting’ conformation of CtMot1CTD is very similar to the crystallographic conformation of the Sulpholobus solfataricus Rad54 Swi2/Snf2 domain (Dürr et al., 2005) (Figure 1D and Figure 1—figure supplement 1B). The functional relevance of the SsoRad54 Swi2/Snf2 domain conformation remained unclear because the crystallized and functionally analyzed fragment of SsoRad54 comprised only the isolated Swi2/Snf2 domain.

Although the precise orientation of lobe 2 might be additionally determined by the bridge element that is missing in the structure, our structural data suggest that the auto-inhibited resting state of CtMot1 is stabilized by the interactions between subdomain 2A and HR1/2. To test this, we mutated ion pairs (R4-D1720, R45-D1716) and a hydrophobic loop (L1658A/Y1659A) to destabilize the resting state (Figure 2A and B). Basal ATPase activity of the point mutants was greatly increased compared to wild-type CtMot1 (WT) and was not further stimulated by DNA and TBP (Figure 2C and Figure 2—figure supplement 1A). CtMot1∆C did not show increased basal ATPase rates and was not activated by TBP alone. However, its ATPase activity in the presence of DNA-containing complexes exceeded that of the WT enzyme. To find out whether this elevated ATPase activity of the mutants translates into productive disruption of the substrate complexes, we performed remodeling assays. Notably, despite an increase in the ATP hydrolysis rate, the ability of CtMot1∆C to dissociate TBP:DNA complexes was impaired (Figure 2D). Assays performed under less efficient dissociation conditions that allowed the TBP:DNA complexes to persist confirmed that all other tested mutants (L1658A/Y1659A, R4D and D1720R) indeed behaved as the WT (Figure 2—figure supplement 1B and C). This shows that the bridge element acts in response to TBP binding and, similarly to SnAC in Snf2 (Xia et al., 2016), ensures productive coupling of ATP hydrolysis to the remodeling reaction.

Figure 2 with 1 supplement see all
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Analysis of CtMot1 mutants.

(A) View at the interface between RecA2A (green cartoon), HR1/2 (yellow surface), and lobe 1 (orange/blue surface). Residues analyzed in this study are shown as sticks and labelled accordingly. (B) Cartoon model showing the positions of mutations in the Mot1NTD (red spheres on yellow surface) and in lobe 2 (green spheres on green surface). Left: orientation as in the CtMot1 crystal structure. Right: CtMot1 with ATPase modeled as in the S. cerevisiae Chd1:nucleosome complex, that is the ATP hydrolysis-competent conformation (Farnung et al., 2017). (C) ATPase activity of the mutants. Error bars represent standard deviations from three technical replicates. CtMot1WT is labelled as WT, CtMot1ΔC as ΔC. (D) Electrophoretic mobility shift assay showing ATP-dependent dissociation of Mot1:TBP:DNA and TBP:DNA complexes. All CtMot1 constructs form ternary complexes with labelled DNA and TBP (M:T:D). In the presence of ATP and unlabeled competitor DNA (DNA*), wild-type CtMot1 (WT), L1658/Y1659, R4D, and D1720R mutants fully disrupt M:T:D and T:D complexes (lanes 5, 9, 11, and 13, respectively), whereas CtMot1ΔC (ΔC) is less efficient (lane 7).

https://doi.org/10.7554/eLife.37774.005
Figure 2—source data 1

Raw data from the ATPase activity assay used for Figure 2C and Figure 2—figure supplement 1A.

https://doi.org/10.7554/eLife.37774.007
Figure 2—source data 2

Raw data from quantification of electrophoretic mobility shift assay used for Figure 2—figure supplement 1B.

https://doi.org/10.7554/eLife.37774.008

Taken together, our data show that Mot1 adopts a resting state with low catalytic activity by stabilizing lobe 2 of the Swi2/Snf2 domain in an inactive conformation relative to lobe 1. Mobilization of lobe 2 from its auto-inhibited state explains the activation of Mot1’s ATPase by TBP and TBP:DNA complexes.

Model of the Mot1:TBP:NC2:DNA complex

The new structure of the near full-length CtMot1 protein together with prior structures enables us to provide a model for the DNA path in the Mot1-bound protein:DNA complex (Figure 3). The EcMot1NTD:TBP:NC2:DNA complex can be readily superimposed with CtMot1 through the conserved structure of the HEAT repeats. Likewise, superimposing SsoRad54:DNA with CtMot1∆C via lobe 1 visualizes how the CtMot1 ATPase could initially contact duplex DNA since in all Snf2/Swi2 protein:substrate structures contacts between nucleic acid and the RecA1 subdomain are preserved. In the resulting model, localization and orientation of the Swi2/Snf2 domain is in full agreement with prior EM and CX-MS analyses (Butryn et al., 2015) and with biochemical studies showing that Mot1 covers two helical turns upstream from the TATA box (Darst et al., 2001; Sprouse et al., 2006; Moyle-Heyrman et al., 2012). Notably, the superimposed DNA segment bound by the ATPase is an almost direct continuation of the promoter DNA fragment from the EcMot1NTD:TBP:NC2:DNA crystal structure. Assuming the generally proposed directionality of ATP dependent translocation of Swi2/Snf2 motor domains on dsDNA (Zofall et al., 2006; Saha et al., 2002; Whitehouse et al., 2003), the structure of CtMot1 and the specific orientation of lobe 1 now suggests that the Swi2/Snf2 motor translocates ‘towards’ the TATA box and TBP along the nucleic acid scaffold.

Figure 3
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Model of the Mot1:TBP:NC2:DNA complex.

CtMot1ΔC was superimposed onto EcMot1NTD:TBP:NC2:DNA via the HEAT repeats (yellow surface). The path of the upstream DNA was determined by superimposing SsoRad54:DNA onto CtMot1ΔC via lobe 1. The TATA box strand from EcMot1NTD:TBP:NC2:DNA as well as the corresponding strand from SsoRad54:DNA are marked in gray. The non-TATA box strands are in red. Substrate proteins TBP and NC2 are represented as dark and light gray surfaces, respectively. The black arrow represents the direction in which Swi2/Snf2 domain is proposed to translocate along the DNA scaffold. CtMot1CTD is color-coded as in Figure 1.

https://doi.org/10.7554/eLife.37774.009

Our model of the Mot1:TBP:NC2:DNA complex suggests where the Swi2/Snf2 domain of Mot1 might engage with upstream DNA and provides new insight into how ATP hydrolysis-associated events are coupled to dissociation of protein:DNA substrates. Since processive ATP-dependent translocase activity has not been observed in biochemical studies, Mot1 could exploit short-range tracking toward TBP. Given the immediate vicinity of the Swi2/Snf2 domain to TBP, very few translocation steps could lead to the displacement of TBP by steric collision (Darst et al., 2001; Auble and Steggerda, 1999; Butryn et al., 2015). In addition, Mot1 could simply displace TBP from DNA by overwinding or introducing other small distortions into upstream DNA (Moyle-Heyrman et al., 2012; Butryn et al., 2015). Similar effects have been observed for other transcription factors, for which not only binding but also dissociation rates depend on the structure of their recognition sites affected by the presence of other factors bound nearby (Luo et al., 2014; Kim et al., 2013). This allosteric effect can be accounted for by local changes to the major and minor groove width (Kim et al., 2013). Such a scenario is plausible since changes two helical turns upstream from the TBP binding site could have an immediate allosteric effect on severely bent and widened TATA box (Tora and Timmers, 2010).

Interestingly, while Mot1’s ATPase orientation suggests that it ‘pulls’ DNA from TBP and overwinds DNA at the substrate, the reverse architecture is seen for the multisubunit INO80 remodeler: here the motor appears to pump DNA into the nucleosome and to underwind DNA at the substrate (Eustermann et al., 2018). Thus, our results suggest that Swi2/Snf2 proteins can use DNA translocation in different ways to disrupts protein:DNA interfaces.

Materials and methods

Key resources table
Reagent type
(species) or resource		DesignationSource or referenceIdentifiersAdditional
information
Gene
(Chaetomium thermophilum)		CtMot1this paperUniProtKB:
G0S6C0_CHATD		gene cloned from a cDNA library
Cell line
(Escherichia coli)		Rosetta(DE3)NovagenMerck: 70954
Cell line
(Escherichia coli)		B843(DE3)NovagenMerck: 69041
Recombinant
DNA reagent		pETDuet-1NovagenMerck: 71146used to express
full-length CtMot1 (1–1886) and its
point mutants
Recombinant
DNA reagent		pET21bNovagenMerck: 69741used to express
CtMot1 (1–1836)
and CtMot1(1–1836, E1434Q)
Chemical
compound, drug		L(+)-SelenomethionineAcros OrganicsAcros Organics:
259960010		42 mg/mL final
concentration
Chemical
compound, drug		SelenoMethionine
Medium Base plus
Nutrient Mix		Molecular DimensionsMolecular Dimensions:
MD12-501
Chemical
compound, drug		Adenosine 5′-
triphosphate disodium
salt hydrate (ATP)		Sigma-AldrichSigma: A2383-10G
Chemical
compound, drug		β-Nicotinamide adenine
dinucleotide reduced
disodium salt hydrate
(NADH)		Sigma-AldrichSigma: 10107735001
Chemical
compound, drug		Phospho(enol)pyruvic
acid monopotassium
salt (PEP)		PanReac AppliChemAppliChem: A2271
Chemical
compound, drug		Pyruvate kinase/lactic
dehydrogenase enzymes
from rabbit muscle		Sigma-AldrichSigma: P0294
Software,
algorithm		XDSKabsch, 2010,
doi: 10.1107/S0907444909047374
Software,
algorithm		PHENIXAdams et al., 2010,
doi:10.1107/S0907444909052925
Software,
algorithm		CootEmsley et al., 2010,
doi: 10.1107/S0907444910007493
Software,
algorithm		UCSF ChimeraPettersen et al., 2004, doi: 10.1002/jcc.20084http://www.rbvi.ucsf.edu/chimera/
Software,
algorithm		ImageJ 1.51 kSchneider et al., 2012,
doi: 10.1038/nmeth.2089		quantification of electrophoretic shift assay
Software,
algorithm		OriginPro 2015OriginLab, Northampton, MA
Sequence-
based reagent		48 bp dsDNABiomers5′–CAGTACGGCCGGGCGCCCGGCATGGCGGCCTATAAAAGGGGGTGGAAT–3'
Sequence-
based reagent		48 bp 6-FAM labelled dsDNABiomers5′–CAGTACGGCCGGGCGCCCGGCATGGCGGCCTATAAAAGGGGGTGGAAT–3'
Sequence-
based reagent		36 bp dsDNABiomers5′–CGGCCGGGCGCCCGGCATGGCGGCCTATAAAAGGGC–3'

Protein purification

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The sequence of the full-length Mot1 (1 – 1886) was isolated from the Chaetomium thermophilum cDNA library and cloned into pETDuet-1 vector (Novagen, Germany) harboring N-terminal His6 tag followed by TEV cleavage site. CtMot1ΔC(1 – 1836) was cloned into pET21 vector containing PreScission protease cleavage site and C-terminal His6 tag. Both constructs were expressed in Escherichia coli Rosetta(DE3) cells (Novagen) and purified using Ni2+-NTA agarose (QIAGEN, Germany). After proteolytic cleavage of the expression tags, the proteins were further purified using ion-exchange chromatography (HiTrap Q HP, GE Healthcare, Germany) and size exclusion chromatography (HiLoad 16/60 200 pg, GE Healthcare). Proteins were concentrated to ~15 mg/ml in 20 mM Tris pH 7.5, 50 mM NaCl and 15% glycerol and stored at −80°C. Selenomethionine labelling of CtMot1∆C was performed in E. coli B843 (Novagen) using SelenoMethionine Medium Base and Nutrient Mix (Molecular Dimensions, UK) supplemented with L(+)-Selenomethionine (Acros Organics, Germany) at 42 mg/L. Purification of selenium-derivatized protein was performed according to the same protocol as for the native protein.

Crystallization and structure determination

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Crystals of selenomethionine-derivatized CtMot1∆C were grown at 20°C by streak seeding in 0.1 M Tris pH 8.9, 0.2 M ammonium acetate and 13% (w/v) PEG 3350. Plate-like crystals with average dimensions of 700 × 150 × 30 µm appeared after three days and were cryocooled in liquid nitrogen using mother liquor supplemented with butanediol at 25% final concentration.

The data were collected at the European Synchrotron Radiation Facility in Grenoble, France at the peak of Se K-edge at 100K. Images were indexed, integrated, scaled, and merged in space group P21 to 3.2 Å using XDS package (Kabsch, 2010). The initial model was built manually to the experimental electron-density derived from SAD phasing using PHENIX AutoSol wizard (Adams et al., 2010). Alternating cycles of manual building using Coot (Emsley et al., 2010) and refinement with PHENIX yielded the final model (Rwork/Rfree of 19.0/23.8%) covering 87% of all residues.

ATPase assay

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The assays were performed using an NADH-coupled assay as described (Kiianitsa et al., 2003). Reactions were performed using 2 mM phosphoenolpuryvate, 25 U/mL of pyruvate kinase/lactic dehydrogenase mix, 1 mM ATP, and 1 mM NADH at final concentrations. Test samples contained 100 nM dsDNA (5′–CAGTACGGCCGGGCGCCCGGCATGGCGGCCTATAAAAGGGGGTGGAAT–3′ top strand), 100 nM TBP, 100 nM NC2 and 250 nM CtMot1.

Electrophoretic mobility shift assays

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Electrophoretic mobility shifts were essentially performed as described (Darst et al., 2001) with some modifications. In the assay shown in Figure 2D, fluorescently labelled dsDNA (40 nM, 5′–CAGTACGGCCGGGCGCCCGGCATGGCGGCCTATAAAAGGGGGTGGAAT–3′ top strand with 6-FAM label on the 5’ end of the reverse strand) was incubated with TBP (10 nM) and Mot1 (25 nM) for 10 min. Unlabeled dsDNA competitor (800 nM, 5′–CGGCCGGGCGCCCGGCATGGCGGCCTATAAAAGGGC–3’ top strand) was then added directly followed by ATP addition (50 µM) for 10 min. Samples were loaded onto 6% polyacrylamide gels and run at 160 V and 4° for 60 min and imaged using Typhoon FLA 9000 imager. The assays shown in Figure 2—figure supplement 1B and quantified in Figure 2—figure supplement 1C were prepared analogously, but TBP was added at a concentration of 15 nM and ATP was added for 6 min before loading the reactions on the gel.

Accession numbers

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The coordinates and structure factors were deposited in the Protein Data Bank under accession code 6G7E.

Data availability

The coordinates and structure factors are deposited in the Protein Data Bank under accession code 6G7E. All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 2 and Figure 2-figure supplement 1.

The following data sets were generated

References

Article and author information

Author details

  1. Agata Butryn

    1. Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
    2. Gene Center, Ludwig-Maximilians-Universität München, Munich, Germany
    Present address
    Diamond Light Source Limited, Harwell Science and Innovation Campus, Didcot, United Kingdom
    Contribution
    Formal analysis, Investigation, Writing—original draft
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5227-4770

  2. Stephan Woike

    1. Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
    2. Gene Center, Ludwig-Maximilians-Universität München, Munich, Germany
    Contribution
    Investigation
    For correspondence
    woike@genzentrum.lmu.de
    Competing interests
    No competing interests declared

  3. Savera J Shetty

    Department of Biochemistry and Molecular Genetics, University of Virginia Health System, Charlottesville, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared

  4. David T Auble

    Department of Biochemistry and Molecular Genetics, University of Virginia Health System, Charlottesville, United States
    Present address
    Diamond Light Source Limited, Harwell Science and Innovation Campus, Didcot, United Kingdom
    Contribution
    Formal analysis, Writing—review and editing
    Competing interests
    No competing interests declared

  5. Karl-Peter Hopfner

    1. Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
    2. Gene Center, Ludwig-Maximilians-Universität München, Munich, Germany
    3. Center for Integrated Protein Sciences Munich, Munich, Germany
    Contribution
    Conceptualization, Formal analysis, Supervision, Writing—original draft
    For correspondence
    hopfner@genzentrum.lmu.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4528-8357

Funding

National Institutes of Health (GM055763)

  • David T. Auble

European Commission (ERC Advanced Grant ATMMACHINE)

  • Karl-Peter Hopfner

Deutsche Forschungsgemeinschaft (Gottfried Wilhelm Leibniz-Prize)

  • Karl-Peter Hopfner

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank the Max-Planck Crystallization Facility (Martinsried) for crystallization trials and the European Synchrotron Radiation Facility (Grenoble) and the Deutsches Elektronen-Synchrotron (PETRA III, Hamburg) for beamtime and excellent support. AB acknowledges support from the Integrated Analysis of Macromolecular Complexes and Hybrid Methods in Genome Biology (DFG GRK1721) training program.

Copyright

© 2018, Butryn et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Agata Butryn
  2. Stephan Woike
  3. Savera J Shetty
  4. David T Auble
  5. Karl-Peter Hopfner
(2018)
Crystal structure of the full Swi2/Snf2 remodeler Mot1 in the resting state
eLife 7:e37774.
https://doi.org/10.7554/eLife.37774

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https://doi.org/10.7554/eLife.37774

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    1. Structural Biology and Molecular Biophysics

    Structural basis for recognition and remodeling of the TBP:DNA:NC2 complex by Mot1

    Agata Butryn, Jan M Schuller ... Karl-Peter Hopfner
    Research Article Updated

    Swi2/Snf2 ATPases remodel substrates such as nucleosomes and transcription complexes to control a wide range of DNA-associated processes, but detailed structural information on the ATP-dependent remodeling reactions is largely absent. The single subunit remodeler Mot1 (modifier of transcription 1) dissociates TATA box-binding protein (TBP):DNA complexes, offering a useful system to address the structural mechanisms of Swi2/Snf2 ATPases. Here, we report the crystal structure of the N-terminal domain of Mot1 in complex with TBP, DNA, and the transcription regulator negative cofactor 2 (NC2). Our data show that Mot1 reduces DNA:NC2 interactions and unbends DNA as compared to the TBP:DNA:NC2 state, suggesting that Mot1 primes TBP:NC2 displacement in an ATP-independent manner. Electron microscopy and cross-linking data suggest that the Swi2/Snf2 domain of Mot1 associates with the upstream DNA and the histone fold of NC2, thereby revealing parallels to some nucleosome remodelers. This study provides a structural framework for how a Swi2/Snf2 ATPase interacts with its substrate DNA:protein complex.

    1. Structural Biology and Molecular Biophysics

    Unanticipated mechanisms of covalent inhibitor and synthetic ligand cobinding to PPARγ

    Jinsai Shang, Douglas J Kojetin
    Research Advance

    Peroxisome proliferator-activated receptor gamma (PPARγ) is a nuclear receptor transcription factor that regulates gene expression programs in response to ligand binding. Endogenous and synthetic ligands, including covalent antagonist inhibitors GW9662 and T0070907, are thought to compete for the orthosteric pocket in the ligand-binding domain (LBD). However, we previously showed that synthetic PPARγ ligands can cooperatively cobind with and reposition a bound endogenous orthosteric ligand to an alternate site, synergistically regulating PPARγ structure and function (Shang et al., 2018). Here, we reveal the structural mechanism of cobinding between a synthetic covalent antagonist inhibitor with other synthetic ligands. Biochemical and NMR data show that covalent inhibitors weaken—but do not prevent—the binding of other ligands via an allosteric mechanism, rather than direct ligand clashing, by shifting the LBD ensemble toward a transcriptionally repressive conformation, which structurally clashes with orthosteric ligand binding. Crystal structures reveal different cobinding mechanisms including alternate site binding to unexpectedly adopting an orthosteric binding mode by altering the covalent inhibitor binding pose. Our findings highlight the significant flexibility of the PPARγ orthosteric pocket, its ability to accommodate multiple ligands, and demonstrate that GW9662 and T0070907 should not be used as chemical tools to inhibit ligand binding to PPARγ.

    1. Structural Biology and Molecular Biophysics

    Structure of scavenger receptor SCARF1 and its interaction with lipoproteins

    Yuanyuan Wang, Fan Xu ... Yongning He
    Research Article

    SCARF1 (scavenger receptor class F member 1, SREC-1 or SR-F1) is a type I transmembrane protein that recognizes multiple endogenous and exogenous ligands such as modified low-density lipoproteins (LDLs) and is important for maintaining homeostasis and immunity. But the structural information and the mechanisms of ligand recognition of SCARF1 are largely unavailable. Here, we solve the crystal structures of the N-terminal fragments of human SCARF1, which show that SCARF1 forms homodimers and its epidermal growth factor (EGF)-like domains adopt a long-curved conformation. Then, we examine the interactions of SCARF1 with lipoproteins and are able to identify a region on SCARF1 for recognizing modified LDLs. The mutagenesis data show that the positively charged residues in the region are crucial for the interaction of SCARF1 with modified LDLs, which is confirmed by making chimeric molecules of SCARF1 and SCARF2. In addition, teichoic acids, a cell wall polymer expressed on the surface of gram-positive bacteria, are able to inhibit the interactions of modified LDLs with SCARF1, suggesting the ligand binding sites of SCARF1 might be shared for some of its scavenging targets. Overall, these results provide mechanistic insights into SCARF1 and its interactions with the ligands, which are important for understanding its physiological roles in homeostasis and the related diseases.