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. Author manuscript; available in PMC: 2020 Jan 2.
Published in final edited form as: Structure. 2018 Nov 8;27(1):66–77.e5. doi: 10.1016/j.str.2018.10.002

Defining a canonical ligand-binding pocket in the orphan nuclear receptor Nurr1

Ian Mitchelle S de Vera 1,5,#, Paola Munoz-Tello 1,#, Jie Zheng 2, Venkatasubramanian Dharmarajan 2, David P Marciano 2,3, Edna Matta-Camacho 1, Pankaj Kumar Girl 1, Jinsai Shang 1, Travis S Hughes 1,6, Mark Rance 4, Patrick R Griffin 1,2, Douglas J Kojetin 1,2,8,*
PMCID: PMC6318021  NIHMSID: NIHMS1509487  PMID: 30416039

Summary

Nuclear receptor related 1 protein (Nurr1/NR4A2) is an orphan nuclear receptor (NR) that is considered to function without a canonical ligand-binding pocket (LBP). A crystal structure of the Nurrl ligand-binding domain (LBD) revealed no physical space in the conserved region where other NRs with solvent accessible apo-protein LBPs bind synthetic and natural ligands. Using solution NMR spectroscopy, hydrogen/deuterium exchange mass spectrometry, and molecular dynamics simulations, we show that the putative canonical Nurrl LBP is dynamic with high solvent accessibility, exchanges between two or more conformations on the microsecond-to-millisecond timescale, and can expand from the collapsed crystallized conformation to allow binding of unsaturated fatty acids. These findings should stimulate future studies to probe the ligandability and druggability of Nurrl for both endogenous and synthetic ligands, which could lead to new therapeutics for Nurrl-related diseases, including Parkinson’s disease and schizophrenia.

eTOC blurb

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The orphan nuclear receptor Nurrl is thought to lack a canonical ligand-binding pocket for natural ligands. Solution structural analyses of the Nurrl ligand-binding domain (LBD) show that its putative ligand-binding pocket is dynamic, solvent accessible, and expands to bind unsaturated fatty acids.

Introduction

The orphan nuclear receptor (NR) transcription factor Nurr1 (NR4A2) is expressed in the embryonic ventral midbrain and is essential for the development and maintenance of dopaminergic neurons (Zetterstrom et al., 1997). Post-mortem analysis of human brain tissue from neuropathologically verified cases of Parkinson’s disease revealed downregulation of NURR1 expression, indicating a role for Nurr1 in the decreased production of dopamine and degeneration of dopaminergic neurons (Decressac et al., 2013). These observations implicate Nurr1 as a therapeutic target for treating Parkinson’s disease, which is supported by studies showing that synthetic Nurr1 agonists show neuroprotective effects in animal models of Parkinson’s disease (Kim et al., 2015; Kim et al., 2016; Zhang et al., 2012).

Crystal structures have revealed that endogenous/natural and synthetic NR ligands bind to a canonical ligand- binding pocket (LBP) within the core of the ligand-binding domain (LBD). However, crystal structures of the LBDs of Nurr1 and two related NR4A subclass orphan NRs, Nur77 (NR4A1) and the Drosophila ortholog DHR38 (NR4A4), show no apparent ligand-binding cavity in this same physical space (Baker et al., 2003; Flaig et al., 2005; Wang et al., 2003). Instead, their putative canonical LBPs are filled with bulky hydrophobic residues. These NR4A crystal structures, coupled with the observation that the NR4As display high cellular activation in the absence of added exogenous ligand, have led to their classification as ligand-independent transcription factors—a classification shared with other orphan NRs with no apparent crystallized canonical LBP (Gallastegui et al., 2015).

Structural studies have shown that other orphan NRs with previously unidentified endogenous ligands can undergo large conformational changes to bind ligand. A prominent example involves the NR1D subclass known as the REV-ERBs. Although a crystal structure of ligand-free (apo)-REV-ERBβ revealed a collapsed LBP (Woo et al., 2007), a subsequent crystal structure revealed its putative LBP dramatically expands by 600 Å3 to accommodate binding of the porphyrin heme (Gallastegui et al., 2015; Pardee et al., 2009), which was identified as an endogenous REV-ERB ligand (Raghuram et al., 2007; Yin et al., 2007). Using NMR spectroscopy we showed that the ligand-free REV-ERBβ LBβ is dynamic (Matta-Camacho et al., 2014), confirming the crystallography observations that its apo-LBP has the ability to expand. Furthermore, NMR studies have revealed dynamic apo-LBPs for other non-orphan NRs with large crystallized LBPs that bind endogenous ligands, including PPARγ(Hughes et al., 2012; Johnson et al., 2000), RXRα (Lu et al., 2006), and VDR (Singarapu et al., 2011).

Of the few synthetic Nurr1 agonists reported to activate Nurr1 transcription (Dubois et al., 2006; Hintermann et al., 2007), one class of compounds, which includes amodiaquine and chloroquine, was shown to bind the Nurr1 LBD using NMR chemical shift footprinting to map the ligand binding epitope (Kim et al., 2015). Furthermore, although no endogenous ligands are known to regulate Nurr1 activity in vivo, mass spectrometry-based metabolomics in vitro identification studies showed that unsaturated fatty acids present in brain tissue bind to the LBDs of Nur77 (Vinayavekhin and Saghatelian, 2011) and Nurr1 (McFedries, 2014). Using NMR chemical shift footprinting, we showed that one of the most enriched brain unsaturated fatty acids in the metabolomics studies, docosahexaenoic acid (DHA), binds to the Nurr1 LBD (de Vera et al., 2016) with a similar NMR-detected binding epitope as the synthetic Nurr1 agonist amodiaquine (Kim et al., 2015).

A preliminary NMR study of the Nurr1 LBD indicated that residues comprising the putative LBP had shorter T2 relaxation times, suggesting flexibility or dynamics on the μs-ms timescale (Michiels et al., 2010). We were therefore interested in the structural mechanism that could potentially allow Nurr1 to accommodate a bound natural ligand given the apparent collapsed Nurr1 LBP conformation captured by crystallography. Here, using NMR spectroscopy and hydrogen/deuterium exchange coupled to mass spectrometry (HDX-MS), which are sensitive to protein dynamics in solution, we show that the putative Nurr1 LBP is dynamic, solvent accessible, and exchanges between two or more conformations. Using NMR chemical shift footprinting, we show that the binding epitope for several unsaturated fatty acids identified by metabolomics in vitro identification studies to bind Nurr1 (McFedries, 2014) colocalizes with the dynamic putative LBP. Using conventional and accelerated molecular dynamics simulations, we show that the collapsed Nurr1 LBP captured by crystallography can expand to a conformation similar to other fatty acid-bound NRs. Finally, differential HDX-MS reveals that unsaturated fatty acid binding to the Nurr1 LBD causes a conformational change that results in increased solvent accessibility of the LBP.

Results

The putative LBP exchanges between two or more conformations in solution

Solution NMR peak lineshape analysis reports on dynamic processes including binding events and conformational changes (Kleckner and Foster, 2011). When we compared NMR peak intensities and lineshapes of backbone amides and side-chain methyl groups in the Nurr1 LBD (Figure 1A-C), residues within structural regions comprising the putative LBP showed reduced NMR peak intensities and broad lineshapes compared to residues in regions remote from the LBP. These features are indicative of dynamics on the microsecond-to-millisecond (μs-ms) time-scale, such as exchange between two or more structural conformations in solution. To confirm this directly, we performed constant-time Carr-Purcell-Meiboom-Gill relaxation dispersion (CT-CPMG RD) experiments that directly probe chemical exchange (Rex) processes on the ps-ms timescale. Elevated Rex rates were observed for 34 residues in the Nurr1 LBD (Figure 1D), which primarily map to regions that show reduced NMR peak intensities; Rex rates for 10 other residues could not be determined because these residues show very broad lineshapes. Other residues within the Nurr1 LBD also show non-zero Rex rates, which indicates a global conformational change between two or more conformations but for these residues the difference in chemical shift (Δω) between the two states is likely smaller than residues with more elevated rates.

Figure 1. NMR detected ps-ms timescale dynamics within the putative Nurr1 LBP.

Figure 1.

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(A-C) NMR lineshape analysis of the Nurr1 LBD in (A) extracted 1D slices of 2D [1H,15N]-TROSY-HSQC NMR data of select backbone amide groups (broad resonances labeled red), (B) 1D [1H]-NMR data of methyl groups (broad resonances labeled blue), and (C) peak intensities of 3D TROSY-HNCO NMR data (residues with peak intensities < 15 colored red; black line represents a two residue running average). (D) Chemical exchange (Rex) rates measured at 18.8 T (800 MHz). Residues with Rex values > 5.6 s−1 (2 s.d. above the trimmed mean Rex value) are colored purple; residues with peaks too broad to measure are noted with light purple bars. (E) 1D [19F]-NMR data of BTFA-labeled cysteine residues of Nurr1 LBD (broad resonances labeled green). (F) NMR data mapped onto the Nurr1 LBD (PDB 1OVL; chain B) including backbone amide groups with sharp (black) or broad (red) peaks in (A), methyl groups (light blue) with broad peaks in (B), HNCO peak intensities < 15 (light pink) in (C), Rex values > 5.6 s−1 or too broad to measure (purple) in (D), and BTFA-cysteine resonances with broad peaks (light green) in (E); select residues displayed in the other panels are labeled. See also Figure S1.

We also performed 19F-NMR by covalently labeling the five cysteine residues present in the Nurr1 LBD with 3- bromo-1,1,1-trifluoroacetone (BTFA), a small molecule compound containing an NMR-observable trifluoromethyl group (-CF3) that is highly sensitive to environment (Kitevski-LeBlanc and Prosser, 2012). 19F-NMR signals corresponding to BTFA labels attached to C465 and C475, which are located within the putative LBP, show much lower intensity relative to C534 and C566 located outside of the LBP (Figure 1E, Figure S1). The peak pattern for C465 in particular is complex, with several visible peaks with very broad lineshapes. When plotted onto the crystal structure of the apo-Nurr1 LBD (Wang et al., 2003), the NMR data showing complex and/or broad NMR lineshapes and elevated Rex rates all map to the putative LBP (Figure 1F). Combined, these data show that the putative apo-Nurr1 LBP exchanges between two or more conformations in solution on the μs-ms timescale.

The putative LBP has high solvent accessibility

HDX-MS is sensitive to solution-state conformational dynamics, including the detection of protein “breathing motions” and energetically excited conformational states (Konermann et al., 2011). HDX-MS has shown the LBPs of apo NRs have high solvent accessibility (Bruning et al., 2007; Musille et al., 2012; Zhang et al., 2010). The crystal structure of the Nurr1 LBD, which shows no apparent LBP volume (Wang et al., 2003), suggests the putative LBP would likely have no apparent solvent accessibility. However, our NMR data above indicates the putative LBP is dynamic and may exchange among other conformations not captured in the crystal structure. We therefore performed HDX-MS to assess the relative solvent exposure of the putative LBP relative to other solvent exposed structural regions (Figure 2).

Figure 2. Solvent accessibility of Nurr1 LBD probed by HDX-MS.

Figure 2.

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Deuterium uptake curves show that peptides within the putative LBP have high solvent exchange (blue) relative to the low-to-moderate exchange observed for peptides within the core of the Nurr1 LBD (green) and other solvent exposed peptides on the surface of the Nurr1 LBD (red). Data represent the mean and s.d. of three experimental replicates. Analyzed peptide fragments are colored according to structural location and displayed on the Nurr1 LBD (PDB 1OVL; chain B).

Peptides on the surface of the Nurr1 LBD that would be predicted to have a large exposure to solvent, such as helix 4–5 (h4–5) and h9, showed slight to moderate exchange with solvent. This indicates these regions are structurally rigid although they could be expected to have high solvent exposure based on their locations and surface accessibility in the crystal structure. Other peptides in structural regions within the core of the LBD similarly show either negligible exchange with solvent, including h5 and h8 or moderate exchange in the case of h7. In contrast, peptides within the putative LBP region show a large degree of exchange with solvent suggesting structural flexibility and high solvent accessibility. This includes peptides within h2–3 and the connecting loop, h3, h6, h11, and h12 and the preceding loop. These highly solvent accessible peptides colocalize to structural regions that our NMR studies showed have μs-ms timescale dynamics.

Unsaturated fatty acids bind to the dynamic putative LBP

In vitro metabolomics studies identified unsaturated fatty acids present in metabolic extracts from mouse brain that bind to the LBDs of Nurr1 (McFedries, 2014) and Nur77 (Vinayavekhin and Saghatelian, 2011), an orphan NR evolutionarily related to Nurr1. Using NMR structural footprinting analysis, we previously demonstrated that the binding epitope of one of the most enriched ligands identified in the Nur77 metabolomics studies, docosahexaenoic acid (DHA; C22:6), maps to the putative Nurr1 LBP (de Vera et al., 2016). Several other Nurr1-binding unsaturated fatty acids were identified in the in vitro brain metabolite identification studies, including arachidonic acid (AA: C20:4), linoleic acid (LA; C18:2), and oleic acid (OA; C18:1). We used tryptophan fluorescence spectroscopy to validate binding of these unsaturated fatty acids (Figure 3A), which provided Kd values of 50 ± 5 μM (AA), 101 ± 13 μM (LA), and 59 ± 15 μM (OA). We performed NMR structural footprinting to determine if the binding epitopes of these ligands are similar to DHA (Figure 3B,C). NMR peaks with the largest changes in chemical shift (Figure 3D) and decrease in peak intensities (Figure 3E) map to the putative LBP (Figure 3F). The binding epitope for AA, LA, and OA is similar to NMR detected ligand binding epitopes of Nurr1 binding to DHA (de Vera et al., 2016) and the antimalarial drug amodiaquine (Kim et al., 2015) (Figure S2) within the putative LBP where the N-terminus of h3 meets the h6–7 loop and N-terminus of h11, as well as where h3 and h11 meet h5 and h12, although amodiaquine binding does not affect the conformation of h12 to the same degree as the unsaturated fatty acids. Furthermore, the Nurr1 ligand binding epitopes are similar to the NMR-detected ligand-binding epitopes of PPARγ (Hughes et al., 2012; Hughes et al., 2016; Johnson et al., 2000), RXRα (Kojetin et al., 2015; Lu et al., 2006; Lu et al., 2009), REV- ERBβ (Matta-Camacho et al., 2014), and VDR (Singarapu et al., 2011). Taken together with our dynamic analyses above, these data indicate the putative LBP can expand from the collapsed crystallized conformation to bind ligand.

Figure 3. NMR footprinting maps the binding epitope of unsaturated fatty acids to the putative LBP.

Figure 3.

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(A) Nurr1 tryptophan fluorescence binding assay for unsaturated fatty acids. (B) 2D [1H,15N]-TROSY-HSQC and (C) 1D 1H (methyl region shown) NMR spectra of Nurr1 LBD in the absence or presence of 1.5 molar equivalent of the indicated unsaturated fatty acids with representative residues labeled. (D,E) Quantitation of (D) chemical shift perturbations and (E) peak intensities in the 2D NMR data. Dotted lines and colored circles denote residues with perturbations or intensity changes greater than 2 s.d. from the mean trimmed value; black circles denote residues with NMR peaks that disappeared into the noise due to very broad lineshapes. (F) Residues affected in the NMR data (C-E) displayed on the Nurr1 LBD (PDB 1OVL; chain b). See also Figure S2.

Crystallography likely captured a collapsed LBP

Atomic displacement factors, also called B-factors or temperature factors, report on thermal motion or vibration and static disorder in structural models derived from x-ray crystallography data. High B-factor values are typically observed in the LBPs of NRs known to bind natural or endogenous ligands such as PPARγ (Nagy and Schwabe, 2004), which is supported by dynamic data derived from NMR and HDX-MS studies (Bruning et al., 2007; Hughes et al., 2012; Johnson et al., 2000). In the six Nurr1 LBD molecules within the crystallography asymmetric unit (Wang et al., 2003), higher than average B-factor values (Figure 4A) are observed for residues comprising the putative ligand entry-exit surface formed at the base of the LBD where h3 and h11 meet with the h6–7 loop, as well as the h2–3 loop (Edman et al., 2015). Unfortunately, structure factors were not deposited for the Nurr1 LBD crystal structure, which limits further interpretation of the electron density of this structure. However, electron density for the h2–3 loop within the region of the putative LBP was apparently poor or absent because this loop was not modeled in four of six molecules in the asymmetric unit, which could imply flexibility within this region. To probe this directly, we performed steady-state [1H,15N]-heteronuclear nuclear Overhauser effect (hnNOE) NMR analysis of the apo-Nurr1 LBD (Figure 4B). The hnNOE NMR experiment reports on flexibility on the ps-ns timescale motions; residues with hnNOE values near 1 are highly restricted in motion, and low hnNOE values indicate significant disorder (Kleckner and Foster, 2011). The hnNOE analysis revealed large amplitude mobility or significant flexibility (hnNOE values < 0.65) in the h2–3 loop and other solvent exposed loop regions (Figure 4C). This suggests that the conformation of the h2–3 loop captured by crystallography, which “caps” the entry to the entry-exit site (Edman et al., 2015) of the putative LBP, is likely influenced by crystal contacts with neighboring symmetry related molecules (Figure S3).

Figure 4. Crystallographic B-factors confirm the putative LBP entrance is dynamic.

Figure 4.

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(A) B-factor values colored from blue (low) to red (high) in chains A-F of the Nurr1 LBD crystal structure (PDB 1OVL); hHelical regions are labeled (e.g., h12 = h12) and the LBP entrance surface is marked (green dotted circle). (B) Nurr1 LBD steady- state [1H,15N]-heteronuclear nuclear Overhauser effect (hnNOE) values measured at 18.8 T. Data represent the mean and s.d. of two individual measurements. (C) Residues with hnNOE values < 0.65 in (red spheres) indicate structural regions with disorder or flexibility on the ps-ns timescale, which maps to the h1-h3/β-sheet surface and other regions displayed on Nurr1 LBD (PDB 1OVL; chain B); unassigned residues are colored dark gray. See also Figure S3.

The putative LBP can expand to a conformation similar to other fatty acid-bound NRs

We superimposed the apo-Nurr1 LBD crystal structure (Wang et al., 2003) with several crystal structures of NR LBDs bound to fatty acids, including PPARγ (Itoh et al., 2008), RXRα (Egea et al., 2002; Xu et al., 2004), and HNF4α (Duda et al., 2004; Rha et al., 2009; Wisely et al., 2002) to determine the structural rearrangements that would need to occur in order for the collapsed crystallized apo-Nurrl LBP conformation to expand (Figure 5A, Figure S4). The conformation of the loop connecting h6 and h7 in the apo-Nurr1 LBD, which is tucked into the putative LBP in close proximity to h3, stands out among the other fatty acid-bound NR LBD structures, which show an open conformation where the h6–7 loop is further away from h3. Notably, the h6–7 loop region is a critical structural mediator of the ligand entry-exit site within the NR LBD (Edman et al., 2015).

Figure 5. The putative Nurr1 LBP expands in molecular dynamics simulations to conformations similar to other ligand-binding NRs.

Figure 5.

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(A) Structural superposition of the LBDs of apo-Nurr1 (PDB 1OVL) with DHA-bound RXRα (PDB 1MV9) and DHA-bound PPARγ (PDB 2VV0). C atoms of H402 (h3) and G478 (loop 6–7) are shown as orange spheres; the distance between this atom pair was monitored in the simulations. (B-D) Conventional MD simulations of apo-Nurr1 reveals that (B) loop 6–7 moves away from h3 (red arrow) during the simulation resulting in an open LBP conformation (purple vs. orange; vs. PDB 1OVL in black), as well as a change in (C) secondary structure from helix to turn and (D) backbone dihedral angles. Starting conformations are noted by dotted lines, and running averages of the MD data are shown as solid colored lines. (E-H) Accelerated MD simulations show that the Nurr1 LBP entry-exit site, which (E) crystallized in a closed conformation compared to (F) the open accessible LBP conformation in PPARγ, (G,H) can undergo a large shift in the location of the h2/h2–3 loop to an open conformation that is similar to PPARγ. Dotted lines note the locations of the h2/h2–3 loop regions.See also Figures S4 and S5.

The superposition and NMR analyses indicate the dynamic h6–7 loop in the apo-Nurrl LBD would need to change from the closed crystallized conformation to an open conformation similar to crystal structures of fatty acid- bound NR LBDs to allow the LBP to expand and bind ligand. To explore this possibility, we performed conventional molecular dynamics (MD) simulations, including 9 independent simulation runs ranging independently from 4.5–15 μs in length. In 8 of the 9 simulations, the h6–7 loop moved and extended away from the crystallized starting conformation towards the open conformation observed in the fatty acid-bound nuclear LBD crystal structures (Figure 5B, Figure S5). This conformational change is associated with a change in secondary structure from a helix in the starting crystallized conformation to a turn for residues within the h6–7 loop region (Figure 5C,D). This is consistent with the other fatty acid-bound NR LBD crystal structures where the h6–7 loop adopts a turn type of secondary structure.

The observation that the h6–7 loop changes to and remains in an open conformation in 8 out of 9 simulations indicates that the crystallization conditions may have selected for a collapsed or closed LBP. Furthermore, the crystallized conformation of the apo-Nurrl h2/h2–3 loop surface is unique compared to most NR LBD crystal structures, as it tucks against h6 forming a “clamp” over the LBP entry-exit surface and would prevent ligand access to the LBP (Figure 5E). In contrast, for example, the PPARγ h2/h2–3 loop surface, which is also called the Ω-loop, adopts an open conformation allowing ligands to access the LBP (Figure 5F). As mentioned previously, structure factors were not deposited for the apo-Nurr1 crystal structure (Wang et al., 2003). Although the h2/h2–3 loop surface was modeled into two of the six chains in the apo-Nurr1 LBD crystal structure, the B-factor values for this region, which are the only assessment to the structural quality of the loop conformation, are relatively high. Notably, our NMR data show that the LBP and the entry-exit surface are dynamic and exchange between two or more conformations on the μs-ms timescale.

Conformational changes on the μs-ms timescale are not realistically accessible by conventional MD. We therefore turned to accelerated molecular dynamics (aMD), which allow sampling of conformational changes that occur on the ms timescale and exploration of conformations beyond the energy basin localized around crystallized conformations, which can remain trapped in conventional MD (Pierce et al., 2012). The aMD simulations revealed an open h2/h2–3 loop surface conformation (Figure 5G), representing a large conformational change compared to the crystallized closed conformation (Figure 5H) similar in location to the PPARγ Ω-loop (Figure 5F). This conformational change increases the solvent accessibility of the surface where h3 meets the h6–7 loop, and is consistent with our NMR footprinting data showing the binding epitope of unsaturated fatty acids to the same region. These data, which complement the solution NMR and HDX-MS dynamical studies, further indicate that the crystallized Nurr1 LBP conformation is collapsed, but can change to an open and solvent-accessible conformation similar to other ligand-binding NRs.

A conformational change after ligand binding promotes a solvent accessible ligand bound state

To further explore the binding mechanism of unsaturated fatty acids to Nurr1, we performed differential HDX- MS on the Nurrl LBD ± AA, the unsaturated fatty acid with the highest binding affinity for Nurrl (Figure 3A). AA binding caused enhanced deuterium exchange for peptides within several regions of the Nurr1 LBD (Figure 6A, Figure S6A). The largest increase in deuterium exchange occurred for peptides encompassing the C-terminal region of h 11, the h 11 —12 loop, and N-terminal region of h12. Other regions showing an increase deuterium exchange include the h6–7 loop, the N-terminal and middle of h11, and hi. Inspection of the deuterium uptake curves (Figure 6B) reveals that the h11 —12 loop region showed differences in HDX in the earliest exchange time points. The nearby h6–7 loop, which along with the h11–12 loop forms the ligand entry/exit site, showed differences in HDX in the intermediate time points. Finally, peptides in the middle and N-terminal regions of h11 showed differences in HDX in the longest time points.

Figure 6. HDX-MS reveals increased deuterium uptake for AA-bound Nurr1 LBD.

Figure 6.

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(A) Differential HDX-MS data displayed on Nurr1 LBD (PDB 1OVL; chain B). (B) Deuterium uptake curves for peptides within or near the LBP. See also Figures S6.

Taken together, the deuterium uptake curves indicate that AA binding induces a more solvent accessible LBP whereby the residues near the h11—12 loop and the h6–7 loop show early and intermediate timescale HDX effects, respectively. Regions of h11 that are further away from this surface show increased deuterium uptake but at later timescales, indicating that it takes more time for deuterium to reach this region. Residues in h1 show increased deuterium uptake at all time points analyzed, indicating an allosteric conformational change upon AA binding that may loosen the association of h1 with the core of the ligand binding domain. Notably, in contrast to AA binding to Nurr1 LBD, HDX-MS of amodiaquine binding to the Nurr1 LBD revealed no change in deuterium uptake for any peptides (Figure S6B) despite observing binding to the LBP by NMR (Figure S2).

Most HDX-MS studies have reported protection from deuterium exchange in proteins upon binding ligand, which typically occurs due to hydrogen bond formation between the ligand and protein. However, enhanced deuterium exchange upon ligand binding can occur if a protein conformational change must occur from a ground (lower- energy) state to an excited (higher energy) state to accommodate the bound ligand (Konermann et al., 2014; Offenbacher et al., 2018). These observations, when considered with our differential HDX-MS analysis, indicates that the Nurr1 LBD undergoes a conformational change after ligand binding to facilitate the bound AA ligand. This is supported by our previous steady-state tryptophan fluorescence studies (de Vera et al., 2016). DHA binding caused a right (red) shift of the fluorescence spectrum indicating increased solvent exposure of tryptophan residues, one of which (W482) is located in the LBP in the h6–7 loop region that shows very broad lines in the apo-Nurr1 NMR data and is affected by ligand binding by NMR and HDX-MS. Notably, ligand binding can also result in no significant change in deuterium exchange if the binding event does not affect the energy landscape of certain regions of, or throughout, the protein (Konermann et al., 2011). Similarly, no change in HDX was observed when decanoic acid binds to the PPARγ LBD (Malapaka et al., 2012). Thus, amodiaquine binding to the Nurr1 LBD does not appear to significantly alter the energy landscape of the LBP or overall structure relative to AA binding.

Discussion

Crystallography has revealed that NR LBPs can vary in size from essentially no LBP volume (e.g., the NR4As) up to 1,600 Å3 in volume (Gallastegui et al., 2015). Although the LBPs of NRs are understood to be conformationally flexible (Nagy and Schwabe, 2004) and adaptable to expand beyond previously crystallized ligand-bound conformations (Molnar et al., 2006; Suino-Powell et al., 2008; Togashi et al., 2005), the absence of LBP volumes in crystal structures of Nurr1 and a small cohort of other orphan NRs including other NR4As, COUP-TFII, DAX, PNR, SHP, and TR4 has led to the conclusion that these receptors lack canonical LBPs and only have noncanonical lig-and-independent regulatory mechanisms (Gallastegui et al., 2015).

In contrast to these structural observations from static crystal structures, we employed structural methods capable of detecting protein motions/dynamics in solution. Our data show that putative Nurr1 LBP dynamically exchanges between two or more conformations on the μs-ms timescale and has high solvent accessibility Our NMR- detected fatty acid LBP binding epitope is similar to the NMR-detected binding epitope for the synthetic ligand amodiaquine (Kim et al., 2015), indicating that natural and synthetic ligands bind to the same canonical LBP in the Nurr1 LBD. Using molecular dynamics simulations, we found that regions of the Nurr1 LBD that in the crystallized conformation would prevent ligand access to the LBP can physically change to a conformation similar to other lig-and-bound NRs, which could allow a ligand to access the Nurr1 LBP. It was previously posited that a ligand binding event could potentially induce a conformational change that would open up a LBP for orphan NRs lacking a canonical crystallized LBP (Xu and Li, 2008). Our HDX data shows that AA binding increased the solvent exposure of the Nurr1 LBP, indicating that unsaturated fatty acids binding induces a conformational change and expands the LBP, perhaps after an initial encounter complex ligand binding event.

Our NMR dynamical analyses revealed the putative LBP to be dynamic on the μs-ms timescale, in particular for the h6–7 loop, which had very broad linewidths. This region is implicated as a critical regulatory element in the lig-and binding pathway of NRs (Edman et al., 2015). The h2–3 loop is also implicated in the ligand binding pathway of NRs (Genest et al., 2008; Martinez et al., 2005). Although this region in some chains of the crystal structure was modeled as a helix that forms a lid or cap over the LBP entry site, NMR chemical shift values for residues within this region suggests a lack of secondary structure. Taken together with our hnNOE NMR analysis, this indicates crystal contacts may have stabilized the dynamic h2–3 loop surface into a conformation that blocks access to the LBP. However, our molecular dynamics simulations showed that the crystallized Nurr1 h2–3 loop and h6–7 loop regions can adopt conformations similar to other fatty acid-binding NRs, which in principle could allow ligand access to the Nurr1 LBP.

Conformational adaption and expansion of LBPs after ligand binding may be a general theme for NRs. In addition to the studies mentioned above on REV-ERBβ, crystallography studies on estrogen-related receptor (ERR) also suggest considerable LBP adaptability, as the LBP volumes in ERR crystal structures span 40–1020 Å3 (Gallastegui et al., 2015). In the case of ERRα, which showed a 100 Å3 apo-LBP (Kallen et al., 2004), an expansion would be needed in order to accommodate binding of the recently discovered endogenous ERRα ligand, cholesterol (Wei et al., 2016), which has a molecular volume of ~440 Å3. In another dramatic example, crystal structures revealed that an analog of glucocorticoid receptor (GR) ligand dexamethasone, where the conserved 3-ketone moiety was replaced with a phenylpyrazole group, effectively doubled the LBP volume, which expanded from 540 Å3 in the dexamethasone-bound structure to 1,070 Å3 (Suino-Powell et al., 2008). Expansion of LBPs has also been observed in a Ras GTPase, where an NMR fragment screen identified hits that bind KRas and crystal structures of several of the hits revealed that larger fragments could expand a surface LBP binding site (Maurer et al., 2012).

In vitro metabolomics studies identified endogenous ligands for PPARγ (Kim et al., 2011) and ERRα (Wei et al., 2016) that regulate cellular activities for these receptors. Although unsaturated fatty acids bind to the Nurr1 LBD they do not cause large transcriptional effects in cell-based Nurr1 reporter assay (de Vera et al., 2016; McFedries, 2014). It is possible that unsaturated fatty acids could be exchangeable structural ligands, or other putative natural ligands present in cells could mask the activity of exogenously added lipids. Furthermore, the question remains as to what defines the constitutively active conformation of Nurr1 in cells, and do fatty acids influence the active conformation and coactivator recruitment? Nurr1 is not known to interact with most common NR coactivators (Wang et al., 2003) due in part to an inverted “charge clamp” AF-2 surface (Savkur and Burris, 2004). There could be non- traditional cell-type specific coregulators that activate Nurr1, the binding of which could in principle be regulated in a ligand-dependent manner given our findings here. Although future work is warranted to explore these concepts, our work here shows that natural ligands can indeed bind to the canonical LBP in the Nurr1 LBD. This finding should inspire future work to seek out endogenous ligands that could regulate the physiological activity of Nurr1 and stimulate the development of synthetic Nurr1 ligands.

STAR Methods

CONTACT FOR REAGENT AND RESOURCE SHARING

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact DJK (dkojetin@scripps.edu).

EXPERIMENTAL MODEL AND SUBJECT DETAILS

Escherichia coli BL21(DE3) cells (Invitrogen) were used to express protein that was purified for the structural studies, which were cultured in either autoinduction media or M9 media supplemented with 15NH4Cl (Cambridge Isotope Labs, Inc.).

METHOD DETAILS

Reagents and ligands

Reagents were obtained from Fisher Scientific unless otherwise indicated. TCEP was obtained from Gold Biotechnology. Arachidonic acid, linoleic acid, and oleic acid, obtained from Sigma, and amodiaquine, obtained from Toronto Research Chemicals, were dissolved in ethanol or ethanol-d6 for NMR experiments.

Protein expression and purification

Nurrl LBD (NR4A2; residues 352–598) protein was expressed in Escherichia coli BL21(DE3) cells (Invitrogen) using a pET-46 as a tobacco etch virus (TEV) protease-cleavable N-terminal His-Tag fusion protein and purified as previously described (de Vera et al., 2016) in either autoinduction media or M9 media supplemented with 15NH4Cl (Cambridge Isotope Labs, Inc.). Protein in wash buffer (50 mM Tris pH 7.4, 500 mM NaCl, 7.5 mM imidazole, 5 mM TCEP) was eluted against a 500 mM imidazole gradient through a Ni-NTA column, subsequently incubated with TEV protease at 4 °C overnight to cleave the hexahistidine tag, and loaded anew onto the Ni-NTA column. The flow through containing purified protein was collected, buffer-exchanged into NMR buffer (20 mM KPO4 pH 7.4, 50 mM KCl, 5 mM TCEP and 0.5 mM EDTA), and stored at −80 °C.

NMR spectroscopy

Protein NMR data were acquired on a Bruker 700 MHz NMR spectrometer equipped with a QCI cryoprobe at 298 K, unless otherwise noted. We previously validated (de Vera et al., 2016) the reported Nurrl LBD NMR chemical shift assignments (Michiels et al., 2010) using 2D [1H,15N]-TROSY; 3D TROSY-based HNCO, HNCA, HN(CO)CA, HN(CA)CB, and HN(COCA)CB); and 3D 15N-NOESY-HSQC. For ligand-bound studies, 2D [1H,15N]- TROSY-HSQC data were acquired at 298 K using 100 μM 15N-labeled Nurr1 LBD in the absence or presence of 1.5:1 molar equivalence of unsaturated fatty acid or 5:1 molar equivalence of amodiaquine in NMR buffer containing 10% D2O (Sigma). 15N-CT-CPMG RD and hnNOE experiments were acquired on a Bruker 800 MHz NMR spectrometer equipped with a TCI cryoprobe at 298 K. 19F NMR data were acquired on a Bruker 400 MHz NMR spectrometer equipped with a BBFO probe at 298 K using 100 μM Nurr1 LBD and Cys-to-Ala mutants (C465A, C475A, C505A, C534A, and C566A) incubated with 8 mM of 3-bromo-1,1,1,-trifluoroacetone (BTFA; Sigma) for 12 h at 4 °C Excess BTFA was removed by buffer exchange befo re concentrating the BTFA-labeled protein to ~350 μM in NMR buffer containing 10% D2O. KF in a coaxial glass insert was used as chemical shift reference. All data were processed with Bruker Topspin 3.0 and analyzed with NMRViewJ (OneMoon Scientific, Inc). CT-CPMG RD data were analyzed by subtracting effective relaxation rates (R2eff) at CPMG field strengths of 25 and 1000 Hz. NMR chemical shift changes (perturbations) were calculated using a scaling factor accounting for the difference in the gyromagnetic ratios of 1H and 15N nuclei (Williamson, 2013).

Hydrogen/deuterium exchange mass spectrometry (HDX-MS)

Solution-phase amide HDX experiments were carried out with a fully automated system described previously (Chalmers et al., 2006) with slight modifications. Five μl of 10 μM Nurr1 LBD in the absence or presence of 10:1 molar equivalence of ligand was mixed with 20 μL of D2O-containing NMR buffer and incubated at 4 °C for a range of time points (0s, 10s, 30s, 60s, 900s or 3,600s). Following exchange, unwanted forward or back exchange was minimized and the protein was denatured with a quench solution (5 M urea, 50 mM TCEP, and 1% v/v TFA) at 1:1 ratio to protein. Samples were then passed through an in-house prepared immobilized pepsin column at 50 μL min- 1(0.1% v/v TFA, 15 °C and the resulting peptides were trapped on a C18 trap column (Hypersil Gold, Thermo Fisher). The bound peptides were then gradient-eluted (5–50% CH3CN w/v and 0.3% w/v formic acid) across a 1 mm × 50 mm C18 HPLC column (Hypersil Gold, Thermo Fisher) for 5 min at 4 °C. The eluted peptides were then analyzed directly using a high resolution Orbitrap mass spectrometer (Q-Exactive, Thermo Fisher). The AA HDX experiment was performed in triplicate, and the amodiaquine experiment was performed once because no change in HDX was observed. To identify peptides, MS/MS experiments were performed with a Q-Exactive Orbitrap mass spectrometer over a 70 min gradient. Product ion spectra were acquired in a data-dependent mode and the five most abundant ions were selected for the product ion analysis. The MS/MS *.raw data files were converted to *.mgf files and then submitted to Mascot (Matrix Science, London, UK) for peptide identification. Peptides with a Mascot score of 20 or greater were included in the peptide set used for HDX detection. The MS/MS Mascot search was also performed against a decoy (reverse) sequence and false positives were ruled out. The MS/MS spectra of all the peptide ions from the Mascot search were further manually inspected and only the unique charged ions with the highest Mascot score were used in estimating the sequence coverage. The intensity weighted average m/z value (centroid) of each peptide isotopic envelope was calculated with the latest version of our in-house developed software, HDX Workbench (Pascal et al., 2012).

Tryptophan fluorescence spectroscopy

Sixteen final ligand concentrations ranging from 0.23 μM to 300 μM were prepared in ethanol or DMSO (Sigma) from 2 μL of ligand stock (30 mM) or vehicle control (ethanol) spiked to 2 wells per concentration in a 96-well black quartz microplate (Hellma) containing 200 μL of 2.5 μM protein or 25 μM L-tryptophan. After an incubation of 30 min, the plate was read at 23–25 °C in a Tecan Safire II or Molecular Devices Spectramax M5e microplate reader with excitation and emission wavelengths set to 280 nm and 335 nm, respectively; or with an emission scan from 300 to 500 nm at 5 nm increments at the same excitation wavelength. Fluorescence units were converted to percentage fluorescence quenching with respect to vehicle controls (i.e., Nurr1 or L-tryptophan with 1% ethanol or DMSO) and normalized to L-tryptophan data and fit to a one-site specific binding equation with or without Hill slope (determined by F-test in GraphPad Prism) to calculate Kd values.

Molecular dynamics simulations

Generation of MD input files and the production MD runs were performed using AMBER 14 (http://ambermd.org). The ff14SB force field parameters were used for the protein molecule, and ionsjc_tip3p parameters were used for ions (Joung and Cheatham, 2008). The apo-Nurr1 LBD crystal structure (PDB 1OVL) was used to generate coordinate and parameter files within tleap (AmberTools14). The overall charge was neutralized using Na+ ions; TIP3P (Jorgensen et al., 1983) water molecules were added to a minimum thickness of 10 Å surrounding the protein in a truncated octahedron box; and K+ and Cl- ions were added to a concentration of 50 mM to be the same as the NMR experiments. The builds were made ready for production runs using a script provided by the Cheatham lab (University of Utah), which involves 4 minimization and 5 equilibration steps in explicit solvent. The equilibration steps are all run at 310K using the Berendsen thermostat to control the temperature. First, minimization is carried out for 1000 steps of steepest descent minimization with strong restraints on the heavy atoms without use of the SHAKE algorithm (Miyamoto and Kollman, 1992). In the second step, molecular dynamics using the NTV ensemble was run for 15 ps with strong restraints on heavy atoms using SHAKE with 1 fs time steps. The third step consists of steepest descent minimization with lower restraints on heavy atoms (no SHAKE), and the fourth and fifth steps repeat the third step with minimal and no restraints on the heavy atoms respectively. The 4 remaining equilibration steps used the NTP ensemble with SHAKE. The first two steps consisted of steadily decreasing restraints on heavy atoms, followed by a third step with steadily decreasing restraints on backbone atoms, for lengths of 5–10 ps and 1 fs time steps. The fourth (final) step removes the restraints entirely and runs 200 ps of MD using the NTP ensemble with 2 fs time steps. The restart file from this final step was then used to start the production MD run. Production MD was performed in the NVT ensemble using the GPU-enabled version pmemd (AM- BER14) (Salomon-Ferrer et al., 2013) with hydrogen mass repartitioning (Hopkins et al., 2015) and a 4 fs time step at 310K. Accelerated MD simulations were also performed using AMBER14 using the same minimized restart file with the following flags in the input file as defined in the AMBER manual: iamd = 3, ethreshd=4062, alphad=188.8, ethreshp=−98754, alphap=5511. Trajectory analysis was performed using CPPTRAJ (Roe and Cheatham, 2013).

QUANTIFICATION AND STATISTICAL ANALYSIS

HDX-MS data statistics

Deuterium uptake for each peptide is calculated as the average of % D for all on-exchange time points and the difference in average %D values between the apo and ligand bound samples is presented as a heat map with a color code given at the bottom of the Supplemental Figures HDX peptide display maps (warm colors for deprotection and cool colors for protection). Peptides are colored by the software automatically to display significant differences, determined either by a >5% difference (less or more protection) in average deuterium uptake between the two states, or by using the results of unpaired t-tests at each time point (p-value < 0.05 for any two time points or a p-value < 0.01 for any single time point). Peptides with non-significant changes between the two states are colored grey. The exchange at the first two residues for any given peptide is not colored. Each peptide bar in the heat map view displays the average D %D values, associated standard deviation, and the charge state. Additionally, overlapping peptides with a similar protection trend covering the same region are used to rule out data ambiguity.

Supplementary Material

1

Highlights.

A previous crystal structure indicated Nurrl lacks a canonical ligand-binding pocket Solution NMR and HDX-MS uncover a dynamic putative pocket NMR exposes a similar binding epitope for fatty acids and a synthetic Nurrl agonist MD simulations and HDX-MS implicates pocket expansion after ligand binding

Acknowledgements

This work was supported in part by National Institutes of Health (NIH) grants R01GM114420 (DJK), F32DK108442 (RB), R00DK103116 (TH); American Heart Association (AHA) fellowship award 16POST27780018 (RB); National Science Foundation (NSF) funding to the Summer Undergraduate Research Fellows (SURF) program at The Scripps Research Institute [Grant 1659594]; and the Academic Year Research Internship for Undergraduates (AYRIU) program at The Scripps Research Institute. A portion of this work was performed at the National High Magnetic Field Laboratory (NHMFL/MagLab), which is supported by National Science Foundation (NSF) Cooperative Agreement No. DMR-1157490 and the State of Florida; we thank Mr. Ashley Blue at the NHMFL for assistance with NMR experiments. NMR data presented herein were collected in part at the City University of New York Advanced Science Research Center (CUNY ASRC) Biomolecular NMR Facility; we thank Dr. James Aramini at the ASRC for assistance with NMR experiments.

Footnotes

DATA AND SOFTWARE AVAILABILITY

Data generated in this manuscript are available upon request. The crystal structure of the Nurr1 LBD was previously deposited (Wang et al., 2003) at the Protein Data Bank (PDB) under accession code 1OVL. The NMR chemical shift assignments for the Nurr1 LBD was previously deposited (Michiels et al., 2010) at the Biological Magnetic Resonance Data Bank under entry number 16541.

Declaration of interests

The authors declare no competing financial interests.

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Supplementary Materials

1
NIHMS1509487-supplement-1.pdf (4MB, pdf)

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