Atomic-level structural information for HSPB1 ACD and CTR regions is available from crystallographic and NMR studies (Baranova et al., 2011; Rajagopal et al., 2015b; Hochberg et al., 2014; Alderson et al., 2017). Given the crucial yet enigmatic role of the disordered NTR in sHSP oligomerization and function, we sought to expand structural studies to include the NTR. Although oligomeric forms of HSPB1 are too large to analyze by traditional solution-state NMR and are too heterogeneous to crystallize, our previously reported HSPB1_dimer is amenable to solution-state NMR approaches (Baughman et al., 2018). A construct in which the CTR is truncated to the same position as the end of our B1-ACD construct (residue 176) is also dimeric in the phosphomimic context (termed ‘NTR-ACD’, Figure 1A). Thus, HSPB1_dimer and its truncated form provide the first opportunity to obtain residue-level information of a sHSP with its NTR in solution. The simplest model for HSPB1_dimer would be a structured ACD dimer with flexible, disordered NTRs and CTRs that behave independently of other domains. Such a species would give rise to an NMR spectrum that would resemble that of the isolated ACD plus resonances that correspond to ‘random coil’ positions. The NMR spectrum of an HSPB1 construct that lacks both its NTR and CTR (‘B1-ACD’, Figure 1A) and forms a well-structured dimer has been assigned (Rajagopal et al., 2015b). Remarkably, few peaks overlap perfectly in overlays of ¹H-¹⁵N HSQC spectra of B1-ACD and NTR-ACD (black versus blue, respectively; Figure 2A), Therefore, the model in which the ACD behaves independently of its flanking domains is inaccurate, demanding investigation of interactions between domains.

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The NMR spectrum of NTR-ACD overlays well with that of full-length HSPB1_dimer, confirming that the CTR lacking its IXI motif does not interact detectably with the rest of the protein in the otherwise phosphomimetic context (Figure 2—figure supplement 1). NTR-ACD was therefore used for subsequent NMR studies to limit spectral overlap of CTR peaks with NTR peaks in the central portion of the HSQC spectrum corresponding to disordered residues. Most peaks corresponding to ACD residues could be identified and assigned from standard heteronuclear triple resonance NMR spectra collected on NTR-ACD (Figure 2—figure supplement 1, assignments deposited in BMRB). Despite the lack of precise overlap between the spectra of NTR-ACD and B1-ACD, the majority of residues in both contexts give rise to similar chemical shifts (¹H, ¹⁵N, and ¹³C). The similarities of the chemical shift ‘fingerprints’ indicate that the ACD structure is retained in the two contexts. Therefore, the widespread ¹⁵NH chemical shift perturbations (CSPs) observed for ACD resonances indicate differences in environment due to proximity of the NTR.

The largest perturbations in ACD peaks are for residues in the β3, β4, and β8 strands and loop L3/4 (Figure 2B). These structural elements compose two grooves on the ACD dimer, known as the β4/β8 groove and the β3/β3 or ‘dimer interface’ groove (Figure 1B). In some cases, resonances appear to be absent in the NTR-ACD spectrum altogether: we were unable to identify peaks for several residues despite having assignments for these in the B1-ACD spectrum (gray squares in Figure 2B). Analysis of 3D-heteronuclear spectra used to assign the NTR-ACD spectrum failed to identify peaks with similar ¹³C_α/¹³C_β chemical shifts for these residues, implying that resonances for these residues are not observed and are likely undergoing intermediate exchange between different conformations and/or chemical environments. As there is no evidence of slow chemical exchange in the context of B1-ACD, the broadening is likely due to dynamics and/or heterogeneity arising from the NTR. Altogether, the large perturbations indicate that the NTR interacts with the β4/β8 grooves at either end of the ACD dimer and the dimer interface at the center of the ACD.

Some ACD residues have multiple peaks in the NTR-ACD spectrum, indicating that they populate different chemical environments that interconvert slowly. The smallest difference in frequencies observed between multiple peaks from a single residue is 2 Hz, indicating a lifetime greater than 500 milliseconds. Notably, no peak doubling is observed in the B1-ACD spectrum. We could assign multiple peaks for four residues: positions 110 and 114 (flanking the β4 strand), position 148 (in loop L7/8, preceding the β8 strand), and residue 123 (in loop L5/6 between β5 and β6+7 strands). These positions are all near regions that exhibit CSPs and/or exchange broadening described above.

To investigate sources of the perturbations in ACD peaks in the NTR-ACD context, we examined NMR spectra of a mixture of ¹⁵N-B1-ACD and unlabeled NTR-ACD. Under reducing conditions (to inhibit disulfide bond formation at the dimer interface), mixed dimers composed of one ¹⁵N-B1-ACD subunit and one NTR-ACD subunit can form. This allowed us to observe perturbations in an ACD due to the NTR of its dimeric binding partner. As shown in Figure 2A, new peaks are observed in the spectrum of the mixture (red spectrum) that align with peaks in the NTR-ACD spectrum (blue spectrum). Peaks that exhibit this behavior nearly all correspond to residues in the β4/β8 groove and its flanking loops, predominantly the same residues whose peaks exhibit the largest CSPs compared to their position in the ACD-only spectrum (Figure 2B). The congruence of certain peaks in the mixed-dimer spectrum with those in the NTR-ACD spectrum provides unambiguous evidence that their resonance positions arise from an interaction involving the NTR of the other subunit within a dimer. The identity of these peaks reveals that a site of interaction is the β4/β8 groove and its flanking loops. Additionally, many other peaks in the mixed-dimer spectrum lose intensity, shift, and/or change shape as compared to the B1-ACD spectrum. Such behavior is indicative of increased heterogeneity caused by multiple processes. Peaks that undergo such changes correspond to residues in the β3 strand and preceding residues and at the end of the β9 strand and start of the CTR (Figure 2—figure supplement 2). Overall, the extent of perturbations observed in this mixing experiment reveals wide scale NTR-ACD contact and establishes that NTR/ACD interactions can occur between subunits within a dimer.

To ask whether specific NTR regions interact with the ACD, we subdivided the NTR sequence into six sub-regions, referred to here as the distal region, aromatic region, conserved motif, tryptophan-rich region, insertion, and boundary region (Figure 1C). We used NMR to test the ability of peptides from each region to bind to B1-ACD. As described below, peptides from the distal, aromatic, conserved motif, and boundary regions all caused distinct changes (either chemical shift perturbations, loss of peak intensity, or both) in the ¹⁵N-HSQC spectrum of B1-ACD, implying specific interactions of these regions with the ACD. No perturbations were observed for the Trp-rich peptide, providing confidence that the effects observed for other peptides are specific.

We also performed paramagnetic relaxation enhancement (PRE) experiments to detect interactions between the NTR and the ACD within the context of the NTR-ACD construct. PRE from a spin label broadens resonances of residues proximal to the label. The sole native cysteine at position 137 was substituted with serine and multiple constructs were created in which a single cysteine residue was introduced at NTR sites to which the spin label MTSL was conjugated. Spin label positions were selected to probe each NTR region, as shown in Figure 1C. Before performing NMR experiments, each Cys mutant was assessed for alterations in secondary structure and oligomeric properties (by CD and SEC, respectively, data not shown). Only variants that retain HSPB1_dimer-like properties were investigated further. ¹⁵N and spin label were incorporated into these species and HSQC spectra of labeled dimers were collected with the active spin label and with the MTSL quenched by ascorbate. Quenched spectra were compared to the NTR-ACD spectrum to confirm that MTSL conjugation did not significantly perturb the constructs. PREs were quantified as ratios of peak intensity in active vs. quenched MTSL spectra. A range of behaviors was observed, depending on the spin label position: 1) strong, discrete effects, 2) smaller but still localized effects, or 3) widespread general effects.

As summarized in Figure 3, the results from the PRE experiments agree well with the peptide-binding studies, confirming that the interactions observed in the peptide-binding studies are recapitulated in the full-length protein. Results from individual peptides and spin-labels are described below and are provided in Figures 4, 5 and Figure 4—figure supplement 1.

Figure 3

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Figure 5

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Addition of the distal peptide to ¹⁵N-B1-ACD caused distinct chemical shift perturbations (CSPs) in the ¹⁵N-HSQC spectrum that map to the β4/β8 groove (Figures 3, 4A and B). Importantly, peaks shift along a trajectory between their positions in the spectra of B1-ACD and NTR-ACD (as indicated by the arrows in Figure 4A). This is strong evidence that the large perturbations observed for these peaks in the NTR-ACD spectrum (relative to B1-ACD) are due to binding of the NTR distal region to the β4/β8 groove. The peptide binding was not saturated in the NMR experiments, thereby producing smaller CSPs relative to the NTR-ACD spectrum. The effective concentration of the peptide when attached to the ACD in its native form will be in the millimolar range, so the effects observed when the peptide is added in trans are relevant to the native state. The results add clarity to the ¹⁵N-B1-ACD/NTR-ACD mixing experiment described above (Figure 2A). The new peaks that appear in positions that correspond to those observed in the NTR-ACD spectrum are due to distal region binding to the β4/β8 groove of the other subunit of the dimer in a ‘domain swap’ relationship. We cannot rule out the possibility of a similar intra-chain interaction, but if it occurs it must be essentially identical to the inter-chain one observed in the mixed dimer.

The spectrum of NTR-ACD with MTSL at position two has strong peak intensity loss in two distinct sequence regions that correspond to loops L7/8 and L4/5, both of which lie near one entrance to the β4/β8 groove (Figures 3 and 5A). Other peaks in the spectrum are largely unaffected by the spin label (i.e., I_para/I_dia ~ 1.0). This remarkably discrete PRE effect from a spin label at the extreme N-terminus of HSPB1 indicates that when the region is near the ACD, it inhabits a highly localized position. Furthermore, the PREs are consistent with only one of the two possible orientations of distal region binding in the groove, namely aligned parallel to the β8 strand and antiparallel to the β4 strand such that position two only contacts residues near the beginning of β8 and the end of β4. This is the opposite orientation from that observed for the CTR IXI motif bound in the β4/β8 groove observed in a crystal structure of HSPB1-ACD (4MJH) (Hochberg et al., 2014). Distal region binding to the β4/β8 groove has been observed in crystals of HSPB6 (Sluchanko et al., 2017) and an HSPB2/3 (Clark et al., 2018) complex, but those sHSPs contain canonical IXI motifs in their NTRs. HSPB1 does not contain such a motif in its distal region; we propose instead that alternating hydrophobic residues in the HSPB1 segment ⁶VPFSLL¹¹ bind, supporting the idea that other hydrophobic amino acids can participate in β4/β8 binding. Our results thus identify a novel interaction and indicate that motifs from both the NTR and CTR of HSPB1 can bind in the β4/β8 groove but are oriented in opposite directions within the groove.

Peptide binding and PRE results indicate that the conserved motif binds at the ACD dimer interface groove (i.e. strands β3 and β6+7 from each subunit), and that the aromatic sub-region spans between the β4/β8 groove and the dimer interface, connecting the distal region to the conserved motif. The aromatic peptide (residues 12–27) caused distinct chemical shift perturbations in peaks corresponding to one side of the ACD, primarily loops L3/4 and L5/6, as well as some residues in β6+7 (Figures 3, 4C and D). This is consistent with the orientation inferred for the distal region, and shows that as the NTR exits the β4/β8 groove, it contacts the side of the ACD via loops L3/4 and L5/6 (Figure 3). The aromatic region contains one of the three HSPB1 phosphorylation sites (Ser15, Figure 1C), so we also tested binding by a version of the aromatic peptide containing phospho-serine at position 15. For the most part, the CSPs are absent or markedly reduced (Figure 4—figure supplement 1A), indicating that phosphorylation of Ser15 serves to release the aromatic region from the ACD. We note that both L3/4 and L5/6 are enriched in negatively-charged amino acids (¹⁰⁰DVNHFAPDE and ¹²⁴HEERQDEHG, respectively), suggesting an electrostatic mechanism by which phosphorylation at Ser15 could regulate HSPB1 structure and activity.

Conserved motif binding at the ACD dimer interface is indicated by intensity loss in peaks of residues in β3, β6+7, and in loop L3/4 upon addition of the conserved peptide to ¹⁵N-B1-ACD (Figures 3 and 4E). A spin label at the aromatic region/conserved motif boundary (position 26) causes subtle peak intensity loss localized to loops L3/4 and L8/9 leading out of the interface groove, as would be expected if the conserved motif is bound in the groove and the aromatic region runs along one side of the ACD between the β4/β8 groove and the dimer interface groove (Figure 5B). Conserved motif binding at the dimer interface groove has been observed in HSPB6 and in HSPB2/3 hetero-tetramer crystals (Clark et al., 2018; Sluchanko et al., 2017). Given the high conservation of this motif and of residues that compose the dimer interface, it is likely that a similar relationship exists in HSPB1.

Although the Trp-rich peptide showed no detectable interactions with B1-ACD (Figure 4F), we were able to obtain some insights regarding its behavior. While attempting to introduce spin labels, we found the Trp-rich region to be highly sensitive to mutagenesis and were unable to probe its interactions through PRE experiments. Cysteine substitutions of S43 and V55 within the NTR-ACD construct yielded species larger than dimers and not amenable to NMR. An S50C NTR-ACD construct was dimeric but conjugation of MTSL at this position produced larger species. While the altered oligomeric propensity rendered these mutants unsuitable for PRE experiments, the ability of changes in this region to override the dimer-promoting mutations implicates the Trp-rich region in maintaining the delicate balance among oligomeric species. This sub-region harbors the disease-associated mutation P39L, which also affects HSPB1 oligomeric properties (see later section).

The insertion region was probed by a spin label at position 65, near the center of the region. Intermediate intensity loss was observed in most B1-ACD peaks, with some stronger effects in β9. The widespread but modest PREs suggest that the insertion region does not make sustained contact with any specific region of the ACD but rather ‘hovers’ over all faces of the ACD (Figure 5C). The region was omitted from peptide binding experiments due to its hydrophobicity and insolubility in aqueous buffer. Nevertheless, we were able to obtain backbone assignments for this region, which gave further insight into its behavior (discussed in next section).

Finally, the boundary region appears to interact at the dimer interface groove, as evidenced both by peptide binding and PRE results (Figures 3, 4G, 5D and E). Addition of the boundary region peptide (residues 74–91) to B1-ACD caused intensity loss and CSPs in peaks corresponding to residues near the dimer interface (Figures 3 and 4G). We were able to incorporate spin labels at two boundary region positions, 83 and 91, which flank the predicted β2 strand (residues 86–88 in PDB 4MJH). The spin label at position 91 (S91-SL) caused distinct intensity loss in peaks from loops L5/6 and L7/8 and from residues at the beginning of β3 and end of β9 (Figures 3 and 5E). The highly localized PREs on one side of the ACD are congruent with residue 91 being near the beginning of the β3 strand. In contrast, the spin label at position 83 caused wide-spread intensity loss in peaks corresponding to the dimer interface and the opposite side of the ACD (Figures 3 and 5D). In this case, the ACD residues that are not affected by the spin-label are more informative: the data clearly show that position 83 does not approach loops L4/5, L7/8, and L9/CTR, all of which are localized on one side of the ACD dimer (the side affected by S91-SL). The fact that spin labels at positions flanking the putative β2 strand hit opposite sides of the ACD provides strong evidence that the boundary region spans the dimer interface groove in an antiparallel direction to strand β3, consistent with formation of a β2 strand. Whether this interaction is mutually exclusive with binding of the conserved motif to the dimer interface or can occur simultaneously cannot be deciphered from these experiments.

A boundary-region peptide that contains phospho-serine residues at positions 78 and 82 caused nearly identical perturbations to the non-phosphorylated peptide (Figure 4—figure supplement 1B), indicating that phosphorylation neither directly disrupts nor enhances the interaction between the boundary region and the dimer interface groove. However, the PRE results from S83-SL indicate that this region approaches residues that are perturbed by the aromatic peptide (Figures 3 and 5D), suggesting that the boundary region phosphorylation sites may be close in space to the S15 phosphorylation site, despite being over 60 residues apart in sequence.

Peaks that are not assigned to ACD residues in the ¹H-¹⁵N HSQC spectrum of NTR-ACD presumably arise mainly from the NTR. Many of these peaks are weak (but reproducible across multiple protein preps) and were not observed in low-sensitivity 3D heteronuclear spectra despite high levels of deuteration and use of high field magnets equipped with cryoprobes. For resonances that were detected in heteronuclear spectra, there was considerable degeneracy in chemical shifts (¹³C_α/¹³C_β), likely due to intrinsic disorder and/or conformational heterogeneity. While these properties hampered unambiguous assignment of NTR peaks, the weak and/or poorly-resolved peaks indicate that a majority of residues in the NTR exist in multiple environments, leading to their peak broadening. There are two clear exceptions: we were able to assign a contiguous stretch of residues that span the insertion and boundary regions, and several residues in the Trp-rich region.

Peaks corresponding to residues 62–79 could be assigned (Figure 6A); these span most of the (non-proline) residues of the insertion region and the start of the boundary region (Figure 1C). All peaks assigned for this stretch have high intensities (similar to CTR, Figure 6C) and chemical shifts consistent with random coil structure (Figure 6A). Only the N-terminal part of the boundary region is assigned, while the latter part of the boundary region that is predicted to form β2 could not be assigned. Notably, residues 64–69 were each assigned to two distinct peak sets, indicating two conformational states that interconvert slowly, despite their both having ‘random coil’ chemical shifts. Based on the observed chemical shift differences and the slow exchange condition, we can estimate an upper limit for the exchange rate of ~25 s⁻¹. The second conformation represents a substantial fraction of the population, as judged from relative peak intensities (>20%). A short contiguous stretch at the C-terminal end of the Trp-rich region (residues 46–49) was assigned, with Ser49 having two peak sets. Thus, both regions exist in at least two distinct conformations and/or environments. The observed behavior may arise from cis and trans conformations of one or more of the numerous proline residues in the insertion and Trp-rich regions. However, ¹³C chemical shifts needed to confirm proline isomerization could not be obtained from the data and we did not pursue mutational analysis to identify the source(s) of heterogeneity due to the abundant prolines in the regions.

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Assignments of NTR HSQC peaks provided information regarding proximities of NTR regions to each other from PREs (Figure 6B). The spin label at insertion region residue 65 yields strong PREs across the assigned NTR peaks, validating their assignments to this region of the sequence. The position two spin label did not yield PREs in assigned NTR residues or in weak unassigned peaks, indicating that the distal region’s locations do not overlap with those of the Trp-rich or insertion regions. A position 26 spin label at the beginning of the conserved region gives modest-to-strong PREs to the Trp-rich and boundary regions but weak PREs to the insertion region. The position 83 spin label gave strong PREs to the boundary and insertion regions, and the position 91 spin label gave moderate PREs to the same regions. Together, this provides evidence for extensive interactions between the sub-regions of the NTR. In particular, the conserved and Trp-rich regions seem to be in close contact with each other, as are the insertion and boundary regions. Only the distal region appears not to make extensive contact with other parts of the NTR. The data support a model in which the NTR exists in a compact state with extensive NTR/ACD and NTR/NTR contacts as well as some residual structure, rather than a more flexible, extended conformation seen for many intrinsically disordered regions.

To investigate the dynamics of assigned NTR residues, we measured transverse relaxation rates (R2) of peaks in the NTR-ACD spectrum (Figure 6D). Residues in the ACD have R2 values of ~40 s⁻¹, consistent with their positions within an ordered region. NTR insertion sub-region residues and the final (unstructured) residues in the ACD have lower R2 values (around 10 s⁻¹), consistent with higher flexibility. The limited number of other assigned NTR resonances have R2 values that range from around 20 s⁻¹ to ~60 s⁻¹, consistent with greater conformational restriction of these NTR regions and/or exchange between multiple states on the millisecond timescale.

We used HDXMS to compare NTR properties in dimeric and oligomeric forms of HSPB1. Backbone amide protons in structured and/or buried regions exchange with deuterons more slowly (i.e., are ‘protected’ from exchange) than those in unstructured or accessible regions. HDXMS can be performed on proteins of any size under a variety of solution conditions, providing an opportunity to obtain peptide-level information on the dynamics of HSPB1 oligomers. Time courses of deuterium exchange from three seconds to one hour were measured at room temperature for C137S-HSPB1 oligomers and C137S-HSPB1_dimer. The sole Cys residue, C137, that resides at the dimer interface was substituted to abrogate the need for reducing agent to inhibit disulfide bond formation across the dimer interface that could confound the analysis. Pepsin digestion yielded peptides across the full length of the protein (peptide statistics are shown in Figure 7—source data 2).

Deuterium uptake levels differed most dramatically across regions of HSPB1 in the earliest (3 s) timepoint, shown in Figure 7 (see Figure 7—source data 1 for full kinetic information). The profile for HSPB1_dimer (middle panel, Figure 7) is congruent with observations and conclusions drawn from NMR presented above. Residues 87–172, which span the ACD, show protection even at the latest time point, which is consistent with its β-sandwich structure. Peptides within the aromatic, insert, and boundary regions in the NTR, along with the far C-terminal region (residues 187–205) of the CTR are all highly deuterated within 3 seconds, consistent with their lacking stable secondary structure. Interestingly, peptides along the distal, conserved, and Trp-rich regions of the NTR showed moderate protection, indicating the presence of some local structure. All peptides covering the distal region displayed a bimodal isotopic mass envelope at the early time points indicating that there are two distinct populations of the distal region: one that exchanges readily with deuterium and one that is protected (see Figure 7—figure supplement 1). The bimodal distributions observed for these peptides may reflect the populations of the NTR that are either bound and sequestered or free and solvent-accessible. A substantial protected population is present through 15 seconds, implying that the lifetime of the bound state is on the order of at least several seconds. All the other peptides outside of the distal region displayed only unimodal spectra in the HSPB1_dimer dataset.

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Figure 7—source data 1

HDXMS profiles for all forms of HSPB1.

The deconvoluted fractions of each population from bimodal distributions are shown in the first two columns for each mutant. The fractional deuterium uptake for each of these populations is shown in the next two columns, followed by a weighted average in the fifth column. All spectra were binomial fit using HX-Express (Guttman et al., 2013).

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Figure 7—source data 2

Statistics for HDXMS experiments on all mutants.

Statistics listed are for C137S forms of the following mutants of HSPB1. Sequence coverage takes into account the lack of observed deuterium uptake in the first two residues of each peptide. Back-exchange is calculated as the % deuterium uptake of the fully deuterated sample divided by the theoretical maximum deuteration for each peptide (89%), excluding prolines and the first two residues.

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The general features of the exchange profile for oligomeric HSPB1 (top panel, Figure 7) are similar to HSPB1_dimer, with the ACD being most protected and the NTR and CTR less protected. As in the dimer, the insertion region, boundary region, and CTR have the highest rate of deuterium uptake, indicating that they remain unstructured and accessible in oligomers. However, the aromatic region shows more protection in oligomers than in HSPB1_dimer, indicating some involvement in the oligomerization. The most striking feature of the oligomeric HSPB1 HDX profile is the large number of bimodal peptides that are observed. The distal region showed similar bimodal spectra as seen with the HSPB1_dimer, except the relative population of the more protected species was diminished (~40% vs. 80%). Beyond the distal peptides, another six peptides have two distinguishable populations in the context of HSPB1 oligomers. Peptides that arise from the conserved, Trp-rich, and boundary regions in the NTR display two populations. The conserved and boundary regions were observed to interact with the ACD in the dimer context from NMR results, so either these interactions are longer-lived within the confines of oligomers, or they may also be directly involved in oligomeric interactions. The presence of a substantial protected population at 4 minutes indicates a lifetime of interaction of several minutes in the oligomeric context. In the CTR, a peptide that contains the IXI motif (mutated to GXG in HSPB1_dimer to inhibit its binding to the β4/β8 groove) has a population (~45%) that is protected from deuterium exchange and one that is completely exchanged. This behavior is consistent with the notion that IXI-containing CTRs exist in both free and β4/β8-bound states.

The populations observed for the bimodal spectra are not sufficiently resolved along multiple timepoints to determine precise rates of slow conformational exchange (‘EX1 kinetics’) or to make quantitative comparisons. Nevertheless, qualitative comparisons at the well-resolved 3 s timepoint can be made between regions or between protein constructs. The weighted average deuteration for each peptide with bimodal exchange in Figure 7 provides a qualitative estimate of the relative populations of ‘bound’ and ‘free’ states. For example, both the distal and IXI-CTR peptides show a substantial proportion (~40–45%) of the region in a protected state. These two regions bind to the same ACD groove, creating a situation where there are twice as many potential β4/β8 groove-binding sequences as there are grooves. The HDX data indicate that enough groove-binding sequences are solvent-protected to imply that the majority of β4/β8 grooves are occupied in HSPB1 oligomers and that, on average, half the distal sub-regions and half the CTR-IXIs are bound.

To get a more complete overall picture of what else is different in the oligomeric form, we compared the deuteration levels for each identical peptide that could be identified between HSPB1_dimer and HSPB1 oligomers, using the weighted average percent deuteration for bimodal peptides. In the bottom panel of Figure 7, peptides that appear above the 0% line are more protected in oligomers and peptides that fall below the 0% lines are more protected in dimers. Overall, the ACD displays similar deuterium uptake regardless of whether it is in an isolated dimer or a large oligomer. This indicates that the extent of deuterium exchange at this timepoint is predominantly dictated by the β-sandwich structure. All peptides arising from the distal region show more protection in HSPB1_dimer, with the weighted deuteration skewed to the more protected state (larger proportion) consistent with a larger fraction of the N-terminus bound in the dimer. This may be due to the CTR mutations introduced to generate the dimer, as the lack of IXI motif in the CTR removes it from competition for the β4/β8 groove. Two other contiguous peptides (33–41 and 42–62) that span conserved and Trp-rich regions show greater protection in the dimer. The overlapping 29–41 peptide is more protected in the oligomer, indicating that protection occurs in the first few residues of the peptide in oligomers, whereas protection occurs in later residues in the dimer, consistent with distinct interactions or structural features. The NMR results identified an interaction between the conserved region and the ACD dimer interface, but provided no evidence for an interaction involving the Trp-rich region and the ACD. According to secondary structure predictions, the Trp-rich region has the highest propensity of any NTR region to adopt secondary structure. Intriguingly, the CD spectrum of HSPB1_dimer, but not HSPB1 oligomer, has an unusual positive peak at 230 nm (Baughman et al., 2018). Similar features have been attributed to exciton couplets that can arise from interactions between Trp rings. Together, the HDX and CD data suggest that the Trp-rich region adopts specific structure in dimeric species that is not populated to a detectable degree in the oligomer. The two regions that are less protected in HSPB1_dimer are the aromatic and boundary regions, which contain the phosphorylation-mimicking mutations. The NMR data indicate that the conserved and boundary regions both interact with the dimer interface groove. The increased accessibility of the aromatic region is consistent with the reduced binding of the phosphorylated form of the aromatic peptide. It is possible that release of the aromatic region, which neighbors the conserved region, is coupled to decreased protection of the boundary region. Formation of a new structural feature involving the Trp-rich region might only be possible when neighboring regions are released from their ACD contacts.

The results presented above indicate that the HSPB1 NTR contains distinct regions that reside in specific locations relative to the well-defined ACD dimer and that, in many cases, make direct contact with the ACD. We sought to combine the information garnered from the peptide-binding, PRE, and HDXMS experiments into a set of structural models. Our goals for this modeling process were two-fold. First, the models aid in visualization of the NTR-ACD interactions described above. Second, the modeling process allows us to determine which combinations of NTR-ACD interactions can generate physically realistic structural models, and thus whether any combination of NTR-ACD interactions may not be physically possible. These structures are intended neither as a complete sampling of HSPB1 conformational space nor as atomic-level models of specific interactions.

We produced a homology model by combining the crystal structure of the HSPB1 ACD (PDB 4MJH) with peptides from the crystal structure of the HSPB2/3 hetero-tetramer (PDB 6F2R) using the PyMOL Molecular Graphics System. The starting model included all of the possible NTR-ACD interactions identified above: two copies of the β2 strand, a distal motif bound in each β4/β8 groove, and a copy of the conserved motif bound in the dimer interface groove. Clashes between the HSPB1 ACD structure and the peptides from the HSPB2/3 crystal structure were eliminated and missing loops were modeled in using PyRosetta. In total, there are four possible ways to connect the fragments in the homology model: either copy of the distal motif in the β4/β8 grooves could be connected to the conserved motif at the dimer interface, and the conserved motif could be connected to either copy of the β2 strand (Figure 8A). In total, we created four ensembles of 100 structural models, which each sampled a different way of connecting the NTR peptide fragments in our initial homology model (Figure 8B, C and Figure 8—figure supplement 1). We found that it was possible to develop realistic structural models that contain all types of NTR-ACD interactions identified above, as well as all possible loop connectivities. The dimer interface groove is able to accommodate two copies of the β2 strand in addition to the conserved motif, and both β4/β8 grooves are able to bind a copy of the distal motif. These peptide fragments can be connected to each other in all four conceivable ways.

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If the conserved motif is connected to the distal motif oriented in the opposite direction, the aromatic region forms a loop along the side of the ACD containing loops 3/4, 5/6, and 8/9 to connect these regions (Figure 8B and C). Given our peptide-binding NMR results for the aromatic region, we believe that this configuration is favored, particularly in the non-phosphorylated state. Nevertheless, it is also physically possible for the aromatic region to connect the conserved motif to the distal region in the opposite groove, in which case it spans across the top of the ACD β-sandwich, contacting the β8, β9, and β3 strands (Figure 8—figure supplement 1). While we do not observe experimental evidence for this interaction, we cannot rule it out, particularly in the phosphorylated state in which the aromatic region has low affinity for the ACD. In either case, the conserved motif can then be connected to either one of the β2 strands, while the distal motif not connected to the bound conserved motif can be connected to the other. These connections consist of ~50–75 residues and contain part of the boundary region, the insertion region, the Trp-rich region, and (for the chain with an unbound conserved motif) the conserved and aromatic regions. Given their length, they can adopt multiple conformations and orientations relative to the ACD. We have omitted these regions from the models shown in Figure 8 and its supplement for clarity and to avoid over-interpretation of the structural aspects of regions for which we have limited experimental data. Surface representations of these models show that the locations of the NTR sub-regions are in good agreement with the ACD surfaces perturbed in the peptide binding and PRE NMR experiments (comparing Figure 3 with Figure 8E).

Overall, the results from the modeling suggest that any combination of the NTR-ACD interactions defined in our study is physically feasible. While we only created models containing the maximum NTR-ACD interactions supported by our experimental data, any of the interacting motifs we have modeled could dissociate from the ACD and adopt a more disordered conformation. The results from our NMR and HDXMS experiments indicate that most of these NTR regions occupy both ACD-bound and ACD-unbound conformations, so it is likely that multiple combinations of NTR/ACD interactions occur in solution. Additionally, the similarity of protected regions in the HDXMS profiles of HSPB1 dimers and oligomers indicate that the interactions depicted in these models also occur within higher-order oligomers. The peptide fragments depicted in our dimeric models could conceivably be connected to other ACD dimers or monomers within an oligomer (Figure 8D). The array of possible interactions within sHSP oligomers is depicted in Figure 8F. Many regions can form intra-chain, intra-dimer, and inter-dimer interactions. The possibility for multiple combinations of interactions and connectivities contributes to the high degree of plasticity and heterogeneity observed for HSPB1 in NMR and HDXMS experiments.

To leverage new insights regarding local structure in the NTR and interactions with the ACD, we sought to assess how reported disease-associated mutations in the HSPB1 NTR affect structure and/or dynamics. We chose two NTR disease mutants whose effects on oligomer size (larger than WT) and chaperone function have been previously characterized: G34R and P39L mutants in the conserved and Trp-rich regions, respectively (Muranova et al., 2015). To identify localized effects of the mutations, we performed peptide binding experiments with mutant peptides and B1-ACD. To detect potential global effects, we performed HDXMS analysis on mutant proteins. Glycine at position 34 is the final conserved residue in the ‘conserved motif’ found among orthologs and paralogs; its mutation to Arg is expected to reduce flexibility of the backbone and/or alter the region’s interactions. At the start of the Trp-rich region, Pro39 is conserved among orthologs but not among paralogs. Notably, Pro39 is two residues before a region that is predicted to have helical propensity (residues 41–46, Figure 1—figure supplement 1). Mutation of Pro39 to leucine might change the structural propensity of this region by increasing flexibility and/or favoring helical formation. As presented below, we found highly divergent local structural effects of the two mutations, despite their close proximity in sequence.

As presented above, the conserved motif peptide (residues 25–37) causes peak broadening in dimer interface groove residues (i.e., β3 and β6+7) in ¹⁵N-B1-ACD spectra. An otherwise identical peptide that contains the G34R substitution yields greatly reduced perturbations (Figure 9B), indicating that the bulky, charged Arg sidechain disrupts the ability of the conserved region to interact with the dimer interface groove. Consistent with the notion that the environment of the mutated conserved region is altered, increased deuterium uptake for the fragment that contains the G34R substitution in otherwise wild-type HSPB1 (oligomer) is observed by HDXMS (Figure 9A, top and bottom panels). The increase in deuterium uptake was sufficiently large (as high as 25% in early time points), that it likely reflects a true increase in local flexibility, rather than a consequence of the G to R substitution altering the intrinsic exchange rate of this peptide (Wales et al., 2016). Unexpectedly, the NTR boundary region also shows enhanced deuterium uptake in G34R-HSPB1 and, strikingly, no longer shows bimodal behavior. All other portions of oligomeric HSPB1 are not significantly changed in their exchange profiles.

Figure 9 with 1 supplement see all

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The non-local effect of the G34R mutation on the boundary region suggested to us that there could be coupling between the two NTR regions, similar to possible coupling between phosphorylation sites in the aromatic and boundary regions mentioned earlier. We took advantage of another identified disease-associated mutation that similarly substitutes arginine for glycine, but in the boundary region (G84R) to test this hypothesis. Intriguingly, HDXMS data on the mutant (Figure 9A) revealed enhanced deuterium uptake in both the boundary region and in the conserved region, confirming that the dispositions of these two non-contiguous NTR regions are inter-dependent. The NMR data indicate that both regions interact with the ACD dimer interface-groove, and our modeling suggests that both interactions can occur simultaneously. While the NMR data do not address whether the binding of one influences the other, the HDXMS data provide some clarity on this question. The observed inter-dependence of protection from deuterium exchange implies that each region enhances the ability of the other to contact the groove and that the conformation that leads to protection in the HDXMS experiment has both a conserved region and a boundary region present in the groove. Whether the two bound sub-regions are on the same HSPB1 polypeptide or different ones within a dimer or oligomer cannot be ascertained from these experiments, but our modeling implies that both cases are likely to occur.

As mentioned above, we did not detect evidence of binding for the Trp-rich region peptide to ¹⁵N-B1-ACD. We obtained similar results for a peptide that contains the P39L substitution, indicating that the mutation does not lead to a gain of binding function (Figure 9C). Nevertheless, introduction of P39L into otherwise wild-type HSPB1 has a marked effect on the HDXMS profile (Figure 9A). The first four NTR sub-regions are substantially more protected from deuteration in mutant oligomers, while the ACD is essentially unaffected. The largest increase in protection is observed in the fragment that contains the mutation (29-41). The comparison to WT is complicated by the addition of an extra amide from the P39L mutation, but the massive decrease in exchange (25%) is consistent with the predicted increased helical propensity in this region and an observed increase in helicity revealed in the CD spectrum (Figure 9—figure supplement 1). Altogether, the observations suggest that the P39L mutation increases local secondary structure and/or promotes NTR-NTR contacts. In further support of this conclusion, the subunit exchange rate for P39L-HSPB1 is three-fold slower than for either wild-type or G34R oligomers (Figure 9—figure supplement 1).

Altogether, the results highlight a variety of alterations that occur in the HSPB1 NTR under situations such as phosphorylation or disease mutation. These alterations can have profound non-local effects on other sub-regions of the NTR, leading to global changes in HSPB1 structure and oligomerization.

Human HSPB1 (accession # P04792) had previously been cloned into pET23a and pET151d vectors (ampicillin resistant). The B1-ACD construct had previously been optimized to truncation of the full-length sequence from Gln80 to Ser176. Site-directed mutagenesis using the QuikChange protocol was used to introduce substitution mutations throughout the sequence.

Several protocols for protein expression were used to obtain different isotopically labeled samples. In almost all cases (unless otherwise specified) for full-length HSPB1, BL21(DE3) E. coli cells were used and a final concentration of 1.0 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) was added to induce protein expression. For the B1-ACD construct, 0.5 mM IPTG was used.

For natural abundance (or non-isotopically labeled) protein, cells were grown in 0.5 L of lysogeny broth (LB) with 100 μg/mL ampicillin. Cells were grown at 37°C in a shaking incubator until OD₆₀₀ ~0.6. IPTG was then added to a final concentration of 1.0 mM and the temperature reduced to 22°C. Protein was expressed in a shaking incubator for ~22 hr. Cells were harvested by centrifugation and resuspension in lysis buffer (50 mM Tris, pH 8.0, 100 mM NaCl, 1 mM ethylenediaminetetraacetic acid [EDTA]).

For ¹⁵N-labeled (no deuteration) protein, MOPS minimal media was used. Per 1 L culture, 1 g of ¹⁵NH₄Cl was used for isotopic labeling. 4 g/L of glucose was used. Growth and expression steps were identical to those used for natural abundance protein.

For ²H¹⁵N¹³C-labeled (partial deuteration,~75%) protein, cells were grown in stages to acclimate the cells to deuterated minimal media. For all ¹³C-labeling, 3 g/L of ¹³C-glucose was used. One colony was grown in 3 mL of LB for ~5 hr and then centrifuged to pellet the cells. These cells were resuspended and grown in 50 mL of H₂O-based M9 minimal media to an OD₆₀₀ ~0.6. Cells were again pelleted, resuspended, and grown in 100 mL of D₂O-based M9 minimal media to an OD₆₀₀ ~0.6. Cells were then transferred directly to a larger 500 mL (total) D₂O-based M9 minimal media and grown to an OD₆₀₀ ~0.6. After induction and reduction of temperature, protein was expressed for ~48 hr.

For ²H¹⁵N¹³C-labeled (perdeuteration) protein, cells were grown in additional stages to acclimate the cells to deuterated minimal media. 3 g/L of ²H¹³C-glucose was used for the final culture. Stocks for deuterated M9 minimal media were also prepared in D₂O. One colony was grown in 3 mL of LB for ~5 hr and then centrifuged to pellet the cells. These cells were resuspended and grown in 50 mL of H₂O-based M9 minimal media to an OD₆₀₀ ~0.6. Cells were again pelleted, resuspended, and grown in 100 mL of D₂O-based M9 minimal (non-deuterated glucose) media for 1–2 hr. Cells were again pelleted, resuspended, and grown in 200 mL of D₂O-based and ²H-glucose-based M9 minimal media to an OD₆₀₀ ~0.6. Cells were then transferred directly to a larger 1 L (total) D₂O-based and ²H-glucose-based M9 minimal media. Cells were grown to an OD₆₀₀ ~0.6 and induced. After induction and reduction of temperature, protein was expressed for ~48 hr.

Cells containing full-length HSPB1 were lysed by freeze-thaw and incubation with lysozyme and protease inhibitors in lysis buffer on ice for 20 min. Generally 0.5 L cultures of cells were lysed in one tube to maximize yield and purity. Deoxycholate was added to the lysed cells and placed on a shaking incubator at 37°C for 15 min. DNase, RNase, and magnesium chloride were then added and shaking incubation continued for 15 min. Cell lysate was centrifuged at high speed at 4°C. Ammonium sulfate was added to the supernatant to 40% saturation on a slow shaker at room temperature and allowed to equilibrate for 30 min. Ammonium sulfate precipitate was centrifuged at high speed at 20°C and the supernatant discarded. Pellets were used immediately or stored at −80°C for up to 1 week.

Ammonium sulfate pellets were resuspended in anion exchange buffer (AEX- 20 mM Tris, 10 mM MgCl₂, 30 mM NH₄Cl, pH 7.6) at room temperature. Remaining solids were pelleted briefly at 20°C. Protein was desalted using a G25 column in AEX buffer at room temperature. Precipitated material was pelleted briefly at 20°C. Desalted protein was separated by an anion exchange DEAE column with a step gradient of AEX buffer with increasing sodium chloride at room temperature. Protein fractions were analyzed for purity by SDS-PAGE, pooled, and concentrated for SEC. Protein was separated on Superdex 200 or 75 columns in 50 mM sodium phosphate (NaPi), 100 mM NaCl, 0.5 mM EDTA, pH 7.5 buffer. Oligomeric proteins (WT, disease mutants) were separated on a Superdex 200 column, while smaller constructs (HSPB1_dimer, NTR-ACD) were separated on a Superdex 75 column. SEC separated fractions were analyzed for purity by SDS-PAGE, pooled, and concentrated to the desired concentration. Concentration was determined by 280 nm absorbance (extinction coefficient of 40,450 M⁻¹cm⁻¹).

For cysteine-free proteins or proteins containing only the native cysteine at position 137, no reducing agent was added during purification. With the native cysteine present, the resulting protein was generally >95% oxidized at the dimer interface as seen by dimer formation by non-reducing SDS-PAGE. For proteins containing non-native cysteines (for fluorophore or spin labeling), the reducing agent dithiothreitol (DTT) was included at each stage of purification to avoid disulfide formation.

All NMR experiments were carried out on either 600 or 800 MHz Bruker spectrometers equipped with cryoprobes. All samples were prepared in 50 mM sodium phosphate (NaP_i), 100 mM NaCl, 0.5 mM EDTA, pH 7.5 buffer. Spectra were collected at 30°C.

Several TROSY-based triple resonance experiments were implemented to assign peaks in the NTR-ACD spectrum to particular residues- HNCO, HN(CA)CO, HNCA, HN(CO)CA, HNCACB, HNCB, and HNCOCANNH (‘NNH’) experiments. Non-uniform sampling (NUS) at a sampling rate of 25% was used for longer experiments. For NUS datasets, an iterative soft threshold algorithm was used to reconstruct full spectra (Hyberts et al., 2012). A maximum protein concentration of 600 μM was used as inter-dimer interactions were evident at higher concentrations (many peaks broadened). Both ~75% deuterated and perdeuterated ²H¹⁵N¹³C-labeled forms of NTR-ACD (expression in previous section) were used for almost all triple resonance experiments, with modest improvements observed for higher deuteration levels. For the HNCB and R2 experiments, only perdeuterated protein was used.

To measure intensities and positions of peaks in 2D spectra, NMRViewJ was used (Johnson, 2004). The following equation was used to calculate chemical shift perturbations (CSPs) between peaks in two spectra:

Spin-label constructs were made with a NTR-ACD/C137S background and cysteines introduced at positions throughout the NTR, one at a time. Samples were prepared in 50 mM NaPi, 100 mM NaCl, 0.5 mM EDTA, pH 7.5 buffer and 10 mM DTT to fully reduce all cysteines. Reducing agent was then removed from protein samples using a desalting column. Immediately after removing reducing agent, 5-fold molar excess (1-oxyl-2,2,5,5-tetramethylpyrroline-3-methyl)-methanethiosulfonate (MTSL) spin label (in DMSO) was added to protein samples and allowed to incubate overnight at 4°C or at room temperature for 2 hr. Excess MTSL was then removed from the labeled protein using a desalting column. The resulting NMR samples contained 300–400 μM protein. 2D ¹H-¹⁵N HSQC-TROSY spectra were collected for each spin-labeled mutant protein. The unpaired spin label was then quenched in each sample by addition of ascorbate (5 mM final concentration). Identical spectra were collected for each quenched sample for intensity comparison between quenched and unquenched spectra.

The distal, aromatic, conserved, and boundary region peptides were purchased from Genscript. The N-terminal residue of all but the distal peptide was formylated, and the C-terminal residue of all peptides was amidated. The Trp-rich peptides were purchased from LifeTein and were acetylated on the N-terminus and amidated on the C-terminus. The distal, conserved, and boundary peptides were dissolved in NMR buffer prior to use. The aromatic and Trp-rich peptides were dissolved in DMSO to a concentration of 100 or 50 mM, then diluted in buffer to a concentration of 1 mM. All peptides were stored at −80°C. A spectrum of ¹⁵N B1-ACD in the presence of 1% DMSO was collected and used as a reference for the aromatic and Trp-rich peptide experiments, although it was highly similar to the ¹⁵N B1-ACD spectrum in the absence of DMSO.

200 μM protein samples were equilibrated at room temperature (22°C) in 50 mM NaPi, 100 mM NaCl, 0.5 mM EDTA at pH 7.5 for several hours. Samples were diluted 10-fold into deuterated buffer (prepared identically but with D₂O) for a final concentration of 20 μM and incubated at room temperature for various periods of time to allow for hydrogen-deuterium exchange. At the desired time point, the deuteration reaction was quenched by adding an equal volume of quench buffer (0.6% formic acid) on ice for a final pH of 2.5. Quenched samples were immediately flash frozen in liquid nitrogen and stored at −80°C. Undeuterated samples were prepared in a similar fashion but replaced addition of D₂O buffer with protonated buffer. Fully deuterated samples were made by first denaturing the protein (3M guanidine HCl and high heat for at least 30 min), making the same dilution into D₂O buffer, incubating for several hours, and quenching the same as all other samples.

Samples were stored in liquid nitrogen until 5 min prior to injection to maintain consistent levels of deuterium loss (back-exchange). Initially, the sample was passed over a custom packed pepsin column (1 × 50 mm) at 200 uL/min in 0.1% formic acid at 1°C for digestion of the protein into peptides. Digested peptides were then captured onto a trapping column (Waters vanguard BEH C18 2.1 × 5 mm 1.7 µm 130 Å) and resolved over a C18 reverse-phase column (Waters BEH 1 × 100 mm 1.7 µm 130 Å) using a linear gradient of 3% to 40% B over 10 min (A: 0.1% formic acid, 0.025% trifluoroacetic acid, 2% acetonitrile; B: 0.1% formic acid in acetonitrile). The LC system was coupled to a Waters SYNAPT G2-Si Q-TOF. The source and desolvation temperatures were 70°C and 130°C, respectively. The StepWave ion guide settings were set to minimize non-uniform deuterium loss during desolvation (Guttman et al., 2016). The pepsin, trap, and resolving columns were washed extensively to reduce sample carryover (Fang et al., 2011; Majumdar et al., 2012). The resulting levels of carryover were below 5% for each peptide analyzed based on blank runs. Peptic peptides of WT and mutant proteins were analyzed by tandem MS (MS^E) analyzed by ProteinLynx Global SERVER.

Most peptides could be directly compared among all mutants. For peptides containing a mutation the comparisons are only qualitative as the change in exchange could be an effect of both intrinsic exchange and local structure. MassLynx software was used to align spectra of various time points at the appropriate retention times, and HX-Express v2 (Guttman et al., 2013) was used to analyze deuterium incorporation and perform bimodal analysis. The deuteration level at each time point was calculated relative to the deuteration levels of the undeuterated and fully deuterated spectra for each peptide. In cases where the fit was very poor due to very broad isotope distributions or clear bimodals, an alternative fitting was used to obtain a bimodal distribution, representing two distinct deuteration states of the peptide at a given time point. For bimodal peptides shown in Figures 7 and 9, their behavior was confirmed across several charge states and in most cases several overlapping peptides. Three biological replicates of WT oligomers and HSPB1_dimer were examined and showed qualitatively similar patterns in HDX (Figure 7—figure supplement 2), with one of these replicates containing the native C137 and including reducing agent to confirm similar behavior to C137S constructs. Disease mutants were examined with single replicates due to limitation of instrument time.

A starting homology model was produced by aligning a crystal structure of the HSPB1 ACD (PDB 4MJH) with a crystal structure of an HSPB2/HSPB3 heterotetramer (PDB 6F2R) and creating a new PDB file with structural elements from both. It contained the HSPB1 ACD, including both β2 strands, a copy of the HSPB2 motif ¹⁵⁷VNEVYISLL¹⁶⁴ bound in each β4/β8 groove, and a copy of the HSPB2 conserved motif ²³RLGEQRFG³⁰ bound in the dimer interface groove. Residues from the HSPB2/HSPB3 crystal structure were mutated to the appropriate sequence for HSPB1 using the PyMOL Molecular Graphics System (Version 2.0, Schrödinger LLC). PyRosetta (Chaudhury et al., 2010) was used to eliminate clashes and model in missing segments. The function FastRelax (Das and Baker, 2008) was used to relax the starting homology model and eliminate clashes between chains. Fragment insertion using the BlueprintBDR (Huang et al., 2011) mover was then used to add the initial residues of the distal region and a segment of the 2/3 loop that was not resolved in the crystal structure. Two distinct connectivities involving the aromatic region were modeled. One structure was generated in which the aromatic region connected the distal region bound in the opposite orientation of the conserved region, such that it wrapped around the edge of the ACD and contacted loops 3/4, 5/6 and 8/9. Another was generated in which it connected the conserved region to the other distal region and crossed over strands β8, β9, and β3. An initial model for each of these two structures was generated using the BlueprintBDR mover, and then subject to 100 rounds of Generalized Kinematic Closure (Bender et al., 2016) to generate two sets of 100 structures that sample the conformational flexibility of the aromatic region. Generalized Kinematic Closure was used to model this longer disordered segment because, unlike the BlueprintBDR mover, it does not explicitly bias structures toward a smaller radius of gyration or use stretches of PDB-derived torsion angles. The missing loops connecting the beginning of the NTR to the β2 strands were then modeled in using the BlueprintBDR mover. Again, there were two possible connectivities for each structure: the conserved motif could be connected to either β2 strand, while the distal region not connected to the conserved motif could be connected to the other β2 strand. For each model, both connectivities were sampled, producing four sets with unique topologies that each contained 100 structures. For each step in which additional residues were introduced in the model, FastRelax was used to relax the new segment and the residues adjacent to it. Finally, FastRelax was used to relax all atoms in the final structures.

Samples were prepared in 25 mM NaPi, 50 mM NaCl, and 0.25 mM EDTA buffer at pH 7.5. Samples were incubated at 20 μM at room temperature for several hours prior to measurement. All measurements were made on a Jasco J-1500 CD spectrometer with Peltier temperature control at 20°C, with 1 nm bandwidth, and averaged over three scans. All data were normalized to units of mean residue ellipticity (MRE). Data were collected at 0.1 nm intervals and then smoothed linearly across 1 nm. All data presented have a high-tension voltage below the recommended cutoff for the detector (800 V).

WT and disease mutant constructs were generated with a cysteine introduced at position 174, analogous to similar fluorescence studies in HSPB5 (Peschek et al., 2013), and the native cysteine at position 137 mutated to serine. Protein was buffer exchanged into 50 mM NaPi, 100 mM NaCl, 0.5 mM EDTA, pH 7.5 buffer with 2 mM tris-(2-carboxyethyl)-phosphine (TCEP). Protein was incubated with 3X molar excess of Alexa Fluor 488 maleimide and incubated at 37°C (to facilitate subunit exchange) for at least one hour. Protein was separated from free dye using gravity desalting columns. Fractions collected from desalting columns were analyzed for separation by comparing absorbances at 495 nm (fluorophore extinction coefficient of 73,000 M⁻¹cm⁻¹) and 280 nm, with an assumed A₂₈₀/A₄₉₅ ratio of 0.11. The resulting pool of fractions was diluted to desired protein concentrations and fluorophore labeling percentages.

Samples were incubated at 37°C at 20 μM and several labeling percentages for at least one hour prior to measurement. For the experiments presented here, samples with 30% fluorophore labeling were mixed 1:2 with unlabeled protein. The resulting dequenching from homo-FRET among fluorophores in oligomers was measured as a function of time. Measurements were collected on a Horiba Fluorolog-3 with double excitation and emission monochromators and Peltier temperature control at 37°C. Excitation and emission wavelengths of 518 nm and 498 nm were used, respectively.

The resulting kinetic data were fitted to the following exponential equation using the R software package to obtain subunit exchange rates:

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

We thank P Rajagopal and L Tuttle for training and discussions of NMR experiments and J Kang for assistance with HPLC. We thank K Reiter and D Wilburn for critical reading of the manuscript and all members of the Klevit lab for useful discussions and suggestions. This work was supported by NIH grant 2R01 EY017370 to REK, R01GM127579 to MG, NIH T32 GM008268 and Hurd Fellowship in Biophysics from the UW School of Medicine to AFC and HERB, and the Hope Barnes Graduate Fellowship from the UW School of Pharmacy to HERB.

© 2019, Clouser et al.

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