Dynamics of the Proline-Rich C-Terminus of Huntingtin Exon-1 Fibrils
Intrinsically disordered protein domains are not only found in soluble proteins but can also be part of large protein complexes or protein aggregates. For example, several amyloid fibrils have intrinsically disordered domains framing a rigid \xce\xb2-sheet rich core. These disordered domains can often be observed using solution NMR methods in combination with modest magic angle spinning and without perdeuteration. But how can these regions be detected using solution NMR methods when they are part of a fibril that is not tumbling isotropically in solution? Here we addressed this question by investigating the dynamic C-terminus of huntingtin exon-1 (HTTex1) fibrils that are important in Huntington\xe2\x80\x99s Disease. We assigned the most dynamic regions of the C-terminus of three HTTex1 variants. Based on this assignment, we measured site-specific residual chemical shifts, peak intensities, and R1, R\xe2\x80\xb22 and R1\xcf\x81 15N relaxation rates. In addition, we determined the residual 1H-15N dipolar couplings of this region. Our results show that the dipolar couplings are averaged to a very high degree resulting in an order parameter that is essentially zero. Together, our data show that the C-terminus of HTTex1 is intrinsically disordered and undergoes motions in the high picosecond to low nanosecond range.
Introduction<!>Protein Production and Fibril Formation<!>NMR spectroscopy<!>Data analysis<!>Results<!>Discussion<!>Conclusions
<p>The importance of intrinsically disordered protein domains (IDDs) is increasingly recognized, and new methods are needed to study the dynamics and residual structure of these domains. While solution NMR has emerged as one of the most important tools to study these domains in soluble proteins, IDDs are more difficult to study when part of a structure too large to be detected using solution NMR.</p><p>One example of such as IDD is found in fibrillar aggregates formed by huntingtin exon-1 (HTTex1) important in Huntington's Disease (HD). HD is a heritable neurodegenerative disease that is caused by a CAG expansion in exon-1 of the huntingtin gene. This CAG expansion leads to an elongated poly-glutamine (polyQ) region in the huntingtin protein, which increases its propensity to aggregate.1 HTTex1, which has been shown to be dominant in cellular huntingtin inclusions in HD and to exist as an alternative splice variant,2 becomes especially aggregation prone when the polyQ expansion exceeds 36 glutamine residues,3 which coincides with the repeat length above which HD is observed.4</p><p>HTTex1 exhibits a highly heterogeneous structure, as shown by circular dichroism (CD), electron paramagnetic resonance (EPR), and solid-state nuclear magnetic resonance (NMR) spectroscopy.5–10 The polyQ domain forms the static core of the fibril, whereas the proline-rich C-terminus is highly dynamic and in a mixed random coil, polyproline II (PPII) helical conformation.5,11 The N-terminus of HTTex1 (N17) has helical propensity in the soluble12,13 and fibrillar state.6 These findings led us to propose a bottle brush model of HTTex1 fibrils where the polyQ domain forms the core and the dynamic, proline-rich domain the bristles of the brush.5</p><p>Similar heterogeneous systems with static, β-sheet-rich cores and highly flexible domains have been observed for several pathologic amyloid fibrils (e.g. α-synuclein14 and hPrP15) as well as functional amyloids (e.g. HET-s,16 Orb217,18). The role of many of the disordered domains found in conjunction with a cross-β fibril core is unclear. However, in the case of HTTex1 the dynamics of the flanking sequences and the C-terminus in particular correlates with the toxicity of recombinant fibrils.9,19 This correlation suggests that the interaction of other cellular components with the C-terminal bristles of the bottle brush might be important for understanding HTTex1 toxicity. Therefore, site-specific assignment and site-specific dynamics are required to measure interactions of the dynamic C-terminus with binding partners.</p><p>We and others have shown that a combination of solid-state and solution NMR techniques allows us to selectively detect IDDs as part of amyloid fibrils,14,16,17,20–22 membrane proteins,23 protein-DNA complexes,24 and chemotaxis receptors.25 We recently showed that site-specific assignments of dynamic amyloid flanking sequences can be obtained using triple resonance NMR experiments developed for protein backbone assignments in solution.18 But how dynamic do regions have to be to allow 1H detected methods to work at moderate MAS frequencies and in the absence of perdeuteration? To answer this question, we not only assigned the most dynamic regions of HTTex1 fibrils, but also measured site-specific relaxation rates and residual dipolar couplings. These measurements allow us to present a detailed model of this dynamic domain.</p><!><p>Uniformly labeled 13C,15N‒HTTex1 Q46 was expressed10 and purified27 as described earlier except that this time we used a BioRad NGC Scout FPLC system for protein isolation. The C-terminally His-tagged HTTex1HT and HTTex1Pro constructs were expressed and purified as previously described.5,10,26</p><p>Fibril formation was instigated by addition of 1% (V/V) unlabeled HTTex1(Q46) seeds to the eluted protein. Samples were incubated with seeds for ~36 hours at 4 °C in 10 mM/100 mM NaCl tris buffer, pH = 7.4 and harvested by ultra-centrifugation with a Beckman Coulter Optima TLX using a Beckman TLA 100.3 rotor at 40,000 rpm for 40 minutes at 4 °C. Pellets were washed with 0.1% triflouroacetic acid (TFA) and ultra-centrifuged again at 40,000 rpm for 40 minutes at 4 °C. Supernatant was reduced to 100 µL for loading the rotor. Electron microscopy (EM) images were taken to ensure fibril quality using a JEOL JEM-1400 Electron Microscope at 100 kV. EM grids were prepared as previously described.10 A 1.6 mm Agilent rotor was loaded by centrifugation of HTTex1(Q46) fibrils directly into the MAS rotor at 10,400 rpm and 4 °C in a Sorvall Surespin 630 rotor for 40 minutes using a home-built packing tool.</p><!><p>All NMR experiments were measured on a 14.1 T Agilent DD2 solid-state NMR spectrometer (1H = 599.78 mHz; 13C = 150.83 MHz; 15N = 60.78 MHz) equipped with a T3 1.6 mm probe. A MAS frequency of 12 kHz was used if not noted otherwise, with 200 kHz and 50 kHz pulses on 1H and 15N, respectively. Temperature was maintained at 0 °C.</p><p>The HSQC, HNCA, and HNcoCA spectra were recorded as described previously with an additional BIRD filter with a water relaxation delay of 260 ms for improved water suppression.18</p><p>R1 rates were measured using the pulse sequence shown in Figure 1A.27 The values of τ and γ were 2.2 and 260 ms, respectively. 1H GARP decoupling with an rf-field strength of 5 kHz was applied during the relaxation delay framed by two 1 ms 10 kHz 1H spin lock pulses for water suppression. 15N WALTZ decoupling with an rf-field strength of 5 kHz was used during acquisition. R′2 rates were measured using the pulse sequence shown in Figure 1B.28 The delays were the same as for the R1 experiments with the additional CPMG delay δ of 0.5 ms, and the spin lock pulses for water suppression lasting for 3 ms with an rf-field strength of 40 kHz. R1ρ was measured with the pulse sequence shown in Figure 1C, which is very similar to the R′2 experiment, except that the CPMG spin echoes were replaced with a 6 kHz spin lock pulse on 15N and 2.5 kHz WALTZ decoupling on 1H. An additional Z-filter of ζ=20ms was added before the spin lock. Residual dipolar couplings were measured using the 1H-15N REDOR pulse sequence shown in Figure 1D.29 This experiment was equivalent to the R1ρ experiment, except that the MAS frequency was 12.5 kHz (τr=80 μs), and the spin lock was replaced with 1H-15N REDOR recoupling in which the rf-field strength of the 1H 180º pulses was 50 kHz to reduce sample heating. S0 was recorded without 1H 180º pulses during the REDOR recouplings. R1, R′2 , R1ρ, and 1H-15N REDOR experiments were recorded with spectral widths of 10 kHz and 3 kHz for 1H and 15N, respectively. 128 acquisitions were co-added for each of the 60 indirect increments. The 1H-13C 2D INEPT HETCOR experiments were recorded as described previously.5 Data were processed using NMRPipe and NMRDraw and plotted with custom scripts written with Python and NMRglue.30</p><!><p>Resonance assignment and peak integration were done using the program CARA. Relaxation curves were fit using custom scripts written in Python.</p><p>Correlation times τc were calculated from the experimentally determined values for R1, R′2, and S. According to Schanda & Ernst,31R1can then be written as
R1=(δD4)2(J(ω1−ωS)+3J(ωS)+6J(ω1+ωS))+δCSA234J(ωS)
where δD/2π is the NH dipolar coupling of 22.954 Hz, ωS the 15N Larmor frequency, ωI the 1H Larmor frequency, and δCSA/2π the chemical shift anisotropy of backbone 15N of 6867 Hz. In a single-timescale, model-free approach under MAS, the spectral density function is described as
J(ω)=25(1−S2)τc1+(ωτc)2</p><p>Similarly, the transverse relaxation rate can be described as
R2=(δD4)2(43J(ωr)+23J(2ωr)+12J(ωI−ωS)+3J(ωI)+32J(ωS)+3J(ωI+ωS))+δCSA2(14J(ωr)+18J(2ωr)+38J(ωS))
where ωr is the MAS frequency. R1 and R2 were solved for τc using the program Mathematica to estimate correlation time of the motion.</p><p>We calculated R2 from the experimental values of R1
and R1ρ using the following equation
R2=[R1ρ−R1cos2(θ)]/sin2(θ)
where θ is the angle between the effective 15N rf-field and the static magnetic field.32</p><!><p>How many residues of HTTex1 Q46 fibrils are so dynamic that they can be detected using 1H detected, J-coupling based, triple resonance spectroscopy? To answer this question, we prepared HTTex1 Q46 fibrils at 4 °C in the presence of 1% (V/V) HTTex1 seeds (see Materials and Methods) and acquired the HSQC (heteronuclear single-quantum correlation) spectrum shown in Figure 2. The lack of a gradient coil in our 1.6 mm Agilent MAS probe required a BIRD (bilinear rotational decoupling) filter element preceding the HSQC experiment to achieve water suppression rather than the pulsed-field gradient based methods used in solution-state NMR spectroscopy. The HSQC spectrum of uniformly labeled 1H,13C,15N‒HTTex1 fibrils has 12 well-resolved amide cross peaks with 1H and 15N linewidths as low as 22 Hz and 24 Hz, respectively. The intensities of the cross peaks vary, with some resonances being well defined and intense and others broadened and barely detectable. The relatively narrow distribution of amide proton shifts around 8.5 ppm indicates intrinsic disorder in this domain.</p><p>To determine the origins of these signals, we recorded HNCA and HNcoCa spectra similar to our previous work on the flexible domains of fibrils formed by the functional amyloid Orb2A.18 As can be seen from the strip plots of the HNCA and HNcoCa in Figure 3, the signal to noise of these spectra is good, which allowed for the resonance assignment of most resonances detected in the HSQC in Figure 2. G102 was assigned based on its unique chemical shift and the fact that there is only one Gly residue in HTTex1.</p><p>The assigned residues are all located at the end of the C-terminus of HTTex1, termed C12, that follows the second polyP stretch P10 (see Figure 4A). To evaluate the (residual) secondary structure of this region, we calculated the secondary Cα chemical shift (ΔδCα), the difference between the measured Cα chemical shift and the random coil chemical shift of the same residue type.33 As can be seen from Figure 4B, the absolute value of ΔδCα is close to or below 1 ppm for most of the residues, indicative of a random coil structure. The larger ΔδCα values observed for E108 and R112 can be explained by the fact that these residues precede Pro residues, known to lower the Cα shift by up to 2 ppm.34 Although prolines could not be detected using amide-proton based spectroscopy, we could observe the Cα chemical shifts of P103 and P109 as Cα−1 of Ala104 and Leu110 in the HNcoCA spectrum. In both cases, the chemical shift of these prolines is about 63 ppm, which is compatible with the Pro type B we described earlier to be in a random coil conformation.5</p><p>Because intrinsically disordered domains are defined as much by their residual secondary structure as by their dynamics, we characterized the dynamics of the C12 region in more detail. Figure 4C shows the sequence-dependent NH peak intensities of the HSQC spectrum in Figure 2. The intensity increases gradually from G102 toward the C-terminus. This increase suggests an increase in mobility resulting in a decrease of R2 relaxation rates, making the HSQC experiment more efficient. To confirm this interpretation, we directly measured the R1 and R′2 relaxation rates of the assigned residues.</p><p>Figure 1A illustrates the pulse sequence we used for measuring 15N R1 rates,27 which includes an additional BIRD filter for improved water suppression. We measured 15N relaxation up to 1.2 s, which could be fit well to mono-exponential decay rates, as illustrated for residue A106 in Figure 5A (Supporting Figure S1 includes the relaxation curves of all assigned residues). We determined R1 rates between 1.4 s−1 and 2.2 s−1, with the highest rates for G102, i.e. the beginning of the C12 region, and from E107 to H111 (Figure 5B).</p><p>We used the Carr-Purcell-Meiboom-Gill (CPMG) pulse sequence shown in Figure 1B to measure 15N R′2 rates of the residues in the C12 region of HTTex1.28 The decay under the CPMG sequence was measured for up to 160 ms, leading to a mono-exponential decay of the peak intensity, as illustrated in blue for residue A106 shown in Figure 6A (Supporting Figure S2 includes the relaxation curves of all assigned residues). The 15N R′2 rates decrease from a value of 12.89 ±1.62 s−1 for residue G102 to R′2=4.40 ±0.43 s−1 for residue R112, indicative of an increase in dynamics toward the C-terminus (Figure 6B).</p><p>A problem with estimating R2 using a CPMG-type experiment in the solid state is that coherent dephasing from incompletely averaged anisotropic interactions can lead to an overestimation of the relaxation rate. R1ρ measurements were shown to be much less affected by this problem while often containing the same information.31,32,35 To test if the CPMG pulse sequence leads to systematically higher rates compared to R1ρ in our case, we measured R1ρ relaxation rates using the pulse sequence shown in Figure 1C. The R1ρ relaxation curve for A106 (Figure 6A) overlaps very well with the R′2 curve of the same residue, and the resulting rate constants are, within error, the same. (Supporting Figure S3 includes the R′2 relaxation curves of all assigned residues). To directly compare R1ρ with R′2, we converted R1ρ to R2 (see Methods section). The resulting R2 was essentially the same as R1ρ for all residues under the conditions used.32
Figure 6B shows that R2 and R′2 are, within error, the same for each residue.</p><p>The fact that we can apply J-based, 1H detected NMR experiments under moderate MAS frequencies in the absence of perdeuteration suggests an almost complete averaging of the dipolar couplings in the C12 region, a surprising result due to the absence of isotropic molecular tumbling in HTTex1 fibrils compared to molecules in solution. To measure the extent of the residual dipolar couplings that are not averaged by molecular motion, we used the 1H-15N rotational echo double resonance (REDOR) pulse sequence shown in Figure 1D, adapted from Schanda and co-workers.29 Measuring residual dipolar couplings this way also allows us to determine the order parameter independent of the relaxation rates.31 In contrast to the original use of the 1H-15N REDOR experiment in rigid, perdeuterated solids, scaling of the dipolar couplings for improved detection was not necessary, and 1H 180° REDOR pulses were applied every half a rotor period for up to a total of 1000 rotor periods (80 ms at 12.5 kHz MAS). The REDOR dephasing curve for residue A104 is shown in Figure 7A. The whole set of REDOR dephasing curves is shown in Supporting Figure S4. We fit the REDOR curves to a series of Bessel functions to estimate the residual dipolar couplings.36
Figure 7B shows these for each of the assigned residues. The residual dipolar couplings continuously decrease from about 350 Hz for residue G102 to 17 Hz at residue R112, with the exception of V105. The order parameter can be calculated as follows S=δexp/δrigid where δrigid=δNH/(2π)=22954 HZ and δexp is the measured dipolar coupling.31 This order parameter is close to zero for all assigned residues, indicating fast motions that lead to substantial reorientation of the NH bond vector for all residues in C12.</p><p>The separate measurement of the order parameter and relaxation rates allows us to estimate the correlation time of the motions using a single-timescale, model-free approach under MAS.29,31 Using the order parameters determined via the REDOR experiment described above, we could estimate τc from both the experimental R1 and R′2 rates (see Supporting Figure S5). The one physical realistic solution for τcR2 is in between the two solutions for τcR1. Generally, the correlation time is estimated to be in the 500 ps to 9 ns range. The slight difference between the correlation time calculated from R1 and R′2 can be either due to systematic errors in the relaxation rates or the simple, single-timescale model-free model used here. We plan to expand the set of relaxation measurements, measure R1 rates at different magnetic fields, repeat our data on perdeuterated samples, and use different models to improve the characterization of this domain in future studies.</p><p>Our previous studies on HTTex1 were performed on a C-terminally His-tagged construct (HTTex1HT).5,10 To test how the C12 region was affected by the His-tag, we collected HSQC, HNCA, and HNcoCA spectra of HTTex1HT. Figure 8A shows the HSQC recorded on HTTex1HT. The assignment showed that only the C12 region was detected in these experiments, very similar to HTTex1 without His-tag. The NH peaks of the His-tag could be assigned in addition to the residues in the C12 region. The secondary chemical shifts ΔδCα of this sample are similar to HTTex1 without the His-tag, and the additional His residues are compatible with a random coil conformation, except for the C-terminal His (Figure 8B).</p><p>The NH peak intensities in Figure 8C show a general increase in intensity toward the C-terminus, with G102 having the lowest intensity and H119 almost twice the intensity compared to the other residues, which might be due to additional conformational freedom at the very C-terminus of the protein.</p><p>HTTex1Pro is another HTTex1 mutant affecting the C12 region. This construct, which was shown not to bind to the PHP1 and PHP2 antibodies due to the absence of the required epitope, has the L17 region replaced with 17 prolines and all prolines in the C12 region replaced with Gly.26 The INEPT-HETCOR spectrum of this mutant confirms the changes to the C-terminus via the absence of the ProB peak, a strong Gly signal, weaker Leu signals, and more well-defined Glu peaks (see comparison of the INEPT HETCOR spectra of the three HTTex1 variants in Supporting Figure S6). Figure 9A shows the HSQC spectrum of HTTex1Pro. The assignment of the residues in this spectrum using HNCA and HNcoCA spectra again showed that all assigned peaks are located in the C12 region. In contrast to HTTex1HT, we were neither able to assign the individual His residues of the His-tag nor the Gly113 preceding the His-tag. The ΔδCα values shown in Figure 9B are all close to zero due to the absence of Pro in the C12 region, with the exception of R112. The HSQC peak intensities shown in Figure 9C are different than those of HTTex1 and HTTex1HT. After an initial increase in intensity from G102 to A104, the intensity decreases toward the C-terminus, rendering the His-tag barely detectable.</p><p>The data on the two HTTex1 variants show that the C12 region is very dynamic irrespective of the presence of the His-tag or prolines in the sequence. Nevertheless, we are able to detect changes in chemical shift and dynamics in this region.</p><!><p>Previously, we and others have shown that the Pro-rich C-terminus of HTTex1 fibrils is dynamic in contrast to the rigid polyQ core of the HTTex1 fibrils.5,8,11 Here, we go a step further by showing that the C12 region of HTT can be assigned using 1H detected, triple resonance, out an and back style solution NMR experiments in the absence of perdeuteration or without high MAS frequencies. But how is this possible in the absence of isotropic tumbling that it is usually required for solution NMR methods to work? To answer this question, we analyzed peak intensities, secondary chemical shifts of these residues, and measured site-specific relaxation rates and residual dipolar couplings.</p><p>G102 is the first residue detectable in our HSQC experiment because the polyP region cannot be detected using amide proton-based spectroscopy. On the other hand, the dynamics of G102 were insufficient to obtain peaks in the HNCA and HNcoCA spectra, indicating that its reduced flexibility limits detection in our approach. This is probably why we could not assign any residues from the L17 region, which is likely to be less dynamic than the C12 region. Nevertheless, the L17 region is still much more mobile than the polyQ domain that forms the core of the HTTex1 fibrils and can, therefore, be detected using straight-though INEPT-HETCOR 2D spectra (as seen in Supporting Figure S6), which are more forgiving to transverse 1H and 13C relaxation. Our R′2 and residual dipolar coupling data support that G102 is the least flexible of the assigned residues. The increase in intensity of the NH cross peaks toward the C-terminus indicates an increase of motions toward the C-terminus. This conclusion is supported by both the R′2 rates and residual dipolar couplings, both of which decrease toward the C-terminus. Furthermore, these conclusions are compatible with EPR data showing that residue 111 is the most dynamic in HTTex1HT fibrils11 and with our bottle brush model of the HTTex1 fibrils.5 In this model, the Pro-rich C-termini form entropic bristles that protrude from the static polyQ core of the fibrils, which allows for maximal motility at the end of the C-terminus.</p><p>Interestingly, our R1 rates do not show a clear trend (Figure 5). Generally, the 15N R1 rates we measured are large compared to what has been observed in solid-state NMR on globular proteins29,37 and more comparable to 15N R1 rates observed in solution NMR of intrinsically disordered or denatured proteins,38,39 indicating a highly dynamic and disordered nature of this region. At R1 rates this high, an error due to 15N-15N spin diffusion is no concern.31 However, a systematic error of up to 20% due to H2O proton-spin saturation similar to what is observed in solution NMR of larger proteins cannot be ruled out.40,41</p><p>Measuring R′2 rates using CPMG-type experiments in solid-state NMR is difficult because of coherent dephasing from incompletely averaged anisotropic interactions even at very high MAS frequencies and with perdeuteration. The fact that we can observe narrow 1H lines in our HSQC spectrum in the absence of perdeuteration suggests that we have favorable conditions to measure R′2 in which dipolar couplings are averaged to a very high degree and the motions are on much faster time scales than the MAS frequency.31 This is confirmed by the R1ρ measurement that, although less affected by coherent dephasing,35 resulted in essentially the same relaxation rates. The R′2 rates are generally similar to what has observed in globular proteins using solid-state NMR as well as intrinsically disordered proteins in solution.32,35,38 Both R′2 and R1ρ decrease continuously towards the C-terminus compatible with a continuous increase in dynamics.</p><p>Because it can be challenging to determine both the correlation time of the motion and the order parameter from relaxation rates alone, additional measurements of S as the scaling of anisotropic interactions like the NH dipolar coupling is a useful method to characterize dynamics in solid-state NMR. Our measurements of the residual NH couplings confirmed that the dipolar couplings in the C12 region are averaged from their theoretical value of 22,954 Hz to only 350–17 Hz. The resulting order parameters are very close to zero, showing that this domain is dynamically disordered. The low order parameter in the C12 region is also compatible with our observation that the secondary chemical shifts ΔδCα are very close to their random coil values. Together these data indicate that the motions of the C12 region of HTTex1 are fast and lead to substantial reorientation of the NH bond vector (at least close to a semi angle θ0 of 90º in a diffusion on a cone model in which S2=[cos θ0(1+cos θ0)/2]2). However, the fact that we were able to measure residual dipolar couplings that decrease toward the C-terminus also shows that the detected residues are not free in solution and still part of the amyloid fibril. Using the order parameter for the NH dipolar coupling measurements, we estimated the motional correlation time from both the R1 and R′2 measurements, which are in the high ps to low ns range. In summary, our data support a model in which the C12 region of HTTex1 is intrinsically disordered and undergoes fast and motions, explaining why they seem to be almost in solution from an NMR perspective.</p><p>The relaxation and residual dipolar coupling data show that the dynamic C12 region of HTTex1 fibrils are, from an NMR point of view, an interesting chimera between solid protein and a protein in solution (see Figure 10). On the one hand, HTTex1 fibrils are not tumbling isotropically in solution, yet the C12 region is so dynamic that homo- and heteronuclear dipolar couplings are averaged to a very high degree. This allows the use of relatively standard solution-state NMR pulse sequences, including CPMG R′2 measurements that suffer from large systematic errors on structured and less dynamic proteins in the solid-state, even at very high MAS frequencies or with perdeuteration.31 On the other hand, we were able to directly observe residual dipolar couplings using dipolar recoupling under MAS similar to less dynamic proteins in the solid-state. For proteins in solution the measurement of residual dipolar couplings usually requires the alignment of the protein with respect to the static magnetic field.42</p><p>Our comparison of three different HTTex1 C-terminal variants shows that the high mobility in this region does not depend on the presence of a His-tag or the presence of the Pro in this region. This is supported by the observation that the His tag is more dynamic in the case of HTTex1HT but less in HTTex1Pro. Therefore, the location of this region at the outer tip of the Pro-rich seems to be more important for its high mobility. Although the majority of the Pro-rich domain is in a poly-proline II helical conformation, as shown by its Pro chemical shifts,5,7 it is nevertheless a highly dynamic structure due to the absence of stabilizing hydrogen bonds.43 This high flexibility makes it possible to detect the Pro-rich domain in our INEPT-HETCOR experiments and allows the C12 region at its tip to be detected using the HSQC based experiments used in this study.</p><p>The results from the all Pro sample clarify the assignment of our original INEPT-HETCOR data.5 Pro B is from those Pro that have non-Pro residues as neighbors. Likewise, the Gln resonances observed in these spectra result from the Gln in L17 and not the end of the polyQ domain. There is very little evidence for signals from the N17 region, which was shown relatively less mobile by EPR and solid-state NMR.6,11</p><p>Our spectra of the three C-terminal HTTex1 variants show that we are able to detect structural changes in these regions due to mutations and potentially detect the interaction of binding partners to this region using the approach presented in this study. This is important because the dynamic C-terminus has been identified to be the site that distinguishes different HTTex1 fibril polymorphs.9 Furthermore, the C-terminus of HTTex1 has been shown to be an important epitope for HTTex1 specific antibodies26 and intrabodies44 that can reduce HTT toxicity. The Pro-rich C-terminus of HTTex1 is also also a major site of functional protein-protein interactions involving HTTex1,45 including profilin46,47 and membrane associated proteins containing SH3 domains.48,49</p><!><p>We assigned the most dynamic domains of HTTex1 using 1H detected J-based triple resonance experiments, moderate MAS frequencies, and fully protonated protein. All assigned residues are located in the C12 region, the very C-terminus of the protein. The assignment allowed us to record residue specific 15N R1, R2, and R1p rates. In addition, we measured residual dipolar couplings of this region. The Cα chemical shifts, relaxation rates, and residual dipolar couplings show a) that the C12 region is in a random coil conformation, with order parameters that are very close to 0, and b) that it undergoes motions in the high picosecond to low nanosecond range. Besides providing a site specific description of the dynamics of the C12 region of HTTex1, we demonstrate that it is not only possible to assign intrinsically disordered domains in the solid-state but also study their dynamics in detail similarly to proteins in solution.</p>