Hearing sensitivity in mammals is sharply tuned by a cochlear amplifier associated with electromotile length changes in outer hair cells (Dallos et al., 2006). These changes are driven by prestin (SLC26A5), a member of the SLC26 anion transporter family, which converts voltage-dependent conformational transitions into cross-sectional area changes, affecting its footprint in the lipid bilayer (Sfondouris et al., 2008). This process plays a major role in mammalian cochlear amplification and frequency selectivity, with prestin knockout producing a 40- to 60-dB signal loss in live cochleae (Liberman et al., 2002). Unlike most molecular motors, where force is exerted from chemical energy transduction, prestin behaves as a putative piezoelectric device, where mechanical and electrical transduction are coupled (Dong et al., 2002). As a result, prestin functions as a direct voltage-to-force transducer. Prestin’s piezoelectric properties are unique among members of the SLC26 family, where most function as anion transporters.

Recent structures determined by cryo-electron microscopy (cryo-EM) have sampled prestin’s conformational space under various anionic environments and located the anion-binding site at the electrostatic gap between the amino termini of TM3 and TM10 helices (Bavi et al., 2021; Ge et al., 2021; Butan et al., 2022). This anion-binding pocket is highly conserved, and is influenced by surrounding hydrophobic residues in TM1 and by a fixed positive charge from residue R399 on TM10. Movements of this binding site are coupled to the complex reorientation of the core domain relative to the gate domainn (Bavi et al., 2021; Ge et al., 2021), reminiscent of the conformational transitions in transporters displaying an elevator-like mechanism (Garaeva and Slotboom, 2020). Prestin exhibits minimal transporter ability yet is structurally similar to the non-electromotive anion transporter SLC26A9 (sequence identity = 34%; Cα root-mean-square deviation (RMSD) = 3.4 Å for the transmembrane domain [TMD], PDB: 7S8X and 6RTC) (Butan et al., 2022; Walter et al., 2019; Chi et al., 2020). Questions remain as to the molecular basis underlying the distinct functions of the two proteins. Importantly, the role of bound anions, which is required for prestin electromotility (Oliver et al., 2001), is still elusive.

Prestin’s voltage dependence is tightly regulated by intracellular anions of varying valence and structure (Rybalchenko and Santos-Sacchi, 2003; Rybalchenko and Santos-Sacchi, 2008), whereas anion affinity is also regulated by voltage and tension (Song and Santos-Sacchi, 2010). These phenomena suggest that anions, rather than behaving as explicit gating charges, may serve as allosteric modulators (Song and Santos-Sacchi, 2010). Incorporating a fixed charge alternative to a bound anion through an S398E mutation preserves prestin’s nonlinear capacitance (NLC) but results in insensitivity to salicylate, a strong competing anionic binder (Butan et al., 2022). Except for residue R399, charged residues located in the TMD distribute toward the membrane–water interface (Bavi et al., 2021; Ge et al., 2021; Butan et al., 2022) and display minimal contributions to the total gating charge estimated from NLCs (Bai et al., 2009). Electrostatic calculations show that R399 has a strong contribution to the local electrostatics at the anion-binding site, by providing ~40% of the positive charge at the bilayer mid-plane (Bavi et al., 2021). However, the existing structural and functional data cannot explain why prestin’s voltage dependence requires close proximity of both a negative charge (the bound anion or S398E) (Oliver et al., 2001) and a positive charge (R399; Bavi et al., 2021; Gorbunov et al., 2014). The resolution of this conundrum will define an essential step toward our understanding of prestin’s unique voltage-sensing mechanism.

Here, we studied the influence of anion-binding on the dynamics and structural changes of prestin as a function of anions (Cl⁻, SO₄²⁻, salicylate, and HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)) via hydrogen–deuterium exchange mass spectrometry (HDX-MS). The HEPES condition was achieved by Cl⁻ removal (dialysis), which inhibits prestin’s NLC (Oliver et al., 2001). In a HEPES-based buffer, prestin’s NLC shifts to depolarized potentials, associated to a more expanded state at 0 mV that is coupled to low anion affinity (Rybalchenko and Santos-Sacchi, 2003; Rybalchenko and Santos-Sacchi, 2008). Based on the above studies and the large size of HEPES anions, we assumed minimal binding of HEPES anions to prestin and hence associated HEPES condition to a putative apo state in this study. By comparing the dynamics of prestin with its close non-piezoelectric relative, the anion transporter SLC26A9, we identified distinct features unique to prestin, including a relatively unstable anion-binding site that folds upon binding, thereby allosterically modulating the dynamics of the TMD. In contrast, the stability and hydrogen-bond pattern of SLC26A9’s anion-binding site were minimally affected by anion binding, albeit displaying high similarities to prestin in both structure and sequence. Prestin reconstituted in nanodisc exhibited indistinguishable dynamics compared to detergent-solubilized prestin, except for a destabilized TM6 which mediates prestin’s mechanical expansion (Bavi et al., 2021). We observed fraying of the helices involved in the binding site whereas cooperative unfolding of multiple lipid-facing helices including TM6, which may explain prestin’s fast and large-scale motions. These results highlight the significance of the anion-binding site’s folding equilibrium in defining the unique properties of prestin’s voltage dependence and electromotility.

We carried out HDX measurements on dolphin prestin and mouse SLC26A9 solubilized in glyco-diosgenin (GDN) at either pD_read 7.1, 25°C or pD_read 6.1, 0°C (Table 1). The observed HDX rates reported on the stability, as exchange occurred mostly via EX2 kinetics (Appendix 1). Employment of the two conditions increased the effective dynamic range of the HDX measurement to span seven log units, allowing us to determine the stability of both the highly and minimally stable regions within the protein (Hamuro, 2021). To properly combine the two datasets, the stability of the protein should be the same under the two conditions, and this was supported by the exchange rates scaling with k_chem (intrinsic exchange rates) (Bai et al., 1993; Nguyen et al., 2018; Appendix 1—figure 1).

The HDX data were presented in terms of the relevant region with the specific sequence and peptides noted in parentheses (Materials and methods), for example, the N-terminus of prestin’s TM10 (Region_394–397: Peptide_392–397). Although we mostly focused on the anion-binding site, we also obtained comparative thermodynamic information throughout the two proteins (Appendix 2).

Prestin’s anion-binding site is less stable than SLC26A9s

To examine the effect of anion binding to the dynamics of prestin and SLC26A9, we dialyzed the proteins purified in Cl⁻ into a HEPES buffer lacking other anions. Cl⁻ removal resulted in distinct stability changes for prestin and SLC26A9, manifested by significant HDX acceleration for prestin while mild HDX slowing for SLC26A9 (Figure 1, Figure 1—figure supplement 1). These HDX effects indicate that anion binding induced global stabilization for prestin while slight destabilization for SLC26A9 (Figure 1).

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Among the observed HDX responses for prestin, the HDX acceleration at the anion-binding pocket appeared to be the most pronounced and indicates local stabilization induced by anion binding (Figure 2A). In detail, HDX accelerated by 20-fold for the N-termini of both TM3 (Region_136–142: Peptide_134–142 + 9 other peptides) and TM10 (Region_394–397: Peptide_392–397) (Figure 2A, Figure 1—figure supplement 2.22–2.31). This HDX change translates to a difference in free energy of unfolding (${\displaystyle \mathrm{\Delta }\mathrm{\Delta }G}$) by at least 1.8 kcal/mol; ${\displaystyle \mathrm{\Delta }\mathrm{\Delta }{G}_{\mathrm{C}\mathrm{I}\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{d}\mathrm{i}\mathrm{n}\mathrm{g}}^{\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{d}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{s}\mathrm{i}\mathrm{t}\mathrm{e}}=1.8\mathrm{k}\mathrm{c}\mathrm{a}\mathrm{l}/\mathrm{m}\mathrm{o}\mathrm{l}}$. At least four residues in the middle of TM1 exhibited faster HDX (Region_90–101: Peptide_88–101 + 10 other peptides), collectively by 350-fold;${\displaystyle \mathrm{\Delta }\mathrm{\Delta }{G}_{\mathrm{C}\mathrm{I}\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{d}\mathrm{i}\mathrm{n}\mathrm{g}}^{\mathrm{T}\mathrm{M}1}=3.5\mathrm{k}\mathrm{c}\mathrm{a}\mathrm{l}/\mathrm{m}\mathrm{o}\mathrm{l}}$ (Figure 1A, Figure 1—figure supplement 2.6–2.16). The TM1 region with accelerated HDX included L93, Q97, and F101, residues that are known to participate in the binding pocket (Bavi et al., 2021; Ge et al., 2021; Schaechinger et al., 2011).

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SLC26A9 exhibited similar stability as prestin in Cl⁻ for the majority of the TMD, except for the N-terminal TM3 (Region_131–134: Peptide_129–134) which exchanged at least 100-fold slower than that of prestin’s (Region_136–142: Peptide_134–142) (Figure 2). This difference in HDX points to a relatively unstable anion-binding site of prestin as compared to SLC26A9; ${\displaystyle \mathrm{\Delta }\mathrm{\Delta }{G}_{\mathrm{S}\mathrm{L}\mathrm{C}26\mathrm{A}9-\mathrm{p}\mathrm{r}\mathrm{e}\mathrm{s}\mathrm{t}\mathrm{i}\mathrm{n}}^{\mathrm{N}-\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{m}\mathrm{i}\mathrm{n}\mathrm{a}\mathrm{l}\mathrm{T}\mathrm{M}3}=2.8\mathrm{k}\mathrm{c}\mathrm{a}\mathrm{l}/\mathrm{m}\mathrm{o}\mathrm{l}}$, and was also seen in the site-resolved protection factors (PFs) that were obtained by deconvoluting the HDX-MS data using PyHDX (Smit et al., 2021; Figure 2—figure supplement 1).

Compared to the 20- to 350-fold HDX acceleration observed at prestin’s binding site upon Cl⁻ removal, HDX of SLC26A9’s binding pocket was only affected mildly (Figure 2B). These included a slight slowing in HDX for the N-termini of TM3 (Region_131–134: Peptide_129–134) and TM10 (Region_392–395: Peptide_390–395) (Figure 2B). The TM1 (Region_72–92: Peptide_83–92 + 10 other peptides) continued to remain undeuterated even after 27 hr (Figure 2, Figure 1—figure supplement 3.4–3.14), emphasizing the intrinsic high stability of SLC26A9’s anion-binding pocket.

Although the anion-binding pocket is highly conserved and structurally similar across members of the SLC26 family and SLC26A5 families (Figure 2C), mammalian prestin is the only member capable of displaying eletromotility (Santos-Sacchi and Navaratnam, 2022). Hence, the distinct stability responses we observe for dolphin prestin and mouse SLC26A9 point to a prestin’s unique adaptation as a motor protein.

In addition to the binding pocket, we observed stability changes in various regions of the TMDs for prestin and SLC26A9 that may explain their distinct functions. For prestin, anion binding resulted in stabilization for the intracellular cavity but destabilization for regions facing the extracellular milieu (Figure 1C, Figure 1—figure supplement 1A). The stabilizing effects for the cytosol-facing regions were manifested by HDX acceleration upon Cl⁻ removal at the linker between TM2 and TM3, and the intracellular portions of TM7, TM8, and TM9 (Region_128–135, Region_284–294, and Region_354–375) (Figure 1—figure supplement 2.22–2.27, 2.58–2.64, 2.82–2.87). In contrast, HDX slowed for the regions facing the extracellular environment, namely the extracellular ends of TM5b, TM6, and TM7 (Region_250–262 and Region_309–316) (Figure 1—figure supplement 2.45–2.51, 2.66–2.68, 2.71). However, for SLC26A9, anion binding destabilized the cytosol-facing regions, as HDX slowed by ~fivefold upon Cl⁻ removal for the intracellular ends of TM8, TM9, and TM12 (Region_351–369 and Region_440–455) (Figure 1C, Figure 1—figure supplement 1B, Figure 1—figure supplement 3.62–3.63, 3.77–3.82). The distinct thermodynamic consequences of anion binding for prestin and SLC26A9 point to a distinct molecular basis underlying their different functions as a motor and a transporter, respectively.

Anion binding drives the folding of prestin’s binding site

For prestin in HEPES, which adopted the putative apo state, the 20-fold HDX acceleration for the binding site (Figure 2A) is consistent with a process of local destabilization, even unfolding, or increased solvent accessibility as the region becomes exposed to the intracellular water cavity (Bavi et al., 2021). To investigate these possibilities, we measured prestin’s HDX in response to a chaotrope, urea, which destabilizes proteins by interacting with backbone amides (Lim et al., 2009). In a background of 360 mM Cl⁻, the addition of 4 M urea accelerated HDX for the N-terminus of TM3 (Region_137–140: Peptide_134–140) by 20-fold (Figure 3A), suggesting that this region was destabilized and accessible to urea in its exchange-competent state. In apo prestin, however, the PF at the N-terminus of TM3 was unaffected by urea (after accounting for the known ~50% slowing of the k_chem; Lim et al., 2009; Figure 3A), arguing that this region was already unfolded prior to the addition of urea (Ramesh et al., 2019).

Figure 3

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We note that in apparent contradiction to our inference that the N-terminus of TM3 was unfolded in the apo state, its HDX was ~100-fold slower than k_chem. Such apparent PF for an unfolded region has been reported when it is located inside an outer membrane β-barrel, rationalized by the region having a lower effective local concentration of the HDX catalyst, [OD⁻], than in bulk solvent (Zmyslowski et al., 2022; Lin et al., 2022). For prestin, we propose that detergent molecules in the micelle can restrict the access of OD⁻ to amide protons, leading to a local effective pD lower than the bulk solvent and hence producing the apparent PF for the unfolded N-terminus of TM3.

The folding reversibility of the anion-binding site was evaluated by tracking the HDX for 5 min after titrating in Cl⁻ to apo prestin. Deuteration levels for the N-terminus of TM3 (Region_137–140) decreased with increasing Cl⁻ concentration (Figure 3B), suggesting reversible folding upon Cl⁻ binding. Similar behavior was seen in the middle of TM1 (Region_86–101: Peptide_84–101) as Cl⁻ binding stabilized the binding pocket (Figure 3B).

We also examined prestin’s stability in its intermediate states, obtained by replacing Cl⁻ anions with SO₄²⁻ and salicylate (Bavi et al., 2021; Ge et al., 2021). When SO₄²⁻ is the major anion, prestin’s HDX was nearly identical as in Cl⁻, except for a slightly faster HDX at the anion-binding site (Region_128–142 and Region_394–397) at labeling times longer than 10³ s (Figure 4). This mild HDX response suggested a slightly destabilized binding site while the remaining regions retained normal dynamics as in Cl⁻. In the presence of salicylate, HDX slowed across the TMD, with the greatest effect seen at the anion-binding site (10-fold; ${\displaystyle \mathrm{\Delta }\mathrm{\Delta }{G}_{\mathrm{s}\mathrm{a}\mathrm{l}\mathrm{i}\mathrm{c}\mathrm{y}\mathrm{l}\mathrm{a}\mathrm{t}\mathrm{e}-\mathrm{C}\mathrm{l}-}^{\mathrm{b}\mathrm{i}\mathrm{n}\mathrm{d}\mathrm{i}\mathrm{n}\mathrm{g}\mathrm{s}\mathrm{i}\mathrm{t}\mathrm{e}}=1.4\mathrm{k}\mathrm{c}\mathrm{a}\mathrm{l}/\mathrm{m}\mathrm{o}\mathrm{l}}$) (Figure 4), indicating that salicylate binding to prestin globally stabilized the TMD, primarily at the anion-binding site. These stability changes provide a thermodynamic context to the cryo-EM structures (Bavi et al., 2021).

Figure 4

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Prestin in a lipid bilayer exhibits a highly dynamic TM6

We chose to measure the HDX of prestin in GDN micelle to match the cryo-EM conditions (Bavi et al., 2021; Ge et al., 2021; Butan et al., 2022). Structures of GDN-solubilized prestin in Cl⁻ obtained by three research groups are indistinguishable (Cα RMSD <1 Å), demonstrating the reproducibility and robustness of the system.

To evaluate the dynamics in a more native membrane environment, we measured HDX of prestin reconstituted in nanodisc (porcine brain total lipid extract). Except for TM6, HDX for prestin in nanodisc highly resembled that in micelles including the anion-binding pocket (Figure 5). Such high agreement between the folding stability in these two membrane mimetics suggest that our findings on prestin’s anion-binding site and its folding equilibrium are pertinent to prestin residing in a lipid bilayer. This is not surprising because structures for human prestin in GDN and nanodisc are shown to be nearly identical (Cα RMSD = 0.2 Å) (Ge et al., 2021).

Figure 5

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Interestingly, nanodisc-embedded prestin displayed a less stable TM6 for the intracellular portion than prestin in micelles, manifested by the 10-fold HDX increase (Region_275–282: Peptide_273–282 + 2 peptides) (Figure 5, Figure 1—figure supplement 2.53–2.55). TM6 defines the interface between prestin and the lipid bilayer, and has been proposed to mediate area expansion through helical bending (Bavi et al., 2021). The exact role of TM6 in regulating prestin’s conformational cycle is currently under investigation.

Incremental unfolding of prestin’s binding site versus cooperative unfolding of the lipid-facing helices

Our broad HDX time range and dense sampling allowed us to observe effects at the residue level. In particular, the binding site of prestin (Region_128–140 and Region_394–397) exhibited a broad deuterium uptake curve in Cl⁻, indicative of helix fraying where exchange of deuterium occurs from multiple states that differ by one hydrogen bond (Figure 6A). Such HDX pattern is consistent with the associated residues undergoing sequential unfolding with distinct PFs (i.e., stability). Site-resolved PFs obtained using PyHDX (Smit et al., 2021) support that the stability increased residue-by-residue for TM3 for amide protons located further away from the substrate (Figure 6—figure supplement 1). This gradual increase in residue stability along the helices is indicative of helix fraying starting from prestin’s binding site.

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In contrast, we observed much more cooperative unfolding in prestin’s lipid-facing helices, with exchange occurring from one or a few high energy states where a set of hydrogen bonds are broken concertedly. Cooperatively exchanging residues have similar PFs and a characteristic sigmoidal deuterium uptake curve for the associated peptide, as seen in prestin’s intracellular portion of TM6 (Region_275–282: Peptide_273–282 + 4 peptides) (Figure 6A, Figure 1—figure supplement 2.53-2.57).

To characterize the degree of cooperativity for the HDX at the N-terminus of TM3 (Peptide_134–140) and the intracellular portion of TM6 (Peptide_273–282), we fit the deuterium uptake curves as a sum of exponentials (Hamuro, 2021), ${\displaystyle D\left(t\right)=\sum _{i=1}^n\left(1-e^{-k_{i}t}\right)}$, where $k_i$ is the exchange rate and $n$ is the number of exponentials, ranging from one to the number of exchange-competent residues. The value of $n$ was determined by the quality of the fit, evaluated by χ² and having a relative error smaller than one (i.e., standard deviation for $k_i$ less than $k_i$ itself). HDX data for the N-terminus of prestin’s TM3 (Peptide_134–140) were fit with four well-separated exponentials with rates spanning five log units for the four residues (Figure 6A). The need to individually fit each site indicates a lack of cooperativity and helix fraying. In contrast, the peptide representing the intracellular portion of TM6 (Peptide_273–282) has eight residues yet it could be fitted with only three rates spanning less than two log units (Figure 6A). This rather concerted deuterium uptake was independent of the anion substrate identity and also observed for TM1, TM5b, the intracellular portion of TM7, and the N-terminus of TM8 (Figure 1—figure supplement 2.6-2.16, 2.42–2.43, 2.45–2.51, 2.58–2.63, 2.78–2.79).

We define a parameter σ_ΔG to quantify the degree of folding cooperativity. The value of σ_ΔG is calculated as the standard deviation for the free energies of unfolding (∆G) for exchange-competent residues comprising the peptide. When a region folds 100% cooperatively, σ_ΔG is zero as all residues have the same ∆G. As the diversity increases (lower cooperativity), the σ_ΔG value becomes larger. The accuracy of the ∆G determination at residue level can be increased by comparing uptake curves for overlapping peptides and/or deconvoluting isotope envelopes (Hamuro, 2021). Here, we assigned exchange rates ($k_i$), obtained from the fitting method mentioned above, to residues based on the directionality of helix fraying, with residues closer to the end of a helix having faster rates. When there is ambiguity on which rate to assign to a given residue, the geometric mean of the rates was used (Materials and methods). We found that prestin’s intracellular portion of TM6 (Peptide_273–282) has σ_ΔG = 1.1, indicating mild cooperativity, whereas the non-cooperative N-terminal TM3 (Peptide_134–140) has a σ_ΔG = 2.9. This significant decrease in folding cooperativity for helices directly involved in the Cl⁻-binding site likely has functional consequences related to prestin’s electromotility, as discussed below.

Using HDX-MS, we provide novel information on the structural dynamics of prestin in its apo state, for which there is not an associated cryo-EM structure. We demonstrate that prestin displays very similar dynamics in nanodisc as in micelles, except for a destabilized lipid-facing helix TM6 that is critical for mechanical expansion. We have explored the energetic and conformational differences between prestin, a voltage-dependent motor, and its mammlian relative SLC26A9, a representative member of the SLC26 family of anion transporters for which a cryo-EM structure is available. Our data point to major differences in the energetics at the anion-binding site of prestin and SLC26A9 despite their structural similarities. This comparison addresses underlying mechanistic questions related to the unique properties of prestin, the origin of its voltage dependence, and the potential mechanisms that couple charge movements to electromotility.

We showed that prestin displays an unstable binding site, regardless of being in nanodisc or micelles (Figure 5). Upon Cl⁻ unbinding, the binding site unfolds by one helical turn at the electrostatic gap formed by the abutting (antiparallel) short helices TM3 and TM10 (Figures 2A and 3). We measured an increase in local ∆∆G = 1.8–3.5 kcal/mol upon anion binding. This energy difference is within the range of the ∆∆G = 2.4 kcal/mol estimated from having a 60-fold excess of Cl⁻ above the EC₅₀ (6 mM) (Oliver et al., 2001). Similar folding events upon anion binding are absent in SLC26A9 (Figure 2B), pointing to a key role of the bound anion as a structural element in prestin, stabilizing the natural repulsion between TM3- and TM10-positive helical macrodipoles. This phenomenon rationalizes the conundrum that prestin’s voltage dependence requires the proximity of a bound anion to R399.

We find that anion binding to prestin mainly stabilizes the interface between the scaffold and the elevator domains (Figure 1C, Figure 1—figure supplement 1A). This phenomenon is consistent with an elevator-like mechanism during prestin’s conformational transition from the expanded to the contracted state (Bavi et al., 2021). Anion binding stabilizes prestin’s intracellular cavity and slightly destabilizes regions facing the extracellular matrix. This effect can result from changes in solvent exposure, as the intracellular water cavity may shrink as prestin contracts. For SLC26A9, the destabilization upon anion binding at the intracellular cavity likely results from a shift from the outward-facing state to the inward-facing state (Figure 1C, Figure 1—figure supplement 1B), supporting the alternate-access mechanism for this fast transporter (Walter et al., 2019; Chi et al., 2020). Similar HDX changes, that is, increased HDX on the intracellular side while decreased HDX on the extracellular side, have been observed in other transporters during their transition from outward- to inward-facing states (Merkle et al., 2018; Martens et al., 2018). Prestin’s distinct HDX response compared to a canonical transporter is consistent with it being an incomplete anion transporter (Oliver et al., 2001; Schaechinger and Oliver, 2007).

The HDX data for prestin in Cl⁻, SO₄²⁻, and salicylate support an allosteric role for the anion binding at the TM3–TM10 electrostatic gap (Rybalchenko and Santos-Sacchi, 2003; Rybalchenko and Santos-Sacchi, 2008; Song and Santos-Sacchi, 2010; Figure 4). SO₄²⁻ binding leads to shifts in the NLC toward positive potentials, thus stabilizing multiple conformations that are on average more expanded than prestin in Cl⁻ (Bavi et al., 2021; Rybalchenko and Santos-Sacchi, 2008; Muallem and Ashmore, 2006). Since the binding of SO₄²⁻ to prestin is weaker than that of Cl⁻⁶, the slight increase in HDX at the binding site likely reflects more prestin molecules adopting the apo state. Salicylate binding inhibits prestin’s NLC and yet the molecular basis of such inhibition remains obscure (Bavi et al., 2021; Gorbunov et al., 2014). Bavi et al., 2021 showed that binding of salicylate occludes prestin’s binding pocket from solvent and inhibits the movement of TM3 and TM10. This is fully consistent with the 10-fold HDX slowing found for the N-termini of TM3 and TM10 upon salicylate binding as compared to the rest of the protein. Our HDX data, together with results from Bavi et al., 2021, suggest that salicylate likely inhibits prestin’s NLC by restricting the dynamics of the anion-binding site.

We identified helix fraying at the anion-binding site of prestin based on its broad deuterium uptake curve in the presence of Cl⁻, consistent with a multi-state landscape (Figure 6A). This fraying suggests that an increase in the anion concentration would promote helical propensity at TM3 and TM10, and is inconsistent with a cooperative (two-state) model involving an equilibrium between an apo state and a single bound state (Figure 6A). Therefore, we suggest that a two-state model of prestin’s conformational changes with a high energy barrier would be insufficient to explain its fast kinetics, whereas charge movement is facilitated by crossing multiple shallow barriers (Rybalchenko and Santos-Sacchi, 2008; Santos-Sacchi et al., 2009).

We propose that having a Cl⁻-binding site that frays can promote prestin’s fast motor response which is thought to have evolved independently of its voltage-sensing ability (Tan et al., 2012; Tang et al., 2013). While the stability for TM10 is similar for prestin and SLC26A9, the latter protein exhibits a more stable, non-fraying TM3 (Figure 2). Notably, the normally highly conserved Pro136 in mammalian prestin is replaced with a Threonine in SLC26A9 and other vertebrates that express non-electromotile prestin (Figure 2—figure supplement 2). This Pro136Thr substitution, based on our Upside MD simulations, results in a hyper-stabilized TM3 that would otherwise have similar folding stability as TM10 (Figure 2—figure supplement 2). A Pro136Thr mutation in rat prestin also leads to a shift of NLC toward depolarized potentials (Schaechinger et al., 2011). These thermodynamic and functional consequences of having a destabilized TM3 with Pro136, which now has similar stability as TM10, lead us to hypothesize that prestin’s fast mechanical activity may be promoted by having simultaneous fraying of the TM3 and TM10 helices.

Helices that exhibit cooperative unfolding all appear to be lipid-facing helices, including TM6–TM7, TM1, TM5b, and TM8. The region with the most pronounced cooperativity, the intracellular portion of TM6, has a series of glycines including the consecutive G274–G275 pair that underlies the ‘electromotility elbow’, a helical bending contributing to the largest cross-sectional area (expanded conformation) and the thin notch in the micelle (Bavi et al., 2021). Importantly, HDX for prestin in nanodisc reveals a significant stability decrease at TM6 while the remaining regions retained similar dynamics as in micelles. This high sensitivity of TM6 stability to membrane environment, together with the structural consequences of cooperativity, speak to the its significance in prestin’s area expansion. We propose that cooperativity allows for long-range allostery (Hilser and Thompson, 2007) so that the lipid-facing helices, particularly TM6, can adopt large-scale structural rearrangements as induced by voltage sensor movements, thereby achieving rapid electromechanical conversions of prestin. The exact mechanism through which cooperativity contributes to prestin’s electromotility remains a key question.

Based on the structural and allosteric role of Cl⁻ binding at the TM3–TM10 electrostatic gap, we propose a model in which prestin’s conformational changes and electromotility are regulated by the folding equilibrium of the anion-binding site (Figure 6B). In our model, anion binding participates in a local electrostatic balance that includes the positively charged R399 and the positive TM3–TM10 helical macrodipoles. In the apo state, the anion-binding site unfolds due to the electrostatic repulsion between these positively charged groups. Being a buried charge, R399 may exit from the electric field concentrated in the lower dielectric environment of the protein, and move into the solvent region as it lacks a neutralizing anion. This event is coupled to the allosteric expansion of prestin’s membrane footprint (Bavi et al., 2021). Anion binding partially neutralizes the positive electric field at the binding site, an event that is coupled to the residue-by-residue folding for the N-termini of TM3 and TM10 as well as the shortening of the electrostatic gap. This folding event results in a more focused electric field and consequent contraction of prestin’s intermembrane cross-sectional area. At physiologically relevant low Cl⁻ concentration (1.5–4 mM) (McPherson, 2018), prestin’s binding site is likely to be only partially folded. Complete folding may be achieved by membrane potential acting on the TM3–TM10 helical dipoles, which leads to the movement of the voltage sensor across the electric field and the rapid areal expansion for the TMDs (i.e., electromotility).

The raw mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD046965. The atomic structure coordinates have been deposited at the RCSB PDB under accession number 8UC1; and the EM maps have been deposited in the Electron Microscopy Data Bank under accession number EMD-42112. All materials generated during the current study are available from the corresponding author under a materials transfer agreement with The University of Chicago.

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   (2023) PRIDE

   ID PXD046965. Folding of prestin’s anion-binding site and the mechanism of outer hair cell electromotility.

   https://www.ebi.ac.uk/pride/archive/projects/PXD046965
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   (2023) RCSB Protein Data Bank

   ID 8UC1. Cryo-EM structure of dolphin Prestin in low Cl- buffer.

   https://www.rcsb.org/structure/8UC1
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   (2023) Electron Microscopy Data Bank

   ID EMD-42112. Cryo-EM structure of dolphin Prestin in low Cl- buffer.

   https://www.ebi.ac.uk/emdb/EMD-42112

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Author details

1. Xiaoxuan Lin

   1. Center for Mechanical Excitability, The University of Chicago, Chicago, United States
   2. Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5356-9135

2. Patrick R Haller

   1. Center for Mechanical Excitability, The University of Chicago, Chicago, United States
   2. Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, United States

   Contribution

   Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

3. Navid Bavi

   1. Center for Mechanical Excitability, The University of Chicago, Chicago, United States
   2. Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, United States

   Contribution

   Validation, Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

4. Nabil Faruk

   Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, United States

   Contribution

   Validation, Investigation, Visualization, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

5. Eduardo Perozo

   1. Center for Mechanical Excitability, The University of Chicago, Chicago, United States
   2. Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, United States
   3. Institute for Neuroscience, The University of Chicago, Chicago, United States
   4. Institute for Biophysical Dynamics, The University of Chicago, Chicago, United States

   Contribution

   Conceptualization, Resources, Software, Formal analysis, Supervision, Funding acquisition, Validation, Visualization, Methodology, Project administration, Writing – review and editing

   For correspondence

   eperozo@uchicago.edu

   Competing interests

   No competing interests declared

6. Tobin R Sosnick

   1. Center for Mechanical Excitability, The University of Chicago, Chicago, United States
   2. Department of Biochemistry and Molecular Biology, The University of Chicago, Chicago, United States
   3. Institute for Biophysical Dynamics, The University of Chicago, Chicago, United States
   4. Prizker School for Molecular Engineering, The University of Chicago, Chicago, United States

   Contribution

   Conceptualization, Resources, Data curation, Software, Formal analysis, Supervision, Funding acquisition, Validation, Visualization, Methodology, Project administration, Writing – review and editing

   For correspondence

   trsosnic@uchicago.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2871-7244

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