Structure and substrate ion binding in the sodium/proton antiporter PaNhaP.

Sodium/proton antiporters maintain intracellular pH and sodium levels. Detailed structures of antiporters with bound substrate ions are essential for understanding how they work. We have resolved the substrate ion in the dimeric, electroneutral sodium/proton antiporter PaNhaP from Pyrococcus abyssi at 3.2 Å, and have determined its structure in two different conformations at pH 8 and pH 4. The ion is coordinated by three acidic sidechains, a water molecule, a serine and a main-chain carbonyl in the unwound stretch of trans-membrane helix 5 at the deepest point of a negatively charged cytoplasmic funnel. A second narrow polar channel may facilitate proton uptake from the cytoplasm. Transport activity of PaNhaP is cooperative at pH 6 but not at pH 5. Cooperativity is due to pH-dependent allosteric coupling of protomers through two histidines at the dimer interface. Combined with comprehensive transport studies, the structures of PaNhaP offer unique new insights into the transport mechanism of sodium/proton antiporters.

Introduction

The Na+/H+ antiporter NhaP from Pyrococcus abyssi (PaNhaP) exchanges protons against sodium ions across the cell membrane. PaNhaP is a functional homologue of the human Na+/H+ exchanger NHE1, which controls intracellular pH and Na+ concentration. NHE1 is an important drug target (Karmazyn et al., 1999), but its structure and detailed mode of action are unknown. Transport mechanisms of eukaryotic membrane proteins are conserved in the more robust prokaryotic transporters from thermophilic bacteria and archaea (Yamashita et al., 2005; Boudker et al., 2007; Lee et al., 2013). High-resolution structures of such homologues are of great value for understanding the mechanisms of cation/proton antiport, provided that (i) the transported substrate ions are resolved, (ii) structures of the same transporter are available in different conformations, and (iii) kinetic data of substrate binding and transport are available. In this paper we report the structure of the electroneutral Na+/H+ antiporter PaNhaP from the hyperthermophilic archaeon P. abyssi in two different conformations at pH 4 and pH 8, with the substrate ion resolved at pH 8. We show that, like NHE1, transport by PaNhaP is cooperative in a pH-dependent manner, indicating a pH-dependent allosteric interaction of protomers in the dimer.

The first structure of a cation-proton antiporter (CPA) revealed that Escherichia coli Na+/H+ NhaA (EcNhaA) is a dimer in the membrane (Williams et al., 1999). The 6 Å map of EcNhaA resolved 12 trans-membrane helices (TMH) in the protomer, arranged in a 6-helix bundle, plus a row of six TMHs at the dimer interface (Williams, 2000). A membrane dimer was also found for the NhaP1 antiporter from Methanocaldococcus jannaschii (MjNhaP1) (Vinothkumar et al., 2005; Goswami et al., 2011; Paulino and Kühlbrandt, 2014). MjNhaP1, PaNhaP and the medically important NHE1 belong to the CPA1 subfamily (Brett et al., 2005) of antiporters, which exchange Na+ and protons with 1:1 stoichiometry and are thus electroneutral. By contrast, EcNhaA and TtNapA from Thermus thermophilus, as well as the eukaryotic NHA1-2 and AtChx1 (Brett et al., 2005), belong to the CPA2 subfamily of electrogenic antiporters, which exchange one Na+ against two protons. Neither the x-ray structure of EcNhaA (Hunte et al., 2005) nor that of TtNapA (Lee et al., 2013) resolved the substrate ion.

Results

Overall architecture of PaNhaP

Crystals of seleno-methionine derivatized PaNhaP grown at pH 8 diffracted isotropically to 3.15 Å resolution. The structure was solved by SAD (Tables 1 and 2). Twelve out of the 14 SeMet positions in the asymmetric unit containing one PaNhaP dimer were identified (Figure 1—figure supplement 1). Seen from the cytoplasm, the PaNhaP dimer is roughly rectangular, with a long axis of 90 Å and a short axis of 53 Å (Figure 1A, Figure 1—figure supplement 2A). Each protomer has 13 TMHs (H1-H13) connected by short loops or helices on the membrane surface. H4-6 and H11-13 form the 6-helix bundle, while H1-3 and H7-10 form the dimer interface. H1-6 and H8-13 are two halves of an inverted 6-helix repeat, connected by H7. Several helices are highly tilted, especially H7 and H8, which include angles of more than 45° with the membrane normal while others, in particular H6 and H10, are bent. H5 and H12 in the 6-helix bundle are discontinuous. Their cytoplasmic and extracellular halves (referred to as H5C, H5E and H12C, H12E respectively) are each connected by unwound stretches with antiparallel orientation, which cross one another in the centre of the protomer (Figure 1, Figure 1—figure supplement 2). The membrane surfaces are marked by three short amphipathic helices connecting H3 to H4 on the cytoplasmic side, H6 to H7 and H10 to H11 on the extracellular side. H10 protrudes by 11 Å on the cytoplasmic surface, and the helix hairpin connecting H12 to H13 protrudes by about 7 Å on the extracellular side. The loops connecting helices H1 to H2 and H8 to H9 are ∼10 Å below the cytoplasmic or extracellular surface (Figure 1B, Figure 1—figure supplement 2B).

Figure 1—figure supplement 1.

Experimental electron density map of PaNhaP.

The structure was solved at 3.15  Å by SeMet-SAD. Twelve of the 14 SeMet positions (grey spheres) in the asymmetric unit were resolved in the anomalous difference map (purple densities at 4σ). SeMet positions werfe used to generate an initial map (blue density at 1σ) and to trace the polypeptide chain. (A) Cytoplasmic view of PaNhaP dimer. (B) Side view with the extracellular side above.

DOI: http://dx.doi.org/10.7554/eLife.03579.006

Figure 1.

PaNhaP at pH 8.

(A) Cytoplasmic view of the PaNhaP dimer. Helices H1 to H13 are color-coded and numbered in one protomer. In the other protomer only the partly unwound helices H5 and H12 are coloured. (B) Side view with the C-terminus of helix H13 on the cytoplasmic side.

DOI: http://dx.doi.org/10.7554/eLife.03579.005

The structure was solved at 3.15  Å by SeMet-SAD. Twelve of the 14 SeMet positions (grey spheres) in the asymmetric unit were resolved in the anomalous difference map (purple densities at 4σ). SeMet positions werfe used to generate an initial map (blue density at 1σ) and to trace the polypeptide chain. (A) Cytoplasmic view of PaNhaP dimer. (B) Side view with the extracellular side above.

DOI: http://dx.doi.org/10.7554/eLife.03579.006

Cartoon representation of PaNhaP with helices shown as cylinders. (A) Cytoplasmic and (B) side view of the dimer colour-coded as in the main figure.

DOI: http://dx.doi.org/10.7554/eLife.03579.007

Side view of the dimer with hydrophilic cavities coloured by surface potential. The substrate-binding site is accessible from the cytoplasmic side through the cytoplasmic funnel and a narrow polar channel. Access from the extracellular side is blocked. One water molecule is trapped in an enclosed polar cavity near the ion-binding site.

DOI: http://dx.doi.org/10.7554/eLife.03579.008

Figure 1—figure supplement 2.

X-ray structure of PaNhaP.

Cartoon representation of PaNhaP with helices shown as cylinders. (A) Cytoplasmic and (B) side view of the dimer colour-coded as in the main figure.

DOI: http://dx.doi.org/10.7554/eLife.03579.007

On the cytoplasmic side of the protomer, a solvent-filled ∼16 Å-deep funnel, lined by H3, H5C, H6, and H10, penetrates to the centre of the protomer between the 6-helix bundle and the dimer interface (Figure 1—figure supplement 3, Video 1). A second, narrow polar channel, lined by the unwound stretches of H5C, H12C and the cytoplasmic halves of H6 and H13, extends from the cytoplasmic surface to the region near the deepest point of the funnel (Figure 1—figure supplement 3, Video 1). On the extracellular side, a deep cavity on the twofold axis of the dimer, lined by interface helices H1, H3, H8 and H10, extends ∼27 Å into the hydrophobic protein interior. Electron density in this cavity indicated bound lipid (Figure 2), which was identified by thin-layer chromatography as phosphatidyl ethanolamine (PE), carried over from the E. coli expression host. The lipid stretches from the 6-helix bundle of one protomer to the interface helices H3 and H10 of the other, providing a hydrophobic link between them. The cavity is large enough to accommodate two lipids, only one of which was resolved in the dimer (Figure 2). The surface potential of the dimer indicates clusters of charged residues on both sides of the membrane (Figure 2—figure supplement 1). The cytoplasmic ends of H10 and H7 are positively charged, carrying a total of seven lysine and arginine residues. The deep funnel on the cytoplasmic surface is lined by negative charges, which would attract positively charged substrate ions.

Figure 1—figure supplement 3.

Hydrophilic cavities in PaNhaP.

Side view of the dimer with hydrophilic cavities coloured by surface potential. The substrate-binding site is accessible from the cytoplasmic side through the cytoplasmic funnel and a narrow polar channel. Access from the extracellular side is blocked. One water molecule is trapped in an enclosed polar cavity near the ion-binding site.

DOI: http://dx.doi.org/10.7554/eLife.03579.008

Video 1.

Movie of PaNhaP monomer with hydrophilic cavities.

DOI: http://dx.doi.org/10.7554/eLife.03579.009

Figure 2.

Hydrophobic extracellular cavity with bound lipid.

(A) One lipid molecule (PE, green) in the cavity between the two protomers in the dimer contributes to the hydrophobic contacts across the dimer interface. The extracellular surface is slightly negatively charged. (B) The alkyl chain of the lipid extends to the center of the molecule. (C) The lipid-facing surface of the central cavity is mainly hydrophobic. The surface potential was calculated at pH 7.0 by APBS.

DOI: http://dx.doi.org/10.7554/eLife.03579.010

The surface potential of the PaNhaP dimer was calculated at pH 4, pH 6, and pH 8 by APBS and visualized in PyMOL. At pH 4 the cytoplasmic surface is strongly positively charged. At pH 6, the extracellular surface and the cytoplasmic funnel are largely neutral. At pH 8 the extracellular surface is predominantly negatively charged and the cytoplasmic funnel is strongly negatively charged, inhibiting the release of substrate ions. The ion-binding site is located at the bottom of the cytoplasmic funnel.

DOI: http://dx.doi.org/10.7554/eLife.03579.011

Figure 2—figure supplement 1.

pH-dependent charge distribution.

The surface potential of the PaNhaP dimer was calculated at pH 4, pH 6, and pH 8 by APBS and visualized in PyMOL. At pH 4 the cytoplasmic surface is strongly positively charged. At pH 6, the extracellular surface and the cytoplasmic funnel are largely neutral. At pH 8 the extracellular surface is predominantly negatively charged and the cytoplasmic funnel is strongly negatively charged, inhibiting the release of substrate ions. The ion-binding site is located at the bottom of the cytoplasmic funnel.

DOI: http://dx.doi.org/10.7554/eLife.03579.011

The ion-binding site

Crystals of PaNhaP grown at pH 8 soaked with thallium acetate diffracted to 3.2 Å (Table 1). Two thallium ions were identified in the dimer by anomalous scattering, one each near the deepest point of the cytoplasmic funnel in the two protomers (Figure 1—figure supplement 3, Video 1, Figure 3A). The Tl+ ions were located ∼14 Å below the cytoplasmic surface and ∼22 Å from the extracellular surface. The ion-binding site is accessible from the cytoplasm but not from the extracellular side, so that the structure shows the inward-open conformation of PaNhaP (Figure 1—figure supplement 3, Video 1). Like Na+ and Li+, but unlike K+, Tl+ is a substrate of PaNhaP (Figure 4). The thallium ions and their surroundings provide a unique view of the ion-binding site and substrate ion coordination in sodium-proton antiporters (Figure 3B,C). Three acidic side chains in three different TMHs contribute to substrate ion-binding. The carboxyl groups of Glu73 in H3 and Asp159 in H6 coordinate the substrate ion directly. Asp130 in the unwound stretch of H5 interacts with the ion via a bound water molecule (Figure 3). The main-chain carbonyl of Thr129, likewise in the unwound stretch of H5, and the hydroxyl side chain of Ser155 in H6 provide two additional ligands, bringing the total up to five. The ion coordination geometry is that of a distorted trigonal bipyramid, with Asp159, Ser155 and the water molecule forming a triangle around the central substrate ion, and the Thr129 main chain carbonyl and Glu73 at the tips of the bipyramid (Figure 3B,C).

Table 1.

Data collection and refinement statistics

DOI: http://dx.doi.org/10.7554/eLife.03579.003

	SeMet @ pH 8	Thallium @ pH 8	Native @ pH 4
Data collection		SLS PXII
Wavelength	0.979	0.979	0.978
Space group	P21	P21	P64
Cell dimensions
a, b, c (Å)	54.5, 107.9, 107.9	54.1, 107.4, 99.8	109.6, 109.6, 209.6
α, β, γ (°)	90.0, 95.2, 90.0	90.0, 96.4, 90.0	90.0, 90.0, 120.0
Resolution (Å)	48.5–3.15 (3.35–3.15)	49.6–3.20 (3.40–3.20)	48.6–3.50 (3.72–3.50)
Rpim	0.033 (0.503)	0.038 (0.622)	0.021 (0.486)
I / σI	11.9 (1.5)	13.4 (1.8)	19.9 (1.9)
CC*	1.000 (0.943)	1.000 (0.936)	1.000 (0.906)
Completeness (%)	99.5 (99.2)	99.6 (99.4)	100.0 (100.0)
Multiplicity	10.8 (10.4)	17.1 (17.4)	9.2 (9.1)
Refinement
Resolution (Å)	48.5–3.15 (3.35–3.15)	49.6–3.20 (3.40–3.20)	48.6–3.5 (3.72–3.5)
Unique reflections	38,952	34,763	33,232
Reflections in test set	2111	1884	1782
Rwork/Rfree (%)	23.8/27.8 (31.8/39.9)	24.8/29.5 (35.9/43.4)	24.1/26.4 (31.8/35.6)
CC(work)/CC(free)	0.843/0.898 (0.842/0.760)	0.861/0.754 (0.813/0.713)	0.791/0.935 (0.749/0.617)
Wilson B-Factor (Å2)	133	81	146
No. atoms in AU	6715	6651	6592
Protein	6582	6560	6560
Ligands	129	81	31
Water	4	10	1
r.m.s. deviations:
Bond lengths (Å)	0.003	0.003	0.009
Bond angles (°)	0.758	0.714	1.002

Figure 3.

Substrate ion coordination in PaNhaP.

(A) Section view of the ion-binding site and interface region of PaNhaP from the cytoplasmic side. Interface helices of the two protomers are shown in blue and beige, respectively. The acidic side chains of Glu73, Asp159, a water molecule held by Asp130, the hydroxyl group of Ser155 and the main-chain carbonyl of Thr129 coordinate the substrate ion. The anomalous density for the Tl+ ion (grey sphere) in the substrate-binding site between helix H3, H6 and the unwound stretch of H5 is shown in magenta at 4σ. The 3σ omit map for the H2O molecule next to Tl+ is green. The water molecule near Glu154 and Asn158 is not directly involved in ion coordination. (B, C) Detailed views of the substrate-coordinating residues from the extracellular and cytoplasmic side, respectively. (D) Side view of core helices and substrate-binding residues in the 6-helix bundle.

DOI: http://dx.doi.org/10.7554/eLife.03579.012

Figure 4.

Ion selectivity of PaNhaP.

Ion selectivity was determined by acridine orange fluorescence at pH 6. Na+, Li+, Tl+ are transported by PaNhaP, K+ is not.

DOI: http://dx.doi.org/10.7554/eLife.03579.013

The second, narrow polar channel next to the cytoplasmic funnel (Figure 1—figure supplement 3, Video 1) leads to an enclosed polar cavity near Asp93, Thr129, Asn158 and the ion pair Glu154/Arg337, which are highly conserved in the CPA1 antiporters (Goswami et al., 2011). A water molecule in the enclosed cavity links the functionally important groups that surround it. The Glu154/Arg337 ion bridge and Thr129 separate the cavity from the narrow polar channel. The ion-binding site at the end of the cytoplasmic funnel is accessible from both the polar cavity and the narrow polar channel via Thr129, Ser155, and Asn158 (Figure 1—figure supplement 3, Video 1, Figure 3).

Conformational changes at pH 4

The structure of PaNhaP crystals grown at pH 4 was determined at 3.5 Å (Table 1) by molecular replacement. As at pH 8, the ion-binding site is accessible from the cytoplasm via the cytoplasmic funnel, but not from the extracellular side. Both structures therefore show an inward-open state. In contrast to the pH 8 structure, the second narrow polar channel is blocked at pH 4 by rearrangements of the surrounding residues Ile151, Phe355, Gly359. The most conspicuous differences to the pH 8 structure are observed near the dimer interface. At pH 8, the His292 sidechains in H10 of the two protomers form a 15 Å chain of hydrogen bonds with the Glu233 residues near the cytoplasmic ends of H8 (Figure 5, Video 2, Figure 5—figure supplement 1). At pH 4, each of the two histidines moves by 6–8 Å, apparently due to electrostatic repulsion upon protonation at acidic pH (Figure 2—figure supplement 1). This pH-induced conformational change disrupts the chain of hydrogen bonds linking the two protomers (Figure 5—figure supplement 1). Other major pH-induced changes are found in the ion bridges linking the protomers across the dimer interface (Figure 5—figure supplement 1). At pH 8, Arg25/Glu228 and Arg26/Asp231 connect the cytoplasmic ends of H8 and H1, while the Glu8/Arg249 bridge links the extracellular ends of these helices. At pH 4, all six ion pairs break, apparently due to partial protonation of the acidic sidechains, so that each protomer tilts away from the dimer interface (Video 2, Figure 5—figure supplement 1).

Figure 5.

pH-induced conformational changes in the PaNhaP dimer.

(A) cytoplasmic view, (B) side view as in Figure 1. At pH 4 (red), helix H4 moves towards the cytoplasm by 1.5 Å. Within the 6-helix bundle, the extracellular ends of helix H5E and H6 move towards H12 by ∼1.5 Å. Helix H11 and H13 tilt by about 2–3° each, such that the cytoplasmic end of helix H11 moves towards H12C, which shifts by a similar amount in the same direction. The extracellular end of helix H12E moves towards helix H3 by ∼3 Å. The rmsd between the structures at pH 8 and pH 4 is 1.57 Å.

DOI: http://dx.doi.org/10.7554/eLife.03579.014

The pH 8 structure (transparent) is superposed on the pH 4 structure. Red and black arrows indicate mainchain and sidechain movements, respectively. (A) At pH 4, His292 at the dimer interface moves by 6–8 Å from its pH 8 position towards the centre of H3. At pH 8, the two His292 in the dimer form a line of hydrogen bonds with glutamates Glu233 on either side (dashed lines). Protonation at pH 4 disrupts the hydrogen bond network, so that the His sidechains and H10 move towards the helix bundle. (B) Six salt bridges between the cytoplasmic end of H1 and H7/8 present at pH 8 break at pH 4. The N-terminus of H1 on the extracellular side becomes more ordered at pH 4 and moves towards H8.

DOI: http://dx.doi.org/10.7554/eLife.03579.015

(A) In protomer A, Asp130 in the unwound stretch of H5 and Glu73 in H3 move towards the ion-coordinating Asp159. The ion-binding site in protomer B changes only slightly from pH 8 to pH 4. (B) At pH 8, Tyr31 is within hydrogen bonding distance of Asp130 in the substrate-binding site. At pH 4, the two residues do not interact. The rmsd between the pH 8 and pH 4 structures is 1.6 Å.

DOI: http://dx.doi.org/10.7554/eLife.03579.016

Video 2.

pH-induced conformational changes in PaNhaP.

A morph between the pH 4 and pH 8 structures reveals only small changes in the 6-helix bundle, but significant rearrangements at the dimer interface. Six ion bridges that lock the two protomers together at pH 8 break at pH 4. As a result, the two protomers tilt away from each other at lower pH. His292 has a pivotal role in the allosteric pH-dependent protomer interaction. At pH 4, the protonated His292 side chains on the cytoplasmic side of the dimer interface repel one another by electrostatic repulsion, resulting in a ∼7 Å movement that disrupts the hydrogen-bonding network with Glu233.

DOI: http://dx.doi.org/10.7554/eLife.03579.017

Figure 5—figure supplement 1.

pH-induced conformational changes at the dimer interface.

The pH 8 structure (transparent) is superposed on the pH 4 structure. Red and black arrows indicate mainchain and sidechain movements, respectively. (A) At pH 4, His292 at the dimer interface moves by 6–8 Å from its pH 8 position towards the centre of H3. At pH 8, the two His292 in the dimer form a line of hydrogen bonds with glutamates Glu233 on either side (dashed lines). Protonation at pH 4 disrupts the hydrogen bond network, so that the His sidechains and H10 move towards the helix bundle. (B) Six salt bridges between the cytoplasmic end of H1 and H7/8 present at pH 8 break at pH 4. The N-terminus of H1 on the extracellular side becomes more ordered at pH 4 and moves towards H8.

DOI: http://dx.doi.org/10.7554/eLife.03579.015

Other significant pH-induced differences occur at the N-terminus of the protomer, where residues 3–6 become ordered at pH 4, so that H1 extends by one turn, and shifts by 3 Å towards the cytoplasmic side (Figure 5—figure supplement 1). In the ion-binding site itself, the sidechain of Asp130 in protomer A moves by 2.7 Å into the space that is occupied by the substrate ion at pH 8 (Figure 5—figure supplement 2A). This movement, which is not observed in the other protomer (Figure 5—figure supplement 2B), could displace a bound substrate ion or prevent ion binding. At pH 4, the conserved Asn158 that interacts with Asp93 at pH 8 moves by ∼2.5 Å towards the ion-coordinating Asp159 in protomer A, forming a H-bond network with Thr129 and the main-chain carbonyls of Glu154 and Ser155. In this way, the reorientation of Asn158 may regulate access to the ion-binding site through the narrow polar channel (Figure 1—figure supplement 3, Video 1).

Figure 5—figure supplement 2.

pH-induced conformational changes in the substrate binding site.

(A) In protomer A, Asp130 in the unwound stretch of H5 and Glu73 in H3 move towards the ion-coordinating Asp159. The ion-binding site in protomer B changes only slightly from pH 8 to pH 4. (B) At pH 8, Tyr31 is within hydrogen bonding distance of Asp130 in the substrate-binding site. At pH 4, the two residues do not interact. The rmsd between the pH 8 and pH 4 structures is 1.6 Å.

DOI: http://dx.doi.org/10.7554/eLife.03579.016

A chain of hydrogen bonds stretches from Glu290 in H10 via His75 near the cytoplasmic end of H3 to Glu73, the only ion-coordinating sidechain from one of the interface helices (Figure 3A). This residue most likely relays allosteric changes from the dimer interface to the ion-binding site. An opening of the ion bridges that link H1 and H8 and the movement of the adjacent H2 in the pH 8 to pH 4 transition is likely to affect substrate binding via Tyr31, which is within H-bonding distance of the substrate-coordinating Asp130 (Figure 5—figure supplement 2). In this way, the conformational changes caused by repulsion of the protonated histidines 292 at the dimer interface are relayed to the ion-binding site to modulate the Na+ binding affinity in a pH-dependent manner (Figure 6).

Figure 6.

Transport activity of PaNhaP.

(A) pH dependence of transport activity determined by 22Na+ uptake with inside-acidic PaNhaP proteoliposomes. The antiporter is active at pH 5 and pH 6; at pH 4 and pH 7 transport drops to background level. (B) Concentration-dependent 22Na+-uptake by inside-acidic PaNhaP proteoliposomes at pH 5 gives a vmax of 87.9 ± 7.5 nmol · min−1 · mg−1 at room temperature, indicating a transport rate of 4.4 Na+ ions per protomer per minute. At pH 5 the Hill coefficient (nh) is 1.1 ± 0.30, indicating non-cooperative transport. (C) At pH 6, transport is cooperative, with a Hill coefficient of 1.9 ± 0.26, indicating allosteric coupling of the two ion-binding sites in the dimer. vmax at room temperature decreases to 16.5 ± 0.5 nmol · min−1 · mg−1.

DOI: http://dx.doi.org/10.7554/eLife.03579.018

(A) Sodium efflux was measured under symmetrical pH conditions by acridine orange fluorescence with PaNhaP reconstituted into proteoliposomes. Transport activity of PaNhaP drops towards pH 7, consistent with 22Na uptake measurements (Figure 4). Transport was not affected by 100 nM valinomycin (B, red curve), indicating that PaNhaP is electroneutral.

DOI: http://dx.doi.org/10.7554/eLife.03579.019

(A) Eadie–Hofstee transformation of kinetic data at pH 5 shows a linear correlation typical for Michaelis–Menten kinetics. The data point at 5 µM sodium concentration was omitted from the linear regression. (B) Eadie–Hofstee transformation of pH 6 data results in a concave curve, indicating homotropic activation.

DOI: http://dx.doi.org/10.7554/eLife.03579.020

pH-dependent cooperativity

22Na uptake into reconstituted PaNhaP proteoliposomes is strongly pH-dependent (Figure 6A). Transport activity was highest at pH 5, dropping to about 75 % at pH 6, 20 % at pH 7, and to background level at pH 8. At pH 4, the activity was about 5 % of the peak value at pH 5, resulting in a roughly bell-shaped pH profile. Sodium uptake measurements performed with reconstituted, inside-acidic proteoliposomes (Figure 6B,C) or sodium efflux measurements under symmetrical pH conditions (Figure 6—figure supplement 1A) showed comparable transport behaviour at basic pH. Valinomycin had no effect on the transport rate (Figure 6—figure supplement 1B), demonstrating that PaNhaP is electroneutral. Measurements of 22Na+ uptake at pH 6 revealed clear positive cooperativity, with a Hill coefficient of 1.9 (Figure 6C, Figure 6—figure supplement 2B). Since PaNhaP forms stable dimers in detergent solution and each protomer binds only one substrate ion at a time, this indicates that the interaction of protomers across the dimer interface is allosteric, such that at pH 6, an ion binding to one protomer increases the binding affinity of the other, as indicated by the K0.5 value of 25 µM (Figure 6C), compared to the Km of 506 µM at pH 5 (Figure 6C). At the pH 5 activity maximum the Hill coefficient was ∼1, indicating non-cooperative transport (Figure 6B, Figure 6—figure supplement 2). Note that the pH-dependent allosteric change of the dimer is different from the inside-open to outside-open transition in the transport cycle of the protomer.

Figure 6—figure supplement 1.

Sodium efflux measurements.

(A) Sodium efflux was measured under symmetrical pH conditions by acridine orange fluorescence with PaNhaP reconstituted into proteoliposomes. Transport activity of PaNhaP drops towards pH 7, consistent with 22Na uptake measurements (Figure 4). Transport was not affected by 100 nM valinomycin (B, red curve), indicating that PaNhaP is electroneutral.

DOI: http://dx.doi.org/10.7554/eLife.03579.019

Figure 6—figure supplement 2.

Eadie-Hofstee plots.

(A) Eadie–Hofstee transformation of kinetic data at pH 5 shows a linear correlation typical for Michaelis–Menten kinetics. The data point at 5 µM sodium concentration was omitted from the linear regression. (B) Eadie–Hofstee transformation of pH 6 data results in a concave curve, indicating homotropic activation.

DOI: http://dx.doi.org/10.7554/eLife.03579.020

Transport activity

At room temperature, vmax of PaNhaP at the pH 5 activity maximum was 87.9 nmol · min−1 · mg−1, giving a transport rate of 4.4 ± 0.4 Na+ ions per minute for each protomer. Between 20°C and 45°C, vmax grew exponentially by a factor of 2.1 for every 5°C rise in temperature (Figure 7A,B) according to the Arrhenius equation. Extrapolation to 100°C, the physiological temperature for P. abyssi, suggests a rate of about 5000 ions per second. Note that temperature affects the transport rate but not substrate binding (Figure 7C,D).

Figure 7.

Temperature dependence of PaNhaP.

(A, B) At pH 6 transport activity increases by a factor of 2.1 for every 5°C rise in temperature, as measured by sodium efflux under symmetrical pH. The slight rise in fluorescence towards longer times at 40°C and above in A is due to increasing proton leakage of the proteoliposomes. (C, D) Effect of temperature on substrate affinity at 25°C (empty dots) and 30°C (filled squares) measured by ΔpH-driven sodium uptake in proteoliposomes using Acridine orange fluorescence. In contrast to vmax, Km does not change much with increasing temperature (1.56 ± 0.11 mM at 25°C; 1.85 ± 0.49 mM at 30°C).

DOI: http://dx.doi.org/10.7554/eLife.03579.021

Residues involved in substrate binding were replaced and the transport activity of mutant proteins was measured in proteoliposomes. Replacement of both ion-coordinating aspartates (Asp130 and Asp159) by serine abolished transport completely (Figure 8A), whereas mutation of Glu73 to alanine increased the activity (Figure 8B), most likely because the substrate ion is released more readily from the binding site. Changing Ser155 to alanine had no significant effect, but mutation of Thr129 to valine that takes this position in eukaryotic CPA1 transporters (Goswami et al., 2011), reduced the activity significantly. This was surprising, because Thr129 coordinates the substrate ion not by its sidechain but by its main-chain carbonyl. However the Thr129 sidechain is a potential interaction partner of the conserved Asn158 that may control access to the ion-binding site through the narrow polar channel. A hydrophobic valine in place of Thr129 would interrupt the local network of hydrogen bonds, which could affect ion binding or proton translocation. A mutant in which His292 was replaced by cysteine migrates as a dimer under oxidizing conditions in SDS-PAGE (Figure 8—figure supplement 1A). The activity of the crosslinked dimer was 35 % of wildtype (Figure 8—figure supplement 1B). Under reducing conditions, when the disulfide bridge between the protomers is broken, activity increases to 150 % of wildtype, highlighting the importance of this position for the regulation of transport.

Figure 8.

Transport activity of binding site mutants.

Sodium efflux from proteoliposomes at pH 6 was measured to investigate PaNhaP mutants. Antiport activity establishes a ΔpH across the membrane, which results in acridine orange fluorescence quenching. (A) Mutation of Asp130 or Asp159 to serine abolishes transport activity. (B) Replacement of Thr129 by valine, as in eukaryotic antiporters, reduces the transport activity. Replacement of Glu73 by alanine increases activity significantly, whereas exchanging Ser155 against alanine has no effect compared to wildtype.

DOI: http://dx.doi.org/10.7554/eLife.03579.022

(A) Mutation of His292 to cysteine results in a prominent dimer band under oxidizing conditions, as protomers are crosslinked by covalent disulfide bonds across the dimer interface both in detergent and in proteoliposomes (PL). Addition of reducing agent (10 mM DTT) breaks the disulfide bonds between crosslinked protomers. Asterisks mark the PaNhaP monomer and dimer on SDS-PAGE. (B) Transport measurements indicate a 60 % drop in activity of cross-linked PaNhaP dimers compared to wildtype. Under reducing conditions the activity of the H292C mutant is 50 % higher than wildtype, due to weaker protomer interactions.

DOI: http://dx.doi.org/10.7554/eLife.03579.023

Figure 8—figure supplement 1.

Interface crosslinks.

(A) Mutation of His292 to cysteine results in a prominent dimer band under oxidizing conditions, as protomers are crosslinked by covalent disulfide bonds across the dimer interface both in detergent and in proteoliposomes (PL). Addition of reducing agent (10 mM DTT) breaks the disulfide bonds between crosslinked protomers. Asterisks mark the PaNhaP monomer and dimer on SDS-PAGE. (B) Transport measurements indicate a 60 % drop in activity of cross-linked PaNhaP dimers compared to wildtype. Under reducing conditions the activity of the H292C mutant is 50 % higher than wildtype, due to weaker protomer interactions.

DOI: http://dx.doi.org/10.7554/eLife.03579.023

Discussion

Ion coordination

The trigonal bipyramidal coordination geometry of sodium ions observed in PaNhaP is not uncommon in membrane transporters (Penmatsa et al., 2013). The same geometry is found in c-rings of Na+-translocating F-type ATPase of I. tartaricus and F. nucleatum (Meier et al., 2009; Schulz et al., 2013), which, like the archaeal CPA1 antiporters, bind and release Na+ in rapid exchange. Although the ion radius of monovalent Tl+ (1.5 Å) is similar to that of K+ (1.44 Å) and larger than that of Na+ (1.12 Å) (Shannon, 1976; Cotton and Wilkinson, 1988), Tl+ is able to replace Na+ in PaNhaP. The same has been found for the Na+-dependent aspartate transporter GltPh (Boudker et al., 2007), the mammalian glutamate transporter EAAC1 (Tao et al., 2008) and fructose-1,6-biphosphatase (Villeret et al., 1995). In GltPh, Na+ but not K+ competes for Tl+ binding, and Tl+ inhibits Na+-driven aspartate transport (Boudker et al., 2007). Coordination geometry and ligand distances for Tl+ in PaNhaP are similar to those typically found for protein-bound Na+ in the PDB (Harding, 2002). The larger ion radius of Tl+ may account for the lower transport rate in PaNhaP. However, Tl+ is a much better substrate than K+, which is not transported at all (Figure 4). The selectivity for Na+ over K+ is reminiscent of the striking selectivity of sodium channels, which is thought to be related to ion solvation (Roux et al., 2011). Presumably, the same principle applies to the Na+/H+ antiporters. The water molecule between the sidechain of Asp130 and Tl+ indicates that the bound substrate ion retains part of its hydration shell, as complete dehydration is energetically unfavourable.

In PaNhaP, all ion-binding residues are found in the first half of the inverted repeat. Interestingly, the structure and interaction of the ion-coordinating Asp159 and Ser155 in H6 resemble those of the inversely oriented Glu408 and Ser404 in H13 in the second half of the inverted repeat (Figure 3A,D). This may imply that an early form of the CPA1 antiporters, which must have arisen by gene duplication of an unknown precursor, had a second, symmetrical ion-binding site that has been lost in the course of evolution. Arg362 in the unwound stretch of H12, which is essential in MjNhaP1 (Hellmer et al., 2003) and completely conserved in the CPA1 antiporters (Hellmer et al., 2003; Goswami et al., 2011), may be a tethered positive charge that takes the place of the Na+ ion in the second half of the inverted repeat, in a way similar to the arginine that replaces the co-transported Na+ in the sodium-independent substrate/product antiporter CaiT (Kalayil et al., 2013).

Regulation of transport activity

The transport activity of PaNhaP is highest at pH 5 and declines at higher or lower pH. The resulting bell-shaped pH profile is explained in terms of the Na+ affinity of the acidic residues in the substrate-binding pocket. The protonation state of these is likely to affect the affinity of the binding site for Na+. At low pH, most if not all of the ion-coordinating carboxyl sidechains (Glu73, Asp130, and Asp159) would be protonated, resulting in reduced affinity for Na+, as has been shown for MjNhaP1 by electrophysiological measurements on solid-supported membranes (Calinescu et al., 2014). At pH 5–7 these carboxyl sidechains would be increasingly deprotonated and able to bind and release Na+ ions, as is necessary for transport. At pH > 7, the ion-binding site is predominantly deprotonated and negatively charged (Figure 2—figure supplement 1), resulting in an increased Na+ affinity. As a result, the transport rate would decrease, as the ions are bound more tightly. This is consistent with the increased transport rate of the E73A mutant, which has one less carboxyl in the binding site, hence releases Na+ more easily (Figure 8B). In addition, the propagation of the pH-induced conformational changes at the dimer interface via Glu73 or Tyr31 would modulate the binding site by changing the coordination geometry for the ions (Figure 5—figure supplement 1A, Figure 5—figure supplement 2). Future structure-based molecular dynamics simulations should show how the protonation state of each of these residues influence the affinity of the binding site for Na+ in a pH-dependent manner.

The pH-dependent transport activity of PaNhaP suggests a self-regulatory mechanism for the binding site rather than regulation by a separate pH sensor as proposed for EcNhaA (Herz et al., 2010; Diab et al., 2011; Schushan et al., 2012). At the pH 5 activity maximum of PaNhaP, transport is not limited by Na+ affinity. Under these conditions, substrate binding of the PaNhaP dimer is non-cooperative, but unexpectedly it becomes cooperative at pH 6. Cooperative ion binding is most likely mediated by Glu73 and may be important for controlling the intracellular pH at neutral or basic pH, where a cooperative increase in Na+ affinity would gradually inhibit substrate release and slow down transport. This may be a safety mechanism to protect the organism against excessive influx of Na+, and hence efflux of protons, at rising pH, which may be critical for survival.

The medically relevant but elusive human Na+/H+ exchanger NHE1 is a dimer (Fafournoux et al., 1994) like PaNhaP. Several other common features, including high sequence homology (Goswami et al., 2011) especially of the unwound stretches in H5 and H12, key residues in the ion binding site such as Ser155, Asp130 and the ND motif, the functionally important Arg337 and Arg362 (Hellmer et al., 2003), as well as pH profiles and transport kinetics suggest that the archaeal and mammalian CPA1 antiporters (Fuster et al., 2008) work essentially in the same way. Remarkably, NHE1 also shows pH-dependent Na+ cooperativity, with a Hill coefficient of 1.8 at pH 6.8 that drops to ∼1 at pH 6 (Fuster et al., 2008). The PaNhaP structure thus serves as an excellent model for the membrane part of NHE1. Molecular details of allosteric regulation in NHE1 are likely to be different, as the His292 that reorients in response to pH in PaNhaP is not conserved (Goswami et al., 2011).

The electrogenic CPA2 antiporters, such as EcNhaA or TtNapA, which exchange two protons against one Na+, have two conserved aspartates in place of the ND motif in H6. In terms of its overall structure, TtNapA (Lee et al., 2013) is more similar to PaNhaP than to EcNhaA (Hunte et al., 2005), especially with respect to the dimer interface. The tertiary structure of the CPA antiporters is thus not a diagnostic of electroneutral or electrogenic transport.

Mechanisms of ion binding and release

In Pyrococcus, the Na+ gradient required for ATP synthesis is maintained by specific antiporters (McTernan et al., 2014). We therefore assume that PaNhaP, like human NHE1, utilizes the Na+ gradient across the membrane (Cohen et al., 2003) for pH homeostasis. Protons, probably in the form of hydronium ions (H3O+), can reach the binding pocket either through the cytoplasmic funnel or through the narrow polar channel (Figure 9). Only small rearrangements of the residues lining this channel would be required for H3O+ to pass. Using the second narrow polar channel for proton translocation would physically separate the routes for Na+ and H3O+ on the cytoplasmic side, which may be an advantage as the two ion currents flow in opposite directions. It would also explain why residues that line this channel, in particular the Glu154/Arg337 ion bridge and Asn158, which do not participate in ion coordination, are so highly conserved in the family. Molecular dynamics simulations and functional analysis of suitable mutants will be required to differentiate between the two proton paths, which both appear equally likely on the basis of the x-ray structures.

Figure 9.

Substrate ion exchange on the cytoplasmic side.

The substrate-binding site of PaNhaP is located between the unwound stretches in the six-helix-bundle and the interface domain. The substrate ion is bound by acidic sidechains and polar groups in the bundle helices H5 and H6, and a glutamate in the interface helix H3 at the deepest point of the cytoplasmic funnel. While the funnel extends between the six-helix bundle and the dimer interface, the narrow polar channel is defined by the bundle helices H5C, H12C, H6 and H13. Protons may approach the binding site either through the cytoplasmic funnel, or through the narrow polar channel (red arrows). A proton displaces the bound substrate ion, which escapes to the cytoplasm (black arrow). Employing the narrow polar channel as the proton path would separate the Na+ ion and proton currents on the cytoplasmic side, which may be advantageous at high transport rates.

DOI: http://dx.doi.org/10.7554/eLife.03579.024

Materials and methods

Cloning, expression and purification

A codon-optimized synthetic gene for the Na+/H+ antiporter from Pyrococcus abyssi (WP_010868413.1) was cloned into a vector with a C-terminal cysteine protease domain fusion as described previously for soluble proteins (Shen et al., 2009). Mutations were introduced by site-directed mutagenesis (Braman et al., 1996). The resulting plasmids were used to transform E. coli C41-(DE3) cells. The protein was expressed for 10 hr at 37°C in ZYM-5052 autoinduction medium (Studier, 2005).

Membranes were isolated from a 12 l culture and resuspended at 15 mg/ml protein in 20 mM Tris pH 7.4, 250 mM sucrose, 140 mM choline chloride. The suspension was diluted 1:3 in solubilization buffer (20 mM Tris pH 7.4, 150 mM NaCl, 30 % Glycerol and 1.5 % Cymal-5). After solubilization overnight at 4°C the solution was clarified by centrifugation at 100,000×g for 1 hr. The supernatant was supplemented with 5 mM imidazole, incubated for 2 hr with Talon resin (Clontech, Mountain View, CA) at 4°C and loaded on a Biorad column. Unspecifically bound proteins were eluted with washing buffer (20 mM Tris pH 7.4, 300 mM NaCl, 10 mM imidazole and 0.15 % Cymal-5). PaNhaP was cleaved off the column by incubating the beads in elution buffer (20 mM Tris pH 7.4, 300 mM NaCl, 0.15 % Cymal-5, 20 µM inositol-hexaphosphate) for 30 min. The eluted protein was concentrated to 5 mg/ml using a concentrator with 50 kDa cutoff and applied to a Superdex 200 size exclusion column equilibrated with 10 mM Na-Citrate pH 4.0, 300 mM NaCl and 0.15 % Cymal-5. Antiporter-containing fractions were pooled and concentrated to 5 mg/ml. The concentrated protein solution was diluted 1:4 with the same buffer containing 100 mM NaCl and re-concentrated as above. Selenomethionine (SeMet) labeled protein was expressed in PASM-5052 autoinduction medium (Studier, 2005) and purified as described for the native protein in the presence of 5 mM β-mercaptoethanol throughout all purification steps. β-mercaptoethanol was exchanged to 1 mM TCEP (Tris-(2-carboxyethyl) phosphine) in the final size-exclusion chromatography step.

Reconstitution

E. coli polar lipids (EPL, Avanti Polar Lipids, Inc., Alabaster, AL) were dried under nitrogen and resuspended in reconstitution buffer. Unilamellar vesicles were prepared by extruding the resuspended lipids using an extruder (Avestin, Inc., Canada) with 400 nm polycarbonate filters. Vesicles were destabilized by stepwise addition of n-octyl-β-D-glucoside (OG). The process was monitored at 540 nm. Addition of OG was stopped at around 1 % final concentration when the absorbance decreased rapidly. Protein was added to the destabilized liposomes at a lipid-to-protein ratio (LPR) of 100:1 and incubated for 1 hr at room temperature. The solution was dialyzed (14 kDa cutoff) overnight at room temperature against detergent-free reconstitution buffer. Biobeads (SM2, Biorad, Hercules, CA) were added to the dialysis buffer to ensure complete removal of the detergent. Proteoliposomes were centrifuged at 300,000×g for 20 min and washed once with reconstitution buffer. Washed proteoliposomes were pelleted again and resuspended at ∼60 mg/ml lipid in reconstitution buffer for further use.

Fluorescence assays

PaNhaP was reconstituted into proteoliposomes in 10 mM choline citrate/Tris pH 6–8, 200 mM NaCl and 5 mM KCl. To start the reaction 2 µl of proteoliposome suspension were diluted into 2 ml reaction buffer (10 mM choline-citrate/Tris at same pH, 5 mM KCl, 2 µM acridine orange). Emission of acridine orange (excitation: 495 nm) was monitored at 530 nm. To determine ion selectivity 5 mM NaAc, LiAc, KAc or TlAc were added to the reaction mixture after the initial sodium efflux reached equilibrium. Addition of substrates for PaNhaP to the reaction buffer results in proton efflux and fluorescence dequenching. Finally, the remaining proton gradient was dissipated by adding 25 mM (NH4)2SO4 in all experiments as a control. Electrogenic transport was investigated by addition of 100 nM valinomycin to the reaction buffer. The temperature was kept constant in a water bath during each experiment. Temperature dependence of transport was measured (triplicates) between 20°C and 45°C by correlating the speed of fluorescence quenching in the mid of the curve drop.

Radioactive 22Na+ uptake assays

PaNhaP was reconstituted in 20 mM choline citrate/Tris pH 4–8, 10 mM (NH4)2SO4. The reaction mixture contained 20 mM of the same buffer, 10 mM choline chloride, 1 µCi/ml 22Na+ and NaCl concentrations ranging from 1 µM to 5 mM. The pH-profile was determined at 200 µM NaCl. For each reaction 2 µl proteoliposomes were diluted in 200 µl reaction buffer to initiate the reaction. The addition of proteoliposomes to the reaction buffer results in NH3 efflux, producing an outward-directed proton gradient (Dibrov and Taglicht, 1993). At the time points indicated, 200 µl samples of the reaction mixture were applied to a 0.2 µm millipore nitrocellulose (Millipore, Billerica, MA) filter and washed with 3 ml 22Na+-free reaction buffer. Filters were transferred to counting tubes and 4 ml liquid scintillation cocktail (Rotiszint, Germany) was added. All measurements were performed at room temperature and repeated at least three times.

Crystallization

Prior to crystallization, the buffer for native protein was exchanged in the final concentrating step to 10 mM Tris/HCl pH 7.4, 100 mM NaCl, 0.15 % Cymal-5. Crystallization was performed in 24-well plates in hanging drops at 18°C. SeMet protein was supplemented with 1 % OG and native protein with 1 % n-octyl-β-thio-maltoside (OTM). The protein solutions were mixed 1:1 with reservoir buffer (native protein: 40 mM Na-Citrate pH 4.0, 100 mM NaCl, 28–33 % PEG 350 MME; SeMet protein: 100 mM Tris/HCl pH 8.0, 100 mM CaCl2/MgCl2, 35–40 % PEG 400). Trapezoidal pH 4 crystals grew up to 200 µm within 7 days. At pH 8, long needle-like crystals grew to full size within 3 months. Crystals were vitrified directly in liquid nitrogen for data collection. For thallium soaks, crystals grown at pH 8 were transferred into a buffer containing 100 mM Tris/acetate, 100 mM MgAc2, 40 % PEG 400, 2 mM K-citrate, 0.15 % Cymal-5 and 1 % OG. After five minutes the crystals were transferred to another drop of the same solution containing 25 mM TlAc. Crystals were incubated overnight and vitrified directly in liquid nitrogen.

Data collection, processing and structure determination

All diffraction data were collected with crystals kept at 100 K at the beamline X10SA of the Swiss Light Source in Villigen, Switzerland. Datasets were processed with XDS (Kabsch, 1993) and scaled with AIMLESS in the CCP4 package (Collaborative Computational Project 4, 1994). Resolution cut-offs were chosen based on CC1/2 (cross correlation of half datasets), completeness and I/σ(I)-values in high resolution shells (Karplus and Diederichs, 2012). Coot (Emsley and Cowtan, 2004) was used for model building and the PHENIX package (Adams et al., 2004) for refinement. Phases were obtained by single-wavelength anomalous dispersion (SAD) using SeMet crystals. Datasets from two crystals were merged to achieve a high multiplicity and to increase the anomalous signal (Liu et al., 2011). The Selenium substructure containing 11 out of 14 possible positions was determined at 5.7 Å using SHELXD (Sheldrick, 2010).

Phasing, hand determination, density modification with Parrot (Zhang et al., 1997) and initial model building with Buccaneer (Cowtan, 2006) was performed with a beta version of CRANK2 (Pannu et al., 2011). The resulting electron density map was used for manual building of an initial backbone model. Selenium positions were used to assign side chains in initial refinement rounds. Molecular replacement was performed using PHASER (McCoy, 2007) with the assigned dimer model to extend the resolution to 3.15 Å. The final pH 8 model was used for molecular replacement to phase the pH 4 structure and the thallium bound structure at pH 8. Superimpositions were performed using secondary structure superimposition (Krissinel and Henrick, 2004) within Coot (Emsley and Cowtan, 2004). Figures were prepared with PyMOL (DeLano and Lam, 2005). The potential surface was calculated with pdb2pqr (Dolinsky et al., 2007) and APBS (Baker et al., 2001). Analysis of transport pathways, channels and cavities was performed with Hollow (Ho and Gruswitz, 2008) and visualized within PyMOL.

Author information

Atomic coordinates and structure factors have been deposited with the PDB under accession codes: 4cz8 for the pH 8 SeMet structure, 4cz9 for the pH 4 structure and 4cza for the thallium-bound structure at pH 8.

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for sending your work entitled ‘Structure and substrate ion binding in the sodium/proton antiporter PaNhaP’ for consideration at eLife. Your article has been favorably evaluated by Michael Marletta (Senior editor), Richard Aldrich (Reviewing editor), and 2 reviewers, one of whom, Rajini Rao, has agreed to reveal her identity. A further reviewer remains anonymous.

The Reviewing editor and the reviewers discussed their comments before we reached this decision, and the Reviewing editor has assembled the following comments to help you prepare a revised submission.

This paper from the Kuhlbrandt group reports a trio of new structures of PaNhaP, an archael homolog of the NhaA family of Na/H exchangers, including the first structure of a Na/H exchanger that includes a cation, in this case a Thallium ion. Together, these structures represent a significant step forward in our understanding of this exchanger family, especially by visualizing a candidate cation binding site (and it's nice that Tl actually can drive transport, suggesting that this is indeed the cation binding site). Mutations of residues involved in cation binding have dramatic effects on transport activity consistent with the proposed role of the identified site. The crystal structures are very nice and the activity assays seem to have been carefully performed. The work is clearly presented and well written. The following comments should be addressed to improve the paper:

1) This manuscript should be combined with the accompanying one as a single revised submission.

2) Role of changes in the dimer interface. The authors report that the low pH form of the crystal primarily shows changes at the dimer interface but the actual structural rearrangements seem quite small. More of a concern though is how to interpret these changes in the context of mechanism. The state of the transporter in the low pH crystal is not at all clear-is it still inward facing? Lacking information about the state, we find it hard to conclude that the changes at the dimer interface ‘relay allosteric changes from the other protomer’. Indeed, the authors' interpretation of the structure implies that the low pH form of the protein should have substantially different Na affinity than the pH 8 form (if indeed they reflect the same overall state), but this prediction is not tested with the experiments shown here. Indeed, the Km for Na of 505 uM at pH 5 seems to shift to ∼200 mM at pH 6 but the structures are at pH 4 and 8, where activity is substantially different. The structures suggest that actual binding affinities could indeed be measured at pH 4 and 8, which would be essential to support the authors' interpretation. In addition, we find the superposition of structures presented in Figure 5–figure supplement 1 to capture the overall comparison of structures much better than the one in Figure 5 itself and would include at least one of these in the primary figure.

3) The acridine orange assay used in both papers to measure proton flux is an excellent assay for qualitative assessment of proton flux. However, the actual mechanism of acridine orange is unknown in detail and it is impossible to quantitatively measure pH change with this assay. Therefore the relative rates as a function of pH in Figure 5 and 6 are unreliable and should be omitted. Na22 flux could be used to measure these rates if desired, or a more quantitative pH probe, like pyranene.

4) The discussion of ‘Self-regulation of transport activity’ is completely disconnected from the evidence presented in the paper. If the authors wish to discuss this, they need to provide some experimental or computational support for their claims. They discuss ‘pH-dependent affinity’ but show no evidence that the affinity is indeed pH dependent beyond Kms at pH 5 and 6. Whether these values actually represent affinity depends on a range of assumptions which may or may not be valid for this protein.

1) This manuscript should be combined with the accompanying one as a single revised submission.

For a number of reasons we prefer to keep the manuscripts separate. One reason is authorship. The x-ray and electron crystallographic structure determination were two separate projects done by two PhD students. Merging the manuscripts would mean an injustice to one of them, and shared first authorship would not reflect the different contributions correctly. The other, more important reason is content. A merged manuscript would become unwieldy, unless important information is omitted, which we do not want to do. We therefore decided to revise the PaNhaP manuscript and to pursue publication of the MjNhaP1 manuscript separately.

2) Role of changes in the dimer interface. The authors report that the low pH form of the crystal primarily shows changes at the dimer interface but the actual structural rearrangements seem quite small.

The structural rearrangements at the PaNhaP interface cannot be described as small. Video 2 and Figure 5 plus supplements clearly show that sidechains in helix H10 move by up to 8A as the pH changes from 8 to 4. Moreover the changes are not confined to the interface but propagate through the whole protomer, including the loops connecting the trans-membrane helices.

More of a concern though is how to interpret these changes in the context of mechanism. The state of the transporter in the low pH crystal is not at all clear-is it still inward facing?

PaNhaP is in an inward-open conformation both at pH8 and at pH4. This is now stated explicitly in the revised manuscript. The substrate-binding site is accessible from the cytoplasm through the cytoplasmic funnel but not from the extracellular side under both conditions. However, the narrow side channel that leads from the cytoplasmic surface to the ion-binding site is blocked at pH4 by side chain rearrangements, as stated in the revised manuscript.

It is important to note that the pH-dependent allosteric change of the dimer is different from the inside-open to outside-open transition in the transport cycle of the protomer. This is now stated explicitly in the revised manuscript. Rather, the allosteric change increases the affinity of the binding site for the substrate ion from Km = 500 µM to K0.5 = 25 µM. This extends the range for high-affinity substrate binding by ∼1 pH unit from acidic towards neutral conditions, as may be necessary for efficient Na+/H+ exchange at physiological pH in this particular organism. The pH-dependent binding affinity will be the subject of a future molecular dynamics study that goes well beyond the scope of the present manuscript.

Lacking information about the state, we find it hard to conclude that the changes at the dimer interface ‘relay allosteric changes from the other protomer’.

It is evident from Video 2 in the revised manuscript (supplementary Video 1 in the original manuscript) that the pH-dependent repulsion of protonated histidines 292 causes conformational changes, and that this mechanism can only work with two protomers next to one another in the dimer. Therefore the changes do indeed relay allosteric changes between protomers. However, we agree that it may be better to say that ‘conformational changes caused by repulsion of the protonated histidines at the dimer interface are relayed to the ion binding site to modulate the Na+ binding affinity in a pH-dependent manner’. This is now stated in the revised manuscript.

To show the pH-induced differences more clearly, we have added supplementary figures 1 and 2 to Figure 5 as stereo images in the revised manuscript.

Indeed, the authors' interpretation of the structure implies that the low pH form of the protein should have substantially different Na affinity than the pH 8 form (if indeed they reflect the same overall state), but this prediction is not tested with the experiments shown here.

Both structures do indeed show the same overall state (inward-open), as explained above. Because the antiporter is only minimally active at pH4 and pH8 (see Figure 4), we are unable to measure the binding affinity under these conditions with the methods available to us. However, the Na binding affinities at pH5 and pH6 are substantially different, as indicated by the K0.5 for the cooperative antiporter at pH6 that we have now added to Figure 4D. Compared to the Km at pH5 in Figure 4C, this indicates a roughly 20-fold increase in Na binding affinity.

In the revised manuscript we have emphasized the probable role of protonation states of acidic residues in the binding site in pH-dependent activity changes. For example, Asp130, which is involved directly in substrate-ion coordination, changes its conformation in response to pH, and this directly affects the coordination geometry. This should now be clearer in the new Figure 5–figure supplement 2, which shows the superposition of the ion-coordinating residues in both protomers in stereo.

Indeed, the Km for Na of 505 uM at pH 5 seems to shift to ∼200 mM at pH 6 but the structures are at pH 4 and 8, where activity is substantially different.

We have no idea how the referees arrive at the conclusion that the Km at pH6 should be 200 mM. There was no Km value or even a K0.5 value for this pH in Figure 4D or anywhere else in the original manuscript. We have added the K0.5 value, which is 25 µM, to Figure 4D of the revised manuscript.

The structures suggest that actual binding affinities could indeed be measured at pH 4 and 8, which would be essential to support the authors' interpretation.

As explained above, PaNhaP is essentially inactive at pH 4 and 8, so these measurements are not feasible by the methods available to us.

In addition, we find the superposition of structures presented in Figure 5–figure supplement 1 to capture the overall comparison of structures much better than the one in Figure 5 itself and would include at least one of these in the primary figure.

In the revised manuscript, we replaced Figure 5 by Figure 5–figure supplement 1 in the original manuscript. Video 1 of the original manuscript is now Video 2, which shows the pH-induced conformational changes very clearly. Stereo pairs of original Figure 5 are now provided as supplements to this Figure, making the conformational changes even clearer.

3) The acridine orange assay used in both papers to measure proton flux is an excellent assay for qualitative assessment of proton flux. However, the actual mechanism of acridine orange is unknown in detail and it is impossible to quantitatively measure pH change with this assay. Therefore the relative rates as a function of pH in Figure 5 and 6 are unreliable and should be omitted.

We do not understand this comment. Figure 5 of the original manuscript shows pH-induced conformational changes. Figure 6 shows the temperature dependence of transport.

Figure 4–figure supplement 1 (both in the original and the revised manuscript) does show acridine orange measurements as a function of pH but these measurements are only qualitative to confirm transport activity under symmetrical pH conditions. All quantitative measurements of pH-dependent activity were performed with radioactive 22Na assays.

Na22 flux could be used to measure these rates if desired, or a more quantitative pH probe, like pyranene.

This is exactly what we have done, as explained in many different places in the original manuscript.

4) The discussion of “Self-regulation of transport activity” is completely disconnected from the evidence presented in the paper. If the authors wish to discuss this, they need to provide some experimental or computational support for their claims. They discuss ‘pH-dependent affinity’ but show no evidence that the affinity is indeed pH dependent beyond Kms at pH 5 and 6. Whether these values actually represent affinity depends on a range of assumptions which may or may not be valid for this protein.

Experimental evidence includes the observation that mutation of His292 to cysteine (Figure 7–figure supplement 1) did not alter the pH dependence, as stated in the revised manuscript. Additional experimental support is provided by SSM measurements with MjNhaP1 as cited in the manuscript.

Table 2.

Data collection and phasing statistics

DOI: http://dx.doi.org/10.7554/eLife.03579.004

	Dataset 1	Dataset 2	Merge
Data collection	SLS PXII
Wavelength	0.979	0.979	0.979
Space group	P21	P21	P21
Cell dimensions
a, b, c (Å)	54.7, 109.0, 110.8	54.6, 108.3, 110.5	54.7, 108.9, 110.7
α, β, γ (°)	90.0, 94.6, 90.0	90.0, 95.0, 90.0	90.0, 94.7, 90.0
Resolution (Å)	49.3–3.8 (3.97–3.8)	49.0–3.8 (3.97–3.8)	49.2–3.8 (3.97–3.8)
Rpim	0.029 (0.470)	0.034 (0.228)	0.036 (0.315)
I / σI	13.8 (2.1)	12.4 (3.9)	14.5 (2.9)
CC*	1.000 (0.929)	0.996 (0.985)	1.000 (0.981)
Completeness (%)	99.7 (99.7)	99.7 (99.6)	100 (100)
Multiplicity	24.5 (16.5)	9.1 (9.5)	33.0 (25.8)
Phasing
CCanom			0.348
Anom slope			1.061
FOM after Phasing (Refmac)			0.230
FOM after DM (Parrot)			0.594