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

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 K_0.5 for the cooperative antiporter at pH6 that we have now added to Figure 4D. Compared to the K_m 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 K_m at pH6 should be 200 mM. There was no K_m value or even a K_0.5 value for this pH in Figure 4D or anywhere else in the original manuscript. We have added the K_0.5 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.