Phosphoinositides (PIPs) are membrane lipids that, together with their corresponding soluble inositol phosphates (IPs), regulate various cellular processes, including membrane recruitment of proteins, actin polymerization, synaptic vesicle trafficking and exo- and endocytosis (Di Paolo and De Camilli, 2006; Balla, 2013; Ueda, 2014). The dynamic control of the membrane distribution and relative abundance of the seven naturally occurring PIPs by a multitude of phosphoinositide kinases and phosphatases forms a versatile signaling mechanism able to tune the spatial and temporal regulation of many crucial events in the cell (Fruman et al., 1998; Anderson et al., 1999; Hsu and Mao, 2015).

The inositol polyphosphate 5-phosphatases (5PPases) form a family of Mg²⁺-dependent enzymes, containing ten members in mammals, that catalyse the hydrolytic removal of the phosphate group on the 5-position of lipid-bound and soluble inositol phosphates (Figure 1, Figure 1—figure supplement 1). Based on their substrate specificity, the mammalian 5PPases can be further subdivided into four groups (Majerus et al., 1999): the type I 5PPase INPP5A (Speed et al., 1996), the type II 5PPases OCRL, INPP5B, INPP5J (or PIPP), SKIP, Synaptojanin1 (Synj1) and Synaptojanin 2 (Synj2) (Lowe, 2005; Trésaugues et al., 2014; Mochizuki and Takenawa, 1999; Ijuin et al., 2000; McPherson et al., 1996; Nemoto et al., 1997), the type III 5PPases, SHIP1 and SHIP2 (Rohrschneider et al., 2000; Le Coq et al., 2017), and the type IV 5PPase INPP5E (or Pharbin) (Asano et al., 1999). Among the type II 5PPases, the closely related Synj1 and Synj2 contain a similar domain arrangement, with in addition to the central 5PPase domain, an N-terminal suppressor of actin 1 (SAC1)-like domain and a C-terminal proline-rich domain (PRD). As such, Synj1 and Synj2 are unique in having two phosphatase activities, where the 5PPase domains can hydrolyse PI(4,5)P₂, PI(3,4,5)P₃, IP_3, and IP₄, while the SAC1-like domain can degrade PI(3)P, PI(4)P, and PI(3,5)P₂ (Hsu and Mao, 2015; Cestra et al., 1999; Whisstock et al., 2002). Both Synaptojanin proteins are implicated in clathrin-mediated endocytosis, where Synj2 is involved in the early stages of this process (Rusk et al., 2003), while the brain-specific 145 kDa splice isoform of Synj1 promotes clathrin uncoating during the late stages of endocytosis (Perera et al., 2006; Ramjaun and McPherson, 1996). In agreement with this role, knock-out studies of Synj1 in mice (Cremona et al., 1999) and Drosophila melanogaster (Verstreken et al., 2003) show endocytic defects at the neuronal synapses, indicating that Synj1 plays a critical role in synaptic function. This key function at the synapse is further illustrated by the implication of Synj1 in several diseases. Brain autopsy of Down syndrome (DS) patients revealed an excessive expression of the Synj1 protein, that is encoded on the triplicated chromosome 21, and it was shown that the overexpression of Synj1 leads to PI(4,5)P₂ deficiency and learning deficits in Down syndrome model mice (Voronov et al., 2008; Arai et al., 2002). Elevated levels of Synj1 are also found in individuals showing a high risk for the development of Alzheimer's disease (AD) (Berman et al., 2008; Martin et al., 2015; Miranda et al., 2018). Based on these findings, Synj1 has been proposed as a potential attractive two-faced target for novel DS and AD treatments (Cossec et al., 2012). Additionally, inhibition of the 5-phosphatase activity of Synj1 has also been found to hold promise toward drug development for TBC1D24-associated epilepsy and DOORS syndrome (Fischer et al., 2016; Lüthy et al., 2019). On the other hand, recessive loss-of-function mutations in Synj1 are associated with either early-onset atypical parkinsonism or refractory epileptic seizures with severe progressive neurodegeneration (Hardies et al., 2016; Xie et al., 2019; Taghavi et al., 2018; Hong et al., 2019; Bouhouche et al., 2017).

Figure 1 with 1 supplement see all

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Out of the ten 5PPase domains present in human, high-resolution structural information has only been published for three representatives: INPP5B, OCRL, and SHIP2 (Trésaugues et al., 2014; Mills et al., 2016). In addition, an unpublished structure of INPP5E has been deposited in the protein data bank (PDB 2XSW). So far no experimentally determined structure is available for the 5PPase domain of either Synj1 or Synj2, although the very first 5PPase structure ever to be determined was the one of a Synaptojanin homolog of the yeast Schizosaccharomyces pombe (Tsujishita et al., 2001).

In addition to the structural data of the 5PPase domains in their apo form, crystal structures in complex with reaction products and inhibitors, together with detailed studies on the structurally and mechanistically related Apurinic/Apyrimidinic endonucleases (APE), have yielded some insights in the catalytic mechanism of the hydrolysis reaction (Whisstock et al., 2000). Indeed, structural and sequence comparison revealed similarities in the active site architecture of the 5PPases and APE1, a Mg²⁺-dependent enzyme that catalyses the cleavage of the phosphodiester bond on the 5’ side of the abasic site in DNA using a conserved aspartate as catalytic base and a conserved histidine that presumably stabilizes the phosphorane transition state. Moreover, leaving group activation has been proposed to occur via a Mg²⁺-bound water molecule, although the exact role of the Mg²⁺-ion in the catalytic mechanism of 5PPases has not been fully established (Trésaugues et al., 2014; Mills et al., 2016; Aboelnga and Wetmore, 2019). A crystal structure of the S. pombe Synaptojanin (SPSynj) homolog in complex with inositol-(1,4)-bisphosphate provided a first snapshot of a potential enzyme-product complex (Tsujishita et al., 2001). However, subsequent structures of the catalytic domain of INPP5B in complex with diC8-PI(4)P and diC8-PI(3,4)P₂ revealed another orientation of these reaction products, suggesting that the placement of the ligand in the SPSynj structure does not correspond to the genuine position of the product (Trésaugues et al., 2014). Further elucidation of the complete hydrolysis mechanism is however hampered by the lack of 5PPase structures in complex with reaction substrates showing directly the exact placement of the 5-phosphate group in the active site.

In this study we report the first structure of the catalytic 5PPase domain of human Synj1 by using Nanobody-aided crystallography. Moreover, we were able, for the first time, to solve a structure of a 5PPase domain with the substrate diC8-PI(3,4,5)P₃ trapped in its active site, revealing the placement of, and the interactions with, the 5-phosphate group. In combination with a detailed kinetic analysis of the hydrolysis reaction catalysed by the Synj1 5PPase domain, these structures provide additional insights in the catalytic mechanism. Finally, we investigated in detail the effect of the three currently described homozygous missense mutations in the 5PPase domain (Y793C, R800C, Y849C) associated with either (young onset) Parkinson’s disease or intractable epilepsy with neurodegeneration (Hardies et al., 2016; Xie et al., 2019; Taghavi et al., 2018) on the reaction rate for different substrates, thus providing insights in the molecular mechanisms underlying these diseases.

In this paper, we report the first structural information on the 5PPase domain of Synj1 (Synj1_528–873) to 2.3 Å resolution, relying on a strategy of Nanobody-aided crystallization to obtain well diffracting crystals. In addition to the Synj1_528–873 structure in the apo state, a short soak with diC8-PI(3,4,5)P₃ followed by flash freezing, also enabled us to trap this substrate in one of the active sites of the protein molecules present in the asymmetric unit, representing the very first structure of any 5PPase in complex with a genuine substrate. This structure thus reveals the interactions with all the important phosphate groups, including the scissile 5-phosphate group. Together with a detailed kinetic analysis of the contribution of these phosphate groups to catalysis, this allows us to propose a refined model for the catalytic mechanism of the 5PPase reaction extending on the previous models based on analogies with the apurinic/apyrimidinic base excision repair endonucleases (Whisstock et al., 2000; Aboelnga and Wetmore, 2019; Dlakić, 2000; Mol et al., 2000; Erzberger and Wilson, 1999), as shown in Figure 6.

Figure 6

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In agreement with its conserved role in AP endonucleases and as previously suggested (Trésaugues et al., 2014), D730 is appropriately positioned to act as a general base by abstracting a proton from an attacking water molecule. Indeed, careful analysis of the electron density in the Synj1_528–873 diC8-PI(3,4,5)P₃-bound structure reveals electron density accounting for such a water molecule located at 3.1 Å from D730 and at 3.5 Å from the P₅-atom. This water molecule is furthermore held in place via a hydrogen bond with the conserved N732 residue (Figure 3; Figure 6). It should be noted that also the catalytically important H859 residue is located close to the nucleophilic water molecule, and, depending on the orientation of the side chain, H859 could either form an interaction with the O_PF-atom of the 5 P group or with this water molecule. The 5 P group is closely surrounded by multiple residues, H689, N732, R734, Y784, and H859, many of them potentially bearing a positive charge. Interactions of these residues with the 5 P group in the ground state contribute to binding, while stabilization of the excess of negative charge building up in the trigonal bipyramidal phosphorane transition state would contribute to catalysis. Previous mutagenesis studies on mouse INPP5A, yeast Inp52P and human INPP5B have shown that mutation of the residues corresponding to H689 and H859 negatively impact the activity, although from these studies it is not clear whether this is due to an effect on binding or catalysis (Whisstock et al., 2000; Jefferson and Majerus, 1996). While multiple interactions are formed with the 5 P group, none of the residues is seen within interaction distance of the oxygen bridging the scissile phosphate to the inositol ring (O₅) and thus in an appropriate position to act as general acid to activate the leaving group. The O₅-atom is however located at a distance of 4.0 Å of the Mg²⁺-ion allowing this distance to be bridged by a Mg²⁺-coordinated water molecule as previously suggested (Trésaugues et al., 2014). Additionally, the O₅-atom is located in our structure at 3.9 Å from one of the oxygens (O_8P) of the 4 P group. This distance could decrease during the reaction path, allowing a direct role of the 4 P group in leaving group activation. We therefore envision two potential scenarios for leaving group activation to occur, indicated as route 1 and route 2 in Figure 6. Route 1 envisions a direct role of the Mg²⁺-ion in leaving group activation by activating a Mg²⁺-bound water molecule to donate a proton to the leaving O₅-atom, although such a water is not observed in our structure. A direct role of Mg²⁺ in catalysis also agrees with its observed large contribution to catalytic turnover (ΔΔG_catalysis = 4.3 kcal/mol, Figure 4). In this scenario, the observed role of the 4 P group in catalysis could for example be due to a stabilizing hydrogen bond to the O₅-atom in the transition state (Figure 6). Route 2, on the other hand, corresponds to a mechanism of substrate-assisted catalysis, with a more direct role of the adjacent 4 P group in leaving group activation, where the O_8P hydroxyl group of the 4 P group would transfer a proton to the leaving O₅-atom. The transferred proton could potentially originally come from the K798 residue in a proton relay mechanism (Figure 6). In our structure, the O_8P-atom from 4 P is located at 3.9 Å from the O₅-atom, but small rearrangements in going towards the transition state could easily bring both atoms in close contact. The residues surrounding the 4 P group could in turn contribute to catalysis by properly orienting the 4-phosphate for this role. Such a scenario is in good agreement with our kinetic data showing that the 4 P group has a large contribution to substrate turnover, with removal of the 4 P moiety (i.e. comparing diC8-PI(3,4,5)P₃ with diC8-PI(3,5)P₂) leading to a 340-fold reduction in k_cat (ΔΔG_catalysis = 3.5 kcal/mol, Figure 4). Moreover, we also showed that the R800 residue contributes to catalysis via its interaction with the 4 P group. Within this scenario, the Mg²⁺-ion could play its observed role in catalysis via a direct interaction with the negative charges building upon the O₅-atom in the transition state. It should of course be noted that the latter mechanism would not apply for the 5PPases that also efficiently catalyse the hydrolysis of PIPs missing the 4 P group (such as SHIP1, SHIP2, and to a lesser extent OCRL). At this point no conclusive distinction between these two scenarios can be made.

Apart from being a potential drug target for Alzheimer’s disease, Down syndrome, and TBC1D24-linked epilepsy, loss-of-function mutations in Synj1 also lead to disease, including epilepsy and Parkinson’s disease (Hardies et al., 2016; Xie et al., 2019; Taghavi et al., 2018; Hong et al., 2019; Bouhouche et al., 2017). This is an important point to take into account in a potential drug design effort. To rationalize such an effort and to find the available therapeutic window for inhibitory drug design, it is of paramount importance to characterize and quantify the effect of the disease-associated mutations on Synj1 activity in detail. At the moment of writing this manuscript, three disease-associated homozygous missense mutations had been described in the 5PPase domain of Synj1: Y793C, R800C, and Y849C (numbering based on Synaptojanin1-145 isoform 2) (Hardies et al., 2016; Xie et al., 2019; Taghavi et al., 2018). Our structural data and detailed kinetic analysis of the mutants in comparison to the wild-type enzyme allow us to gain deeper understanding in the molecular basis underlying the associated diseases. Our data shows that the Y849C mutant does not display any 5PPase activity with any of the tested substrates. The crystal structure shows that Y849 is nearly completely buried in the core of the protein (Figure 5C) and the Y849C mutation leads to severe problems in expressing this protein in a soluble form. Together, this indicates that the Y849C mutant corresponds to a near complete loss-of-function mutation due to disruption of the protein fold. In contrast, the R800 residue is located in the active site where it forms important interactions with the 4-phosphate group of the substrate (Figure 5B). Our kinetic analysis of the R800C mutant shows that this residue contributes to both substrate binding and catalysis, specifically via its interaction with the 4 P group. The contribution to substrate turnover (k_cat) can be explained by a model where R800 acts by orienting the 4 P group in an orientation suitable for subsequent proton transfer to the inositol 5-hydroxylate leaving group, as outlined above (Figure 6, route 2). Interestingly, the magnitude of the overall effect of the R800C mutation depends on the substrate that is being considered, with a decrease in k_cat/K_M of a factor 900, 54, and 12 for IP₃, diC8-PI(4,5)P₂ and diC8-PI(3,4,5)P₃, respectively. Whether this difference in activity toward the different substrates is physiologically relevant and leads to an imbalance in the cellular distribution of PI(3,4,5)P₃, PI(4,5)P₂, and IP₃, remains to be seen. Such an imbalance of the PI(3,4,5)P₃/PI(4,5)P₂ ratio, with a relative increase in PI(4,5)P₂ versus PI(3,4,5)P₃ as expected from the kinetics of the R800C mutant, has previously been linked to the occurrence of Parkinson’s disease (Sekar and Taghibiglou, 2018). Finally, the Y793 residue is located on the same active site loop as R800, and likely contributes to stabilizing the loop conformation rather than being directly involved in interactions with the substrate (Figure 5B). Our kinetic analysis of the Y793C clinical mutant shows that the 5PPase activity of Synj1 is diminished between 5- and 10-fold for all the substrates in a rather indiscriminatory way.

As a general trend, it can be observed that the impact of the mutations on the catalytic efficiency of the Synj1_528–873 enzyme in vitro links with the severity and age of onset of the clinical manifestations in the patients that are homozygous carriers of these Synj1 mutations. Indeed, the very early onset of the severe neurodegeneration observed for the patients carrying a Y849C mutation (Hardies et al., 2016), corresponds well with our observation that this mutation leads to a complete loss of 5PPase activity. On the other hand, the R800C mutation leads to a decrease in overall activity of 54- to 12-fold for the most relevant substrates PI(4,5)P₂ and PI(3,4,5)P₃ and was reported to associate with parkinsonism and seizures at age 24 (Taghavi et al., 2018), while the even smaller effects on activity we observe for the Y793C mutation, with a decrease in activity of 8-fold for PI(4,5)P₂ and 7-fold for PI(3,4,5)P₃, leads to Parkinson’s disease at later age (Xie et al., 2019). Although care should be taken with extending these observations to other mutations, this seems to indicate that the impact of the missense mutations on the kinetics of the 5PPase reaction in vitro can be used to a certain level to predict the severity of the disease outcome in patients who are homozygous for the corresponding mutation. Finally, these observations also have implications for the window that is available to therapeutically target the 5PPase domain of Synj1 in DS, AD, and TBC1D24-associated DOORS syndrome. While it was previously shown that genetic ablation of one Synj allele, corresponding to half of the normal cellular activity, was sufficient to rescue the disease phenotype in TBC1D24-mutant flies (Fischer et al., 2016), we show here that a close to 10-fold reduction of Synj1 activity on the long term could lead to disease in its own respect, thus still leaving a window for inhibitor design.

Diffraction data have been deposited in the PDB under the accession code 7A0V and 7A17. All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figure 4 and Figure 4—figure supplements 1–3.

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   ID 7A0V. Crystal structure of the 5-phosphatase domain of Synaptojanin1 in complex with a nanobody.

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Acceptance summary:

Paesmans and colleagues describe the nanobody-bound crystal structure of the 5-phosphatase domain of human synaptojanin1, an important regulator of clathrin-mediated endocytosis, with and without a trapped substrate. Based on the structure, they kinetically characterize catalysis towards different substrates and come up with two alternative mechanisms how Synj1 mediates catalysis. They finally characterize three disease mutants of synaptojanin1, which show reduced activity or aggregate.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review. Please note that in a second round of review, the reviewers requested the chain order be changed, so that chain E became chain A.]

Thank you for submitting your work entitled "A structure of substrate-bound Synaptojanin1 provides new insights in its mechanism and the effect of disease mutations" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

The reviewers are concerned that the crystallized substrate-bound complex is non-productive. Moreover, the absence of substrate in 2 of the 3 synaptojanin molecules is troublesome, and there are other concerns about this structure (details in reviews below). Thus, in the present form, the paper is not acceptable.

Reviewer #1:

This is a potentially a major contribution towards understanding the molecular mechanism of synaptojanin. The previous structure of the 5-phosphatase domain of S. pombe synaptojanin in complex with inositol‐(1,4)‐bisphosphate (Tsujishita, et al., PDB ID 1i9z) was determined at high resolution but the electron density for the inositol-(1,4)-bisphosphate was ambiguous. As the authors point out, subsequent structures of the catalytic 102 domain of INPP5B in complex with diC8‐PI(4)P and diC8‐PI(3,4)P2 revealed another orientation but these structures lack the 5-phosphate group. In this work the authors present high-resolution structures of the 5-phosphatase domain of synaptojanin, and its complex with the substrate diC8-PI(3,4,5)P3, providing a high-resolution image of a 5-phosphatase with a trapped substrate in its active site. The new structures suggest an enzymatic mechanism and explain how certain disease-associated mutations affect the catalytic rate.

In the substrate-bound structure, electron density for diC8-PI(3,4,5)P3 was only found for 1 of the three molecules in the asymmetric unit. Electron density consistent with phosphate groups was found for the other two molecules. However, the B-factors are perhaps a little higher around the binding site for one of the two molecules without substrate. Is it possible that substrate was partially occupied in that site? Please provide simulated annealing omit maps for all three binding sites.

A cis-peptide bond is found between Y784 and K785, as in all other 5-phosphatase structures except in the deposited structure of S. pombe synaptojanin. It might be instructive to try to fit a cis peptide bond against the S. pombe diffraction data.

As the authors point out, the effect of the Y793C mutant might be indirect by affecting nearby residues at the active site. It is particularly interesting that Y793 is close to the cis-peptide bond between Y784 and K785. Is it possible that the mutation destabilizes this cis-peptide bond, corroborating its functional importance?

Reviewer #2:

Paesmans and colleagues describe the nanobody-bound crystal structure of the 5-phosphatase domain of human Synj1, an important regulator of clathrin-mediated endocytosis, with and without a trapped substrate. Based on the structure, they kinetically characterize catalysis towards different substrates and come up with two alternative mechanisms how Syn1 mediates catalysis. They finally characterize three disease mutants of Syn1, which show reduced activity or aggregate.

The paper is well drafted, the nanobody-aided structure determination of Syn1 is elegant and the kinetic experiments appear overall sound. A question to discuss is whether the novelty of the structural data is sufficient for publication in eLife. The structure of yeast synaptojanin is known for a long time and it seems rather similar to the human counterpart (although this is difficult to judge from this manuscript). Furthermore, many features of substrate binding and catalysis were already deduced from the product-bound yeast synaptojanin structure. I have also some questions/doubts about the substrate-bound structure.

1) Detailed structural comparisons of Syn1 to other 5'-phosphatases (including rmsd) are missing to judge similarities and recognize potential differences between the structures. A structural comparison of the catalytic centers of some 5' phosphatases, including human and yeast synaptojanin with bound ligands, complementing Supplementary file 1, would allow the reader to appreciate (potentially) new mechanistic insights from the Syn1 structure.

2) A major point of the paper is the substrate bound to Syn1. However, it remains unclear why the substrate has not been turned over after an overnight incubation at 20 C. The authors speculate about a missing catalytic water, but this cannot explain missing catalysis since the substrate-binding site is apparently accessible, in particular for water. Is the crystallization condition incompatible with catalysis? Does nanobody-binding reduce catalysis? Is there at all any substrate left after overnight incubation with the crystals? Without answering this, it remains uncertain whether the observed ligand is indeed a non-converted substrate molecule (see also next point).

3) Crystallographic data of the substrate-bound crystals: Why is the signal to noise value so low for the substrate-bound crystals? Why did the authors apply anisotropic corrections to their data? Did the crystals diffracted anisotropically – if yes, provide some more information, e.g. to which resolution along a*,b*,c*. Judging from the spherical completeness, the dataset rather correspond to a 3.x A resolution dataset in terms of reflection numbers. It is astonishing that for such low resolution, 91 water molecules with an average B-factor of only 34 Angstroms could be modelled (I would hardly expect to see any visible water molecules at this resolution). Is the model overfitted? Any explanation for the wide gap of Rwork and Rfree? What was the exact refinement strategy? Overall, the data quality of the substrate-bound crystals is borderline to accurately evaluate ligand binding, in particular if the identity of the bound ligand is not 100% clear.

Reviewer #3:

The authors report crystal structures of the apo and substrate bound versions of the inositol 5-phosphatase domain of human synaptojanin-1 (Synj1), crystallized in the presence of a nanobody. The overall structure is similar to those of several previously solved 5-phosphatase enyzmes from yeast and humans. The main addition to the field in the current structure with respect to past work is that a substrate is present in this case, whereas previous structures were apo or product-bound.

The structure contains 3 catalytic chains per asymmetric unit. The ligand-bound form was obtained by soaking apo crystals in a di-C8 form of the substrate phosphoinositide. The structure of the complex was determined at 2.9 Å resolution. Detail on the catalytic geometry is limited at this resolution. Water and metal ion positions are not visualized at all, or with limited accuracy. Only one of the three catalytic chains binds to substrate. I did not see an explanation for the lack of binding to the other two chains. The 5-phosphate is still present on the substrate, despite that catalytically active wild-type enzyme was used. The authors present a detailed scheme for the catalytic mechanism on the basis of their structure, but I did not see an explanation for the failure of the enzyme to hydrolyze the 5-phosphate. I would speculate that the pH 5.0 of the crystallization could account for this. Have the authors attempted to adapt the crystals to neutral pH? Is a non-hydrolyzable substrate analogue available, or could it be synthesized? (I appreciate that phosphoinositide synthetic chemistry is non-trivial.) It would certainly be helpful for the field to know the geometry of a catalytically productive substrate complex, yet I am not convinced that the authors have as yet succeeded in obtaining such a structure. In my opinion, the conclusions concerning enzyme mechanism are, at this stage, over-interpreted.

The paper is, apart from the omissions noted above, very thorough. It reads like a review article in the Introduction and Discussion. The results includes more detail than is standard. Overall the paper would be more readable if it were edited to about half of the current length.

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review. Please note that in a second round of review, the reviewers requested the chain order be changed, so that chain E became chain A.]

Reviewer #1:

This is a potentially a major contribution towards understanding the molecular mechanism of synaptojanin. The previous structure of the 5-phosphatase domain of S. pombe synaptojanin in complex with inositol‐(1,4)‐bisphosphate (Tsujishita, et al., PDB ID 1i9z) was determined at high resolution but the electron density for the inositol-(1,4)-bisphosphate was ambiguous. As the authors point out, subsequent structures of the catalytic 102 domain of INPP5B in complex with diC8‐PI(4)P and diC8‐PI(3,4)P2 revealed another orientation but these structures lack the 5-phosphate group. In this work the authors present high-resolution structures of the 5-phosphatase domain of synaptojanin, and its complex with the substrate diC8-PI(3,4,5)P3, providing a high-resolution image of a 5-phosphatase with a trapped substrate in its active site. The new structures suggest an enzymatic mechanism and explain how certain disease-associated mutations affect the catalytic rate.

In the substrate-bound structure, electron density for diC8-PI(3,4,5)P3 was only found for 1 of the three molecules in the asymmetric unit. Electron density consistent with phosphate groups was found for the other two molecules. However, the B-factors are perhaps a little higher around the binding site for one of the two molecules without substrate. Is it possible that substrate was partially occupied in that site? Please provide simulated annealing omit maps for all three binding sites.

In our revised manuscript we have carefully re‐analyzed our data processing and refinement of the Synj1 structure in complex with diC8‐PI(3,4,5)P₃, suggesting an even higher resolution for this substrate-bound structure of 2.73 Å. Subsequently, we carefully analyzed the unbiased electron density in the active site of the three protein molecules in the asymmetric unit. The omit map for molecule E very clearly shows electron density for diC8‐PI(3,4,5)P₃, including the phosphate groups on position 3, 4 and 5, allowing us to unambiguously model this compound (see unbiased omit map and 2Fo‐Fc and residual Fo‐Fc maps after refinement in Author response image 1). The omit maps for the other two active sites (chain A and chain C) show some continuous density which might account for the presence of a ligand at low occupancy. The latter is especially true for chain C (Author response image 1). These differences in occupancy could reflect differences in kinetics of access of substrate or release of products from the three active sites, depending on their local environment in the crystal packing, and/or could reflect the substrate being hydrolyzed to different degrees. Since the density in chain A and C is very weak and to avoid overinterpretation, we opted not to model these densities as substrate molecules.

The modelling of the densities in the different active sites is now also being discussed in more detail in the Results section of our revised manuscript.

Author response image 1

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A cis-peptide bond is found between Y784 and K785, as in all other 5-phosphatase structures except in the deposited structure of S. pombe synaptojanin. It might be instructive to try to fit a cis peptide bond against the S. pombe diffraction data.

As suggested by the reviewer, a cis‐peptide bond was introduced between the corresponding Y794 and K795 residues of the SPSynj structure (PDB 1I9Z), after which refinement against the diffraction data was performed and electron densities were compared. Author response image 2 shows a comparison of the resulting 2Fo‐Fc map at 1σ and the Fo‐Fc map at 3σ for a model with a trans‐peptide bond (Author response image 2A) versus a cis‐peptide bond (Author response image 2B). The resulting maps show a clear preference for the cis‐peptide, suggesting that the cis‐peptide between Y784 and K785 (or Y794 and K795 in SPSynj) is conserved in all 5PPases. We now also mention the possibility for the occurrence of a cis‐peptide bond in SPSynj in our revised manuscript.

Author response image 2

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As the authors point out, the effect of the Y793C mutant might be indirect by affecting nearby residues at the active site. It is particularly interesting that Y793 is close to the cis-peptide bond between Y784 and K785. Is it possible that the mutation destabilizes this cis-peptide bond, corroborating its functional importance?

The Y793 residue is indeed located on the large P4IM loop, in between active site residues Y784‐K785 and K798‐R800. Since the Y793C mutant has a rather mild but broad effect on the catalytic activity for all tested substrates, we anticipate a scenario where this mutation has a subtle effect on the conformation of the entire P4IM loop. At this point it is therefore very hard to uncouple the potential effect of this mutation on the cis‐peptide bond from the effect on the other active site residues resulting from changes in loop conformation.

Reviewer #2:

Paesmans and colleagues describe the nanobody-bound crystal structure of the 5-phosphatase domain of human Synj1, an important regulator of clathrin-mediated endocytosis, with and without a trapped substrate. Based on the structure, they kinetically characterize catalysis towards different substrates and come up with two alternative mechanisms how Syn1 mediates catalysis. They finally characterize three disease mutants of Syn1, which show reduced activity or aggregate.

The paper is well drafted, the nanobody-aided structure determination of Syn1 is elegant and the kinetic experiments appear overall sound. A question to discuss is whether the novelty of the structural data is sufficient for publication in eLife. The structure of yeast synaptojanin is known for a long time and it seems rather similar to the human counterpart (although this is difficult to judge from this manuscript). Furthermore, many features of substrate binding and catalysis were already deduced from the product-bound yeast synaptojanin structure. I have also some questions/doubts about the substrate-bound structure.

With regard to the novelty of the data presented in our manuscript we would like to emphasize that this is the first structure of the human Synj1 protein. Considering the importance of this protein as a potential target for drug design, a structure of the real human protein rather than a homologue will be crucial for any structure‐based drug design effort. Also, note that we show in our study that the substrate specificity profile of the yeast homologue, SPSynj, differs from that of the human Synj1, which shows that SPSynj is not an optimal model system, again emphasizing the importance of a structure from the human protein.

We also provide the very first structure of any 5‐phosphatase protein in complex with the real substrate. In the revised document we carefully re‐processed and reanalyzed the diffraction data, suggesting a higher resolution for the substrate‐bound structure of 2.73 Å, and the PDB validation report shows the good quality of the structure compared to structures of similar resolution. Furthermore, the only objective criterion to address the question of data quality in our believe is to look at the electron density around the bound ligand. We have made a figure showing the electron density around the ligand in our structure and that of SPSynj in complex with the product inositol‐(1,4)bisphosphate (Author response image 3). For comparison we provide the 2Fo‐Fc and Fo‐Fc maps around the ligands of the respective structures as well as the omit maps. Both maps were calculated in exactly the same way for straightforward comparison. I hope that the reviewer will appreciate that (i) the position of the diC8‐PI(3,4,5)P₃ substrate in our structure is very clearly and unambiguously defined and also the scissile 5‐phosphate group can be modelled in an unambiguous way without the danger of any overfitting; (ii) the ligand in the SPSynj structure is clearly much less well defined in the density and modelling seems rather ambiguous. As discussed in the revised document, careful analysis of the electron density in our re‐processed structure of human Synj1 in complex with diC8‐PI(3,4,5)P₃ now also reveals density that could account for the nucleophilic water molecule, adding to the information with regard to the mechanism.

Finally, we also performed a very thorough kinetic analysis, showing for the first time the direct contribution of the 4‐phosphate group to catalysis rather than substrate binding. Together with the structural information, this allowed us to propose two potential mechanisms for this enzyme. Additionally, we also performed a first thorough kinetic characterization of disease‐associated mutations showing a clear link between the effect on catalysis and the severity of disease.

Author response image 3

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1) Detailed structural comparisons of Syn1 to other 5'-phosphatases (including rmsd) are missing to judge similarities and recognize potential differences between the structures. A structural comparison of the catalytic centers of some 5' phosphatases, including human and yeast synaptojanin with bound ligands, complementing Supplementary file 1, would allow the reader to appreciate (potentially) new mechanistic insights from the Syn1 structure.

An additional supplementary figure (Figure 2—figure supplement 3) has been added to the revised manuscript, showing the structural superposition of our structure with the 5PPase‐domain of INPP5B, SHIP2, OCRL, INPP5E and SPSynj. Also, an extra supplementary table (Supplementary file 1) is added to the manuscript, displaying rmsd values for structural superposition and sequence identities between the relevant 5PPases.

2) A major point of the paper is the substrate bound to Syn1. However, it remains unclear why the substrate has not been turned over after an overnight incubation at 20 C. The authors speculate about a missing catalytic water, but this cannot explain missing catalysis since the substrate-binding site is apparently accessible, in particular for water. Is the crystallization condition incompatible with catalysis? Does nanobody-binding reduce catalysis? Is there at all any substrate left after overnight incubation with the crystals? Without answering this, it remains uncertain whether the observed ligand is indeed a non-converted substrate molecule (see also next point).

We understand the concern raised by the reviewer that a substrate molecule would still be present in the active site after overnight incubation. This question is in part triggered by a lack of detail in the description of the cryo‐solution in our initial manuscript. In contrast to the apo‐structure, for the diC8PI(3,4,5)P₃‐bound structure the cryo‐solution was also supplemented with 1 mM of the substrate diC8PI(3,4,5)P₃. As we now explain in detail in our revised manuscript, crystals were transferred to the cryosolution for approximately 30 seconds prior to flash‐freezing in liquid nitrogen. So, it is highly likely that during short soaking in the cryo‐liquid a new substrate molecule re‐entered the active site of one of the protein molecules and was then trapped by flash freezing. This substrate is present with full occupancy and has very clearly determined density, as shown in the refined and omit density maps (see Author response image 1). These issues are now clarified and discussed in the revised document. In addition, the orientation of the substrate in the active site is clearly physiologically relevant as witnessed by the orientation of catalytically important residues vis‐à‐vis the substrate and the high correlation between the structure and our mutagenesis/kinetics results.

As suggested by the reviewer, we also addressed the question on whether either the presence of the nanobody and/or the crystallographic condition (pH) could affect catalysis. First, using the same method as described in the Materials and methods section of the manuscript, we compared the Michaelis‐Menten kinetics for the substrate diC8‐PI(3,4,5)P₃ for Synj1_528‐837 in either absence or presence of an excess of Nb15 (Author response image 4A). This analysis shows that binding of Nb15 induces a small decrease in the overall catalytic efficiency (k_cat/K_M) of the enzyme by a factor of 3.5, potentially by affecting the enzyme dynamics. Secondly, we tested the effect of the buffer and pH of the mother liquor used to crystallize the protein, by comparing enzyme catalysis in HEPES buffer with pH 7.5 and citrate buffer with pH 5.5. This analysis shows a decrease in catalytic efficiency by a factor of 12 at the pH used to crystallize the protein. Cumulative this shows that substrate turnover is slowed down under the conditions used to crystallize the protein, which could have aided in trapping the substrate in a non‐hydrolyzed form. However, this also shows that the enzyme is still catalytically competent under the conditions used, thus proving the relevance of our structure.

Author response image 4

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3) Crystallographic data of the substrate-bound crystals: Why is the signal to noise value so low for the substrate-bound crystals? Why did the authors apply anisotropic corrections to their data? Did the crystals diffracted anisotropically – if yes, provide some more information, e.g. to which resolution along a*,b*,c*. Judging from the spherical completeness, the dataset rather correspond to a 3.x A resolution dataset in terms of reflection numbers.

In the revised document we carefully re‐processed and re‐analyzed the diffraction data for the crystal of the substrate‐bound protein. This analysis suggested an even higher resolution for the substratebound structure of 2.73 Å. These resolution limits were automatically set using the autoPROC pipeline (https://www.globalphasing.com/autoproc/manual/autoPROC4.html), using a diffraction cut‐off of I/sigI = 1.4. Since the anisotropy analysis by STARANISO indeed indicated anisotropic diffraction, the anisotropy correcteddata from STARANISO was used, which was automatically generated in the autoPROC pipeline. The anisotropic diffraction is shown in Appendix 1—figure 1, which was generated by STARANISO.

The diffraction limits along the axes of the ellipsoid fitted to the diffraction cut‐off surface used by STARANISO for the Synj1_528‐873‐diC8‐PI(3,4,5)P₃ data are:

2.86 Å along 0.043 a* + 0.999 c*

2.72 Å along b*

3.14 Å along ‐0.974 a* + 0.228 c*

All details about the processing and refinement strategy and the diffraction limits along the axes of the ellipsoid are now provided in the Materials and methods section of the revised document and Appendix 1—figure 1, which shows this information for both structures.

The application of the anisotropy correction by STARANISO provided an outstanding improvement of the data statistics as well as the map quality. Author response table 1 shows that the dataset would correspond to approximately 3.0 Å resolution without anisotropy correction, based on CC(1/2) and on I/sigI = 1.4 cut‐off criteria. Anisotropic correction improved this to 2.73 Å resolution.

^a Values in parentheses are for the high‐resolution shell.

While resolution limits are by default somewhat arbitrary and the optimal cut‐off criteria (if any at all) are being debated in the field, the real criterion for judging the quality of data and for comparing different processing strategies lies in the final density. As can be judged from Author response image 5, the level of detail in the unbiased omit map around the ligand improved after using STARANISO and the quality of this unbiased omit map allowed to place the substrate with high confidence in the active site.

Author response image 5

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It is astonishing that for such low resolution, 91 water molecules with an average B-factor of only 34 Angstroms could be modelled (I would hardly expect to see any visible water molecules at this resolution). Is the model overfitted? Any explanation for the wide gap of Rwork and Rfree? What was the exact refinement strategy? Overall, the data quality of the substrate-bound crystals is borderline to accurately evaluate ligand binding, in particular if the identity of the bound ligand is not 100% clear.

After re‐analysis (as outlined above) and re‐refinement of the data for the Synj1_528‐873‐diC8‐PI(3,4,5)P₃ structure, the obtained resolution was set at 2.73 Å. The refinement strategy consisted of iterative cycles of manual building and refinement with Phenix.Refine. Initial rounds of the refinement strategy included XYZ coordinate, real‐space and individual B‐factors refinement, and towards the final cycles optimization of the X‐ray/stereochemistry weights was included. Finally, the structure was submitted to the PDB‐REDO server (Joosten et al., 2014) for a final optimization. After, critical assessment of all the water molecules, some water molecules were deleted which led to a final amount of 71. This seems a very reasonable amount of water molecules considering the resolution and the large amount of amino acid residues in the asymmetric unit (3 Synj1_528‐873 chains and 3 Nanobody chains). For a comparison of expected water molecules per amino acid residues at different resolution limits we could e.g. refer to Carugo, Oliviero, and Domenico Bordo. "How many water molecules can be detected by protein crystallography?." Acta Crystallographica Section D: Biological Crystallography 55, no. 2 (1999): 479‐483.

In the final refined Synj1_528‐873‐diC8‐PI(3,4,5)P₃ structure the R and R_free values are 19.88 % and 25.74%, respectively, corresponding to a R‐R_free gap of 5.86%. As can be judged from statistics of R, R_free and RR_free gaps from structures of similar resolution (Author response table 2, as given by phenix.r_factor_statistics), this gap is perfectly expected for a good structure at a resolution of 2.73 Å resolution.

We would also like to point out that the electron density maps in the final structures look very convincing (see e.g. Author response image 6 for electron density in representative parts of the structure) and allowed to unambiguously model the substrate diC8‐PI(3,4,5)P₃ as shown in Author response image 1, 3 and 5.

Finally, we also attached the PDB validation reports with this re‐submission, showing the good quality of our structures in comparison to other structures of similar resolution.

Author response image 6

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Reviewer #3:

The authors report crystal structures of the apo and substrate bound versions of the inositol 5-phosphatase domain of human synaptojanin-1 (Synj1), crystallized in the presence of a nanobody. The overall structure is similar to those of several previously solved 5-phosphatase enyzmes from yeast and humans. The main addition to the field in the current structure with respect to past work is that a substrate is present in this case, whereas previous structures were apo or product-bound.

We thank the reviewer for his assessment and comments.

For the comment with regard to the novelty of our findings we would make the following remarks (see also our answer to reviewer 2):

This is the first structure of the human Synj1 protein. Considering the importance of this protein as a potential target for drug design, a structure of the real human protein rather than a homologue will be crucial for any structure‐based drug design effort. Also, note that we show in our study that the substrate specificity profile of the yeast homologue, SPSynj, differs from that of the human Synj1, which shows that SPSynj is not an optimal model system again emphasizing the importance of a structure from the human protein.

We also provide the very first structure of any 5‐phosphatase protein in complex with the real substrate. In the revised document we carefully re‐processed and re‐analyzed the diffraction data, suggesting a higher resolution for the substrate‐bound structure of 2.73 Å. As discussed in the revised document, careful analysis of the electron density in our re‐processed structure of human Synj1 in complex with diC8‐PI(3,4,5)P₃ now also reveals density that could account for the nucleophilic water molecule, adding to the information with regard to the mechanism.

Finally, we also performed a very thorough kinetic analysis, showing for the first time the direct contribution of the 4‐phosphate group to catalysis rather than substrate binding. Together with the structural information, this allowed us to propose two potential mechanisms for this enzyme. Additionally, we also performed a first thorough kinetic characterization of disease‐associated mutations showing a clear link between the effect on catalysis and the severity of disease.

The structure contains 3 catalytic chains per asymmetric unit. The ligand-bound form was obtained by soaking apo crystals in a di-C8 form of the substrate phosphoinositide. The structure of the complex was determined at 2.9 Å resolution. Detail on the catalytic geometry is limited at this resolution. Water and metal ion positions are not visualized at all, or with limited accuracy.

As explained above, in our revised manuscript we have carefully re‐analyzed our data processing and refinement of the Synj1 structure in complex with diC8‐PI(3,4,5)P₃ suggesting an even higher resolution for this substrate‐bound structure of 2.73 Å. Subsequently, we carefully analyzed the unbiased electron density in the active site of the three protein molecules in the asymmetric unit. The omit map for molecule E very clearly shows electron density for diC8‐PI(3,4,5)P₃, including the phosphate groups on position 3, 4 and 5, allowing us to unambiguously model this compound (see unbiased omit map and 2Fo‐Fc and residual Fo‐Fc maps after refinement in Author response image 1). The orientation of the substrate in the active site is clearly physiologically relevant as witnessed by the orientation of catalytically important residues vis‐à‐vis the substrate and the high correlation between the structure and our mutagenesis/kinetics results. The excellent quality of the density in the active site of chain E also revealed additional electron density that could be accounted for by the nucleophilic water molecule. We attached the PDB validation reports with this re‐submission, showing the good quality of our structures in comparison to other structures of similar resolution.

Only one of the three catalytic chains binds to substrate. I did not see an explanation for the lack of binding to the other two chains. The 5-phosphate is still present on the substrate, despite that catalytically active wild-type enzyme was used. The authors present a detailed scheme for the catalytic mechanism on the basis of their structure, but I did not see an explanation for the failure of the enzyme to hydrolyze the 5-phosphate. I would speculate that the pH 5.0 of the crystallization could account for this. Have the authors attempted to adapt the crystals to neutral pH? Is a non-hydrolyzable substrate analogue available, or could it be synthesized? (I appreciate that phosphoinositide synthetic chemistry is non-trivial.) It would certainly be helpful for the field to know the geometry of a catalytically productive substrate complex, yet I am not convinced that the authors have as yet succeeded in obtaining such a structure. In my opinion, the conclusions concerning enzyme mechanism are, at this stage, over-interpreted.

We understand the concern raised by the reviewer that a substrate molecule would still be present in the active site after overnight incubation. This question is in part triggered by a lack of detail in the description of the cryo‐solution in our initial manuscript. In contrast to the apo‐structure, for the diC8PI(3,4,5)P₃ structure the cryo‐solution was also supplemented with 1 mM of the substrate diC8PI(3,4,5)P₃. As we now explain in detail in our revised manuscript, crystals were transferred to the cryosolution for approximately 30 seconds prior to flash‐freezing in liquid nitrogen. So, it is highly likely that during short soaking in the cryo‐liquid a new substrate molecule re‐entered the active site of one of the protein molecules and was then trapped by flash freezing. This substrate is present with full occupancy and has very clearly determined density, as shown in the refined and omit density maps (see Author response image 1, included in this rebuttal letter). These issues are now clarified and discussed in the revised document.

As suggested by reviewers 2 and 3 we also investigated the influence of the Nb and the pH of the crystallization conditions on Synj1 catalytic activity. As detailed in our answer to reviewer 2 (see Author response image 4A), we find that the presence of the Nb induces a small decrease in the overall catalytic efficiency (k_cat/K_M) of the enzyme by a factor of 3.5, potentially by affecting the enzyme dynamics, while a decrease of pH from 7.5 to 5.5 causes a decrease in catalytic efficiency by a factor of 12. Cumulative this shows that substrate turnover is slowed down under the conditions used to crystallize the protein, which could have aided in trapping the substrate in a non‐hydrolyzed form. However, this also shows that the enzyme is still catalytically competent under the conditions used, thus showing the relevance of our structure.

Concerning the presence of clear density for the non‐hydrolyzed substrate in only one of the active sites, as stated above, it is true that the other two active sites (mainly chain C) show only some continuous density which might account for the presence of a ligand at low occupancy (see Author response image 1). These differences in occupancy could reflect differences in kinetics of access of the substrate or release of products from the three active sites, depending on their local environment in the crystal packing, and/or could reflect the substrate being hydrolyzed to different degrees.

This being said, we think it is not unusual that ligands are only found occupying a subset of the binding pockets present in the asymmetric unit, despite being seemingly accessible via solvent channels. A survey at the CCP4 bulletin board and through the literature yielded several similar cases, some of which we mention here:

– Structure of penicillin‐binding protein 2A in complex with a quinazolinone (PDB 6Q9N), where only one of the two protein chains in the active site contained density for the compound (Janardhanan et al., 2019. DOI: 10.1128/AAC.02637‐18).

– Structure of the N‐terminus of a glycosidase from Cellulosimicrobium cellulans in complex with 1deoxymannojirimycin (DMNJ) (PDB 4AQ0), where only one of the two chains in the asymmetric unit contained the ligand DMNJ, while the other contained a solvent molecule (Bis‐Tris propane) (Tiels et al., 2012. DOI: 10.1038/nbt.2427).

– Structure of NADP⁺‐specific isocitrate dehydrogenase from Corynebacterium glutamicum (PDB 3MBC) where the co‐enzyme was present in only one of the two biologically monomeric NCS‐related protein molecules in the asymmetric unit (Sidhu et al., 2011. DOI: 10.1107/S0907444911028575).

– Structure of holo‐transaminase soaked with an acceptor substrate (α‐ketoglutarate) (PDB 5E25), with nice ligand density observed in only one of the three monomers in the asymmetric unit (currently not published).

– Multiple examples in a paper reporting the in crystallo‐screening for discovery of human norovirus 3C‐like protease inhibitors (Guo et al. J Struct Biol. 2020 4:10003. DOI: 10.1016/j.yjsbx.2020.100031), e.g. PDB 6T49, 6T6W, 6T2I.

The paper is, apart from the omissions noted above, very thorough. It reads like a review article in the Introduction and Discussion. The results includes more detail than is standard. Overall the paper would be more readable if it were edited to about half of the current length.

As suggested by reviewer 2 and 3, we significantly shortened the manuscript in certain parts, taking care not to lose information.

Author details

1. Jone Paesmans

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Data curation, Formal analysis, Investigation, Writing - original draft, Writing - review and editing

   Contributed equally with

   Ella Martin

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3292-4609

2. Ella Martin

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Data curation, Formal analysis, Investigation, Writing - original draft, Writing - review and editing

   Contributed equally with

   Jone Paesmans

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-9607-7074

3. Babette Deckers

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Data curation, Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3855-4776

4. Marjolijn Berghmans

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-8699-6915

5. Ritika Sethi

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

6. Yannick Loeys

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

7. Els Pardon

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Supervision, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2466-0172

8. Jan Steyaert

   1. VIB-VUB Center for Structural Biology, Brussels, Belgium
   2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

   Contribution

   Supervision, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3825-874X

9. Patrik Verstreken

   1. VIB-KU Leuven Center for Brain and Disease Research, Leuven, Belgium
   2. KU Leuven, Department of Neurosciences, Leuven Brain Institute, Leuven, Belgium

   Contribution

   Supervision, Methodology, Writing - review and editing

   Competing interests

   Reviewing editor, eLife

10. Christian Galicia

    1. VIB-VUB Center for Structural Biology, Brussels, Belgium
    2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

    Contribution

    Formal analysis, Supervision, Investigation, Writing - original draft, Writing - review and editing

    For correspondence

    Christian.Galicia.Diaz.Santana@vub.be

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0001-6080-7533

11. Wim Versées

    1. VIB-VUB Center for Structural Biology, Brussels, Belgium
    2. Structural Biology Brussels, Vrije Universiteit Brussel, Brussels, Belgium

    Contribution

    Conceptualization, Formal analysis, Supervision, Funding acquisition, Investigation, Writing - original draft, Project administration, Writing - review and editing

    For correspondence

    wim.versees@vub.be

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-4695-696X

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