Mycobacteria use a set of specialized secretion systems called ESX to transport proteins across their complex, diderm cell walls (Gröschel et al., 2016). Originally described as virulence factors in Mycobacteria tuberculosis (Guinn et al., 2004; Hsu et al., 2003; Lewis et al., 2003; Stanley et al., 2003), orthologs of ESX have since been discovered in most Gram-positive bacteria (Bottai et al., 2017), and are more generally referred to as Type VII secretion systems (Bitter et al., 2009). In mycobacteria there are five paralogous ESX operons (ESX 1–5) each of which encodes an inner membrane translocon complex consisting of three conserved Ecc proteins: EccB, EccC, and EccD. A fourth protein, EccE is conserved in all ESX operons except the ancestral ESX-4 operon and is also considered a part of the ESX translocon complex as it copurifies with EccB, EccC, and EccD (Houben et al., 2012). All Type VII secretion systems translocate proteins in the WXG100-superfamily, which share a common two-helix hairpin structure and are found as homo- or heterodimers (Poulsen et al., 2014) and are mutually dependent for secretion with other substrates (Fortune et al., 2005). In contrast to the general secretory apparatus (Sec), ESX substrates have been shown to be secreted in their folded, dimeric state (Sysoeva et al., 2014).

Structural and functional information has been reported for truncated and isolated, soluble domains of the ESX translocon complexes and their homologs (Korotkova et al., 2015; Korotkova et al., 2014; Renshaw et al., 2005; Rosenberg et al., 2015; Strong et al., 2006; Wagner et al., 2016; Wagner et al., 2014; Wagner et al., 2013; Zhang et al., 2015; Zoltner et al., 2016). A low resolution, negative stain electron microscopy structure of ESX-5 shows a translocon complex assembled into a hexamer (Beckham et al., 2017). Structures of other proteins encoded in ESX operons including secreted substrates (Ilghari et al., 2011), substrate chaperons (Ekiert and Cox, 2014), and the protease MycP (Solomonson et al., 2013) have been solved. Despite revealing important functional information about ESX, structures of overexpressed and isolated proteins are insufficient to understand the regulated secretion of fully folded substrates. We therefore undertook structural studies of an endogenously expressed ESX-3 complex from the model organism M. smegmatis using cryo-electron microscopy (cryo-EM). During the preparation of this work for publication, a similar structure of the ESX-3 system expressed from a plasmid was published by another group (Famelis et al., 2019).

The ESX-3 translocon complex is important for iron acquisition (Serafini et al., 2013; Siegrist et al., 2009), cell survival (Tinaztepe et al., 2016), and virulence in pathogenic mycobacteria (Tufariello et al., 2016), and its role in iron homeostasis is conserved in the model system, M. smegmatis (Siegrist et al., 2009). The ESX-3 translocon complex proteins are transcribed in a single operon (Li et al., 2017), and expression of the ESX-3 operon is dependent on the transcriptional regulator IdeR, which controls iron metabolism (Rodriguez et al., 2002) and is required for growth in the human pathogen M. tuberculosis (Pandey and Rodriguez, 2014). The ESX-3 operon is 67% identical between the non-pathogenic model organism M. smegmatis and the pathogen M. tuberculosis over the 4354 amino acids of the ESX-3 operon. This high degree of conservation and essential role in cell growth makes ESX-3 an important candidate for small molecule inhibition (Bottai et al., 2014), as blockade of ESX-3 will both inhibit virulence in M. tuberculosis and kill a broad range of pathogenic mycobacteria.

A major innovation made possible by the dramatic improvements in cryo-EM (Cheng, 2018) is the ability to examine challenging protein samples at atomic resolution, even when samples are only available at low concentrations. When coupled with recent genetic manipulations that allow for facile insertion of chromosomal epitope and purification tags (Murphy et al., 2018), cryo-EM now holds the promise of routine structural characterization of many endogenously expressed protein complexes not previously tractable by structural techniques. We undertook the purification of the ESX-3 complex from the native host without the need for overexpression. To facilitate purification of the endogenous translocon complex, a cleavable EGFP tag was inserted into the chromosome of M. smegmatis mc(2)155 (wild type) and ΔideR (Dussurget et al., 1996) strains at the C-terminus of EccE₃ via the ORBIT method (Murphy et al., 2018) (Figure 1A, Figure 1—figure supplement 1). EccE₃ is the final gene in the 11 gene long ESX-3 operon making insertion at this site less likely to disrupt regulation and expression of the operon. Deletion of the gene for the iron acquisition regulator IdeR greatly increases chromosomal expression of ESX-3 from negligible amounts of protein to a yield sufficient for purification and structure determination (Figure 1—figure supplement 2A). Components of ESX-3 were pulled down using an anti-GFP nanobody (Rothbauer et al., 2008) and the EGFP tag was proteolytically cleaved. After size exclusion chromatography, the peak fractions were pooled and analyzed biochemically and by cryo-EM .

Global structure of the ESX-3 dimer

Four components of the ESX-3 translocon complex EccB₃, EccC₃, EccD₃ and EccE₃ were stably affinity-purified as a large molecular weight species of about 900 kDa (Figure 1B and C, Figure 1—figure supplement 2B). The sample was imaged by cryo-EM and reconstructed revealing a dimeric structure, which can be divided into four areas: the flexible periplasmic multimerization domain, the stable transmembrane region, the stable upper cytoplasmic region, and the flexible lower cytoplasmic motor domain (Figure 1D, Table 1). While the peak fraction does not contain particles of a larger size consistent with higher order oligomers, thorough examination of the void volume revealed a small number of particles in a higher oligomeric state (Figure 1—figure supplement 3A–E). The resolution of the ESX-3 dimer varies substantially in different parts of the electron microscopy map and this heterogeneity was partially resolved through data processing (Figure 1—figure supplement 4 and Figure 1—figure supplement 5). Initially, the entire ESX-3 dimer was reconstructed to 4.0 Å resolution (Figure 1—figure supplement 6A). Using symmetry expansion, and focused classification and refinement techniques, the resolutions of targeted regions of the ESX-3 complex were improved to 3.7 Å for the transmembrane region and upper cytoplasmic region (Figure 1—figure supplement 6B–D), 5.8 Å for the periplasmic region (Figure 1—figure supplement 6E), and ~7 Å resolution for the lower cytoplasmic region (Figure 1—figure supplement 6F). The highest resolution maps for each region were combined and filtered to the threshold of the lowest resolution map to form an overall 10 Å combination map for the entire ESX-3 dimer.

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Table 1

Data Collection
Collection	Initial Screening	Collection 1	Collection 2	Void peak
Microscope	Talos Arctica	Titan Krios	Titan Krios	Talos Arctica
Voltage (kV)	200	300	300	200
Detector	Gatan K2	Gatan K2	Gatan K2	Gatan K3
Pixel size (Å/pixel)	1.14	0.82	0.82	0.9
Exposure Time (s)	9	10	10	11.7
Electron dose (e-/Å2)	63	80	67	58
Defocus range (µm)	1.5-2.5	0.4-1.2	0.6-1.4	1.5-2.5
Number of micrographs	2,499	2,705	4,632	1,215
Consensus Reconstruction
Data Set	Initial Screening	Collection 1 & 2		Void peak
Software	Relion 2.1, cisTEM, and cryosparc	Relion 3.0		cisTEM
# of particles, picked	240,000	778,149		259,333
# of particles post, Class2D	138,000	554,901		640
# of particles post, Class3D	46,830	362,438		NA
# of particles post, skip align Class3D	NA	90,479		NA
Symmetry	C1	C1		NA
Map sharpening B-factor (Å2)	NA	-160		NA
Final resolution (Å)	4.7 Å	4.0 Å		NA
Focused Refinements
Location of focus	# of particles	Resolution
Protomer i	76,967	3.8
Protomer ii	90,479	3.8
Symmetry expanded protomer	52,067	3.7
Periplasmic EccB3	70,000	5.8
ATPase 1, 2, and 3	30,000	7

The ESX-3 dimer is comprised of ten total proteins, two copies each of EccB₃, EccC₃, and EccE₃ and four copies of EccD₃. Two pseudo-symmetric protomers referred to as i and ii, combine to form the ESX-3 dimer. Each protomer contains one copy of EccB₃, EccC₃, and EccE₃ and two conformationally distinct copies of EccD₃, referred to as EccD_3-bent and EccD_3-extended (Figure 1E) based on their highly asymmetric conformations. At 3.7 Å resolution, it was possible to build de novo atomic models for all observable amino acids in the transmembrane and upper cytoplasmic regions, except the two transmembrane helices of EccC₃ (Figure 1E and Supplementary file 1). The lower resolution regions of density, the EccC₃ transmembrane helices, EccC₃ ATPase 1, 2, and 3 domains and the EccB₃ periplasmic domain, were flexibly fit using homology models. Using this hybrid approach, a model of the entire ESX-3 dimer has been produced (Figure 1F).

EccD₃ forms a homodimer that encloses a large hydrophobic cavity

There are two copies of EccD₃ in each ESX-3 protomer (Figure 2A). The ubiquitin-like N-terminal domain of each EccD₃ molecule interacts with EccE₃ and EccC₃ in the cytoplasm, and a long linker joins the soluble domain of EccD₃ to 11 transmembrane helices (Figure 2B and Figure 2—figure supplement 1A–F). The four EccD₃ molecules account for 44 of the total 54 transmembrane helices observed in the ESX-3 dimer. A distinct transmembrane cavity is formed by dimerization of the two copies of EccD₃ in each protomer with a cross-sectional diameter of ~20×30 Å without significant regions of constriction. Transmembrane helices 1, 9 and 10 interact across the cavity dimer interface in a tight bundle making passive lipid transport into the membrane from the cavity unlikely (Figure 2C). The inner surface of the periplasmic half of the cavity is composed primarily of hydrophobic residues and in our maps, eight extended densities consistent with hydrophobic lipid tails or detergent molecules line the periplasmic inner face of the cavity (Figure 2C). In contrast, the cytoplasmic face of the cavity has several polar residues and ordered hydrophobic densities are not visible.

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In the cytoplasm below the membrane, a stable, upper cytoplasmic region is formed by interactions between the soluble domains of EccD_3-bent, EccD_3-extended, EccE₃, and EccC₃. The linker joining the cytoplasmic ubiquitin-like domain to transmembrane helix 1 (residues 100–127) of EccD₃ conserves a high sequence identity throughout evolution (Ashkenazy et al., 2016), yet it adopts two distinct secondary structures resulting in the asymmetric placement of the cytoplasmic domains of EccD₃ (Figure 2D and Figure 2—figure supplement 1A–G). In EccD_3-bent, residues 100–127 are bent, folding into an α-helix and forming a nexus of stabilizing contacts with EccB₃ and EccC₃ (Figure 2—figure supplement 1H). In EccD_3-extended, residues 100–127 are extended and fold into a shorter α-helix that interacts with EccE₃ and the cytoplasmic domain of EccD_3-bent (Figure 2—figure supplement 1I). This conformational flexibility suggests that if residues 100–127 were released from their associations with EccC₃ and EccE₃ they could rearrange into the alternative bent or extended conformation with little energetic barrier.

EccC₃ and EccE₃ make extensive, stabilizing interactions with the asymmetric, cytoplasmic domains of EccD₃

The next component of the stable upper cytoplasmic region is EccE₃. EccE₃ is positioned at the front of the ESX-3 dimer (Figure 3A), where the conserved transmembrane helix 1 of EccE₃ interacts with helix 11 of EccD_3-bent in the membrane. Helix 1 is followed by a second EccE₃ transmembrane helix, a linker helix, and then extends into the cytoplasm (Figure 3B and C and Figure 3—figure supplement 1A–D). The anti-parallel β-sheets of the cytoplasmic domain of EccE₃ have weak structural homology to glycosyl transferase proteins, however, the nucleotide binding pocket is absent in EccE₃, leaving it incapable of performing this function (Figure 3—figure supplement 1E and F, Supplementary file 2). EccE₃ does not have another obvious ligand or catalytic site. Two conserved helices in the cytoplasmic region of EccE₃ between amino acids 133 and 163 form extensive stabilizing interactions with both subunits of EccD₃ (Figure 3D, Figure 3—figure supplement 1G). These interactions hold the flexible linker of EccD_3-extended in the extended conformation and sterically hinder EccD_3-extended from assuming the bent conformation (Figure 3—figure supplement 1H). EccE₃ does not form direct protein-protein interactions with either EccB₃ or EccC₃ suggesting the contacts with EccD_3-extended and EccD_3-bent are extremely stable as EccE₃ was the tagged protein used to immunoprecipitate the ESX-3 dimer.

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The final component of the stable upper cytoplasmic region is the domain of unknown function (DUF) of EccC₃. EccC₃ extends from the membrane into the upper and lower cytoplasmic regions (Figure 4A). Amino acids 1–33 and 94–403 of EccC₃ were built de novo into the higher resolution region of the electron microscopy map revealing the structure of the DUF domain (Figure 4B, Figure 4—figure supplement 1A–C). The de novo model of the DUF has the typical Rossman fold of a nucleotide hydrolysis domain (Figure 4—figure supplement 1D and E) and its closest homolog by Dali search is the ATPase 1 domain of EccC of T. curvata. It is linked to the transmembrane domains by a long helical bundle making extensive contacts with the flexible linker region of EccD_3-bent (Figure 4C). The DUF makes additional stabilizing contacts with the ubiquitin-like domains of EccD_3-bent and EccD_3-extended in the cytoplasm (Figure 4D).

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When the transmembrane and upper cytoplasmic regions are compared between protomers, only the transmembrane helices of EccC₃ and the N-terminal tail of EccB₃ differ (Figure 4—figure supplement 2A and B), otherwise the protomers are superimposable. All four EccC₃ transmembrane helices were modeled at 6 Å resolution through a combination of homology modeling and molecular dynamics. In protomer i, transmembrane helix 2 forms lipid mediated hydrophobic interactions with the transmembrane helix of EccB₃ in protomer i, and transmembrane helix 1 interacts with transmembrane helix 2 of EccC₃ in protomer ii. Transmembrane helix 1 of EccC₃ in protomer ii is shifted relative to the protomer i conformation and does not directly interact with other proteins.

EccB₃ extends into the periplasm and stabilizes dimer formation

The ESX-3 dimer is stabilized by cross-protomer interactions formed by the two EccB₃ proteins. EccB₃ begins in the cytoplasm with a flexible N-terminal tail leading into a linker helix, followed by a single-pass transmembrane helix, and an extended periplasmic domain (Figure 5A and B, Figure 5—figure supplement 1A and B). The N-terminal tail of EccB₃ from protomer i forms extensive cross-protomer contacts with EccB₃, EccC₃, EccD_3-bent and EccD_3-extended from protomer ii (Figure 5C, Supplementary file 3). The linker helix of EccB₃ forms further protein-protein interactions with EccC₃ and EccD_3-bent. The transmembrane helix of EccB₃ interacts with transmembrane helix 11 of EccD_3-extended. Two hydrophobic tails consistent with a lipid or detergent molecules link the transmembrane helix of EccB₃ to transmembrane helix 2 of EccC₃ (Figure 5D). The two EccB₃ periplasmic domains share a large interaction interface across the protomers further stabilizing dimerization. Homology models of two EccB₃ proteins can be docked into the periplasmic domain (Figure 5—figure supplement 1C); however, this region is not resolved sufficiently to identify specific interactions. The majority of cross-protomer interactions involve EccB₃, suggesting the periplasmic domain is essential for oligomerization.

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The ESX-3 structure presented here is purified without the addition of substrates or nucleotide. It is therefore likely to be in a conformation representing the end of the translocation cycle, awaiting either the direct binding of substrates, the binding of nucleotide or both, to reset a substrate-binding competent state. By fitting our dimer structure into a prior low resolution envelope we suggest a model of the oligomeric state of the complex, in close agreement with Famelis et al. However, even allowing for major rearrangements in the more flexible regions of EccC₃ and EccB₃, a model built by the static trimerization of the experimentally determined dimer structure cannot itself explain the mechanism of action of ESX-3 secretion. The existence of an R-finger catalytic site for ATPase 1 of EccC₃ (Rosenberg et al., 2015) requires the R-finger of one protomer to insert into the active site of another protomer. Given the ~65 Å distance we observe between ATPase 1 domains, the completion and activation of the catalytic site of ATPase 1 by an R-finger will necessitate an extremely large rearrangement of the position of the ATPase domains. How might this rearrangement occur? We propose movements in the highly flexible EccD₃ linker lead to the release of EccE₃ and EccC₃ from their rigid positions, thus allowing for a rearrangement of the ATPase domains into an active conformation (Figure 6).

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Once EccC₃ assembles into an oligomeric state, the substrate proteins will need to translocate through the inner membrane. We have considered two models for how pore formation and transit might occur: 1) through a pore created by the oligomerization of EccC₃ and EccB₃ or 2) through the large cavity created by the dimerization of EccD₃. In the first model (Figure 6A), the resting state of an ESX translocon complex is a hexamer, with disordered EccC₃ ATPase domains free in the cytosol, stabilized by interactions with proteins not seen in the structures presented here (van Winden et al., 2016). It is possible the rare multi-dimer oligomeric state we see in the void volume, and also seen by Famelis et al., represents this state. In a hexamer model, the center of the multimer is formed by the transmembrane helices of EccC₃ and EccB₃, which create a cavity that could serve as a pore for translocation of substrates. These transmembrane helices are largely hydrophobic and do not contain obvious residues that would allow for the conductance of hydrated substrates. Thus the production of a protein transit channel would require either a large, conformational change in the transmembrane helices, likely facilitated by movements in the cytoplasmic domains of EccC₃, EccD₃ and EccE₃, or a novel mechanism of action for transit through the central pore.

A hexameric pore created by EccC₃ agrees well with the documented mechanism of action for motor ATPases in the additional strand catalytic E (ASCE) division of P-loop NTPases (Erzberger and Berger, 2006), which includes EccC₃. A hexameric pore also agrees with the proposed mechanism of action for other bacterial secretion systems, such as the Type IV secretion system VirD4 coupling protein (Gomis-Rüth et al., 2001; Hormaeche et al., 2002), which is related evolutionarily to EccC₃ (Iyer et al., 2004). The hexamer model is thus firmly grounded in the motor ATPase and bacterial secretion systems literature, although the oligomeric state of VirD4 has recently been called into question (Redzej et al., 2017) and remains controversial (Llosa and Alkorta, 2017).

In a second, more speculative model, EccD₃ dimers form a channel for translocation of substrates (Figure 6B). The large cavity found in the EccD₃ dimer is striking and by structural homology, is unlike any other membrane protein in the Protein Data Bank. In our density maps, the EccD₃ dimer cavity appears capped on the periplasmic side by a dense layer of lipids. In contrast, on the cytoplasmic side the cavity does not exhibit bound lipids due to the polar residues lining the lower half of the cavity. The large cavity is of sufficient diameter to transit a folded EsxG/H dimer, however given the strong hydrophobicity of the cavity the mechanism would not be mediated by water and would require a novel mechanism of secretion that has not been seen in other bacterial secretion systems. It is also possible that the cavity exists to transit a non-protein substrate such as a specific mycobacterial lipid. The ability to transport non-protein substrates could resolve some of the mysteries that remain about the relationships between ESX systems, cell wall stability, lipid content, and nutrient acquisition (Barczak et al., 2017; Bosserman and Champion, 2017; Siegrist et al., 2014; Tufariello et al., 2016).

As each protomer contains an EccD₃ cavity, the second model, proposing translocation through EccD₃, does not require hexamerization. However, this model is also not incompatible with hexamerization, which would not block a substrate path through EccD₃. Further, the role of the hexamer may not be to form a central channel for substrate transit. Rather, hexamerization could serve some other purpose. For example, it may tether functional dimers together, facilitate localization, or increase local concentration and allosteric control of enzymatic activity (Kuriyan and Eisenberg, 2007).

Although the ESX-3 structure presented here allows for mechanistic hypotheses about the transit of substrates across the inner-membrane, it does not provide sufficient information to allow for a structural model of transit across the outer mycomembrane. The EccB₃ periplasmic domain (Wagner et al., 2016) has been found to have similarity to the peptidoglycan binding phage protein PlyCB, which forms a ring inside the bacterial cell wall facilitating phage entrance into the cell. A hypothesis is that EccB₃ is anchored to a larger outer membrane complex, but the purification conditions we have employed remove proteins required for its stabilization.

The description of ESX-3 presented here agrees in both protein composition and structure with the work recently published by Famelis et al. Given the high sequence conservation among the ESX systems in mycobacteria and related actinobacteria, these structures likely represent an excellent framework for the structural modeling of the other ESX systems. Together, these structures provide a wealth of information about protein-protein interaction interfaces and ESX complex architecture, which can be used to guide structure-based drug design and to generate hypotheses for further mechanistic investigations.

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Acceptance summary:

This manuscript by Poweleit et al. describes the cryoEM structure of a natively purified ESX-3 (Type VII) secretion system from Mycobacterium smegmatis. Our understanding of the type VII secretion mechanism has been limited by the lack of high-resolution structural information of the fully assembled, ~900 kDa machine. Here the authors isolate the native ESX-3 complex from M. smegmatis and solve the structure by single-particle cryoEM in detergent. This is a beautiful and informative structure, and the analysis is rigorous. The work is significant because it corroborates the structural observations recently published by a competing group (Famelis et al., 2019), and provides a framework for understanding how this secretion machine transports its folded clients across the cytoplasmic membrane.

Decision letter after peer review:

Thank you for submitting your article "The native structure of an ESX translocon" for consideration by eLife. Your article has been reviewed by four peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Richard Aldrich as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This manuscript by Poweleit et al. describes the high-resolution structure of an ESX-3 complex of M. smegmatis obtained by cryo-electron microscopy. A virtually identical structure of the same complex has been published in Nature by Famelis et al. as the authors were preparing to submit this paper. In contrast to the structure published by Famelis et al., which was obtained by overexpressing the ESX-3 system from a plasmid, Poweleit et al. tagged the complex chromosomally, on the last gene (EccE) of the operon, to bypass potential transcriptional defects. While the presented work therefore represents a native structure, as opposed to the already published structure by Famelis and colleagues, the observed structural features/composition are the same for both structures. It was felt that the eLife policy on "scoops" protects the authors of this manuscript and that it therefore should be considered further after suitable revisions.

Essential revisions:

1) The authors fail to see any higher order oligomers, which have been previously reported in native (Houben et al., 2012, van Winden et al., 2016) and also plasmid expressed conditions (Famelis et al., 2019; Beckham et al., 2017). These publications have reported higher order oligomers in multiple mycobacterial species (M. bovis BCG, M. marinum and M. smegmatis) and from different type VII secretion systems (ESX-1, ESX-3 and ESX-5). Higher order multimerization has also been suggested/observed by the corresponding author in previously published work (Rosenberg et al., 2015) for the EccC ATPase. Importantly, Beckham et al. and Famelis et al. showed that these higher order oligomers are only present in the void peak fraction of the SEC analysis. In the current study it is unfortunately not mentioned which SEC fractions were used for the cryo-EM analysis, but it is suspected that the void peak fractions were not analyzed. To confirm that the native ESX-3 complex forms higher order oligomers, the authors should more thoroughly investigate the presence of these multimers, e.g. by including the void peak fractions in the analysis and/or analyzing the ESX-3 membrane complex by BN-PAGE analysis before SEC (including crosslinking).

2) The authors propose an alternative and contentious model for protein translocation through the membrane complex, i.e. through a hydrophobic cavity formed by two EccD subunits, which is (partially) filled with lipids. Indeed, the presence of such a large cavity in between these two subunits is striking. However, the surface of this cavity consists, except for a number of polar residues, solely of hydrophobic residues. In agreement with a hydrophobic interface, the structure reveals extra densities at the periplasmic part of the cavity suggesting the presence of lipids or detergent. The cytoplasmic half of the cavity does not reveal these extra densities, which could be caused by the presence of several polar residues here, as also discussed by the authors. However, lipids in this part of the cavity could have been removed during solubilization of the complex. To show that partially lipid-filled cavities have been observed also in other membrane complexes, the authors refer to a publication describing the presence of a similar lipid plug in the central cavity of a reconstituted rotor cylinder of the ATP synthase (Meiers et al. Febs Lett 2001). However, it is explicitly mentioned in this article that "As the detergent-purified c cylinder is completely devoid of phospholipids, these are incorporated into the central hole from one side of the cylinder during the reconstitution procedure". It therefore remains unclear whether such lipid-plugged membrane pores actually exists in vivo. In addition, Poweleit et al. propose a "lipid mediated" mechanism of protein translocation through this hydrophobic pore. It is difficult to envision though how lipids would be able to mediate translocation of folded substrate dimers with mostly hydrophilic surfaces. This would be a completely new mechanism of protein translocation not seen for any other protein secretion machineries. In addition, the authors speculate even further that besides (hydrophilic) proteins also hydrophobic molecules, such as lipids, could be transported through this cavity, while there is no clear evidence that type VII secretion systems are able mediate lipid transport. In conclusion, while it is indeed striking that such a large cavity is present in the ESX-3 complex, the authors should more cautiously discuss this model, by also mentioning the major problems with this model as discussed above. The translocation model of substrates through the central pore of a hexameric complex is far less speculative, as it is supported by experimental data, published by several groups (as also mentioned in the Discussion). This model should therefore get more emphasis.

3) The regulatory role of the "cytoplasmic bridge" is speculative and should be more carefully discussed. e.g. "multimeric cytoplasmic domain is poised to regulate the translocation of ESX substrates" and "EccC₃ and EccE₃ collaborate to buttress the cytoplasmic domains of EccD₃ and form a stable regulatory bridge".

Essential revisions:

1) The authors fail to see any higher order oligomers, which have been previously reported in native (Houben et al., 2012, van Winden et al., 2016) and also plasmid expressed conditions (Famelis et al., 2019; Beckham et al., 2017). These publications have reported higher order oligomers in multiple mycobacterial species (M. bovis BCG, M. marinum and M. smegmatis) and from different type VII secretion systems (ESX-1, ESX-3 and ESX-5). Higher order multimerization has also been suggested/observed by the corresponding author in previously published work (Rosenberg et al., 2015) for the EccC ATPase.

We simply do not have definitive proof of the “physiological” multimeric state of the protein complex and so decided not to speculate extensively about this important issue in our first submission. Prior work, well characterized in the editor’s review, and including our published work, has strongly suggested higher order structural and functional oligomerization. In particular, the requirement for a an “R-finger” mechanism for EccC ATPase activity – a hallmark of multimer formation – makes it clear that at some point in the secretion cycle, the ATPase 1 domains act as oligomers. In the structure we present in this manuscript the ATPase 1 domains, which have been presumed to be the active motor domains based on structural, biochemical and genetic data, would require a total of 65 Å of movement to come into an active position where the R-finger from one protomer is positioned to stimulate catalysis in the next subunit. The requirement for this extraordinarily large movement of the ATPase domains makes it difficult to presuppose the path of the ATPase domains during the secretion cycle. In our revision we comment on this issue in the Discussion and alter the model figure (revised Figure 6) to reflect the need for these molecular motions.

Importantly, Beckham et al. and Famelis et al. showed that these higher order oligomers are only present in the void peak fraction of the SEC analysis. In the current study it is unfortunately not mentioned which SEC fractions were used for the cryo-EM analysis, but it is suspected that the void peak fractions were not analyzed. To confirm that the native ESX-3 complex forms higher order oligomers, the authors should more thoroughly investigate the presence of these multimers, e.g. by including the void peak fractions in the analysis and/or analyzing the ESX-3 membrane complex by BN-PAGE analysis before SEC (including crosslinking).

In order to address this important concern, we have repeated the purification and collected a new cryo-EM data set of of the “void” fractions. These micrographs contain a majority of aggregated particles and are challenging to analyze, which is why we left them out of the first submission. However, there is a small population of monodispersed particles, which we have analyzed. The majority of these particles appear to be in the dimer state we have described, however, additionally, we see a small set of side views that are consistent with a larger particle of approximately 280 Å and that look similar to particle described in Famelis et al. In terms of the size, these particles could be interpreted as a hexameric form, but given the excellent distribution of views we observe in the dimer form we were surprised not to observe any evidence in the 2D class averages that would suggest symmetry higher than 2. It is certainly possible that these particles do not represent a fixed state but rather misincorporation of several dimers into a single micelle. We present these data in a new supplemental figure (Figure 1—figure supplement 4).

We have also attempted to crosslink the purified protein before running on the SEC and we do not see evidence of additional, significant crosslinking between dimers, in the purified preparation. We include this gel in Author response image 1 but not in the resubmitted manuscript.

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2) The authors propose an alternative and contentious model for protein translocation through the membrane complex, i.e. through a hydrophobic cavity formed by two EccD subunits, which is (partially) filled with lipids. Indeed, the presence of such a large cavity in between these two subunits is striking. However, the surface of this cavity consists, except for a number of polar residues, solely of hydrophobic residues. In agreement with a hydrophobic interface, the structure reveals extra densities at the periplasmic part of the cavity suggesting the presence of lipids or detergent. The cytoplasmic half of the cavity does not reveal these extra densities, which could be caused by the presence of several polar residues here, as also discussed by the authors. However, lipids in this part of the cavity could have been removed during solubilization of the complex.

As the reviewers note, the presence of a large cavity in the inner membrane, formed by the EccD dimer, is very unusual. In searching the literature and doing extensive homology searches, we have discovered no clear examples of similar membrane protein structures. In our structure, the periplasmic side of the EccD cavity is completely filled with detergent or lipids carried over from the purification, which would provide a mechanism for the “hole” to be sealed. Although we do believe that this cavity will eventually be shown to be of functional importance, as the reviewers appear to recognize, to fully understand the native state and function of EccD is beyond the scope of the current work. We have thus rewritten the Results and Discussion section to be more descriptive and less speculative.

To show that partially lipid-filled cavities have been observed also in other membrane complexes, the authors refer to a publication describing the presence of a similar lipid plug in the central cavity of a reconstituted rotor cylinder of the ATP synthase (Meiers et al. Febs Lett 2001). However, it is explicitly mentioned in this article that "As the detergent-purified c cylinder is completely devoid of phospholipids, these are incorporated into the central hole from one side of the cylinder during the reconstitution procedure". It therefore remains unclear whether such lipid-plugged membrane pores actually exists in vivo.

We have removed the reference to the c cylinder as we do not want to confuse the reader with a loose analogy that may not provide useful evidence.

In addition, Poweleit et al. propose a "lipid mediated" mechanism of protein translocation through this hydrophobic pore. It is difficult to envision though how lipids would be able to mediate translocation of folded substrate dimers with mostly hydrophilic surfaces. This would be a completely new mechanism of protein translocation not seen for any other protein secretion machineries. In addition, the authors speculate even further that besides (hydrophilic) proteins also hydrophobic molecules, such as lipids, could be transported through this cavity, while there is no clear evidence that type VII secretion systems are able mediate lipid transport. In conclusion, while it is indeed striking that such a large cavity is present in the ESX-3 complex, the authors should more cautiously discuss this model, by also mentioning the major problems with this model as discussed above. The translocation model of substrates through the central pore of a hexameric complex is far less speculative, as it is supported by experimental data, published by several groups (as also mentioned in the Discussion). This model should therefore get more emphasis.

We have restructured and rewritten the Discussion to further emphasize the relationship between the prior literature and the oligomer model.

3) The regulatory role of the "cytoplasmic bridge" is speculative and should be more carefully discussed. e.g. "multimeric cytoplasmic domain is poised to regulate the translocation of ESX substrates" and "EccC₃ and EccE₃ collaborate to buttress the cytoplasmic domains of EccD₃ and form a stable regulatory bridge".

Our intension with this language was to point out the interactions between the cytoplasmic domain of EccD₃ and the other two proteins with significant cytoplasmic domains, EccE₃ and EccC₃. EccD₃ contains a highly conserved region (116-125) in the linker connecting the ubiquitin-like domain at the N-terminus to the transmembrane region. Despite the strong conservation, the interface made by this region each copy of EccD₃ in the homodimer is strikingly different. In one case, the region interacts extensively with the other cytoplasmic domain of EccD₃ as well as with EccE, anchoring EccE in a rigid conformation. In contrast, this region has a completely different conformation in the symmetry related copy of EccD, where it reaches in a different direction to form a nexus of interactions with EccB, and EccC. Given the need for the rearrangement of EccC to accommodate the R-finger mechanism of the ATPase 1 domain, we speculate that flexibility of the EccD₃ linker could allow for the release of EccE and EccC from their rigid positions, thus allowing for a rearrangement of the ATPase domains into an active conformation. To respond to the comments. we have clarified this idea of a “regulatory bridge” and separated the observations of the structure from the speculation about the functional relevance, which is now in the Discussion. We have updated and expanded Figures 3C-D, Figure 4C-D and Figure 5C to highlight these interactions.

Author details

1. Nicole Poweleit

   1. Department of Medicine, Division of Infectious Diseases, University of California, San Francisco, San Francisco, United States
   2. Chan-Zuckerberg Biohub, University of California, San Francisco, San Francisco, United States

   Contribution

   Investigation, Formal analysis, Validation, Visualization, Methodology, Writing – Original Draft Preparation, Writing – Review & Editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2700-0241

2. Nadine Czudnochowski

   1. Department of Medicine, Division of Infectious Diseases, University of California, San Francisco, San Francisco, United States
   2. Chan-Zuckerberg Biohub, University of California, San Francisco, San Francisco, United States

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3146-4110

3. Rachel Nakagawa

   Department of Medicine, Division of Infectious Diseases, University of California, San Francisco, San Francisco, United States

   Contribution

   Resources, Investigation, Methodology, Writing – Review & Editing

   Competing interests

   No competing interests declared

4. Donovan D Trinidad

   1. Department of Medicine, Division of Infectious Diseases, University of California, San Francisco, San Francisco, United States
   2. Chan-Zuckerberg Biohub, University of California, San Francisco, San Francisco, United States

   Contribution

   Investigation, Methodology, Validation

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1439-9927

5. Kenan C Murphy

   Department of Microbiology and Physiological Systems, University of Massachusetts Medical School, Worcester, United States

   Contribution

   Resources, Investigation, Methodology

   Competing interests

   No competing interests declared

6. Christopher M Sassetti

   Department of Microbiology and Physiological Systems, University of Massachusetts Medical School, Worcester, United States

   Contribution

   Resources, Validation, Methodology, Supervision, Writing – Review & Editing

   Competing interests

   No competing interests declared

7. Oren S Rosenberg

   1. Department of Medicine, Division of Infectious Diseases, University of California, San Francisco, San Francisco, United States
   2. Chan-Zuckerberg Biohub, University of California, San Francisco, San Francisco, United States

   Contribution

   Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Validation, Visualization, Methodology, Project administration

   For correspondence

   oren.rosenberg@ucsf.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-5736-4388

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