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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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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