Biophysical characterization of Eag chaperones suggests the mechanism of effector transmembrane domain release

Domain: neuroscience
Published: 2026-01-22
Editorial outcome: major_revision

Abstract
The type VI secretion system (T6SS) is a dynamic protein nanomachine found in Gram-negative bacteria that secretes toxic effectors into prey-cells. For secretion, effectors require chaperones or adaptors for proper loading onto the T6SS. Effector associated genes (Eags) are a family of T6SS chaperones that stabilize N-terminal transmembrane domains (TMDs) found in thousands of effectors. Eags are essential for secretion and inhibit effector TMDs from prematurely adopting a membrane-penetrative conformation. However, the mechanism of TMD release from its cognate Eag chaperone is unknown. Here, we take a biochemical and biophysical approach to probe the mechanism of TMD binding and dissociation from Eag chaperones. Using steady-state fluorescence, stopped-flow measurements, and bacterial competition assays, we compare the thermodynamics, kinetics, and in vivo chaperone function of wild-type and point variant Eag-TMD complexes. Additionally, we solve an X-ray crystal structure of an Eag-TMD point variant complex that captures an intermediate state of TMD release. Our data reveals the molecular features and specific residue contacts necessary for TMD binding and demonstrates the Eag conformational change required to initiate rapid release of the TMD. Overall, our work details the stability of Eag-TMD complexes and the energetic pathway for the dissociation of effector TMDs from their Eag chaperones.

Introduction
The ability to secrete proteins into the environment is a central part of bacterial communication and virulence1. Secreted proteins often mediate bacterial antagonistic behaviors that shape microbial communities to form complex cooperative and/or competitive networks2,3. As such, bacteria have evolved diverse general and specialized secretion mechanisms. These include large membrane-spanning complexes designed specifically for the translocation of proteins into the extracellular environment or even directly into rival cells4–6. In Gram-negative bacteria, one such complex is the type VI secretion system (T6SS). Generally, the T6SS provides bacteria a competitive advantage in an ecological niche7.

The T6SS is a contractile nanomachine that forcibly injects toxic protein effectors into nearby cells in a contact-dependent manner2,8,9. The T6SS is structurally related to the T4 bacteriophage tail10,11 and consists of a needle, a baseplate, and a membrane-spanning complex. The needle is made of hexameric Hemolysin co-regulated proteins (Hcp) surrounded by a contractile protein sheath that is polymerized starting from the baseplate10–12. The membrane-spanning complex orients the needle for ejection into the environment13,14. Within the baseplate, a trimeric valine glycine repeat protein G (VgrG) binds the top of the Hcp needle12,15. VgrG forms the tip of the T6SS and serves as a modular platform for effector loading16,17. VgrG can also be “sharpened” by a proline-alanine-alanine-arginine (PAAR) domain-containing protein16,18 or a PIPY domain protein, which both bind VgrG to create a structural spike19–21. Both PAAR and PIPY domains can serve to load VgrG with cargo effectors22–26 and PAAR domains can be covalently linked to an effector to create a specialized or evolved effector9,27,28. The T6SS then secretes the entire Hcp-VgrG-PAAR-effector complex. Finally, the effectors inhibit a wide range of essential processes that kill or inhibit the growth of neighbouring prokaryotic and/or eukaryotic cells6,29.

A critical step in the secretion of many effectors is the selective recruitment or piloting of effectors to the T6SS by accessory proteins that act as chaperones or adaptors. Chaperones stabilize effectors in a “secretion-competent” state and affect protein folding, whereas adaptors typically only aid in the binding of effectors to T6SS machinery. The terms chaperones and adaptors are often used interchangeably in T6SS literature, especially as some chaperones also aid in effector loading27,30. Despite the ambiguity, chaperones and adaptors are essential for the proper delivery and toxicity of numerous T6SS effectors17,21,27,31. Several T6SS chaperone and adaptor families have been described, including DUF2169, Type VI adaptor proteins (Tap) (DUF4123), and Effector associated genes (Eag) (DUF1795). DUF2169 forms complexes with an effector and a PIPY protein to facilitate loading onto VgrG19,22,32, Tap adaptors pilot effectors to VgrGs that contain a helix-turn-helix at their C-terminus17, and Eag chaperones facilitate the binding of prePAAR containing membrane protein effectors to VgrG27,33,34.

The basic architecture of a prePAAR containing effector is an N-terminal prePAAR motif followed by a transmembrane domain (TMD), a PAAR domain, and a C-terminal toxin domain27. prePAAR-containing effectors can be further divided into two classes. Class 1 effectors belong to the Rhs family of proteins and possess a single TMD, while Class 2 effectors possess two TMD regions directly before and after the PAAR domain27. The toxin domains of prePAAR effectors are known to localize to the cytoplasm and inhibit competitor cell growth by acting in the cytoplasm of prey cells, such as by ADP-ribosylation of RNA or protein35–37 or rapid depletion of the electron carriers NAD+ and NADP+33,38. The TMDs are thought to facilitate the translocation of the effector across the prey cell cytoplasmic membrane31. Prior to secretion, the TMDs are shielded from the aqueous milieu of the producing cell cytoplasm by their associated Eag chaperone. To promote secretion, Eag chaperones bind the effector TMDs and drastically distort their conformation to inhibit TMD folding27. This creates a pre-insertion state to both pilot the prePAAR effector to the T6SS and prevent erroneous membrane insertion before secretion. Additionally, the Eag allows the prePAAR motif to complete a zinc binding motif necessary for the effector PAAR domain to recognize VgrG27,39. Notably, Eags are specific for their cognate TMDs and exhibit different effector binding modes.

Given that a bound Eag chaperone would inhibit the proposed membrane penetrating function of the effector TMD, the Eag is presumably removed prior to secretion by the T6SS. Consistent with this hypothesis, Eag chaperones are not secreted alongside their associated effectors22,23,31,40. However, there is no current data showing how the Eags are separated from their cognate TMDs. To address this mechanistic question, we took a biophysical and biochemical approach to determine how a T6SS effector TMD is released by an Eag chaperone.

Here, we measure the thermodynamic properties and unfolding kinetics of wild-type Eag-TMD complexes compared to several site-specific point variant Eag-TMD complexes. Our data yields the binding energies and dissociation kinetics of Eag-TMD complexes, which shows that although Eag-TMDs are remarkably stable only a small perturbation of the Eag is required to promote TMD dissociation. Specifically, TMD binding to an Eag depends upon a crucial knob-in-hole packing interaction (large against small side chain packing) and/or a hydrogen bond that, if removed, results in drastically reduced complex stability and rapid effector release. Furthermore, we show that the disruption of these interactions between an Eag and its cognate TMD reduces effector secretion and bacterial fitness, directly correlating our mechanistic findings to biological function. Finally, an Eag-TMD variant X-ray co-crystal complex captures a TMD release intermediate providing key mechanistic details for TMD binding and dissociation from a chaperone. Our work emphasizes the essential role of Eags in TMD folding inhibition and collectively enables us to propose the mechanism for TMD release from Eag chaperones.

Results
Effector TMD binding drastically increases Eag chaperone stability
Effector TMDs are highly insoluble and rapidly degrade unless they are bound and stabilized by an Eag chaperone31,38. Given this, it is not possible to express and purify the TMDs without also co-expressing the Eag chaperone. The inability to separately purify Eag and TMD proteins precludes the ability to determine the mechanistic details of their interaction using common binding assays41,42. To circumvent this problem, we can study the mechanism of effector TMD binding and dissociation by assaying the stability and thermodynamic properties of Eag-TMD complexes compared to unbound Eags. For this study, we use the Eag-TMD complexes SciW-Rhs(1–59) from Salmonella Typhimurium and EagT6-Tse6(1–61) from Pseudomonas aeruginosa27. For simplicity, the complexes will be referred to as SciW-TMD, EagT6-TMD, or Eag-TMD generally. Notably, the Eag chaperones contain tryptophan residues, whereas the TMDs lack tryptophan. As such, by monitoring the change in tryptophan fluorescence during denaturation, we can determine how TMD binding affects the energetics and folding behavior of the Eag chaperones to extract TMD dissociation data.

Using this approach, we first determined the thermostability of the Eag-TMD complexes compared to unbound or apo-Eags. This was done to gauge the stability of the Eag upon TMD binding. When an Eag binds its cognate TMD, there is a large increase in the thermostability of the chaperone. Nano differential scanning fluorimetry (nanoDSF) reveals an increase in melting temperature (Tm) of 30°C for SciW and 12.2°C for EagT6 (Fig. 1A, B and Supplementary Table 1). This demonstrates that upon TMD binding, the chaperones become as stable as proteins from extremophiles (>65°C)43. The purified protein materials used in these experiments are shown in Supplementary Fig. 1.

Because the TMDs are invisible in nanoDSF due to their lack of tryptophan residues, we also compared our results to thermal melting by differential scanning calorimetry (DSC). This was done to observe if the TMDs have melting transitions distinct from their Eag chaperones. The DSC experiments gave nearly identical results to nanoDSF (Fig. 1C–F). Furthermore, there were no additional melting transitions in the DSC relative to the nanoDSF for the EagT6-TMD complex indicating that the TMD melts cooperatively with the EagT6 chaperone (Fig. 1D). In contrast, the SciW-TMD complex melts in two transitions (Fig. 1E). Two transitions observed by DSC but only one by nanoDSF suggests that the TMD changes conformation while still bound to SciW before the complex melts (first transition, Fig. 1E). However, in agreement with the nanoDSF data the observed changes in the enthalpy of unfolding (ΔH) for both pairs of Eag chaperones indicates that substantial energy is required to dissociate an Eag-TMD complex.

Effector TMD binding significantly increases the free energy of Eag chaperone unfolding
We next probed Eag chaperone resistance to chemical denaturation. By monitoring Eag and Eag-TMD unfolding by denaturants, we will be able determine the free-energy (ΔG) of Eag stabilization by the TMD and the binding energy of the Eag-TMD interaction. Specifically, the ΔG of unfolding measured at various denaturant concentrations is used for extrapolation to 0M denaturant. This yields native denaturant-independent ΔGs of unfolding and TMD binding energy in a physiologically relevant buffer44. Therefore, the data reveals the binding energy that must be overcome during secretion to remove the Eag. We denatured Eag and Eag-TMDs with both urea and guanidine hydrochloride (GdmCl), then monitored steady-state Eag unfolding by tryptophan fluorescence. Shown in Fig. 2A, B and Supplementary Table 1, the Eag-TMD complexes are highly resistant to denaturation by urea. Notably, SciW-TMD could not be unfolded with urea (Fig. 2A). In contrast, the Eag-TMD complexes could be readily unfolded with GdmCl (Fig. 2C, D). The C50 (concentration of denaturant at midpoint) values for Fig. 2A–D are summarized in Supplementary Table 1. In agreement with the thermal denaturation data, the chemical denaturation experiments demonstrate significantly increased Eag stability upon TMD binding.

Now that we have an appropriate denaturant (GdmCl), we can determine the free energy of Eag-TMD unfolding. First, we measured the intrinsic fluorescence spectra of the Eags and the Eag-TMDs as a function of increasing GdmCl concentrations (Supplementary Fig. 2). This was done to experimentally find the wavelength maxima (λmax) for the unfolding transitions. Measuring the change in λmax is a more accurate way to monitor protein unfolding relative to our initial experiments using the Prometheus. The Prometheus compares the intensities measured at the fixed wavelengths of 330 nm and 350 nm, which allows for rapid screening45,46. Using the data from Supplementary Fig. 2, the fluorescence emission maximum wavelength λmax as a function of denaturant concentration was determined (Supplementary Fig. 3). If we assume that the Eag chaperones unfold through a two-state mechanism (Eq. (1)), the fraction of tryptophan residues in the unfolded state “α” can be related to the measured λmax47 to plot the transition (Eqs. (2) and (3)). All equation derivations are detailed in “Methods”.

Using Eq. (2), we have plotted α (fraction unfolded) as a function of denaturant concentration for SciW and EagT6 in their apo and TMD-bound forms (Fig. 2E, F and Supplementary Fig. 3). Notably, the experimentally determined λmax steady-state fluorescence data is in close agreement with the fixed 330/350 nm wavelength data (Fig. 2C, D). However, given the slight variation all quantitative thermodynamic and kinetic data will be derived from the experimental λmax values.

Next, the data in Fig. 2E, F was used to calculate the change in unfolding free energy (ΔGunfolding) at various denaturant conditions and determine the native ΔG°unfolding at 0M denaturant (Eqs. (4)–(6), “Methods”)44. From fitting the data in Fig. 2E, F to Eq. (5), the values of ΔG°unfolding and m (slope of the linear extrapolation) for the Eag and Eag-TMD complexes were obtained (Table 1). Figure 2G, H shows the plot of ΔG as a function of denaturant and the fitted linear extrapolation to 0M GdmCl44. Both Eags have a similar native ΔG°unfolding of ~103 kJ/mole and each gains significant stability as an Eag-TMD complex. When bound to their cognate TMD, the ΔG°unfolding for SciW increases by ~71 kJ/mole and the ΔG°unfolding for EagT6 increases by ~39 kJ/mol. For context, the ΔG°unfolding for lysozyme in a native buffer is ~30 kJ/mol48.

The effector TMD binding energy (ΔG°binding) and association constant (Kbinding) can then be obtained from comparing Eag ΔG°unfolding to Eag-TMD ΔG°unfolding. This calculation requires the following assumptions: (1) the TMD is a small ligand compared to the Eag, (2) TMD binding does not significantly alter the Eag fold, (3) all free TMD in solution results from complex dissociation, (4) the chaperone folds by a 2-state model, and (5) the TMD is assumed to selectively bind the folded form of the Eag with an association constant KTMD. Given these, the effect of TMD binding upon the standard unfolding free energy for the chaperone can be estimated using Eq. (7) (“Methods”)49,50. ΔG°binding is the standard free energy change of TMD binding the chaperone. This directly relates the change in Eag unfolding standard free energy to that of TMD binding at native conditions.

The unfolding data allows us to estimate a ΔG°binding of approximately −176 kJ/mol for the SciW-TMD complex and a ΔG°binding of −116 kJ/mol for the EagT6-TMD complex (Table 1). Antibodies with pico to femto-Molar (fM, 10−15) dissociation constants have a ΔG°binding near −70 kJ/mol51. The calculated binding constants for the SciW-TMD and EagT6-TMD interactions are on the order of ~7×10−32 M and ~5×10−21 M, respectively. Our thermodynamic data (Figs. 1 and 2) clearly shows that the Eag chaperones bind their TMDs extremely tightly and gains significant stability when in complex with a TMD. This in turn raises the question of the biochemical mechanism needed to overcome the extreme binding energy for TMD release.

Eag chaperone α-helical packing residue mimics act to inhibit TMD folding
Using the thermodynamic properties of the Eag-TMD complexes combined with their known molecular interactions, we can probe the mechanism of TMD dissociation. Co-crystal structures of SciW-TMD and EagT6-TMD show that Eags bind their cognate TMDs by mimicking alpha helical membrane packing27. This is accomplished by the Eag dimer folding like a claw around the TMD to create a hydrophobic environment. Additionally, the Eags harbor conserved residues within the pocket of the claw that make hydrophilic bifurcated hydrogen bonds (H-bonds) and knob-in-hole packing interactions with the TMD (Fig. 3 and Supplementary Fig. 4). The net effect is to inhibit the TMD from making H-bond and knob-in-hole packing interactions with itself to properly fold and thus remain bound to the Eag.

To determine which Eag molecular interactions are important for TMD binding and complex stability, we performed site-directed mutagenesis on several conserved residues found in both SciW and EagT6 (Fig. 3). The positions of each residue and their specific interactions with the TMD are shown in Fig. 3A, B. Note that due to the domain-swapped architecture of the dimeric Eag chaperones, these contacts are not equivalent in each Eag monomer. The interaction surfaces of each point variant in the opposing chain are also depicted in Supplementary Fig. 4C.

Eag residue I25 (SciW)/I24 (EagT6) was changed to phenylalanine to interfere with the TMD properly packing against the base of the hydrophobic Eag claw (Fig. 3B). The Eag hydrophilic residues S41, Q58, and Q106 (SciW)/Q102 (EagT6) were substituted to alanine because the residues make bifurcated hydrogen bonds to the TMD peptide backbone (Fig. 3B). Side-chain to main-chain bifurcated H-bonds are crucial for transmembrane helices to properly adopt their tertiary structure52–54. The conserved Eag residue L66 was substituted to L66A because it is a mimic of “knob-in-hole” packing interactions. Knob-in-hole packing is when a large hydrophobic residue on one TM-helix acts as a knob to fill the hole created by a small residue, such as glycine or alanine on another TM-helix with a GxxxG/A motif55. As shown in Fig. 3B Eag L66 acts as a knob to fill a hole with TMD alanine residues. A shorter side-chain (leucine to alanine) removes the knob. Eag L66 was also of interest because it is on the outer part of the Eag claw. Specifically, the loop containing L66 undergoes a conformational change that moves the Eag claw structure over the TMD upon binding27.

After residue substitution, each apo-Eag variant and Eag-TMD variant was purified and its fold assayed by dynamic light scattering (DLS). As shown in Supplementary Fig. 5 and Supplementary Table 2, all point variants purified similar to wild-type and exhibited the same DLS profile. This demonstrates that like the wild-type Eags each Eag variant is dimeric in solution and not aggregated due to misfolding. Furthermore, all Eag-TMD variants still co-purify bound to their cognate TMD, showing they are all still functional. Given that we know the ΔG°unfolding and ΔG°binding for wild-type Eag-TMDs (Fig. 2), we can now determine how much binding energy is contributed by each Eag residue contact with the TMD and determine mechanistic details of TMD binding and dissociation.

Eag α-helical packing residue mimics contributes significantly to TMD binding
To probe the effect of Eag variants on TMD binding, we screened their resistance to both thermal (Supplementary Fig. 6) and chemical denaturation (Supplementary Fig. 7 and Supplementary Table 3). Both thermal and chemical denaturation show a similar pattern of lower complex stability for the selected point variants. However, the chemical denaturation data was used to assay stability for ΔG measurements. For SciW-TMD, complex stability was reduced by the residue substitutions but was still significantly more stable than the apo-SciW variants (Supplementary Fig. 7A, C, E, G, I and Supplementary Table 3). When comparing the change in C50 between the apo-EagT6 and TMD bound variants, we observe that all substitutions greatly weaken the complex stability (Supplementary Fig. 7B, D, F, H, J and Supplementary Table 3). However, EagT6 I24F, L66A, and Q102A appear to have the greatest effect (small or no change in bound to unbound variant C50). This shows that EagT6-TMD binding is severely weakened by either the loss of knob-in-hole (L66A) or bifurcated H-bond interactions (Q58A) (Fig. 3 and Supplementary Fig. 4). Additionally, steric hindrance of proper TMD packing by a larger hydrophobic residue (I24F) is also severely disruptive. For the SciW-TMD complex, the loss of bifurcated H-bonds (Q58A) or knob-in-holes (L66A) is substantial but less pronounced (Supplementary Fig. 7 and Supplementary Table 3). Additionally, Q106/2 was important for EagT6-TMD binding but had almost no effect on SciW-TMD binding.

The variants Q58A and L66A represent two distinct modes of α-helix membrane protein packing and affect both SciW-TMD and EagT6-TMD complex stability. Given this, we selected these variants for ΔGunfolding quantification by intrinsic protein fluorescence spectral analysis. The tryptophan fluorescence wavelength maxima (λmax) for the Q58A and L66A point variants were determined and used to plot α (fraction unfolded) as a function of denaturant concentration using Eq. (6) (Supplementary Figs. 3 and 8, Fig. 4A–D, and “Methods”). The ΔGunfolding for the point variants was calculated from fitting α (fraction unfolded) with Eqs. (1)–(6) and ΔG°binding from Eq. (7), similar to the wild-type. Because the “m” parameter in Eq. (6) only depends upon the change in surface accessible area upon unfolding56, the α (fraction unfolded) values were fitted globally, sharing the “m” parameter in these subsets: (1) apo-SciW point-variants, (2) SciW-TMD point-variants, (3) apo-EagT6 point-variants, (4) EagT6-TMD point-variants. The resulting parameters from the fits are listed in Table 1. The dependence of ΔGunfolding on denaturant concentration with fitted extrapolations to native buffer conditions at 0M denaturant (ΔG°unfolding) are shown in Fig. 4E, F.

Compared with the wild-type ΔG°unfolding, the point variants Q58A and L66A have little effect on the ΔG°unfolding for apo-SciW. However, both residue substitutions significantly lowered the ΔG°unfolding of the SciW-TMD complex, indicating a reduction in TMD binding affinity (Table 1 and Fig. 4A, C, E). The ΔG°binding values for SciW Q58A and L66A with the TMD are −142 kJ/mol and −154 kJ/mol, respectively, compared to −176 kJ/mol for wild-type. We can conclude that the bifurcated H-bonds provided by Q58 contribute ~34 kJ/mol to TMD binding, and the knob-in-hole interactions of L66 contribute ~22 kJ/mol. Within experimental error, these are roughly 15–20% of the total SciW-TMD binding energy. This also increases the dissociation constant for Q58A and L66A relative to wild-type by several orders of magnitude to ~1×10−25 M and ~7×10−28 M, respectively.

For EagT6, the point variants Q58A and L66A have a more pronounced effect on complex stability and TMD binding. The folding free energy of apo-EagT6 is minimally affected by the Q58A substitution, but the stability of EagT6 is reduced by L66A (Table 1 and Fig. 4B, D, F). However, the EagT6 L66A variant chaperone is still properly folded and functional as it co-purifies bound to the TMD (Supplementary Fig. 5). Moreover, we observe drastic reductions in the ΔG°unfolding for the EagT6-TMD variant complexes showing that Q58 and L66 play critical roles in TMD binding. For the Q58A, the ΔG°binding to the TMD has reduced in magnitude from −116 kJ/mol to −60 kJ/mol. Thus, we can conclude that the bifurcated H-bonds and packing interactions of Q58 with the TMD contribute at least 56 kJ/mol or about 1/2 the binding energy. This increases the dissociation constant by several orders of magnitude to ~3×10−11 M. For EagT6 L66A, the value of ΔG°unfolding is similar for the variant apo and TMD bound. This indicates that minimal TMD binding to the EagT6 L66A variant exists under our assay conditions. Our fluorescence experiments were performed under dilute conditions (1 μM chaperone) and in the absence of crowding agents, making it likely that the TMD dissociated prior to the assay. This shows that the binding constant KTMD for the L66A substitution has been reduced to less than 106 M−1 putting an upper limit of ~−35 kJ/mol upon the ΔG°binding. Our data show that the knob-in-hole packing of residue L66 is crucial for maintaining the EagT6-TMD complex.

Minor variations to the Eag molecular surface accelerate the rate of TMD release
Our biophysical data thus far has been measured at equilibrium conditions. However, T6SS assembly and firing is a dynamic process that can occur within several seconds57–59. Given this, how quickly the TMD can be released prior to secretion is of central importance to the function of the Eag chaperone. To determine the rate of TMD release, we investigated the unfolding kinetics of wild-type and variant Eag chaperones using stopped-flow spectrofluorimetry. Unfolding fluorescence trace data as a function of time for wild-type Eag and Eag-TMD complexes is shown in Supplementary Fig. 9 and point variant traces in Supplementary Fig. 10. The relaxation profiles can be described with a mono-exponential decay and the native unfolding rate constants (k°unfolding) calculated using chevron plot analysis by extrapolation to 0M GdmCl44 (Fig. 5 and Table 1, and Eqs. (8) and (9), “Methods”).

Wild-type SciW and the Q58A and L66A variants have unfolding rate constants (k°unfolding) that are indistinguishable within the limits of experimental uncertainty (Fig. 5A and Table 1). The k°unfolding constants reveal that the unfolding/folding lifetime for SciW is on the order of ~5 min. Again, this highlight that the point variants have no effect on the biophysical properties of the SciW chaperone itself. However, both Q58A and L66A significantly increase the rate of unfolding for the SciW-TMD complex (Fig. 5A). The wild-type SciW-TMD complex has a lifetime on the order of ~15 h but is reduced to ~2 h for SciW L66A and ~16 min for SciW Q58A. The disruption of a knob-in-hole (L66) or bifurcated H-bond (Q58) increases the rate of TMD dissociation by 7.5× and 60×, respectively, but only at the cost of 15–20% binding energy (Fig. 4F and Table 1).

The wild-type EagT6 chaperone has a folded state lifetime of ~5 s, which increases to ~18 s when bound to TMD1 of Tse6 (Fig. 5B and Table 1). The extreme difference in the folded lifetime of apo EagT6 compared to apo SciW was unexpected, as both chaperones have the same ΔG°unfolding (Table 1) and domain-swapped dimeric structure (Fig. 3)27,60. Additionally, we find that both Q58A and L66A increase the rate of unfolding for the apo EagT6 chaperone to ~1 s. Interestingly, this differs from the steady-state data where EagT6 Q58A is indistinguishable from wild-type. However, similar to SciW, both EagT6 Q58A and EagT6 L66A greatly increase the rate of EagT6-TMD complex unfolding. EagT6 Q58A reduces the lifetime of the EagT6-TMD complex to ~5 s. This is a ~4 s difference in lifetime for the Q58A variant as compared to ~13 s for wild-type, demonstrating that the increased TMD dissociation rate is not primarily due to the faster unfolding of the apo-variant. For EagT6 L66A we observe no kinetic difference with the TMD (Fig. 5B and Table 1). Again, this is likely because TMD release by EagT6 L66A is faster than we can measure. In agreement with the steady-state data, the knob-in-hole interactions provided by EagT6 L66 are essential to keep the TMD bound to the Eag.

The transition state intermediate of TMD release from Eags differs by effector class
The effect of Eag variants upon the unfolding rate (k°unfolding) and energy (ΔG°unfolding) also reveals mechanistic details about the TMD dissociation transition state. By applying a “φ-value analysis” to the measured Eag thermodynamic and kinetic parameters we gain insight into the rate-limiting step of the unfolding transition state. The φ-value is the ratio in change of the transition state free energy for unfolding (ΔG‡unfolding = −RT(lnk°unfolding)) to the equilibrium free energy of unfolding (ΔG°unfolding) between wild-type and variants (Eq. (10)). This gives a φ-value between 0 and 1, where 0 represents a dynamic or unstructured transition state and 1 an ordered or structured transition state61.

The φ-value analysis of the Eag-TMD complexes yields 0.7±0.4 for SciW Q58A, 0.5±0.2 for SciW L66A, and 0.1±0.3 for EagT6 Q58A. A φ-value calculation for EagT6 L66A was not possible because the TMD had already dissociated. The small φ-value for EagT6 Q58A indicates a dynamic or disordered conformation at the unfolding transition state of the EagT6-TMD complex. This could be due to that the TMD is conformationally dynamic at the transition state and/or that the TMD has already been released. Moreover, release of the TMD from EagT6 and the unfolding of EagT6 are uncoupled processes. Given the inability to determine a φ-value for EagT6 L66A a similar disordered transition state is likely. For SciW Q58A and L66A, the φ-values are both 0.5 or higher, which is more characteristic of a structured unfolding transition state. Additionally, the φ-values suggest that the TMD is bound to SciW during the unfolding transition state. This indicates that the release of the TMD and unfolding of SciW are coupled. Overall, our biophysical characterization shows that the exact mechanistic details and transition state of TMD release differ between SciW (class 1 effector TMDs) and EagT6 (class 2 effector TMDs). First, that disruption of the H-bonds provided by Q58 or Q102, or a knob-in-hole provided by L66 is sufficient to dissociate the TMD from EagT6, but TMD release from SciW requires disruption of more than one contact. Second, that the dissociation of the TMD from SciW involves a stable conformation intermediate of either the TMD, SciW, or both.

EagT6-TMD stability variants inhibit effector secretion and decrease bacterial fitness
To observe how our biophysical characterization relates to biological activity, we next determined the contribution of our Eag point-variants to the function of the chaperones in vivo using growth competition assays. EagT6-Tse6 was chosen for our model because our point variants have a significantly greater effect on Tse6 TMD release from EagT6 as compared to SciW and the TMD from Rhs1 (Figs. 4 and 5). For example, in EagT6 L66A is sufficient to rapidly release the TMD in vitro. Moreover, EagT6 is known to be essential to T6SS-dependent bacterial competition in P. aeruginosa31,38. Additionally, the T6SS of P. aeruginosa is a highly tractable model system since deletion of the negative T6SS regulator retS results in strains with constitutive T6SS activity17,31,33. In contrast, the S. Typhimurium T6SS requires induction by addition of bile-salts to the media62. We were able to show T6SS-dependent killing of Escherichia coli by Salmonella; however, a ΔsciW deletion strain only had an intermediate loss of function phenotype (Supplementary Fig. 11). This is in agreement with past results that showed ΔsciW had little effect on S. Typhimurium fitness in a mouse model63. Given the moderate phenotype of the ΔsciW deletion strain and that the SciW point variants do not abrogate binding like EagT6, assays in Salmonella were not pursued.

Competition assays were performed with a P. aeruginosa donor and an isogenic Tse6-susceptible recipient that lacks Tse6 and the immunity protein Tsi6 (Δtse6Δtsi6)27,31. The donor strains encode the wild-type EagT6 chaperone (parent), no chaperone (ΔeagT6), or one of the EagT6 point variants expressed from their native chromosomal locus. When co-cultured on a solid surface, the wild-type donor readily outcompetes the recipient strain (Fig. 6A). Consistent with our biophysical data, any of the eagT6 alleles encoding variants of the chaperone display attenuated fitness against the Tse6-susceptible recipient. Specifically, EagT6 L66 strongly contributes to the function of EagT6 in vivo and therefore the proper secretion of Tse6. Furthermore, the donor strain encoding eagT6 (Q58A) displays intermediate fitness against the recipient strain. This is consistent with our biophysical finding that Q58 contributes less significantly than L66 to the stability of the EagT6-TMD complex. The agreement between these genetic results and our biophysical data suggest that the determinants of complex stability identified in vitro closely reflect the function of the EagT6-TMD complex in P. aeruginosa.

Since Tse6 requires EagT6 binding for cytosolic accumulation in P. aeruginosa31, we reasoned that Tse6 levels may be reduced in strains harboring our mutant eagT6 alleles. Indeed, expression of the EagT6 variants decreased intracellular Tse6 levels (Fig. 6B). Furthermore, the relative abundance of Tse6 in the EagT6 Q58A and L66A variant backgrounds correlates with the contribution of those residues to competitive fitness and in vitro TMD binding energy (Figs. 4–6 and Table 1). Notably, the relative abundance of Tse6 is high for EagT6 Q58A as compared to other the mutants and wild-type (Fig. 6B). Supporting this, EagT6 Q58A has a ΔG°binding of −60 kJ/mol, which is still indicative of a stable complex. Regardless, in agreement with our biophysical data the loss of TMD binding affinity by Q58A significantly decreases Tse6 secretion (Fig. 6A). Overall, these findings demonstrate that the knob-in-hole packing provided by EagT6 L66 with the TMD of Tse6 is critical for the stability of the EagT6-Tse6 complex in vivo and proper secretion of the effector.

A crystal structure of SciW-TMD L66A suggests a TMD release intermediate
To further probe the molecular details of Eag-TMD complex stability, we attempted to crystallize a point variant Eag-TMD complex. We were able to obtain diffracting crystals of SciW-TMD L66A and obtain phases by molecular replacement (Fig. 7). Two SciW-TMD complexes were present in the asymmetric unit and all chains exhibited very high B-factors suggesting conformational dynamics (Supplementary Fig. 12A and Table 2). Specifically, one SciW-TMD L66A complex (chains D–F) showed several areas of missing density and the TMD was not confidently modelled. However, the other SciW-TMD L66A complex (chains A–C) showed strong density for the TMD (Supplementary Fig. 12B). Strikingly, the conformation of the bound TMD is significantly different from the wild-type complex (Figs. 7 and 3A). Given the high B-factors, lack of strong TMD density for one complex, and the observed alternate TMD conformation, the captured SciW-TMD L66A structure likely represents a TMD release intermediate.

SciW-TMD L66A retains a dimeric structure similar to wild-type; however, the loop containing L66A at the tip of each claw is in a different conformation (Figs. 7A and 3). There is no observable electron density for the loop between SciW residues K64 and G67 of chain B (Fig. 7A, right). Furthermore, the TMD is also no longer in a twisted asymmetric conformation. In wild-type SciW, the TMD (blue) adopts an asymmetric binding mode, making non-equivalent contacts with the chaperone dimer (Fig. 3, Supplementary Fig. 4, and Fig. 7B)27. Instead, the TMD folds as two symmetric helices packed similarly into the bottom of the L66A chaperone claw (Fig. 7A, right).

When comparing the SciW-TMD L66A structure to the wild-type complex, we observe that the L66A variant is unable to form critical knob-in-hole and hydrophobic packing interactions with the TMD (Fig. 7B). First, L66A in chain B no longer forms a knob-in-hole with Rhs1-TMD A24 in helix α1 or packs against residue I56 in helix α3 due to the shorter side chain of the substituted alanine. This in turn frees SciW loop K64 to G67 and releases the C-terminus of the TMD from the chaperone, which causes the TMD to open outwards from its normal twisted conformation (Fig. 7B, chain B). Simultaneously in chain A, L66A no longer packs against TMD residues L44 or L47 in helix α3. The loop containing A66 moves ~9 Å away, also releasing the TMD to open in a hinge-like fashion, forming an α-helix that fills the chaperone binding pocket. This is in part due to TMD residue L47 moving into the space that the SciW L66 side chain previously occupied. Additionally, the bound N-terminal prePAAR region (residues 8–13) of the TMD is pushed out and released from SciW. We note that the prePAAR must be free to bind the PAAR for secretion27. A movie demonstrating the conformational change is provided (Supplementary Movie 1).

The captured SciW-TMD L66A dissociation intermediate likely represents a biologically relevant conformation, particularly when compared to wild-type SciW and EagT6 complexes. Figure 7C shows the conformational changes involving residue L66 that are required for effector TMD binding. In both Eags, the position of L66 is shifted significantly inward to interact with the TMD. However, only in EagT6 does L66 in both chains appear to move. Given the importance of L66 for TMD binding and dissociation, the repositioning of L66 upon TMD binding may reflect a reversible conformational switch. Here, inward movement of L66 facilities TMD binding and outward movement of L66 disrupts the knob-in-hole packing interaction to promote TMD dissociation.

Our structural data also shows why L66 is important for SciW-TMD complex stability and in agreement with our φ-value analysis, suggests a partially ordered transition state during TMD dissociation. Given that residue L66 is critical for EagT6-TMD complex stability and that L66A allows the visualization of SciW-TMD unfolding intermediate, a conformational change to move L66 away from the TMD is likely central to the mechanism of TMD release for both classes of Eag chaperones.

Methods
Molecular cloning of protein expression constructs
Wild-type Eag chaperones SciW from Salmonella Typhimurium (Eag: SL1344_0285) and EagT6 from Pseudomonas aeruginosa (Eag: PA0094) and their respective point variants were commercially synthesized from Twist Bioscience in the vector pET-29b(+). The Eag point variant SciW Q106 was commercially synthesized in multiple cloning site 2 of pETDuet-1 (NdeI/XhoI) (Genscript). A C-terminal VSV-G tag for detection by western blot was added using restriction sites NdeI and Xho1. Eag-TMD complexes SciW-Rhs1 (Eag: SL1344_0285, Rhs1(1–59)NT TMD: SL1344_0286) and EagT6-Tse6 (Eag: PA0094, Tse6(1–61)NT TMD: PA0093), were created previously in the vector pETDuet-1 in Ahmad et al.27. C-terminal VSV-G-tagged Eag chaperones were both cloned into multiple cloning site 2 (Nde1/Xho1) and N-terminal domains of the effectors (TMDs) were cloned into multiple cloning site 1 with a C-terminal His6-tagged (Nco1/HindIII). Eag-TMD point variants were created using Q5 Site-directed mutagenesis kit (New England Biolabs (NEB)) by amplifying the full-length construct using primer pairs outlined in Supplementary Table 4. Primers were generated from Integrated DNA Technologies (IDT) and constructs were confirmed by Sanger sequencing (Sick Kids Toronto). The complete set of strains and plasmids used in this study are outlined in Supplementary Tables 5 and 6.

Protein expression and purification of Eag chaperones
Eag chaperones and their point variants were overexpressed in E. coli BL21(DE3)-Gold cells using Lysogeny Broth (LB) culture medium and supplemented with 50 μg/mL kanamycin. Cells were grown at 37°C in a shaking incubator overnight and the next day diluted in 3 liters of LB broth and grown to OD600 of 0.6. Protein expression was induced by the addition of 1 mM isopropyl β-d-1-thiogalactopyranoside (IPTG) and cells were further incubated for 20 h overnight at 20°C. Next, the cells were harvested by centrifugation at 4200×g for 30 min and resuspended in Eag lysis buffer (50 mM Tris pH 8). PMSF and MgCl2 were added at final concentrations of 1 mM and 10 mM, respectively, along with a small amount of DNase. The cells were then lysed using an Emulsiflex-C3 High Pressure Homogenizer (Avestin) and centrifuged at 17,000×g for 25 min at 4°C. The supernatant was passed over a Q-sepharose gravity column (GE Healthcare) equilibrated with 2× column volumes of Eag lysis buffer for purification by ion exchange. The column was washed with 3× column volumes of lysis buffer before being subjected to a stepwise gradient of 1× column volumes of Eag lysis buffer containing increasing concentrations of NaCl as follows: 50 mM NaCl, 100 mM NaCl, or 1 M NaCl. Fractions from the 50 mM elution step (50 mM Tris pH 8.0, 50 mM NaCl) were concentrated to 2 mL by centrifugation using a 3 kDa concentrator (Amnicon). The concentrated elution was further purified by size-exclusion chromatography (SEC) using a HiLoad (16/600) Superdex 75 column (Cytvia) with an AKTApure (Cytvia). The SEC column was equilibrated with gel filtration buffer (50 mM Tris pH 8.0, 250 mM NaCl, 1 mM 2-Mercaptoethanol (BME)) before samples were injected. All SEC purifications were performed at 4°C. To confirm purity, fractions were run on a 13% sodium dodecyl sulfate-polyacrylamide electrophoresis (SDS-PAGE) gel and visualized with Coomassie Brilliant Blue. Selected fractions were further concentrated and flash frozen in liquid nitrogen for further use.

Protein expression and purification of Eag-TMD complexes
Eag-TMD complexes and their point variants were expressed in BL21(DE3)-Gold cells using LB culture medium supplemented with 100 μg/mL ampicillin. Eag-TMD complexes and point variants were lysed and harvested as described above. For Eag-TMD complexes, the Eag-TMD lysis buffer consisted of 50 mM Tris pH 8.0, 500 mM NaCl, and 20 mM imidazole. Protein complexes were purified by affinity chromatography using a nickel-NTA agarose gravity column (Goldbio) equilibrated with 1× column volume of lysis buffer. The bound proteins were washed with 3× column volumes of lysis buffer before being eluted by 1× column volume of elution buffer (50 mM Tris pH 8, 500 mM NaCl, 500 mM Imidazole). Following metal-affinity chromatography, Eag-TMD complexes and point variants were further concentrated and purified by SEC using the same protocol as the Eag chaperones. For all Eag-TMD complexes and point variants, the gel filtration buffer was 50 mM Tris pH 8.0, 250 mM NaCl, 1 mM BME. Finally, pure fractions as assayed by Coomassie-stained SDS-PAGE gel were concentrated and flash frozen in liquid nitrogen for further crystallization and biophysical experiments.

Dynamic light scattering (DLS)
DLS experiments were performed with a Prometheus Panta (NanoTemper) in DLS buffer (50 mM Tris pH 8.0, 250 mM NaCl, 1 mM BME) at 25°C and a protein concentration of 0.5 mg/ml. Each measurement lasted for 15 s and was repeated 10 times in triplicate thus providing accumulation of 30 measurements. The data were analyzed with the PR.Panta Analysis software version 1.10.2 (Nanotemper) using the intensity distributions for all subsequent evaluations.

Nanodifferential scanning fluorimetry (nanoDSF)
Thermal unfolding experiments of purified wild-type and variant Eag chaperones and Eag-TMD complexes were performed in a Prometheus NT.48 instrument (NanoTemper Technologies Inc.). Protein samples were diluted in gel filtration buffer (50 mM Tris pH 8.0, 250 mM NaCl, 1 mM BME) to 0.5 mg/mL and loaded in 10 μL nanoDSF Grade Standard capillaries in triplicate. Samples were heated from 20°C to 95°C with a temperature ramp of 1.0°C/min. Fluorescence intensity was monitored at 330 nm and 350 nm after excitation of tryptophan at 280 nm. Melting temperatures (Tm) were calculated from the first derivative of the 350/330 nm emission ratio. Data was analyzed with the PR.ThermControl software (NanoTemper Technologies Inc.) and GraphPad Prism version 10.0.2, GraphPad Software, Boston, MA, USA.

Differential scanning calorimetry (DSC)
Purified wild-type Eag chaperones and Eag-TMD complexes were dialyzed overnight into buffer containing 50 mM PBS pH 7.5, 250 mM NaCl, and 0.1 mM Tris(2-carboxyethyl)phosphine hydrochloride. For DSC measurements, an equal volume of 0.5 mg/mL protein solution in DSC buffer as well as a reference solution containing DSC buffer without the addition of protein, were prepared. Both the protein and reference solutions were degassed for 10 min under vacuum to ensure removal of air bubbles prior to loading of DSC. Experiments were performed using a NanoDSC (TA Instruments, New Castle, DE) with a 0.3 mL capillary cell volume. Samples were equilibrated at 20°C for 600 s and heated to 95°C with a scan rate of 1°C per minute at a constant pressure of 3 atm. A background scan was performed by loading both sample and reference cells with reference solution prior to the sample scan. Data was analyzed using NanoAnalyze (TA Instruments, New Castle, DE). The sample background was subtracted, and a sigmoidal baseline was applied to the data. The data was fit to a two-state scaled model for the transition.

Steady-state tryptophan fluorescence spectral analysis
For chemical denaturation experiments using the Prometheus NT.48 (fixed 330 and 350 nm), purified proteins were denatured in increasing concentrations of either urea of guanidine hydrochloride (0.08 M to 7.6 M urea, Δ dilution=0.3 per capillary; 0.6 M to 5.6 M guanidine hydrochloride, Δ dilution=0.24 per capillary). For each experiment, capillaries containing 24 different concentrations of denaturant and 0.5 mg/mL of protein were loaded in triplicate into a Prometheus NT.48 instrument (NanoTemper Technologies Inc.). Data was analyzed with the PR.ChemControl software (NanoTemper Technologies Inc.) and GraphPad Prism version 10.0.2 GraphPad Software, Boston, MA, USA.

For quantitative spectral analysis of wild-type Eags and both Q58A and L66A variants, intrinsic protein fluorescence was measured in the presence of increasing concentrations of guanidine hydrochloride (GdmCl) (1.50 M, 2.0 M, 3.0 M, 3.5 M, 4.0 M, 4.5 M, 5.0 M, 5.5 M, 6.0 M). Concentrated protein was dialyzed in 1 L of buffer (25 mM Tris pH 7.0) for 1.5 h to ensure adequate buffer exchange. The protein was then diluted to the appropriate stock concentration. Steady-state fluorescence spectra were measured on a Fluorolog-3 Horiba Jobin Yvon spectrofluorometer (Edison, NJ) using a 10×2 mm2 quartz cuvette to hold the sample. The samples were excited at 280 nm, the excitation and emission slits were set to a 2 nm bandpass. A series of 500 μL samples with a final concentration of 1 μM protein dimer (2 μM monomer), were prepared at various GdmCl concentrations. All samples were prepared in triplicate. The samples were equilibrated at room temperature for 1 h prior to the scan. The final denaturant concentrations were determined by refractive index using C10 Abbe Refractometer (VEE GEE Scientific) post data acquisition. The data were analyzed with SigmaPlot version 12.3 (2013) Systat Software Inc, (San Jose, CA) software and OriginPro Version 2025 (OriginLab Corporation, Northampton, MA, USA).

Calculation of free energy of Eag unfolding and TMD binding from steady-state tryptophan fluorescence
If it is assumed that the chaperones SciW and EagT6 unfold through a two-state mechanism with an equilibrium constant of Kunfolding:

dimer,folded ⇌ 2 monomer unfolded (1)

The fraction of tryptophan residues in the unfolded state “α” can be related to the measured λmax through the equation47:

α = (λmax − λf) / (λu − λf) (2)

The denaturant concentration dependencies of λf and λu are given in Supplementary Fig. 3. Allowing us to define the equilibrium constant:

Kunfolding = 2c0 α2 / (1 − α) (3)

Where c0 is the total concentration of Eag monomer in solution, λf is the emission maximum wavelength of the folded protein and λu is maximum wavelength of the unfolded protein.

Next, Kunfolding is related to the change in free energy by:

ΔGunfolding = −RT ln Kunfolding (4)

Which allows the determination of the standard change in unfolding free energy in buffer ΔGo unfolding, assuming a linear free energy dependence upon denaturation concentration44:

ΔGunfolding = ΔGo unfolding + m[GdCl] (5)

Based upon this, the denaturation concentration dependence of α (fraction unfolding) is given by:

α = [−Kunfolding + √(Kunfolding^2 + 8c0 Kunfolding)] / 4c0 (6)

In which Kunfolding = e^(−(ΔGo unfolding + m[GdCl]) / RT).

If the TMD is assumed to selectively bind the folded form of the Eag with an association constant KTMD, the effect of TMD binding upon the standard unfolding free energy for the chaperone can be estimated using the relationship49,50:

ΔGo unfolding,TMD bound = ΔGo unfolding,apo + RT ln(1 + KTMD[TMD]) (7)

Where KTMD = [chaperone-TMD] / ([chaperone][TMD]), if the TMD binds the chaperone tightly, the majority of the chaperone is bound to the TMD and [chaperone-TMD] ≈ 0.5c0, [TMD] ≈ 0.5c0 and KTMD[TMD] is significantly larger than one. We can then simplify Eq. (7) to:

ΔGo unfolding,TMD bound ≈ ΔGo unfolding,apo + RT ln(0.5c0 × KTMD) (7a)

Rearranging this equation we obtain:

ΔGo unfolding,TMD bound − ΔGo unfolding,apo = 0.5RT ln 0.5c0 + 0.5RT ln KTMD = 0.5RT ln 0.5c0 − 0.5ΔGo binding (7b)

Stopped-flow kinetics
Unfolding kinetics were performed using Applied Photophysics SX-20 (Surrey, UK) stopped-flow fluorescence instrument (dead time < 2 ms). To ensure the appropriate dilution of denaturant, asymmetric mixing was set up using 2.5 mL and 250 μL drive syringes purchased from Deltaphotonics (Ottawa, CA). Purified wild-type and variant Eag chaperones and complexes were first diluted to a concentration of 10 μM in gel filtration buffer (50 mM Tris pH 8.0, 250 mM NaCl, and 1 mM BME). Purified protein was diluted in a 1/10, v/v, mixing ratio with guanidine hydrochloride (GdmCl) in 50 mM Tris, 250 mM NaCl, 1 mM BME of the desired concentration. The excitation wavelength was set to 280 nm, and the emission was monitored using a 330 nm Bandpass filter (FWHM 10 nm). All data was collected at 20°C. At least 12 unfolding traces were collected at each desired concentration of GdmCl and averaged. The final denaturant concentrations were determined by refractive index using C10 Abbe Refractometer (VEE GEE Scientific) post data acquisition. All data was analyzed using Sigma Plot 12.3 (2013) Systat software (San Jose, CA).

Calculation of unfolding rate constants and psi-value analysis from stopped-flow kinetics
The EagT6 and SciW relaxation profiles can be described as a mono-exponential decay:

F = F0 e^(−kobs t) (8)

Where F is the measured fluorescence, F0 is the fluorescence at time zero and kobs is the measured rate constant; at high denaturant concentrations kobs ≈ kunfolding, kunfolding being the apparent unfolding rate constant. Using a chevron-type analysis, it can be assumed:

ln kunfolding = ln k°unfolding + m0[denaturant] (9)

Where kunfolding is the unfolding rate constant measured at a given concentration of denaturant, ln k°unfolding is the unfolding rate constant in pure buffer, and m0 is the “kinetic m-value”. We have measured time-dependent unfolding traces of all Eag proteins (WT and variants), in the apo and TMD-bound states and have plotted ln kunfolding as a function of denaturant concentration in Fig. 5; analyzing this data via Eq. (9) yields the kinetic parameters ln k°unfolding and m0 as shown in Table 1. For “φ-value analysis”, if the free energy of the unfolding pathway’s rate-limiting transition state is represented by ΔG‡unfolding = −RT ln k°unfolding + RT ln A in which A is the pre-exponential factor defined by transition state theory and the φ-value for each mutant is defined as:

φ = (ΔG‡unfolding,WT − ΔG‡unfolding,mutant) / (ΔG°unfolding,WT − ΔG°unfolding,mutant) (10)

P. aeruginosa strains and growth conditions
P. aeruginosa strains were derived from the sequenced strain PAO1. Cultures were grown in LB medium (10 g/L tryptone, 5 g/L yeast extract, 10 g/L NaCl), shaking at 37°C. Solid media containing 1.5% or 3% (w/v) agar, as indicated. E. coli strains XL1 blue and SM10 were used for plasmid maintenance and conjugative transfer, respectively. E. coli strains were grown in LB at 37°C in a shaking incubator. Cultures were supplemented with 15 mg/mL gentamicin (E. coli), 30 mg/mL gentamicin (P. aeruginosa), and 25 mg/mL irgasan (P. aeruginosa).

DNA manipulation, plasmid construction, and strain generation for bacterial competition and secretion assays
All primers were synthesized by IDT. Phusion polymerase, restriction enzymes, and T4 DNA ligase were obtained from NEB. Sanger sequencing was performed by the Centre for Applied Genomics, The Hospital for Sick Children in Toronto, Ontario. Chromosomal mutations in P. aeruginosa were generated by double allelic exchange as previously described69. Approximately 500 bp flanks upstream and downstream of the region to be mutated were amplified by PCR and spliced together by overlap-extension PCR. Primers were designed such that the desired point mutation was contained within the overlap. The resulting amplicon was ligated into the pEXG2 allelic exchange vector, transformed into E. coli SM10, and introduced to P. aeruginosa by conjugative transfer. Merodiploids were selected on LB agar containing 30 mg/mL gentamicin and 25 mg/mL irgasan and streaked on LB agar lacking NaCl containing 5% (w/v) sucrose for sacB counter-selection. Strains that grow on sucrose but are gentamicin sensitive were screened by Sanger sequencing the PCR amplicon of the region of interest to confirm the presence of point mutations. A complete list of strains used in this study is outlined in Supplementary Table 6. Primers for the creation of Pseudomonas mutant strains are shown in Supplementary Table 7.

Bacterial competition assays in P. aeruginosa
Assays were performed in co-culture as previously described70. Recipient strains harboured a constitutively expressed lacZ gene at a neutral site in the chromosome to distinguish them from donor strains upon plating on LB containing 40 mg/mL 5-bromo-4-chloro 3-indolyl-β-D-galactopyranoside (X-gal). Overnight cultures of indicated donor and recipient strains were diluted to an OD600 of 1.0 and combined in an initial donor/recipient ratio of 5:1 (v/v), and the initial mixture was serially diluted to enumerate colony-forming units (CFU) by plating on media containing X-gal. 10 μL of this mixture was spotted onto a nitrocellulose membrane on LB containing 3% (w/v) agar and cultured at 37°C for 20 h. Co-cultures were resuspended in 1 mL LB and serially diluted to enumerate CFU. Data are presented as the fold change in the ratio of predator to prey at the end of the experiment relative to the beginning. Statistical significance was calculated by a one-way ANOVA on the log-transformed competitive indices using Tukey’s correction for multiple comparisons between each mean. Western blot was done using a custom polycolonal antibody against Tse631 (1:3000 dilution) and a commercial anti-RNAP antibody (BioLegend) (1:5000 dilution).

Bacterial competition assays in S. Typhimurium
Briefly, cultures of S. Typhimurium SL1344 (streptomycin-resistant) and E. coli DH5α carrying pET29b (kanamycin-resistant) were grown overnight in LB broth supplemented with the appropriate antibiotic. The following day, overnight cultures were washed three times with PBS and adjusted to an OD600 of 0.4. The strains were then mixed at a 1:1 (vol/vol) ratio of wild-type or mutant S. Typhimurium to competitor E. coli DH5α. To determine the initial CFU ratio, the input mixture was serially diluted and plated on LB agar containing either kanamycin or streptomycin. Interbacterial competition assays were performed by spotting 10 μL of the mixed cultures onto 0.22 μm nitrocellulose on LB agar plates overlaid with either 0.05% ox-bile (Sigma) or PBS (control). After 48 h, cells were recovered from the membranes with an inoculating loop, resuspended in PBS, serially diluted, and plated on selective LB agar to enumerate final CFU counts. The competitive index was calculated as the fold change in the ratio of S. Typhimurium to E. coli between the initial and final CFU counts. Statistical significance was assessed using an unpaired t-test. A complete list of strains used in this study is outlined in Supplementary Table 4. The wild-type SL1344, ΔsciW, and ΔclpV mutant strains were a generous gift from Brian Coombes (McMaster University)63.

Protein crystallization
Purified SciW-Rhs1(1–59) L66A was screened for crystallization utilizing commercially available screens (Nextal) and a crystal Gryphon robot (Art Robbins Instruments). Crystals of SciW-Rhs1(1–59) L66A grew in gel filtration buffer and 0.1 M MES pH 6, 0.2 M MgCl2, and 20% (v/v) PEG 6000 20 mg/mL protein in a 1:1 mixture at 4°C. Crystals were further optimized by micro-seeding using a Seed Bead kit (Hampton Research). Crystals were cryo-protected in 30% PEG 6000 before flash freezing directly in liquid nitrogen. Data was processed using XDS71 and CCP472. Initial phases were obtained from Phenix73 using SciW-Rhs1(1–59) structure (PDB id 6XRR) as a model for molecular replacement. The structure was further built using Coot74 and refined using Phenix, and TLS refinement75. Molecular graphics were generated using ChimeraX and GraphPad Prism version 10.0.2.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The bacterial type VI secretion system (T6SS) is a potent antimicrobial weapon used by Gram-negative species. The T6SS can deliver numerous toxins with a broad range of biochemical activities, including enzymatic such as peptidoglycan hydrolases or lipases, as well as membrane pore forming. Targeting and loading of effectors onto the T6SS machine can involve a range of chaperones of which the Eag family has been involved in protecting hydrophobic and transmembrane domain (TMD) regions in toxins, such as nicely described between EagT6 and the Pseudomonas aeruginosa toxin Tse6. Here the authors studied in very fine detailed the molecular interactions between Eag and two T6SS effectors, T6SS and SciW from Salmonella Typhimurium. They performed in depth thermodynamic and biophysical characterization to monitor the high stability of the Eag-TMD interaction that prevents transmembrane domains to prematurely insert into membranes as well as the subsequent effective release of this interaction. To support their data, they investigate structural features of domain interactions and identify key residues essential for the stability of the complex and which are found in a knob-in-hole packing arrangement features. This is further validated by genetic studies in vivo demonstrating that lack of stability in the Eag-TMD by substituting these key residues results in absence of killing during interbacterial competition. Overall, the study is very technical and methodological, but very convincing although the gain in knowledge is rather incremental\nSpecific comments\nThe difference in lifetime and stability between the complex EagT6-Tse6 and Eag-SciW is astonishing, 18s vs 15h, respectively. Although the authors put these two comp0lexes in different classes, it seems that the key residues involved in stability are the same in both complexes. Could this mean that other domains in the T6SS effector may still strongly influence the stability of the complex and possibly that additional partners (line 486) strengthen and weaken the interaction. For example, if Eag-SciW1 separation requires disruption of more than one contact (lines 369-370) then this may be catalyzed by additional partner. Do the Eags have other partners that could be identified for example through in vivo pull-down. Are there mutations that could be identified that would dramatically increase the lifetime of Eag-Tse6 or dramatically decrease lifetime of Eag-SciW1.\nThe authors suggest that the contact disruption within the Eag complexes occur by conformational change once the effector is in the T6SS baseplate (line 472). The firing and contraction mechanism associated with T6SS dynamic, involves a drastic conformation change in the baseplate from dome to starshaped. Would this contraction event contributing to the energy requires to disrupt Eag complexes. Could the Eag complex loaded in the T6SS nanomachine be modelled in the context of an extended or contracted version of the T6SS?\nIn figure 6 the most drastic impact for interbacterial competition is the Q102A substitution which is at least as impactful than the L66A substitution. The equivalent of Q102A substitution in the Eag-SciW1 complex was unsuccessful (lines 231-232). As such there is very little data or discussion on that particular residue in the main text. Yet could it be a difference that may contribute explaining the lifetime differences between Eag-Tse6 and Eag-SciW1. This may be worth to investigate and discuss further.", "reviewer_2": "Since my experience with the various biophysical techniques used in this manuscript is limited, I will only comment on the phenotypes related to T6SS function and impact of this paper on the field. This is overal a well written manuscript that addresses an important aspect of T6SS effector loading. The insights are interesting and such a detailed analysis is important for the field. I see no missing controls for the experiments testing phenotypes of the mutations generated based on the biochemical analysis. I suggest to add a figure with a cartoon like model for effector loading on the T6SS spike and removal of the chaperon (synthesis of the presented data and previous knowledge). This would help readers to imagine the whole sequence of events during T6SS assembly and effector loading.", "reviewer_3": "Effectors of the type VI secretion system (T6SS) are usually delivered with the ejected needle (Hcp) or its tip (VgrG/PAAR), either as direct fusions to these proteins or bound to them. Some effectors require chaperone proteins that can take various roles, but are usually required for delivery of their cognate effector, without being exported themselves. One such class of chaperones are the Eag chaperones, which in particular bind transmembrane domain (TMD) effectors. The authors of this manuscript had investigated the binding of Eag chaperones to a conserved motif found in TMD-containing effectors in eLife in 2020. In this manuscript, they follow up on this publication by addressing the obvious question how the Eag chaperones are removed from the effectors prior or during secretion. To this aim, the authors measure the stabilization of two Eag chaperones by the presence of the TMD of their respective effectors. They complement this analysis by structural analysis and a set of competition and expression experiments on a set of mutants.\nThe manuscript investigates an interesting question and the results appear solid and are well-presented (excluding the points raised in the minor comments). However, the reasoning and logical connections are hard to follow in various places, and it is unclear if the experiments (largely unfolding by chaotropic agents) allow for the conclusions on the T6SS assembly or export process. It is, therefore, difficult to assess whether the authors’ model and conclusions are justified by the data.\nMajor comments:\n1. In particular, it remains unclear at which time point the authors think the Eag chaperones are removed from the TMH effectors. The scheme in the authors’ 2020 eLife paper shows the Eag chaperones as parts of the assembled T6SS prior to firing. This assumption is also retained throughout the manuscript, until, suddenly, in the description of the model (lines 463 onwards), Eag-TMD release is hypothesized to occur upon binding to the baseplate (which is not analyzed or even mentioned once in the results). The two scenarios are vastly different, as removal upon firing would likely imply mechanical stress, which is quite different from the scenario to which the measurements likely apply (GdmCl-induced unfolding).\n2. The authors state that they have been unable to express TMD effectors without their chaperones and understandably, therefore, choose indirect methods to analyze the binding and unbinding. However, it is not immediately obvious that these methods allow to draw the conclusions made in the manuscript. To the best of my knowledge, changes in the Tm provide qualitative, but not quantitative binding affinities. Perhaps more importantly, this applies to equilibrium kinetics, but not to mechanical stress, as possible relevant for the removal of the T6SS chaperones. If the methods do allow for these conclusions, the underlying reasoning needs to be made clearer to the reader.\n3. Negative controls, such as the addition of generic TM or of the respective non-cognate TMD, are missing for the key experiments.\nMinor comments:\n1. Why does Rhs1(1-59) aggregate on SDS-PAGE gels (Fig. S1 and S5), but not in the DLS measurements (Fig. S5)? Tse6(1-61) behaves different in this respect. Additional controls, beyond the fact that Rhs1(1-59) still binds to SciW, would provide evidence for its native binding, and the respective conclusions.\n2. Is there a difference between chaperones and adapters? If so, it should be explained, if not, this should be mentioned when mentioning adaptors (e.g. Tap adaptors).\n3. The reason to focus on the first TMD of Tse6 (or a reference showing binding of this TMD only to EagT6) should be provided.\n4. Lines 66/67: This statement does not apply to all effectors, and is therefore misleading in its current state.\n5. The role of chaperones should be explained more clearly – lines 67-69 are rather vague and unspecific.\n6. Line 123: Rhs should be called Rhs1, as elsewhere in the manuscript.\n7. The dimer formation of Eag chaperones, one of their defining features, is not taken into consideration in the denaturation experiments (Fig. 1, 2), although likely to influence the results. Likewise, it is not taken into account in the equilibrium calculations described in lines 692-698. The authors seem to assume an essentially complete dimerization of the chaperones, but no evidence for this is provided.\n8. How exactly does the difference in denaturation by GdmCl and Urea allow for the determination of the free energy of Eag-TMD unfolding (statement in lines 163-164)? Is urea used afterwards and what for?\n9. Are lines 212-218 new results (as implied by the inclusion in the results part and lack of reference)? If they are instead a recapitulation of the results in ref. 27, they should be moved from the results part to the introduction, or this fact should be at least made clear.\n10. The cellular concentration of Tse6 (which the authors, conceivably, assume to be determined by the binding to its chaperone) does not correlate well with the competitive index. For example, the Q58A mutant, in which the level is similar to the WT, has a strongly decreased fitness compared to the WT, and S41A, which retains expression, is doing worse than I24F, where almost no Tse6 could be detected. Can the authors explain these discrepancies?\n11. What is meant by “the Eag chaperone catalyzes completion of the PAAR domain with the prePAAR motif” (lines 463-465, in the description of the model)? The expression is not contained in the listed ref. 27, and the mechanics described by it remain unclear.\n12. There is a certain inconsistency and confusion in the gene identifiers given by the authors (lines 547-552 and elsewhere):\n- SL1344_0268, assigned to Salmonella SciW in line 547, encodes instead for TssC, the large subunit of the T6SS contractile sheath\n- SL1344_0286, assigned to SctW elsewhere, seems to encode for an RHS protein instead\n- SL1344_0285, assigned to Rhs1, encodes for a much smaller protein\n- Likewise, the two protein identifiers may have been mixed up for the Pseudomonas genes", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_5": "The manuscript by Van Schepdael et al. explores the mechanism of effector transmembrane domain (TMD) release from Eag chaperones, crucial for Type VI secretion system (T6SS) function. The authors use a combination of biophysical techniques and bacterial assays to demonstrate that small perturbations in Eag-TMD interactions, such as disrupting hydrogen bonds or knob-in-hole packing, can trigger TMD release. Their work underscores the essential role of conformational changes in Eag chaperones for the dissociation of TMDs, a critical step for effector secretion and bacterial fitness. Additionally, the X-ray co-crystal structure of the Eag-TMD complex captures an intermediate state of TMD release, offering a novel perspective on this process. This study contributes significantly to our understanding of how T6SS chaperones regulate effector folding and secretion. The insights provided in this study are important and should be of broad interest. While, to strengthen the work, the authors may need to address my concerns listed below.\nMajor concerns:\n1.The study investigates the mechanism of Eag chaperone dissociation from effector proteins, but the extent to which the in vitro-derived dissociation mechanism reflects the actual dissociation process in bacterial cells is questionable. For example, the manuscript primarily examines the impact of different mutations on bacterial secretion but does not confirm whether these mutations specifically affect Eag dissociation, which then influences bacterial secretion and survival. The authors should include in vitro dissociation kinetics experiments and in vivo/in-cell binding assays to demonstrate the role of chaperones in regulating effector protein folding and their involvement in T6SS under physiological conditions.\n2.The SciW-TMD L66A crystal structure presented in the study captures an intermediate state of Eag chaperone dissociation from TMD and supports the proposed dissociation model. However, this structure was obtained by introducing the L66A mutation to stabilize the dissociation intermediate. It is unclear whether this structure fully reflects the dissociation process under physiological conditions in bacterial T6SS, as wild-type chaperones in cells may dissociate differently. Could the structure of the wild-type chaperone dissociating from TMD under natural conditions be obtained by optimizing parameters such as temperature?\n3.The study attributes the dissociation pathways of Class 1 (SciW-TMD) and Class 2 (EagT6-TMD) effector proteins to “the different roles of Q58/L66 contacts” and “structural differences in transition states.” However, the TMD sequences, overall conformations, and coupling with VgrG/PAAR differ significantly between the two effector classes, suggesting that other factors may regulate chaperone dissociation. The authors should include further experiments to confirm that the chaperone-TMD interactions are responsible, rather than other factors playing a key role in regulating T6SS.\n4.The authors determined the crystal structure of SciW L66A in complex with its substrate, while the functional assays in Figure 6 were conducted using EagT6. Given the clear differences between the two proteins, additional cell-based functional validation experiments using the SciW mutant should be included to support the structural findings.\n5.The dissociation rates between SciW-TMD and EagT6-TMD appear to differ significantly, suggesting that their dissociation mechanisms may not be identical. Therefore, the intermediate dissociation state of SciW-TMD captured in this study may not be universally representative. It would be valuable for the authors to further elaborate on this point in the discussion, addressing the potential differences in dissociation mechanisms and the broader relevance of the observed intermediate.\nMinor concerns:\n1.In Figure 6B, the expression levels of Tse protein are analyzed, and the authors suggest that different mutations in Eag affect bacterial competition activity by altering Tse protein stability. However, the figure lacks a quantitative analysis of Tse protein expression. From the data, the Tse protein expression in the L66A group is lower than in the Q102A group, yet the L66A group shows higher competition activity. The authors should provide a quantitative analysis or offer an alternative explanation for this discrepancy.\n2.There is an extra scale bar in the lower right corner of the eagT6 label in Figure 6B. It is recommended to remove this unnecessary scale bar.\n3.The panel numbers (A, B, C) in different figures are inconsistently sized. It is recommended to adjust them to a uniform size for better presentation.\n4.For the Eag variants, the authors only performed unfolding assays to assess their resistance to chemical denaturation. I recommend that the revised manuscript include NanoDSF measurements to evaluate the stability of the Eag variants both individually and in complex.\n5.In Figure S6H, the unfolding rate of the bound state of the EagT6 L66A mutant appears to be faster than that of the unbound state. This seems counterintuitive. Is the TMD still bound to the EagT6 L66A protein under these conditions? Further clarification or experimental validation may be needed to confirm the binding status."}, "author_response": "Response: We thank the reviewer for their time and suggestions to improve our\nmanuscript, especially their positive comments on the rigor and high quality of our\nexperimentation. With our manuscript edits, we hope we have more clearly emphasized\nthe significance and broad interest of our work. Including the methodological impact to\nstudy interactions with integral membrane proteins.\nSpecific comments\n>The difference in lifetime and stability between the complex EagT6-Tse6 and Eag-SciW is\nastonishing, 18s vs 15h, respectively. Although the authors put these two complexes in\ndifferent classes, it seems that the key residues involved in stability are the same in\n2\nboth complexes. Could this mean that other domains in the T6SS effector may still strongly\ninfluence the stability of the complex and possibly that additional partners (line 486) strengthen and weaken the interaction. For example, if Eag-SciW1 separation requires disruption of more than one contact (lines 369-370) then this may be catalyzed by additional partner. Do the Eags have other partners that could be identified for example\nthrough in vivo pull-down.\nResponse: We agree with the reviewer that another binding partner likely catalyzes\nrelease. Our work details the key interactions and conformational changes of the Eag\nrequired to bind/release the TMD. In the discussion we hypothesize that another T6SS\nrelated protein is required to induce a conformational change in the Eag. Our results\nsupport that the conformational change includes moving L66 away from the TMD to initiate\nTMD dissociation. We have emphasized this to make it clearer for the reader. This\nincludes edits to Figure 7 showing that L66 is part of the native conformational change\nupon TMD binding.\nAttempts to discover additional Eag binding partners has been an ongoing line of study in\nour lab and has already been previously attempted. Notably, pull-down experiments with\nEags have already been performed in Pseudomonas but only revealed the interaction with\nthe cognate effector Tse6 and the VgrG.\nWhitney et. al. Cell. 2015 Oct 22;163(3):607-19. doi: 10.1016/j.cell.2015.09.027.\nEpub 2015 Oct 8. PMID:26456113\nGiven that the previous pulldown results only found (Eag-TMDeffector-VgrG) any\nadditional binding partner is likely to be transient which required challenging timing and/or\nadvanced cross-linking to capture in a pull-down.\nEven after this complex optimization, potential hits must be cloned and purified to test\nbinding with Eag and Eag-TMD complexes. Mass-spectrometry identification of proteins\nfrom pull-downs commonly gives a list with many false-positives, and optimization of\nproteins for over-expression for binding assays is non-trivial. We have not yet been able\nto prove another Eag-T6SS interaction. Regardless, the results of a binding-partner\ndiscovery study when complete would constitute an entire manuscript of work on its own.\nHowever, in the discussion we have stated that additional experimentation would be\nrequired to discover other Eag T6SS protein interactions.\n>Are there mutations that could be identified that would dramatically increase the lifetime\nof Eag-Tse6 or dramatically decrease lifetime of Eag-SciW1.\nResponse: We agree that this is an interesting question. We do note that we have\nalready discovered mutations that dramatically decrease the lifetime of SciW-Rhs1\n(Q58A and L66A), albeit not to the level of Eag-Tse6.\nprograms (RFdiffusion etc.). Furthermore, the aim of the current work is to determine the\nnative molecular features of the chaperones required for effector TMD binding and\ndissociation. A study to redesign the Eags for increased TMD binding by introducing\nartificial stabilizing interactions is beyond the scope of the study.\n>The authors suggest that the contact disruption within the Eag complexes occur by\nconformational change once the effector is in the T6SS baseplate (line 472). The firing\nand contraction mechanism associated with T6SS dynamic, involves a drastic\nconformation change in the baseplate from dome to starshaped. Would this contraction\nevent contributing to the energy requires to disrupt Eag complexes. Could the Eag\ncomplex loaded in the T6SS nanomachine be modelled in the context of an extended or\ncontracted version of the T6SS?\nResponse: We agree with the reviewer. A mechanism of Eag-TMD dissociation could be\nthat a T6SS baseplate protein binds the Eag and the conformational change of the\nbaseplate is tied to removal. The energy could be used to induce an Eag conformational\nchange at the time of contraction to dissociate the TMD. Our data shows that the wild-type\nEag-TMD complexes are incredibly stable, but the mechanical stress of T6SS firing would\nlikely provide enough energy to break the complex (PMID: 25822993). Or at least provide\nenough energy to induce a conformational change that disrupts the interaction with L66\nand/or Q58. We have added this as a potential model (discussion, Figure 8).\n>In figure 6 the most drastic impact for interbacterial competition is the Q102A substitution\nwhich is at least as impactful than the L66A substitution. The equivalent of Q102A\nsubstitution in the Eag-SciW1 complex was unsuccessful (lines 231-232). As such there\nis very little data or discussion on that particular residue in the main text. Yet could it be a\ndifference that may contribute explaining the lifetime differences between Eag-Tse6 and\nEag-SciW1. This may be worth to investigate and discuss further.\nResponse: We agree with the reviewer. As we were unable to make Q106A in SciW we\nhave ordered it from Genscript. The results of these assays have been added to all other\nSciW variant figures (Figure 3, Table S2, Table S3, Table S5, and Figures S4-7).\nQ106A had little to no effect on the interaction between SciW and Q106A. This point has\nbeen added to the discussion of chaperone mechanistic differences.\n>Overall, the study is very technical and methodological, but very convincing although the\ngain in knowledge is rather incremental\nResponse: We thank the reviewer for the positive review but would like to stress and\nclarify the broad importance and impact of our work.\nare removed from effectors as part of the T6SS secretion process. To our knowledge, our\nwork is currently the only study that provides energetic and detailed mechanistic data of\nhow any T6SS chaperone is dissociated from its cargo. Because of this, our biophysical\ndata is a significant step forward in the field of T6SS.\nSecond, a significant scientific advancement of our work was the development of an\nexperimental strategy and model (eq. 1-9) to overcome the obstacles of studying the\ninteraction of a soluble chaperone and an insoluble, unstable membrane protein.\nSpecifically, because the TMD can only be produced already bound to the Eag. We\nsucceeded and have extracted detailed mechanistic binding/dissociation data on Eags\nand their effectors that was previously unattainable.\nThird, our findings are of interest not only for T6SS chaperones, but in the field of protein\nchaperones in general. This is in part because Eags are chaperones that unfold cargo\nand stabilize a mis-folded conformation to inhibit native folding until the proper time. The\nmechanistic and energetic details of how the Eag accomplishes this are relevant to\nunderstand other secretion chaperones and general cellular chaperones that work to\nunfold and then refold proteins.\nFinally, our experimental approach can also be a model system for other groups studying\nchaperones or proteins that require co-expression with a binding partner for solubility,\nespecially membrane protein chaperones.\nReviewer #2 (Remarks to the Author):\n>Since my experience with the various biophysical techniques used in this manuscript is\nlimited, I will only comment on the phenotypes related to T6SS function and impact of this\npaper on the field. This is overal a well written manuscript that addresses an important\naspect of T6SS effector loading. The insights are interesting and such a detailed analysis\nis important for the field. I see no missing controls for the experiments testing phenotypes\nof the mutations generated based on the biochemical analysis. I suggest to add a figure\nwith a cartoon like model for effector loading on the T6SS spike and removal of the\nchaperon (synthesis of the presented data and previous knowledge). This would help\nreaders to imagine the whole sequence of events during T6SS assembly and effector\nloading.\nResponse: We thank the reviewer for their positive assessment of our work, the writing\nof the manuscript, and the significance of our findings. We have added a mechanistic\nfigure as suggested (Figure 8).\nReviewer #3 (Remarks to the Author):\n>Effectors of the type VI secretion system (T6SS) are usually delivered with the ejected\nneedle (Hcp) or its tip (VgrG/PAAR), either as direct fusions to these proteins or bound\n5\nto them. Some effectors require chaperone proteins that can take various roles, but are\nusually required for delivery of their cognate effector, without being exported themselves.\nOne such class of chaperones are the Eag chaperones, which in particular bind\ntransmembrane domain (TMD) effectors. The authors of this manuscript had investigated\nthe binding of Eag chaperones to a conserved motif found in TMD-containing effectors in\neLife in 2020. In this manuscript, they follow up on this publication by addressing the\nobvious question how the Eag chaperones are removed from the effectors prior or during\nsecretion. To this aim, the authors measure the stabilization of two Eag chaperones by the\npresence of the TMD of their respective effectors. They complement this analysis by\nstructural analysis and a set of competition and expression experiments on a set of\nmutants.\nThe manuscript investigates an interesting question and the results appear solid and are\nwell-presented (excluding the points raised in the minor comments). However, the\nreasoning and logical connections are hard to follow in various places, and it is unclear if\nthe experiments (largely unfolding by chaotropic agents) allow for the conclusions on the\nT6SS assembly or export process. It is, therefore, difficult to assess whether the authors’\nmodel and conclusions are justified by the data.\nResponse: We thank the reviewer for their overall positive assessment of our work and\ntheir suggestions to improve our study. We have made edits to both improve readability\nand to make the reasoning behind our line of investigation clear to the reader.\nMajor comments:\n>1. In particular, it remains unclear at which time point the authors think the Eag chaperones\nare removed from the TMH effectors. The scheme in the authors’ 2020 eLife paper shows\nthe Eag chaperones as parts of the assembled T6SS prior to firing. This assumption is also\nretained throughout the manuscript, until, suddenly, in the description of the model (lines\n463 onwards), Eag-TMD release is hypothesized to occur upon binding to the baseplate\n(which is not analyzed or even mentioned once in the results).\nResponse: We apologize as we did not intend to suggest a model at odds with our past\npublication. The exact words used were:\n“Next, we hypothesize that the Eag is removed from the TMD within the\nbaseplate/membrane complex. ... Within the baseplate, a conformational change in Eag\nis induced to disrupt the Eag-TMD complex.”\nTo agree with our past publication, we only meant to suggest that dissociation occurs within\nthe baseplate. We currently do not know the exact time of removal or if the Eag-TMD does\nin fact directly bind another T6SS protein for dissociation. As also addressed for reviewer\n1, pull-down experiments to find and subsequently prove another Eag interaction partner\nhave been tried but have yet to produce substantial results.\nWe have further clarified this point in the model discussion section for the reader.\n6\n>The two scenarios are vastly different, as removal upon firing would likely imply\nmechanical stress, which is quite different from the scenario to which the measurements\nlikely apply (GdmCl-induced unfolding).\nResponse: We would again like to emphasize that although we used chaotropic agents\nto study Eag-TMD unfolding and stability, all the thermodynamic and kinetic\nparameters reported in table 1 are native values in buffer without any denaturant.\nOur measurements and model yield free-energy and kinetic values in a standard in\nvitro physiologically relevant experimental buffer.\nBy studying Eag and Eag-TMD folding at different concentrations of chaotropic agents we\ncan then fit the data with our model (equations 1-9) and extrapolate to 0 M GdmHCl. None\nof the reported binding or dissociation values are influenced by GdmHCl.\nThe data shows how much energy is required and the molecular contacts that are critical\nfor Eag-TMD complex formation and thus are likely disrupted to promote TMD\ndissociation. The energy for disruption and TMD dissociation may then come from the\nmechanical stress of T6SS firing (PMID: 25822993). Or at least provide\nenough energy to induce a conformational change that disrupts the interaction with L66\nand/or Q58. We have added this as a potential model (discussion, Figure 8).\nAdditionally, the biophysical data are supported by our current SciWL66A structure that\ndemonstrates a partial release of the TMD. Furthermore, our previous X-ray structures\nshow that the region containing L66 is part of the Eag conformational change induced\nupon TMD binding for both EagT6 and SciW. Thus, a hypothesis that disruption of the\ninteraction of L66 with the TMD is part of the dissociation mechanism is supported by our\nprevious structural data and all mechanistic data presented in this manuscript.\nHowever, expanding upon the details of a firing-coupled model where mechanical stress\nmay help provide the energy for Eag removal was also suggested by reviewer 1, and we\nhave expanded our discussion.\nWe have also included additional figure panels illustrating known Eag conformational\nchanges from our various structures in Figure 7. Also as suggested by reviewer 2, we\nhave added dissociation model in Figure 8.\n>2. The authors state that they have been unable to express TMD effectors without their\nchaperones and understandably, therefore, choose indirect methods to analyze the\nbinding and unbinding. However, it is not immediately obvious that these methods allow\nto draw the conclusions made in the manuscript. To the best of my knowledge, changes\nin the Tm provide qualitative, but not quantitative binding affinities. Perhaps more\nimportantly, this applies to equilibrium kinetics, but not to mechanical stress, as possible\nrelevant for the removal of the T6SS chaperones. If the methods do allow for these\nconclusions, the underlying reasoning needs to be made clearer to the reader.\nResponse: We have not used Tms (melting temperatures) for the derivation of our\nthermodynamic and kinetic parameters. All ΔG and rate constants (k ) are\nunfolding\n7\nmeasured from chemical denaturation then extrapolated to physiological buffer using our\nmodel in equations 1-9 (PMID: 11013396).\nAll ΔG° unfolding, ΔG° binding and k° in table 1 are native values in a physiologically\nunfolding\nrelevant buffer without any denaturant present.\nTo make this clear to the reader, we now state this immediately in the first chemical\ndenaturation section:\n“We next probed Eag chaperone resistance to chemical denaturation. By measuring Eag\nand Eag-TMD unfolding by denaturants, we will be able determine the free-energy (ΔG) of\nEag stabilization and the binding energy of the Eag-TMD interaction. Specifically, the ΔG of\nunfolding measured at various denaturant concentrations is used for extrapolation to 0 M\ndenaturant. This yields native denaturant-independent ΔGs of folding and TMD binding\nenergy in a physiologically relevant buffer (PMID: 11013396).”\nAdditionally, we have edited the manuscript in other locations to also better explain to the\nreader how and why our thermodynamic and kinetic parameters are physiologically\nrelevant.\nWe do agree with the reviewer that the chemical denaturation binding calculations can be\nviewed as estimates as they depend upon the assumptions in our model (equations 1-9).\nWe have clearly stated the assumptions inherent in the model for the reader:\n“The effector TMD binding energy (ΔG° ) and association constant (K ) can then be\nbinding TMD\nobtained from comparing Eag ΔG° to Eag-TMD ΔG° . This calculation requires\nunfolding unfolding\nthe following assumptions: 1) the TMD is a small ligand compared to the Eag, 2) TMD\nbinding does not significantly alter the Eag fold, 3) all free TMD in solution results from\ncomplex dissociation, 4) the chaperone folds by a 2-state model, and 5) the TMD is\nassumed to selectively bind the folded form of the Eag with an association constant K .”\nTMD\nHowever, we note that our mechanistic details are based on the differences in energy\nbetween Eag, Eag-TMD, and the point variants. As we have made the same assumptions\nfor all calculations, any error inherent in these simplifications is subtracted.\nWe also note that we have stated the binding values are estimates:\n“This directly relates the change in Eag unfolding standard free energy to that of TMD\nbinding at native conditions. The unfolding data of Table 1 allow us to estimate a ΔG°\nbinding\nof approximately -176 kJ/mol for the SciW-TMD complex and a ΔG° of -116 kJ/mol for\nbinding\nthe EagT6-TMD complex.”\n8\n>3. Negative controls, such as the addition of generic TM or of the respective non-cognate\nTMD, are missing for the key experiments.\nResponse: We again emphasize that the TMDs are unstable and insoluble unless they\nare co-expressed already bound and stabilized by the Eag (PMIDs: 33320089, 30177742,\n26456113 and current Figure 6). We cannot add a purified TMD to a purified Eag. These\nfacts are why we pursued denaturation of Eag-TMD complexes to extract dissociation data.\nThe suggested experiments are simply not possible given the system.\nAdditionally, we strongly disagree that suggested experiments with a generic TM or a non-\ncognate TMD are appropriate controls. Comparison of Eag binding to another protein will\nneither prove or disprove effector binding or the measured binding energies of the Eags\nto their known and proven cognate TMDs.\nSupporting our work, there is extensive data and proof showing that Eags bind their\ncognate effector TMD and are specific to their effector class (Ahmad et. al, elife 2020\nFigures 1-5 and associated supplemental figures PMID: 33320089). Several other\npublications show similar findings (PMIDs: 30177742, 26456113, 33323487, 34853317).\nHowever, we have provided extensive controls for Eag-TMD binding and to support our\nresults. First, the binding controls are internal for this study. Namely, Eag-TMD complexes\nrelative to Eag alone. And subsequently, wild-type proteins compared to point-variants.\nFurther controls include TMD-affinity tagged purifications that select for Eag-TMD\ncomplexes, DLS that shows the variants are properly folded, and additional nanoDSF data\n(Table S2, Figure S5 and Figure S6). These all show that we have in fact purified the\nrelevant Eag-TMD complexes of wild-type and point-variants.\nMinor comments:\n>1. Why does Rhs1(1-59) aggregate on SDS-PAGE gels (Fig. S1 and S5), but not in the\nDLS measurements (Fig. S5)? Tse6(1-61) behaves different in this respect.\nResponse: On denaturing SDS-PAGE gels membrane proteins (like the TMDs)\nsometimes do not migrate as expected. This is due to their highly hydrophobic nature and\ninteraction with SDS micelles. Alpha-helical membrane proteins (like the TMDs) often\naggregate or oligomerize on SDS-page gels. The reason that the Tse6-TMD runs as\nexpected could be due to sequence differences relative to the Rhs1-TMD that might\npromote oligomerization. These observations are usually empirical and are difficult to\npredict.\nDetergent binding explains anomalous SDS-PAGE migration of membrane\nproteins Proc Natl Acad Sci U S A. 2009 Feb 10;106(6):1760-5. doi:\n10.1073/pnas.0813167106 PMID: 19181854\nSDS-induced oligomerization of Lys49-phospholipase A2 from snake venom\n30787342\nThe reason no TMD aggregates are observed by DLS is because:\n1) The material is co-purified SciW-Rhs1TMD. The Rhs1-TMD is tightly bound to\nSciW. Our binding data indicates antibody level affinity (at least pico molar). When\nit is bound to SciW, the TMD is packed within the SciW claw and stabilized. There\nare likely no free TMDs above the detection threshold.\n2) If any TMD has separated from SciW it may have precipitated or degraded (PMIDs:\n30177742, 26456113). All DLS samples must be centrifugally filtered.\n> Additional controls, beyond the fact that Rhs1(1-59) still binds to SciW, would provide\nevidence for its native binding, and the respective conclusions.\nResponse: Our past and current work has provided extensive data to prove that SciW\nnatively binds Rhs1(1-59).\n1) SciW and Rhs1 (full-length) binding was shown in our previous eLife paper by pull-\ndown (Figure 1 in eLife PMID: 33320089).\n2) In the previous eLife paper, SciW-Rhs1(1-59) was purified by metal affinity (TMD\nHis-tagged) followed by size exclusion chromatography (SEC). As only the TMD is\nselective for Ni-NTA column binding, we will only purify SciW if SciW is tightly bound\nto the TMD. Additionally, SEC co-elution requires stable complexes.\n3) We have solved a co-crystal structure of SciW bound to Rhs1(1-59) (6XRR). To\ncapture a co-crystal structure between proteins requires specific native binding.\n4) Purifications of all Eag-TMD complexes in the current work also use nickel affinity\nchromatography followed by SEC where only the TMD is His -tagged. This\n6\nspecifically selects for the SciW-Rhs1(1-59) complex. This is a strong control for\nnative binding. (Figure S5)\n5) All biophysical experiments of SciW-Rhs1 (1-59) and point variants show a drastic\nincrease in stability relative to apo-SciW. This includes Tms by nanoDSF and\nDSC, chemical denaturation experiments, and stopped-flow measurements. Note\nthat the nanoDSF and DSC are in a standard physiological buffer. (Figures 1-2,\n45 and Figures S6)\n6) We have shown in this study that the binding affinity of SciW to Rhs1 is at least\npico molar. This is indicative of a specific biologically relevant complex.\n7) We have solved an X-ray crystal structure of a SciW point variant bound to the\nTMD of Rhs1. (Figure 7). Even weakening the binding affinity still allows for the\ncapture of the TMD bound to SciW.\nAdditionally, there are many examples in the literature that show that the adjacently\nencoded Eag and Rhs effector are native binding partners.\n1. Elife. 2020 Dec 15:9:e62816. doi: 10.7554/eLife.62816 PMID: 33320089\n2. Proc Natl Acad Sci U S A 2020 Dec 29;117(52):33540-33548. doi:\n10.1073/pnas.1919350117 PMID: 33323487\n10\n3. Nat Commun. 2021 Dec 1;12(1):6998. doi: 10.1038/s41467-021-27388-0. PMID:\n34853317\n4. PLoS Pathog. 2022 Jan 5;18(1):e1010182. doi: 10.1371/journal.ppat.1010182\nPMID: 34986192\n>2. Is there a difference between chaperones and adapters? If so, it should be explained, if\nnot, this should be mentioned when mentioning adaptors (e.g. Tap adaptors).\nResponse: Yes, there is a difference. We have clarified this in the introduction text:\n“A critical step in cargo effector secretion is the selective recruitment or piloting of effectors\nto the T6SS by accessory proteins that act as chaperones or adaptors. Chaperones\nstabilize effectors in a “secretion-competent” state and affect protein folding, whereas\nadaptors typically only aid in the binding of effectors to T6SS machinery. The terms\nchaperones and adaptors are often used interchangeably in T6SS literature, especially as\nsome chaperones also aid in effector loading (PMID:34092034, 33320089). Despite the\nambiguity, chaperones and adaptors are essential for the proper delivery and toxicity of\nnumerous T6SS effectors.”\n>3. The reason to focus on the first TMD of Tse6 (or a reference showing binding of this\nTMD only to EagT6) should be provided.\nResponse: The reason we focus on TMD1 is that we lack a crystal structure of EagT6-\nTMD2. Additionally, the AlphaFold3 prediction poorly predicts the conformation of the\nTMD. The highest confidence region of the TMD only ranges from 65-80 pLDDT with most\nof the TMD in the 0-50 range (Figure R1). Given this, we cannot make any detailed\nmechanistic conclusions about EagT6 and TMD2.\nFigureR1: AlphaFold3 model of EagT6 and putative TMD2 region residues 167-222\nWe have made this clear to the reader in the discussion:\n11\n“It is important to note that we lack a crystal structure of EagT6-TMD2 and AlphaFold3\nfails to predict a confident model (pLDDT < 65 for TMD2). Our mechanistic dissociation\ndata currently only applies to EagT6-TMD1. However, L66A is sufficient to inhibit secretion\nin vivo strongly supporting our TMD1 dissociation model and importance of TMD1\nbinding.”\n>4. Lines 66/67: This statement does not apply to all effectors, and is therefore misleading\nin its current state.\nResponse: We have changed the language from ‘cargo effectors’ to be ‘for many\neffectors’ to be more general.\n>5. The role of chaperones should be explained more clearly – lines 67-69 are rather\nvague and unspecific.\nResponse: We have clarified the differences between chaperones and adaptors and\nadded that chaperones typically affect protein folding. Additionally, we provide several\nexamples of known chaperones and adaptors for detail.\n>6. Line 123: Rhs should be called Rhs1, as elsewhere in the\nmanuscript. Response: Thank you, these have been corrected.\n>7. The dimer formation of Eag chaperones, one of their defining features, is not taken\ninto consideration in the denaturation experiments (Fig. 1, 2), although likely to influence\nthe results. Likewise, it is not taken into account in the equilibrium calculations described\nin lines 692-698. The authors seem to assume an essentially complete dimerization of the\nchaperones, but no evidence for this is provided.\nResponse: We agree with the reviewer. We need to provide data demonstrating the\noligomeric state of the Eags. In Figure S5 we performed extensive DLS on all Eag samples\nin the study.\nWe have added the wild-type DLS curves and measurements to Figure S1. Additionally,\nwe have added a table listing the measured particle size and polydispersity index (PDI) of\nwild-type and all point variants in Table S2.\nAs demonstrated Eag and Eag-TMDs have a radius of ~2.8 to 3.2 nm, which based on\nthe crystal structure is the correct radius for a dimer. This is also in agreement with the\naccepted sizes of globular protein standards that indicates a MW of near 35 kDa, or a\ndimer.\nBiophys Rev. 2016 Oct 6;8(4):409–427. doi: 10.1007/s12551-016-0218-6 PMID:\n28510011\nsignificantly increase the size of the protein in solution. Additionally, we only observe one\ntype of particle in solution shown by one primary peak and a low PDI. The crystal\nstructures and cryoEM data of Eags show that Eags are dimers when bound to the TMD\nor when unbound, further supporting that Eags are exclusively dimers in solution.\nNat Microbiol. 2018 Oct;3(10):1142-1152. doi: 10.1038/s41564-018-0238-z. PMID:\n30177742\nElife. 2020 Dec 15:9:e62816. doi: 10.7554/eLife.62816 PMID: 33320089\nFinally, we note that K is defined in equation 1 as dimer to monomer:\nunfolded\n𝑐h𝑎𝑝𝑒𝑟𝑜𝑛𝑒 2 𝑚𝑜𝑛𝑜𝑚𝑒𝑟 𝐾\ndimer,folded → ← unfolded unfolding\nThe relationship between K and free energy in equation 4 is then the basis for the\nunfolded\nderivation of all other equations:\n∆𝐺 = −𝑅𝑇𝑙𝑛𝐾\nunfolding unfolding\n>8. How exactly does the difference in denaturation by GdmCl and Urea allow for the\ndetermination of the free energy of Eag-TMD unfolding (statement in lines 163-164)? Is\nurea used afterwards and what for?\nResponse: Figure 2A shows that SciW-TMD is immune to denaturation by urea, so no\nfolding information or free energy can be calculated. Instead, we found that GdmCl is\nsufficient to denature all Eags and complexes in the study. Urea is thus not used again in\nthe study. We have clarified this for the reader.\n>9. Are lines 212-218 new results (as implied by the inclusion in the results part and lack of\nreference)? If they are instead a recapitulation of the results in ref. 27, they should be moved\nfrom the results part to the introduction, or this fact should be at least made clear.\nResponse: Although, they are past results it is important to keep this information in the\nsection for the reader to clarify the selection of point variants. We have clarified that these\nare already known results and further emphasized their origin to the eLife reference.\n>0. The cellular concentration of Tse6 (which the authors, conceivably, assume to be\ndetermined by the binding to its chaperone) does not correlate well with the competitive\nindex. For example, the Q58A mutant, in which the level is similar to the WT, has a strongly\ndecreased fitness compared to the WT, and S41A, which retains expression, is\n13\ndoing worse than I24F, where almost no Tse6 could be detected. Can the authors explain\nthese discrepancies?\nResponse: We thank the reviewer for their attention to these details; however, this\ncomment is an over-interpretation of the data presented in figure 6. The experiment\npresented in figure 6A demonstrates that any of the mutations introduced at the EagT6-\nTse6 binding interface reduce the competitive fitness of the donor over the recipient\nrelative to wild-type EagT6. We cannot make claims about the significance of these\nmutations relative to one another, as this competition assay lacks the statistical power to\ndetect the differences between individual mutants, which are very subtle as the reviewer\npoints out. The Western blot suggests that Q58A is less disruptive to the Eag-Tse6\ncomplex than other mutations such as L66A, which is consistent with our biophysical data.\nWe have clarified the text.\n>1. What is meant by “the Eag chaperone catalyzes completion of the PAAR domain with\nthe prePAAR motif” (lines 463-465, in the description of the model)? The expression is not\ncontained in the listed ref. 27, and the mechanics described by it remain unclear.\nResponse: The Eag positioning the prePAAR motif to complete the PAAR domain fold was\ndiscussed and experimentally tested in the elife paper (Figure 6 and supplement). This\nmechanism is also included in the proposed model Figure 8 (elife PMID: 33320089).\nWe have made this clearer for the reader in the text.\n>2. There is a certain inconsistency and confusion in the gene identifiers given by the\nauthors (lines 547-552 and elsewhere):\n- SL1344_0268, assigned to Salmonella SciW in line 547, encodes instead for TssC,\nthe large subunit of the T6SS contractile sheath\n- SL1344_0286, assigned to SctW elsewhere, seems to encode for an RHS protein\ninstead\n- SL1344_0285, assigned to Rhs1, encodes for a much smaller protein\n- Likewise, the two protein identifiers may have been mixed up for the Pseudomonas\ngenes\nResponse: We thank the reviewer for finding these typos. 0268 is a typo for 0286.\n0285 is SciW and 0286 is Rhs1. They were mistakenly reversed in the text and tables.\nWe have also fixed the mistaken PA identifiers. Furthermore, as much of the T6SS work in\nSalmonella has used the LT2 strain we note what the equivalent LT2 genes are to help\nclarify for the reader.\nReviewer #4 (Remarks to the Author):\n>I co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review\n14\nand to provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\nResponse: We are happy to help support ECR training and we thank the reviewer for\ntheir time.\nReviewer #5 (Remarks to the Author):\n>The manuscript by Van Schepdael et al. explores the mechanism of effector\ntransmembrane domain (TMD) release from Eag chaperones, crucial for Type VI\nsecretion system (T6SS) function. The authors use a combination of biophysical\ntechniques and bacterial assays to demonstrate that small perturbations in Eag-TMD\ninteractions, such as disrupting hydrogen bonds or knob-in-hole packing, can trigger TMD\nrelease. Their work underscores the essential role of conformational changes in Eag\nchaperones for the dissociation of TMDs, a critical step for effector secretion and bacterial\nfitness. Additionally, the X-ray co-crystal structure of the Eag-TMD complex captures an\nintermediate state of TMD release, offering a novel perspective on this process. This study\ncontributes significantly to our understanding of how T6SS chaperones regulate effector\nfolding and secretion. The insights provided in this study are important and should be of\nbroad interest. While, to strengthen the work, the authors may need to address my\nconcerns listed below.\nResponse: We thank the reviewer for their overall positive assessment of our work. We\nhave made both edits to the manuscript and performed additional experimentation as\nrecommended.\nMajor concerns:\n>1.The study investigates the mechanism of Eag chaperone dissociation from effector\nproteins, but the extent to which the in vitro-derived dissociation mechanism reflects the\nactual dissociation process in bacterial cells is questionable. For example, the manuscript\nprimarily examines the impact of different mutations on bacterial secretion but does not\nconfirm whether these mutations specifically affect Eag dissociation, which then\ninfluences bacterial secretion and survival. The authors should include in vitro dissociation\nkinetics experiments and in vivo/in-cell binding assays to demonstrate the role of\nchaperones in regulating effector protein folding and their involvement in T6SS under\nphysiological conditions.\n> The authors should include in vitro dissociation kinetics experiments... under\nphysiological conditions\nResponse: We thank the reviewer, but the manuscript already explicitly addresses the\nreviewer’s concerns. The presented data (Table 1, Figure 2-5, Figures S2-3, and Figures\nS8-S10) are the requested in vitro binding and dissociation kinetics.\nconstants in physiological buffer independent of challenge with denaturant as requested.\nFigure 2G-H, Figure 4E-H and Figure 5 shows how the denaturation data was used to\nobtain the native values for wild-type Eags and the two variants of SciW and EagT6 in an\nin vitro physiologically relevant buffer (no denaturant).\nAgain, the TMDs are insoluble and degrade unless they are bound and stabilized by an\nEag. It is not possible to add purified TMD to Eag in a physiological buffer then perform\nexperiments such as ITC, MST, or BLI. We can only make Eag-TMD complexes for in\nvitro experiments. This limitation is why we denatured the Eag and Eag-TMDs then\ncreated a model to extrapolate to physiological conditions to obtain native in vitro binding\nenergies and dissociation kinetics.\n> vivo/in-cell binding assays to demonstrate the role of chaperones in regulating effector\nprotein folding and their involvement in T6SS under physiological conditions\nResponse: We have already performed experiments to correlate our in vitro binding data\nto in vivo function. In Figure 6 we measure the effect each point variant has on the ability\nto stabilize Tse6 (which contains the TMD) (Figure 6B) and promote secretion of Tse6 to\nkill other cells (Figure 6A). These experiments were performed in the bacteria\nPseudomonas aeruginosa that natively carries the T6SS apparatus, EagT6 and Tse6.\nAdditionally, significant past work has shown that EagT6 binds Tse6 in vivo and stabilizes\nthe protein. We now show that are binding point variants can no longer bind and stabilize\nTse6 as efficiently as wild-type and reduces T6SS mediated killing.\nCell. 2015 Oct 22;163(3):607-19. doi: 10.1016/j.cell.2015.09.027. PMID: 26456113\nNat Microbiol. 2018 Oct;3(10):1142-1152. doi: 10.1038/s41564-018-0238-z. PMID: 30177742\n>2.The SciW-TMD L66A crystal structure presented in the study captures an intermediate\nstate of Eag chaperone dissociation from TMD and supports the proposed dissociation\nmodel. However, this structure was obtained by introducing the L66A mutation to stabilize\nthe dissociation intermediate. It is unclear whether this structure fully reflects the\ndissociation process under physiological conditions in bacterial T6SS, as wild-type\nchaperones in cells may dissociate differently. Could the structure of the wild-type\nchaperone dissociating from TMD under natural conditions be obtained by optimizing\nparameters such as temperature?\nResponse: It is near impossible to predict how a protein will crystallize or what\nconformation will be captured. It is also near impossible to alter crystallization conditions\nto selectively force a protein into a different conformation. Typically, changing\ncrystallization conditions from the known buffer results in no crystals. For example, to\n16\nobtain the structures we have previously published and present here required screening\nhundreds of conditions (including temperature) to even acquire diffracting crystals.\nHowever, to help address the reviewer’s concern of relevance we note that the region of\nthe Eag that changes conformation upon binding the TMD includes residue L66.\nSpecifically, in both SciW and EagT6 residue L66 changes conformation moving over the\nTMD to make the critical knob-in-hole interactions (elife PMID: 33320089). We have\nadded an overlay of bound and unbound Eag structures highlighting the movement of L66\nupon binding in the wild-type proteins to Figure 7. It should be clearer to the reader that\nthis comparison shows how our point variant structure disrupts this essential interaction\nand partially reverses the induced L66 conformational change upon binding. Furthermore,\nL66 is highly important for TMD binding in all our interaction studies. Thus, we believe it\nis a reasonable hypothesis that the structure represents a model of an intermediate of\nTMD dissociation where the L66 knob-in-hole has been disrupted.\n>3.The study attributes the dissociation pathways of Class 1 (SciW-TMD) and Class 2\n(EagT6-TMD) effector proteins to “the different roles of Q58/L66 contacts” and “structural\ndifferences in transition states.” However, the TMD sequences, overall conformations,\nand coupling with VgrG/PAAR differ significantly between the two effector classes,\nsuggesting that other factors may regulate chaperone dissociation. The authors should\ninclude further experiments to confirm that the chaperone-TMD interactions are\nresponsible, rather than other factors playing a key role in regulating T6SS.\nResponse: We agree that other factors regulate chaperone dissociation. This was\noutlined in the discussion of the manuscript. The potential models are that a T6SS ATPase\nin the baseplate catalyzes dissociation or that a baseplate protein binds the Eag and\nforces the TMD out of the Eag coupled to T6SS firing (added in the revision). A model\nschematic has also been included in Figure 8.\nAt no point do we propose that the Eag can self-catalyze the dissociation of the TMD. Our\nbiophysical data provides a model for how the Eag can be manipulated to release the\nTMD. We show what contacts are critical to TMD binding, show that they linked to the\nconformation change that the Eag undergoes when it binds the TMD, and hypothesize\nanother protein and/or force manipulates the Eag to disrupt the interaction of L66\nand Q58 with the TMD. This then results in TMD dissociation.\nFurthermore, attempts to discover Eag binding partners have been performed in\nPseudomonas but only found the EagT6-Tse6-VgrG complex (PMID: 26456113). We\nhave attempted to find additional Eag partners but have been so far unsuccessful. Any\nbinding partner that catalyzes Eag-TMD dissociation is likely to be transient which requires\nchallenging timing and/or advanced cross-linking to capture in a pull-down.\nPotential hits must be cloned and purified to test binding with Eag and Eag-TMD\ncomplexes which typically include many false-positives. Regardless, the results of a\n17\nbinding-partner discovery study when complete would constitute an entire manuscript of\nwork on its own.\n>4.The authors determined the crystal structure of SciW L66A in complex with its substrate,\nwhile the functional assays in Figure 6 were conducted using EagT6. Given the clear\ndifferences between the two proteins, additional cell-based functional validation\nexperiments using the SciW mutant should be included to support the structural findings.\nResponse: We agree with the reviewer that additional assays in Salmonella Typhimurium\nwould enhance the current study. We have performed bacterial competition assays in S.\nTyphimurium with wild-type, a /iclpV deletion (T6SS negative control), and a /isciW deletion\n(Figure S11). Although we are able to show T6SS dependent killing by S. Typhimurium,\nthe /isciW deletion only shows an intermediate phenotype. We note that this is in agreement\nwith past results showing that deletion of sciW has a limited effect on bacterial fitness in a\nmouse model (PMID: 22493086).\nAlthough wild-type does have a significant killing phenotype relative to the /iclpV deletion,\nthe overall phenotype for Salmonella is very low compared to Pseudomonas and is barely\nsignificant from the /isciW mutant (~1.6x ratio wild-type vs ~1.3x ratio /isciW, p-value\n0.035). Given this and that the SciW variants do not drastically disrupt TMD binding it is\nhighly unlikely that any statistically significant differences in function can be detected\nbetween the wild-type and SciW point variants. Compounding this is the high experimental\nvariability due to bile-salt induction (see error bars in Figure S11). As such we have not\nfurther pursued experiments in S. Typhimurium, but we have included the obtained data\nas supplemental Figure S11.\n>5.The dissociation rates between SciW-TMD and EagT6-TMD appear to differ\nsignificantly, suggesting that their dissociation mechanisms may not be identical.\nTherefore, the intermediate dissociation state of SciW-TMD captured in this study may\nnot be universally representative. It would be valuable for the authors to further elaborate\non this point in the discussion, addressing the potential differences in dissociation\nmechanisms and the broader relevance of the observed intermediate.\nResponse: We agree. We have included additional dissociation mechanistic models as\nsuggested. We had originally explicitly stated that we believe the mechanisms are different\nin the discussion, but we have made this clearer and expanded.\nMinor concerns:\n>1.In Figure 6B, the expression levels of Tse protein are analyzed, and the authors\nsuggest that different mutations in Eag affect bacterial competition activity by altering Tse\nprotein stability. However, the figure lacks a quantitative analysis of Tse protein\nexpression. From the data, the Tse protein expression in the L66A group is lower than in\nthe Q102A group, yet the L66A group shows higher competition activity. The authors\n18\nshould provide a quantitative analysis or offer an alternative explanation for this\ndiscrepancy.\nResponse: As discussed in our response to reviewer 3 above, we are not drawing\nconclusions regarding the relative contributions of different mutants to the function of\nEagT6 in vivo - we are simply concluding that each of the mutations we introduced at the\nEagT6-Tse6 binding interface disrupt the competitive fitness of the donor strain against\nthe recipient strain relative to wild-type EagT6. As evidenced by the overlapping error bars\nbetween the L66A and Q102A group, the competition assay lacks the statistical power to\ndetect potential differences between these two mutations, which are very subtle if they\nexist at all.\n>2.There is an extra scale bar in the lower right corner of the eagT6 label in Figure 6B. It\nis recommended to remove this unnecessary scale bar.\nResponse: The figure has been fixed.\n>3.The panel numbers (A, B, C) in different figures are inconsistently sized. It is\nrecommended to adjust them to a uniform size for better presentation.\nResponse: We thank the reviewer for spotting this to be corrected. We have made sure\nthat our figures have all been made in the same professional graphics program with\nconsistent font sizes. They have been designed to export as 2-column or 1-column of the\nappropriate size. We hope that automatic figure resizing by placing figures in Word or\nother programs has not re-created the problem.\n>4.For the Eag variants, the authors only performed unfolding assays to assess their\nresistance to chemical denaturation. I recommend that the revised manuscript include\nNanoDSF measurements to evaluate the stability of the Eag variants both individually and\nin complex.\nResponse: These have been added to the supplemental as Figure S6. The data shows\na similar pattern of Eag and Eag-TMD stability changes as chemical denaturation.\nHowever, unlike chemical denaturation we cannot derive binding energies from the\nnanoDSF which is why we did not originally include them.\n>5.In Figure S6H, the unfolding rate of the bound state of the EagT6 L66A mutant appears\nto be faster than that of the unbound state. This seems counterintuitive. Is the TMD still\nbound to the EagT6 L66A protein under these conditions? Further clarification or\nexperimental validation may be needed to confirm the binding status.\nResponse: We respectfully point out that this analysis is incorrect. Figure S6H is the\nsteady state denaturation (not unfolding rate) using fixed 330/350 ratios. Additionally, the\ndata for both the apo-EagT6 L66A and EagT6-TMD L66A are within experimental error.\nFurthermore, this experiment is repeated in Figure 4D using wavelength maxima\n19\nmeasurements as outline in the text. As clearly shown the apoEagT6L66A and TMD\nbound are essentially the same. The TMD-bound is not lower.\nWe demonstrate clearly in Figure S5 that EagL66A-TMD co-purifies, and thus the TMD is\nbound. The requested nanoDSF also shows a Tm shift for EagL66A-TMD relative to apo-\nEagT6 but interestingly shows the appearance of a shoulder at the apo-Tm (Figure S6H).\nThis indicates that the TMD is likely dissociating from the Eag. We state clearly in the text\nthat we believe the TMD has dissociated prior to our unfolding experiments:\n“This indicates that minimal TMD binding to the EagT6 L66A variant exists under our\nassay conditions. Our fluorescence experiments are performed under dilute conditions\nand in the absence of crowding agents, it is likely that the TMD has dissociated prior to\nthe assay.”\nWe thank the reviewers for their positive response to our revisions and for their time to\nhelp us improve our work.\nREVIEWERS' COMMENTS\n>Reviewer #1 (Remarks to the Author):\nThis is a paper that I earlier reviewed and I have to commend the authors to address all\nmy points. These were addressed by including points of discussion and updated figures\n(e.g fig.8) as well as experimental data whenever feasible (e.g Q106A in SciW). The\nauthors also further detailed the importance of their work the advance the T6SS field.\nResponse: We thank the reviewer for their time and help, and we appreciate that we\nhave addressed all their concerns.\n>Reviewer #3 (Remarks to the Author):\nThe authors have replied to all comments and made several changes, mostly to the\ndiscussion and explanation of the results. This has made different parts of the\nmanuscript clearer and more accessible.\nResponse: We thank the reviewer for their time and positive assessment of our\nrevisions.\nThe main question that remains open to me, related to the previous major point 2, is\nhow the delta-G values and binding energy measurements, calculated from chemical\nand thermal denaturation, relate to the physiological events causing the\ndissociation of the interacting proteins, which are most likely based on mechanical transitions. In other\nwords, can the chemical / thermal denaturation experiments give relevant insights into\nthe events the authors aim to study? If yes, this connection (and possible limitations)\nneeds to be pointed out more clearly.\nResponse: As explained throughout the manuscript and in detail in our previous\nresponse to review, we used chemical denaturation as a tool to obtain the\nphysiologically relevant binding and dissociation energies and rates for the Eag-\nTMD complexes. The thermodynamic and kinetic parameters reported in table 1\n(ΔGº ΔGº kº ) are obtained from extrapolating to 0 M denaturant\nunfolding, binding unfolding\nconcentration, and thus are the native physiological values. None of the reported values\ndepend upon denaturant (ΔGº ΔGº kº ).\nunfolding, binding unfolding\nThe reported ΔGº (energy of TMD binding) is the physiological energy (as if we had\nbinding\nused conventional methods such as ITC, MST, BLI) that the cell must overcome during\nthe secretion process to remove the TMD from the Eag. The ΔGº for each variant\nbinding\nallows to see how much energy that specific interaction contributed to the complex. We\nalso outline in detail the assumptions inherent in our model to calculate the physiological\nenergy of Eag-TMD binding.\nIn the discussion we also highlight that previous studies have shown that T6SS firing\nproduces enough energy to remove the TMD from the Eag. Furthermore, the sum of our\nbiophysical, structural, and in vivo data shows that a mechanism to catalyze TMD release\nand lower the energy of Eag-TMD dissociation is to induce a conformational change in\nthe Eag that moves L66 away from the TMD. These possibilities have been outlined in\nthe discussion.\nHowever, we have made further edits to clarify the significance of our data to the reader.\nIntroduction:\n“Here, we measure the thermodynamic properties and unfolding kinetics of wild-type Eag-\nTMD complexes compared to several site-specific point variant Eag-TMD complexes. Our\ndata yields the binding energies and dissociation kinetics of Eag-TMD complexes, which\nshows that although Eag-TMDs are remarkably stable only a small perturbation of the\nEag is required to promote TMD dissociation.”\nAnd we’ve also clarified in the secretion lines 196-213\n“Specifically, the ΔG of unfolding measured at various denaturant concentrations is used\nfor extrapolation to 0 M denaturant. This yields native denaturant-independent ΔGs of\nfolding and TMD binding energy in a physiologically relevant buffer. Therefore, the data\nreveals the binding energy that must be overcome during secretion to remove the Eag.”\n>In addition, the previous minor point 10 relating to the competition assay shown in\nFigure 6 has not been addressed in my view. This is the only experiment that directly\nlinks the results of the study to physiological effects in the bacteria, and the results\ndo not match.\nResponse: We respectfully disagree with the reviewer in their assessment of the data.\nSpecifically, the statement that ‘the results do not match’. The biophysical data and the in\nvivo data in fact do match and agree.\nNotably, we quantify EagT6 and EagT6-TMD wild-type, the Q58A variant, and the L66A\nvariant. In table 1 and data figures (2-5), show that relative to the wild-type (-116 kJ/mol),\nQ58A reduces the magnitude of the ΔGº (-60 kJ/mol) and L66A reduces the ΔGº\nbinding binding\nfurther to a level that we cannot quantify in our assay (weaker than 1 μM Kd, or worse\nthan -35 kJ/mol).\nWe state this explicitly in the text:\n“This shows that the binding constant K for the L66A substitution has been reduced to\nTMD\nless than 106 M-1 putting an upper limit of ~ -35 kJ/mol upon the ΔGº .”\nbinding\nWhen the biophysical data (binding energies) of wt (-116), Q58A (-60), L66A (worse than\n-35) are compared to the in vivo data in Figure 6, we see exactly the same trend. A\ncompetition ratio of ~6 for wild-type, ~4 for Q58A, and ~2 for L66A. Additionally, all results\nhave statistical significance relative to each other.\nWe again note that L66A does bind the TMD (Supplemental Figures 5-7) but at the dilute\nconcentrations required for the quantitative denaturation assays the complex has\nalready dissociated.\nWe also state this explicitly in the text:\n“Our fluorescence experiments were performed under dilute conditions (1 μM chaperone)\nand in the absence of crowding agents, making it likely that the TMD dissociated prior to\nthe assay.”\n>This is not a detail, but a core result. Downplaying these strong and statistically highly\nsignificant changes as \"very subtle\" and \"details\" does not appear appropriate. The\nauthors claim to have \"clarified the text\" in this respect, but it is not clear where and\nhow. This central point needs to be addressed.\nResponse: To help clarify and provide additional analysis, we have calculated the\nstatistical differences between the mutants in Figure 6a with further interpretation in the\ntext. The statistics support that L66 is more important to TMD binding than Q58, in perfect\nagreement with the biophysical data.\n“Furthermore, the donor strain encoding eagT6(Q58A) displays intermediate fitness\nagainst the recipient strain. This is consistent with our biophysical finding that Q58\ncontributes less significantly than L66 to the stability of the EagT6-TMD complex. The\nagreement between these genetic results and our biophysical data suggest that the\ndeterminants of complex stability identified in vitro closely reflect the function of the\nEagT6-TMD complex in P. aeruginosa.”\nAlso edited:\n“Furthermore, the relative abundance of Tse6 in the EagT6 Q58A and L66A variant\nbackgrounds correlates with the contribution of those residues to competitive fitness and\nin vitro TMD binding energy (Figures 4-6 and Table 1). Notably, the relative abundance of\nTse6 is high for EagT6 Q58A as compared to other the mutants and wild-type (Figure 6B).\nSupporting this, EagT6 Q58A has a ΔGº of -60 kJ/mol which is still indicative of a\nbinding\nstable complex. Regardless, in agreement with our biophysical data the loss of TMD\nbinding affinity by Q58A significantly decreases Tse6 secretion (Figure 6A).”\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review\nand to provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\nResponse: We again thank the reviewer for their time and help to improve our study.\nReviewer #5 (Remarks to the Author):\nI don't have further question.\nResponse: We thank the reviewer for their time and help to improve our work. We are\nglad we were able to address all concerns."}]

Decision letter