1. Microbiology and Infectious Disease
  2. Structural Biology and Molecular Biophysics

Structural intermediates observed only in intact Escherichia coli indicate a mechanism for TonB-dependent transport

  1. Thushani D Nilaweera
  2. David A Nyenhuis
  3. David S Cafiso  Is a corresponding author
  1. Department of Chemistry and Center for Membrane Biology, University of Virginia, United States
https://doi.org/10.7554/eLife.68548
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  1. Thushani D Nilaweera
  2. David A Nyenhuis
  3. David S Cafiso
(2021)
Structural intermediates observed only in intact Escherichia coli indicate a mechanism for TonB-dependent transport
eLife 10:e68548.
https://doi.org/10.7554/eLife.68548
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Abstract

Outer membrane TonB-dependent transporters facilitate the uptake of trace nutrients and carbohydrates in Gram-negative bacteria and are essential for pathogenic bacteria and the health of the microbiome. Despite this, their mechanism of transport is still unknown. Here, pulse electron paramagnetic resonance (EPR) measurements were made in intact cells on the Escherichia coli vitamin B12 transporter, BtuB. Substrate binding was found to alter the C-terminal region of the core and shift an extracellular substrate binding loop 2 nm toward the periplasm; moreover, this structural transition is regulated by an ionic lock that is broken upon binding of the inner membrane protein TonB. Significantly, this structural transition is not observed when BtuB is reconstituted into phospholipid bilayers. These measurements suggest an alternative to existing models of transport, and they demonstrate the importance of studying outer membrane proteins in their native environment.

eLife digest

Bacteria must obtain nutrients from their surrounding environment in order to survive. In Gram-negative bacteria, proteins in the outer membrane surrounding the cell actively transport carbohydrates and trace nutrients like iron into the cell’s interior. Although the structures of many of these transport proteins have been determined, the mechanism they use to move molecules across the membrane is poorly understood.

To better understand this process, Nilaweera, Nyenhuis and Cafiso examined the structure of BtuB, a transport protein found in the outer membrane of Escherichia coli that is responsible for absorbing vitamin B12. Previous experiments analyzing the structure of BtuB, and other similar transporters, have been carried out on purified proteins that were extracted from the outer membrane. However, these isolated proteins fail to replicate the transport activity observed in bacterial cells. Nilaweera, Nyenhuis and Cafiso therefore wanted to see how the structure of BtuB changes when it is still enclosed in the membrane of E. coli.

This revealed that BtuB undergoes large structural changes when it binds to vitamin B12, suggesting that this is an important part of the transport process. However, when purified BtuB was placed into an artificial membrane, these structural changes did not occur. This indicates that the cellular environment in the bacteria is needed for BtuB to carry out its transport role, and explains why previous experiments using purified proteins struggled to see this structural shift.

This work highlights the importance of studying bacterial membrane proteins in their native cell environment. BtuB and similar transporters represent a large family of proteins unique to Gram-negative bacteria that have an impact on human health. Since these proteins are structurally alike, the results of this study may help resolve the transport mechanisms of other proteins, ultimately leading to new ways to control bacterial growth.

Introduction

The passive permeation of low molecular weight solutes across the outer membrane (OM) of Gram-negative bacteria is typically facilitated by porins. However, many higher molecular weight solutes and trace nutrients, including carbohydrates, iron siderophores, cobalamin, copper, and nickel, are bound and transported across the OM by a family of active transporters that are TonB-dependent (Bolam and van den Berg, 2018; Noinaj et al., 2010). These TonB-dependent transporters (TBDTs) derive energy from the bacterial inner membrane by interacting with TonB, a transperiplasmic protein that interacts with the inner membrane proteins ExbB and ExbD (Celia et al., 2016; Maki-Yonekura et al., 2018). This family of transporters has two distinct domains: a β-barrel formed from 22 anti-parallel β-strands, and a core or hatch domain that fills the interior of the barrel. The β-strands on the extracellular surface of the protein barrel are often connected by long loops, while short turns join the strands on the periplasmic interface (see Figure 1B). TonB interacts with the transporter through a conserved motif on the N-terminal side of the core termed the Ton box (Pawelek et al., 2006; Shultis et al., 2006).

Figure 1
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Electron paramagnetic resonance (EPR) spectra obtained in vivo from spin labeled core sites on extracellular face of the BtuB.

The spin labeled side chain (A) R1 was attached to sites on the extracelluar core of BtuB shown in (B). BtuB is shown in both extracellular (top) and side views (PDB ID: 1NQH), with the core in yellow and barrel in light blue. The Cα atom on site 188 on the second extracellular loop is shown, which is used as a reference point for measurements to the core. Labeled Cα sites in the core are shown in magenta, along with substrate binding loops SB1, SB2, and SB3. In (C) EPR spectra are shown in the absence (apo state in blue) and presence of substrate (vitamin B12 bound state in red). No change in the spectrum is observed at sites 65 and 90 upon the addition of substrate. The spectra are characterized by well-defined hyperfine extrema ((i) and location of arrow), where the difference between these points is approximately 69 Gauss. This value is close to 2Azz, twice the value of the hyperfine tensor, and indicates that the label is immobilized on the ns time scale. Spectra are the sum of 10–100 Gauss scans.

The mechanism of transport in TBDTs is presently poorly understood; however, the large size of most substrates and the absence of any obvious pathway for substrate permeation (Faraldo-Gómez et al., 2003) have led to proposals that transport is mediated by a significant conformational event that involves a partial rearrangement or full removal of the core domain from the surrounding barrel (Chimento et al., 2005). Although many high-resolution structures are available for TBDTs, there is no direct evidence for a major structural change within the core of TBDTs that might indicate a transport mechanism. In the Escherichia coli vitamin B12 (cobalamin) transporter, BtuB, EPR spectroscopy shows that substrate binding unfolds the Ton box at the N-terminus and extends it into the periplasm, an allosteric event that may facilitate the binding of TonB to BtuB (Kim et al., 2007; Xu et al., 2006); however, no other significant structural changes have been observed in the core.

High-resolution crystal structures have been obtained for a C-terminal fragment of TonB in complex with BtuB, the ferrichrome transporter FhuA, and the ferrioxamine B transporter FoxA (Pawelek et al., 2006; Shultis et al., 2006; Josts et al., 2019). When TonB binds, the Ton box extends from the core and interacts with the β-sheets of TonB in an edge-to-edge manner. Except for the Ton box, the remainder of the core remains folded and is essentially unchanged. Because TonB binding does not alter the core of BtuB in the BtuB-TonB structure, it has been proposed that TonB alters the core by exerting a mechanical force on the transporter, and current models for transport favor a mechanism where TonB acts by pulling the Ton box thereby unfolding the core (Gumbart et al., 2007; Hickman et al., 2017; Sverzhinsky et al., 2015). Models involving a rotation of TonB have also been proposed (Klebba, 2016); however, in FhuA there are four to five unstructured residues between the Ton box and core when TonB is bound, making the transfer of torque from TonB to the core unlikely (Sarver et al., 2018). Pulling models have been explored using steered molecular dynamics (MD) (Gumbart et al., 2007) as well as single-molecule AFM (atomic force microscopy) pulling experiments (Hickman et al., 2017), and these studies indicate that an N-terminal region of the core (up to residue 73) is preferentially unfolded to permit the movement of vitamin B12 into the periplasm. This work concludes that the C-terminal region of the core is static and does not unfold during transport, a result that is consistent with denaturation experiments on BtuB (Flores Jiménez and Cafiso, 2012).

An important caveat to almost all the structural work on BtuB is that it has been carried out on purified or partially purified protein where the native OM environment is no longer present. Since transport in this family of transporters has never been reconstituted, it has never been established that the isolated, purified, and membrane reconstituted BtuB is capable of transport. Recently, we developed an approach to attach spin labels to either extracellular or periplasmic sites on BtuB in intact cells, thereby permitting EPR measurements to be made under conditions where the protein is known to be functional (Joseph et al., 2019; Nilaweera et al., 2019). Preliminary measurements made on BtuB indicate that it behaves differently in the intact cell than it does in a purified reconstituted phospholipid system. For example, a substrate-dependent change in the core domain of BtuB involving substrate binding loop 3 (SB3) is observed in situ but is not seen in a detergent-treated OM preparation (Nilaweera et al., 2019). Moreover, the extracellular loops of BtuB are also highly constrained in the intact cell, and substrate-induced structural changes and structural heterogeneity that is observed for BtuB in proteoliposomes are not observed for BtuB in situ (Nyenhuis et al., 2020a).

In the present work, we perform double electron-electron resonance (DEER) on BtuB in intact cells to determine whether structural changes take place in the core domain that are associated with substrate binding and transport. Upon substrate binding, a movement of the core is observed involving sites 90 and 93 in SB3. No other movements in the core are detected. When the ionic lock is broken between site R14 on the C-terminal side of the Ton box and site D316 in the barrel, long-distance components appear upon substrate binding, indicating that SB3 can assume a state where it has moved into the BtuB barrel toward the periplasmic side of the protein. Under these same conditions, other sites on the N-terminal side of the core remain static. Since this ionic interaction would normally be broken upon TonB binding, this structural transition is likely to take place during transport. This result suggests that transport may involve allosteric changes in the C-terminal side of the core upon TonB binding. Remarkably, these substrate-induced structural changes are not observed for purified, membrane reconstituted BtuB, which may be due to the absence of lipopolysaccharide (LPS) or other periplasmic components in the reconstituted system. The importance of the native OM environment provides an explanation for why this structural transition has not been previously observed.

Results

Multiple sites in the core region of BtuB may be spin labeled in intact E. coli

To investigate movements that might occur in the BtuB core region in situ, pairs of spin labels were placed into the extracellular region of BtuB, with one label located at an outer loop site that is known to be relatively fixed in the cell environment and a second label located at a site in the BtuB core. Previous work has demonstrated that several sites on the extracellular surface of BtuB may be spin labeled in vivo using site-directed cysteines and a standard methanethiosulfonate reagent to produce the side chain R1 (Figure 1A). These included multiple sites on the extracellular loops of BtuB (Nilaweera et al., 2019; Nyenhuis et al., 2020a; Joseph et al., 2016) as well as two sites in the core region (Nilaweera et al., 2019).

Recent work has also shown that the efficient incorporation of pairs of spin labels to make distance measurements using DEER required the use of a strain deficient in the disulfide bond formation (Dsb) chaperone system (Nilaweera et al., 2019). We tested several additional single cysteine mutants in BtuB using a DsbA- strain to determine whether spin labeling of additional sites in the core was possible. Shown in Figure 1c are spectra from site 90, which was previously labeled, as wells as four additional sites in the core region. Sites 63 and 65 lie in substrate binding loop 1 (SB1), site 72 lies in substrate binding loop 2 (SB2), and sites 90 and 93 are positioned in SB3. These spectra arise from label having more than one motional component but are dominated by a broad feature that is characteristic of a population of label with hindered motion on the ns time scale, consistent with the confined environment in the extracellular region of the core. For sites 63, 72, and 93, the addition of vitamin B12 alters the spectra and increases the population of the immobile component indicating that incorporation of the label at these sites has not prevented the binding of substrate. No significant changes with substrate are seen for sites 65 and 90. At site 90, substrate does bind (see below) and the lack of a change in the EPR spectrum may reflect the fact that in the apo state the label is already highly immobile. Site 65 is also highly immobile, but we cannot exclude the possibly that incorporation of R1 at this site has blocked the binding of vitamin B12.

The apex of the SB3 loop in the BtuB core undergoes a substrate-dependent conformational change

For distance measurements using pulse EPR, each set of spin pairs included a label at position 188 on the 3/4 extracellular loop (the second loop connecting β-strands 3 and 4). This site was chosen as a reference point because previous work in whole cells demonstrated that this loop assumed a well-defined position and exhibited minimal or no movement upon substrate addition (Nyenhuis et al., 2020a).

All data were analyzed using LongDistances. Positions of both components were held constant throughout and were set to the average of an initial round of fits where distance was varied. Width and area for the two components were allowed to vary freely. Error ranges were taken from the output of the fitting routine.

Shown in Figure 2 are the results for measurements on the V90R1-T188R1 spin pair in cells. Preliminary results from this pair were presented in a previous study demonstrating the use of disulfide chaperone mutants to achieve double labeling of BtuB in whole cells (Nilaweera et al., 2019). The background corrected DEER data and resulting distance distributions are shown in Figure 2b,c. Both the apo (blue) and vitamin B12 bound (red) distributions yield two main intramolecular peaks at 2.4 and 3.2 nm, with a substrate-dependent shift observed toward the shorter component. Predicted distance distributions were generated from the apo and vitamin B12 bound in surfo crystal structures of BtuB (PDB IDs: 1NQG and 1NQH) using the program MMM (Jeschke, 2018). The distributions generated from these structures also show a shift toward a shorter distance in the substrate bound state, which is due to the unfolding of a helical turn in the SB3 loop (Chimento et al., 2003). However, the magnitude of the predicted shift is smaller than that observed by DEER. Because the position of site 188 in the 3/4 extracellular loop of BtuB is not altered with substrate addition in situ (Nyenhuis et al., 2020a), this structural change must involve a movement of the SB3 apex or a change in rotamers assumed by R1 at position 90.

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Substrate-dependent shifts are detected at the apex of substrate binding loop 3 (SB3) in whole cells.

A side view of BtuB (A) with the locations of site 90 in SB3 of the core domain (yellow) and site 188 near the apex of the 3/4 extracellular loop (light blue). (B) Background corrected double electron-electron resonance (DEER) data for the apo (blue) and substrate bound states (red) of the V90R1-T188R1 spin pair, where the red traces represent the fits to the data. The resulting distance distributions are shown in (C) where the histograms represent predicted distance ranges obtained from the in surfo crystal structures 1NQG (blue) and 1NQH (red) using the software package MMM (Jeschke, 2018). (D) This structural change titrates between two states (labeled 1 and 2) with the addition of vitamin B12. The conversion between states saturates at concentrations above 60 µM. For the distributions shown in (C), data were analyzed using LongDistances v932 using the model-free fitting mode, whereas for distributions in (D), data were fit to a two-Gaussian model where position, width, and amplitude were variables in the fit. Errors in the fit to the distributions in (D) are shown in Supplementary file 1.

We also titrated this structural change by measuring the distance distribution with increasing concentrations of the vitamin B12 substrate, where the result is shown in Figure 2D. For this analysis data were processed using a model-based approach with two-Gaussian components where the position, width, and amplitude were varied. As seen in Figure 2D, there is a strong progressive response to increases in substrate until saturation is reached in the range of 30–60 μM vitamin B12. This titration likely reflects substrate loading. Because substrate concentrations greatly exceed the affinity of vitamin B12 to BtuB (Bradbeer et al., 1986), the saturation point likely reflects the concentration of BtuB in our sample and is roughly consistent with the spin concentrations expected from the EPR signal intensity.

Substrate-dependent shifts in the extracellular face of the hatch domain are localized to the SB3 loop

To determine whether the structural change observed in the apex of SB3 is limited to this site or part of a broader conformational change across the core domain, we tested additional spin pairs on the extracellular face of the protein using the core sites shown in Figure 1B. The spin pairs examined are shown in Figure 3A,B and include site 93, which also lies in SB3, as well as sites 63 and 65 in SB1 and site 72 in SB2. The distance distributions that result from these spin pairs are shown in Figure 3C, along with the V90R1-T188R1 spin pair. It should be noted that the distance distributions in Figure 3, as well as subsequent figures, have been truncated at 5 nm. As seen in the raw data (Figure 2—figure supplement 1 and Figure 3—figure supplement 1), a longer distance component is apparent in some of these dipolar evolutions. This is due to the presence of BtuB-BtuB interactions in the OM that leads to a 6.5 nm distance component (Nyenhuis et al., 2020b).

Figure 3 with 4 supplements see all
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Substrate-dependent conformational shifts are limited to substrate binding loop 3 (SB3) and are altered by mutation of the R14-D316 ionic lock.

(A) Top view of BtuB (PDB ID: 1NQH) showing the locations of the hatch sites relative to the reference site, 188, in the 3/4 extracellular loop. In (B) the location of the R14-D316 ionic interaction between the core and the barrel is also shown along with the Ton box (red). (C) Distance distributions obtained for hatch: barrel pairs in the apo state (blue) and with substrate (red). (D) Distance distributions obtained for hatch: barrel pairs in the apo state (blue) and with substrate (red) in the presence of the R14A mutation. Both V90R1-T188R1 and S93R1-T188R1 spin pairs show additional distances at 3.8 and 4.5 nm in the presence of R14A. Data were analyzed using LongDistances v932 and the model-free fitting regime. The shaded error bands in (C) and (D) represent variation due to background noise, start time, dimensionality, and regularization. Histograms are predicted distances generated from the in surfo crystal structures PDB ID: 1NQG (light blue) and PDB ID: 1NQH (light red) using the software package MMM (Jeschke, 2018).

As seen in Figure 3C, distance distributions for sites located outside SB3 show little evidence for any structural change upon the addition of substrate, with spin pairs involving sites 63, 65, and 72 having nearly identical distributions for both apo and vitamin B12 bound conditions. This lack of a shift in these spin pairs is consistent with work showing that site 188 does not show a substrate-dependent shift in its position in situ (Nyenhuis et al., 2020a). In the SB3 loop, however, site 93 at the edge of the loop shows a substrate-dependent change in position along with site 90 at the loop apex, although the change is much larger for site 90 (about 8 Å) than for site 93 (about 4 Å) and in an opposite direction. The structural changes measured by DEER are qualitatively consistent with the predictions from the crystal structures, which show a loss in helical structure and change in position of the loop. For the sites examined here, substrate-dependent changes in the core appear to be confined to the region of the SB3 loop. Interestingly, as we demonstrate below, this substrate-dependent change does not occur when the protein is removed from its native environment.

Breaking an ionic lock between the BtuB core and barrel triggers a large substrate-dependent change in SB3

As indicated above, BtuB is expected to bind to both substrate and TonB during transport, which will break the ionic lock between R14 in the core and D316 in the barrel (Shultis et al., 2006). However, under the conditions of our experiment, BtuB is in large excess relative to TonB, perhaps by a factor of 10–20 or more. As a result, only a small portion of the BtuB would be bound to TonB at any time during our distance measurement.

To determine whether there might be a connection between the structure of the core and this internal ionic lock, we examined the effect of disrupting the R14-D316 interaction on the core by introducing the R14A mutation into the existing pairs of labels between site 188 in the 3/4 extracellular loop and the core. Distance measurements for the apo and vitamin B12 bound states made in the presence of this mutation are shown in Figure 3D. For distance distributions involving sites 63, 65, and 72 in SB1 and SB2, the core remains largely unchanged in response to substrate and unchanged by the R14A mutation. However, distance distributions involving sites 90 and 93 in SB3 are altered by breaking the D316/R14 ionic lock.

For distances measured to sites 90 and 93, the R14A mutation has two main effects. First, for the V90R1-T188R1 pair, the substrate-dependent conversion between the 2.4 and 3.2 nm distance components is absent, and the shorter distance now dominates in both apo and vitamin B12 bound conditions. Second, for both the V90R1-T188R1 and S93R1-T188R1 pairs, additional distance components are observed in the presence of substrate centered at 3.8 and 4.5 nm. These long components result in a distribution that is substantially broader than that predicted by the crystal structures, and they indicate the formation of a novel conformation of the SB3 loop and an altered substrate binding mode.

The longer distance components that appear for the V90R1-T188R1 and S93R1-T188R1 spin pairs represent a substantial movement of the SB3 loop toward the periplasmic side of BtuB. A movement of SB3 toward the extracellular surface is highly unlikely, in part because movement in this direction would require a major unfolding of the core that we do not observe. For measurements to SB3, there were relatively few positions that were both accessible and within the range of the pulse EPR measurement, but we made measurements to site 90 from site 237, which is located near the apex of the 5/6 extracellular loop. In the apo and vitamin B12 bound states, the predicted distances from this site are shorter than 2 nm and are not within a range that can accurately be measured by DEER. The results are shown in Figure 3—figure supplement 2. In the absence of the R14A mutation, no clear substrate-dependent shift in the position of SB3 is observed, which is likely due to the short distance involved. However, in the presence of the R14A mutation, a new distance component appears with substrate addition around 3 nm that is beyond the distance range predicted by the crystal structure. This is shorter than the 4.5 nm observed from position 188, which likely reflects differences in the side chain direction and the relative positions of the 3/4 and 5/6 loops.

These ionic lock mutations generate structural states that are not seen in the wild-type protein. However, disrupting the ionic lock does not appear to abolish transport. The BtuB R14A mutant is functional in transport as determined by a growth assay (Figure 3—figure supplement 3). This is consistent with earlier work where a mutant containing a dipeptide insertion into one of the barrel strands in BtuB, which should have disrupted the D316-R14A ionic lock, was shown to support transport of vitamin B12 (Lathrop et al., 1995).

It should be noted that we examined the stability of the substrate-induced conformation of SB3 for times as long as 60 min before freezing and preparing the cells for DEER. The data are shown in Figure 3—figure supplement 4, and indicate that the conformations are stable over time, indicating that the label is stable and not being reduced, and that conversion of the transporter back to the apo state does not occur under the conditions of this experiment.

Substrate-dependent changes in the SB3 loop require a native environment

Two conformations are observed by crystallography for SB3. In the apo structure (PDB ID: 1NQG), SB3 has a single helical turn and shorter conformation that we speculate may be associated with the distance observed by EPR at 3.2 nm for the V90R1-T188R1 spin pair (labeled 1 in Figure 2D). In the vitamin B12 bound structure (PDB ID: 1NQH), SB3 is more extended, and this state may be associated with the distance at 2.4 nm (labeled 2 in Figure 2D). Computational work suggests that the extended state of SB3 requires the interaction of BtuB with LPS, whereas the shorter helical state of SB3 occurs in the presence of phospholipid (Balusek and Gumbart, 2016), suggesting that environment, specifically LPS, may be important in controlling the configuration of SB3.

To test for an environmental effect on SB3, we reconstituted four spin pairs of BtuB into 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC) proteoliposomes. These included the V90R1-T188R1 and S93R1-T188R1 spin pairs both in the absence and presence of the R14A mutation. The dipolar evolution data and distance distributions from DEER measurements on these reconstituted BtuB samples are shown in Figure 4 (the raw time domain data are presented in Figure 4—figure supplement 1). In the absence of the R14A mutation, the substrate-dependent conversion to the shorter distance that was seen for the V90R1-T188R1 spin pair in whole cells (Figure 2C and Figure 3C) is now much more limited, with only a minor shoulder appearing around 2.5 nm. For the S93R1-T188R1 spin pair, the change seen in Figure 3C with substrate is largely absent. The 0.4 nm substrate-dependent shift to a longer distance is absent, but the small 2.2 nm distance component is still present.

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The substrate-induced changes in substrate binding loop 3 (SB3) are altered or absent in proteoliposomes.

Background corrected double electron-electron resonance (DEER) signals (left) and distance distributions (right) for the V90R1-T188R1 and S93R1-T188R1 pairs involving SB3 in the absence and presence of the R14A mutation, where the labeled BtuB have been reconstituted into 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC) vesicles. Data were analyzed using LongDistances v932 and the model-free fitting mode. The shaded error bands represent variation due to background noise, start time, dimensionality, and regularization. Histograms are predicted distances generated from the in surfo crystal structures 1NQG (blue) and 1NQH (pink) using the software package MMM (Jeschke, 2018).

The behavior of SB3 in the presence of the R14A mutation is also altered in the phospholipid reconstituted system. Rather than increasing the population of the shorter distance component, adding the R14A mutation to the V90R1-T188R1 pair in the reconstituted system results in a single short distance, where the resulting distribution aligns almost perfectly with the predicted distribution from the 1NQH structure. For the S93R1-T188R1 pair, R14A causes a small increase in the short distance component at 2.2 nm. But significantly, for neither spin pair are the longer substrate-induced shifts that were seen in Figure 3D observed in the reconstituted system. Thus, when removed from the native OM environment, the structure of SB3 is altered and the large substrate-induced movement of SB3 toward the periplasmic surface in the presence of the R14A mutation is no longer seen.

It should be noted that in our initial work on the V90R1-T188R1 spin pair, we failed to observe a substrate-dependent conformational change in SB3 using an isolated OM preparation where the preparation includes a sarkosyl treatment (Nilaweera et al., 2019). This suggests that this detergent treatment of the OM to remove inner membrane components is sufficient to alter the behavior of BtuB. These observations provide an explanation for why these changes in the conformation of SB3 have not been previously observed.

Mutating either R14, D316, or both alters SB3 conformations and populates a state where SB3 is moved toward the periplasmic surface

In earlier work, we demonstrated that breaking the ionic lock between D316 and R14 altered a conformational equilibrium in the Ton box and promoted its unfolded state (Lukasik et al., 2007). In this work, the effect of the R14A mutation on the Ton box equilibrium was comparable to that of a D316A mutation, but slightly enhanced for the dual R14A/D316A mutant.

Figure 5 shows a result of mutating one or both of R14 and D316 on the substrate-dependent changes in SB3 as measured using the V90R1-T188R1 spin pair (the time domain data are provided in Figure 5—figure supplement 1). In the apo state, the distributions for both the R14A and D316A mutants are similar, with peaks falling in the same positions, although the D316A mutant yields more residual area under the 3.2 nm peak than is observed for R14A. In the presence of substrate, both show significant peaks around 3.8 and 4.5 nm, and a significant peak at 2.4 nm in both the apo and vitamin B12 bound samples. The double R14A-D316A mutation yields a distance distribution that is more perturbed than either single mutant, with a single broad peak centered around 2.4 nm in the apo state and an increase in the longer distance components in the substrate bound state. Thus, disrupting the D316-R14 ionic lock by mutating one or both residues has a dramatic effect on the conformation of the apex of SB3 and populates a state where this binding loop has moved a significant distance toward the periplasmic interface. The data indicate that this ionic interaction plays a role in mediating allosteric changes within the BtuB core, affecting not only the Ton box equilibrium but the substrate binding loop SB3 on the extracellular surface.

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R14A, D316A, or D316A-R14A have similar effects on the conformation of SB3.

In (A) is shown the structure of BtuB highlighting the positions of the R14 and D316 side chains and the location of the Ton box (from PDB ID: 1NQG). Background corrected double electron-electron resonance (DEER) data are shown in (B) and distance distributions in (C) for the V90R1-T188R1 spin pair in whole cells in the presence of the R14A, D316A, or the combined R14A-D316A mutants. Data are shown for both apo (blue) and vitamin B12 bound (red) states. Lines through the DEER data represent the fits for the distributions shown on the right. These data were analyzed using LongDistances v932 and the model-free fitting mode. The shaded error bands in (C) represent variation due to background noise, start time, dimensionality, and regularization. Histograms represent predicted distances generated from the in surfo crystal structures for PDB ID: 1NQG (light blue) and PDB ID: 1NQH (pink) using the software package MMM (Jeschke, 2018).

Discussion

The pulse EPR measurements made here in intact E. coli indicate that SB3, which includes residues 82–96 in the core of BtuB, undergoes a substrate-induced structural change. When the ionic interaction between R14 in the core and D316 in the barrel is broken, an alternate and more dramatic structural change in SB3 occurs upon substrate binding, where SB3 is displaced approximately 2 nm toward the periplasmic surface of BtuB. Remarkably, these structural changes do not occur when the protein is removed from the native cell environment and reconstituted into a phospholipid bilayer, indicating that features in the intact cell environment modulate the energetics of the conformational states in BtuB.

Earlier experimental work provides evidence for an allosteric coupling between the substrate binding site, the R14-D316 ion pair, and the Ton box in BtuB. Measurements made by EPR in isolated OM or reconstituted phospholipid membranes demonstrated that substrate binding partially unfolded the Ton box (Xu et al., 2006; Merianos et al., 2000), and shifted the energy of the folded and unfolded Ton box states by about 2 kcal/mol (Freed et al., 2010). When the ionic interaction between R14 in the core and D316 in the barrel was broken, the Ton box was also observed to unfold and the coupling between substrate binding and the Ton box was broken (Lukasik et al., 2007). In addition, a connection between the Ton box and SB3 was seen by scintillation proximity assays where both the Ton box and SB3 were found to be necessary for a TonB-dependent retention of vitamin B12 (Mills et al., 2016).

The connection between these sites in BtuB is also suggested by computational studies. When LPS is included in MD simulations, the interaction between R14 and D316 is weakened and the energy to unfold the Ton box reduced (Balusek and Gumbart, 2016). The inclusion of LPS also alters the state of SB3. In a symmetric phospholipid bilayer, SB3 assumes the more helical form, whereas in an asymmetric membrane containing LPS, SB3 assumes an extended form. These simulations are consistent with the results presented here, except that fully populating the extended form of SB3 in our whole cell measurement (state 2 in Figure 2d) requires substrate binding. Although the role of periplasmic components such as the peptidoglycan cannot be ruled out, the computational results indicate that interactions made by LPS with the extracellular loops of BtuB may alter conformational equilibria in the protein and provide an explanation for the differences in the behavior of BtuB when EPR measurements are made in cells versus reconstituted phospholipid bilayers. It should be noted that the interconversion between helical and extended forms for SB3 was also absent or diminished when the V90R1-T188R1 spin pair was examined by EPR in an isolated OM preparation (Nilaweera et al., 2019); as a result, the procedure to produce this OM preparation, which includes the use of sarkosyl, is apparently sufficient to modify the behavior of the protein.

An unexpected observation made here is that mutation of the R14-D316 ion pair alters the structure of the SB3 loop (Figure 3D and Figure 5). In the apo state of BtuB, this mutation enhances the extended form of SB3 (state 2 in Figure 2B), and upon substrate binding an alternate conformational state is generated where sites 90 and 93 are extended as much as 4.5 nm from site 188 on the 3/4 extracellular loop. Among the core sites examined, this more dramatic structural change involves only SB3 as labels in the first and second substrate binding loops (SB1 and SB2) at sites 63, 65, and 72 do not exhibit any significant structural changes. Such a large structural change involving SB3 has not been previously observed, and it appears to be localized to the C-terminal side of the core.

A high-resolution model obtained by crystallography for a fragment of TonB in complex with BtuB (Shultis et al., 2006) shows TonB interacting with the Ton box in an edge-to-edge manner (Figure 6A). Since the core is largely unaltered by TonB binding, models for transport have focused on the idea that TonB alters the core structure by exerting a mechanical force on the Ton box. In particular, TonB has been proposed to function by pulling on the Ton box, which then results in an unfolding of the N-terminal region of the core (Hickman et al., 2017). Single-molecule pulling experiments (Hickman et al., 2017) as well as steered MD simulations (Gumbart et al., 2007) suggest that pulling the Ton box will extract the N-terminal side of the core and eventually open a pore sufficient to allow substrate to pass. One difficulty with this model is that both experimental and computational approaches indicate that the extraction of an extended polypeptide chain longer than the width of the periplasm is required to open this pore.

Figure 6
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Conformational shifts in substrate binding loop 3 (SB3) with release of R14-D316 ionic lock.

(A) View of BtuB (PDB ID: 2GSK) in complex with the C-terminal domain of TonB (purple) showing the core (yellow) and barrel (light blue) with the substrate binding loop SB3 (magenta), and the Ton box (red), where the R14-D316 ion pair has been broken. (B) An Xplor-NIH simulation showing positions of the Cβ carbon (magenta mesh) on site 90 in SB3 for 300 structures that are consistent with the distributions obtained by double electron-electron resonance (DEER) for V90R1-T188R1-R14A and V90R1-S237R1-R14A in the presence of vitamin B12 (see Materials and methods). TonB binding extracts the Ton box from the core to create an edge-to-edge interaction with TonB, thereby breaking the R14-D316 ionic interaction. Breaking this interaction promotes the movement of SB3 toward the periplasmic interface of the transporter (red arrow) and may facilitate passage of vitamin B12.

We do not observe any significant structural changes in the N-terminal side of the core in cells (sites 63, 65, and 72), suggesting that the N-terminus may not move during transport. Rather, a significant substrate-induced structural change is found to take place in SB3 on the C-terminal side of the core when the D316-R14 ionic lock is broken. At present, we have limited restraints to generate a model and we do not know the positions of many segments in the core, but a movement in SB3 that satisfies the EPR restraints is shown in Figure 6B. In this model, substrate binding moves SB3 into the barrel and toward the periplasm. We do not presently know whether the movement of SB3 is accompanied by the movement of substrate; however, it is interesting to note that the C-terminal side of the core has a lower side chain density than the N-terminal side, suggesting that there is more space for structural rearrangements within this region.

Presently, the precise sequence of steps that take place during transport are not known; however, both substrate and TonB are expected to be bound to BtuB at some point during transport, which should break the ionic lock between D316 and R14 (Figure 6A). This suggests that the large substrate-induced structural change observed here on the C-terminal side of the core (Figure 6B) will occur during transport. The energy to promote this structural change and disrupt core-barrel electrostatic interactions could be provided by the free energy of binding of TonB to BtuB, which is significant and characterized by a Kd in the nM range (Freed et al., 2013). To complete the transport cycle, TonB must be disengaged from BtuB and the inner membrane complex of ExbB and ExbD may perform this function and restore the apo state. Whether this involves a mechanical action of TonB, such as a pulling or rotational motion, or another process such as the exchange of the strand-to-strand TonB-Ton box interaction for a strand-to-strand interaction within a TonB dimer (Freed et al., 2013; Gresock et al., 2015) remains to be determined.

In summary, EPR spectroscopy in whole cells provides evidence for a substrate-induced structural transition in BtuB involving SB3 on extracellular apex of the core. The introduction of a mutation to break the R14-D316 ionic lock acting between the core and the barrel of BtuB produces an alternate structural state upon substrate addition so that SB3 is displaced as much as 2 nm into the barrel toward the periplasmic side of the protein. Under these same conditions, no movement of the N-terminal side of the core is detected. This ionic lock will be broken upon TonB binding and mutating this ionic lock may partially mimic the TonB bound state. As a result, this substrate-induced structural transition likely represents a structural state that occurs during the transport process. Remarkably, when BtuB is reconstituted into a phospholipid bilayer, these structural changes in SB3 are no longer observed, indicating that features in the native OM environment, such as the LPS, are required to populate conformational states that are important for BtuB function.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Escherichia coli)RK5016 (A derivative of MC4100 with the genotype araD139 Δ(argF-lac)169 flbB5301 ptsF25 relA1 rpsL150 rbsR22 deoC1 gyrA219 non-9 metE70 argH1 btuB461 recA56)Robert Kadner (University of Virginia)PMID:2982793E. coli strain lacking chromosomal gene for BtuB This strain was authenticated using phenotype assays
Strain, strain background (Escherichia coli)RI90
(araD139 Δ(araABC-leu)7679 galU galK Δ(lac)X74 rpsL thi phoR Δara714 leu+, dsbA:: Kanr)		Coli Genetic Stock Center (Yale University, New Haven, CT)PMID:8917542E. coli DsbA null strain
This strain was authenticated using phenotype assays
Recombinant DNA reagentpAG1
(plasmid)		Robert Kadner (University of Virginia)pUC8 with btuB ORF (2.4 kb) and regulatory regionPlasmid containing WT BtuB gene
Recombinant DNA reagentpAG1 with single point mutations in BtuB (L63C, S65C, N72C, S93C and S237C)Applied Biological Materials (Richmond, BC, Canada)Plasmids used to construct and express BtuB with single mutations
Sequence-based reagentBtuB D316A-FPThis paperPCR primers(5’ – 3’)
GGTGCGGGTGTCGCCTGGCAGAAACAGACTAC
Sequence-based reagentBtuB D316A-RPThis paperPCR primers(5’ – 3’)
GTAGTCTGTTTCTGCCAGGCGACACCCGCACC
Sequence-based reagentBtuB R14A-FPThis paperPCR primers(5’ – 3’)
GTTACTGCTAACGCTTTTGAACAGCCGCGCA
Sequence-based reagentBtuB R14A-RPThis paperPCR primers(5’ – 3’)
TGCGCGGCTGTTCAAAAGCGTTAGCAGTAAC
Sequence-based reagentBtuB V90C-FPNilaweera et al., Biophys. J. 117, 1476–1484.
PMID:31582182		PCR primers(5’ – 3’)
GAATCTGGCGGGGTGTAGTGGTTCTGCCG
Sequence-based reagentBtuB V90C-RPNilaweera et al., Biophys. J. 117, 1476–1484.
PMID:31582182		PCR primers(5’ – 3’)
CGGCAGAACCACTACACCCCGCCAGATTC
Sequence-based reagentBtuB T188C-FPNilaweera et al., Biophys. J. 117, 1476–1484.
PMID:31582182		PCR primers(5’ – 3’)
ACCGGATGCCAAGCGCAGACAGATAACGATGG
Sequence-based reagentBtuB T188C-RPNilaweera et al., Biophys. J. 117, 1476–1484.
PMID:31582182		PCR primers(5’ – 3’)
GCGCTTGGCATCCGGTATTACCATAGGCAACAAC
Chemical compound, drugOG (octylglucoside or n-octyl-β-D-glucopyranoside)Chem-Impex, international (Wood Dale, IL)Cat# 00234Detergent for BtuB reconstitution
Chemical compound, drugVitamin B12(CN-Cbl, Cyanocobalamin)Sigma AldrichCat# V2876Substrate for BtuB
Chemical compound, drug1-Palmitoyl-2-oleoyl-glycero-3-phosphocholineAvanti Polar Lipids, (Alabaster, AL)POPC Cat#8 50457Lipid used for membrane reconstitution of BtuB
Chemical compound, drug(1-Oxy-2,2,5,5-tetramethylpyrrolinyl-3-methyl)methanethiosulfonateCayman Chemical, Ann Arbor MichiganMTSSL
Cat# 16463		Reagent for spin labeling protein cysteine residues
Software, algorithmLongDistances
(v. 932)		Christian Altenbach (UCLA)LabVIEW software routine for the analysis of pulse EPR dataUsed to examine DEER data
Software, algorithmDeerAnalysis
(v. 2019)		Gunnar Jeschke (ETH Zürich)MATLAB routine for the analysis of pulse EPR dataUsed to examine DEER data
Software, algorithmMMM
(v. 2018.2)		Gunnar Jeschke (ETH Zürich)MATLAB routine for the determination of spin label rotamers and predicted label-label distancesUsed in this study to predict distance distributions from crystal structures and in silico BtuB structures for simulated annealing
Software, algorithmXplor-NIH (v. 3.2)Charles Schwieters, Marius Clore (NIH, NIDDK)Used for simulated annealing to generate structures consistent with DEER data
Software, algorithmMATLAB
(v. 2020a)		MathWorks, Inc (Natick, MA)Program needed to execute DeerAnalysis and MMM
Software, algorithmPymol
(v. 2.4.0a0)		Schrödinger, LLC (New York, NY)Program for molecular graphics

Cell lines and mutants

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The pAG1 plasmid with WT btuB gene and the RK5016 strain (araD139 Δ(argF-lac)169 flbB5301 ptsF25 relA1 rpsL150 rbsR22 deoC1 gyrA219 non-9 metE70 argH1 btuB461 recA56) used for growth assays were kindly provided by late professor R Kadner, University of Virginia. The E. coli dsbA null (dsbA-) mutant strain, RI90 (araD139 Δ(araABC-leu)7679 galU galK Δ(lac)X74 rpsL thi phoR Δara714 leu+, dsbA:: Kanr) were obtained from the Coli Genetic Stock Center (Yale University, New Haven, CT). L63C, S65C, N72C, S93C, and S237C btuB mutants were custom produced by Applied Biological Materials Inc (Richmond, BC, Canada). The btuB mutants (L63C-T188C, S65C-T188C, N72C-T188C, V90C-T188C, V90C-S237C, S93C-T188C) with and without the R14A mutation, and V90C, V90C-T188C-D316A, and V90C-T188C-R14A-D316A were engineered using polymerase chain reaction (PCR) mutagenesis. The plasmids were confirmed by sequencing and were transformed into dsbA- cells. Glycerol stocks were prepared and stored at −80°C. The RK5016 strain was authenticated using phenotype assays as described previously (Lathrop et al., 1995). This strain fails to grow in minimal media (MM) that is not supplemented with methionine and vitamin B12. The RI90 strain carried kanamycin resistance and lacks DsbA function (Rietsch et al., 1996). This cell line was authenticated by testing for kanamycin resistance and determining that cells were not able to oxidize pairs of cysteine residues that were expressed on the cell surface (Nilaweera et al., 2019).

Whole cell sample preparation

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dsbA- cells expressing V90C-T188C, L63C, S65C, N72C, V90C, and S93C BtuB were grown in MM supplemented with 200 µg/ml ampicillin, 0.2 % w/v glucose, 150 µM thiamine, 3 mM MgSO4, 300 µM CaCl2, 0.01 % w/v methionine, and 0.01% w/v arginine (Nilaweera et al., 2019). Cells expressing BtuB with the V90C-T188C mutation was spin labeled as described (Nilaweera et al., 2019) and the aliquots of processed cell pellets were mixed with vitamin B12 (0, 1, 5, 20, 30, 60, and 100 µM final concentrations). The cells expressing L63C, S65C, N72C, V90C, and S93C BtuB mutants were processed as described in Nyenhuis et al., 2020b.

Glycerol stocks of dsbA- cells expressing L63C-T188C, S65C-T188C, N72C-T188C, V90C-T188C, S93C-T188C, and V90C-S237C BtuB with and without R14A, V90C-T188C-D316A, and V90C-T188C-R14A-D316A BtuB were used to directly inoculate the pre-precultures (Luria Bertani media with 200 µg/ml ampicillin), grown for 8 hr at 37°C and used to inoculate the MM precultures. The main MM cultures were inoculated with precultures, grown until OD600 ~0.3, and spin labeled (Nyenhuis et al., 2020b). Briefly, the cells were spin labeled with methanethiosulfonate spin label (MTSSL) ((1-oxy-2,2,5,5-tetramethylpyrrolinyl-3-methyl)methanethiosulfonate) (Cayman Chemical, Ann Arbor, MI) in 100 mM HEPES buffer (pH 7.0) containing 2.5% (w/v) glucose with the final concentration of 7.5 nmol/ml of cell culture at OD600 0.3 for 30 min at room temperature (RT). Spin labeled cells were washed by resuspending in 2.5% (w/v) glucose supplemented 100 mM HEPES buffers, first, at pH 7.0 and then, at pD 7.0. During the washing steps, cysteine double mutants without R14A were incubated for 15 min, while 2–5 min for mutants with R14A and 10 min for D316A and for R14A-D316A mutants. For the data shown in Figure 3—figure supplement 4, aliquots of processed V90C-T188C-R14A samples were incubated with vitamin B12 for 0, 30, and 60 min at RT. All other samples were incubated with vitamin B12 for 20 min or less prior to freezing. It should be noted that previous work demonstrated that BtuB can be specifically labeled in these dsbA- cells and that significant labeling of other OM proteins does not occur (Nilaweera et al., 2019).

Reconstituted BtuB sample preparation

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MM main cultures of V90C-T188C and S93C-T188C with and without R14A were grown for 8 hr at 37°C. The harvested cells were used to isolate intact OM (Nyenhuis et al., 2020b). After the second spin at 118,370 × g for 60 min at 4°C, the pellets were resuspended in 5 ml of HEPES buffer. The OMs were solubilized in 100 mM Tris pH 8.0 buffer with 10 mM EDTA and 0.5 g of octylglucoside (OG) (Chem-Impex, Wood Dale, IL). The OM suspension was incubated at 37°C for 10 min and 2 hr at RT and then spun at 64157 × g, 60 min at 4°C. The supernatants were used to spin label BtuB with 12 mM MTSSL at RT, overnight. BtuB was purified using six column volumes (CV) of wash buffer (17 mM OG, 25 mM Tris pH 8.0), 12 CV of 0–100% gradient of elution buffer (1 M NaCl, 17 mM OG, 25 mM Tris pH 8.0) and 6 CV of 100% elution buffer using a Q column and fractions containing BtuB were pooled. POPC (Avanti Polar Lipids, Alabaster, AL) (20 mg/ml) was sonicated in reconstitution buffer (150 mM NaCl, 100 nM EDTA, 10 mM HEPES pH 6.5) with OG (100 mg/ml) until clear, next, 1 ml from micelles mixture was added to each pooled BtuB mutant and incubated at RT, 40 min. BtuB was reconstituted into POPC by dialyzing OG over six buffer exchanges using reconstitution buffer and bio-beads (with minimum of 6 hr dialysis per exchange). Reconstituted BtuB was pelleted by centrifugation at 23425 × g for 40 min at 4°C, resuspended in 200 μl of reconstituted buffer and further concentrated to 50 μl by using Beckman airfuge. The samples were frozen and stored at −80°C.

EPR spectroscopy

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For CW EPR, 6 μl of cell pellet, or 6 μl of cell pellet with 100 μM vitamin B12 were loaded into glass capillaries (0.84 OD, VitroCom, Mountain Lakes, NJ). Capillaries were loaded into a Bruker ER 4123D dielectric resonator (Bruker BioSpin, Billerica, MA) mounted to a Bruker EMX spectrometer. Data were taken at X-band and at temperature with 100 G sweep width, 1 G modulation, and 2 mW of incident microwave power. EPR spectra on live E. coli that did not produce a signal-to-noise ratio of 7–10 or greater with a single 20 s field sweep failed to produce pulse echoes at Q-band pulse of adequate amplitude and were discarded. For pulse EPR, 16 μl of cell pellet, 20% glycerol, and 100 μM CNCbl (vitamin B12) when applicable were combined and loaded into quartz capillaries (1.6 mm OD., VitroCom, Mountain Lakes, NJ). Samples were flash-frozen in liquid nitrogen and loaded into an EN5107D2 resonator (Bruker BioSpin, Billerica, MA). Data were collected on a Bruker E580 at Q-band and 50 K using a 300 W TWT Amplifier (Applied Systems Engineering, Benbrook, TX). The dead-time free 4-pulse DEER sequence was used for all experiments, with rectangular pulses of typical lengths π/2 = 10 ns and π = 20 ns, and a 75 MHz frequency separation. It should be noted that modulation depths obtained from whole cell samples were generally highly variable. We suspect that this is a result of the cells actively metabolizing during the labeling and washing steps in their preparation. Labeled whole cell samples that did not label well or produce sufficiently large modulation depths (approximately 4%) were re-grown and re-labeled. For the instrument settings and specific resonator used in this work, the modulation depth for a well-labeled protein is approximately 20%. Based upon this, labeling efficiencies for most samples were estimated to be approximately 40–60%.

Data processing

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CW EPR spectra were normalized by dividing by the spectral second integral using in-house python scripts. All pulse EPR data, except for data in Figure 3—figure supplement 2, were processed using LongDistances v 932 (Christian Altenbach, UCLA). Data were fit to a variable dimension background, after which the model-free mode was used for distance fitting. The value of the smoothing parameter was selected based on the L-curve, ensuring that the selected value passed through the major oscillations present in the data. Error analysis used 100 variations at the default values for background noise, start time, dimensionality, and regularization error. For data in Figure 2d, the data were instead fit using a model-based mode with two-Gaussian components with free position, width, and amplitude to investigate the dosage dependence of the substrate-dependent shift toward a shorter distance component for the 90–188 pair. Supplementary file 1 contains the error analysis for these data. Data in Figure 3—figure supplement 2 were processed using the DeerNet routine (Worswick et al., 2018) in DeerAnalysis (Jeschke et al., 2006). A folder of all Source Data (raw, unprocessed DEER data) has been provided. All EPR figures were generated using python scripts and the matplotlib plotting library. Protein structure images were generated using Pymol (DeLano, 2002). Simulated distance distributions were generated using the software package MMM v. 2018.2 and the default rotamer library (Jeschke, 2018; Polyhach et al., 2011).

Modeling

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The 90–188 and 90–237 distributions with the 14A mutation and in the presence of vitamin B12 were used in the generation of a model (Figure 6b) for the motion of the apical hatch loop using the software package Xplor-NIH (v. 3.2). The starting structure for modeling was the in surfo structure crystal structure in the presence of cobalamin (PDB ID: 1NQH), which we previously determined to be closest to the native state of the extracellular loops in the native environment (Nyenhuis et al., 2020a). In that work, we found minimal evidence for motion of the extracellular loops, and we assumed that motion was localized to the SB3 loop. The template structure was labeled in silico with the R1 side chain at the 90, 188, and 237 positions using the software package MMM (v. 2017.2) and the default rotamer library. The top three rotamers were selected from the in silico labeling and used to generate three input structures for ensemble calculations, with the relative weights of the three rotamers conserved from the in silico labeling calculation.

During the calculation, the reference R1 sites in the barrel (188 and 237) were held entirely fixed. The R1 side chain at site 90, the underlying SB3 loop element comprising residues 81–104, and the adjoining, unstructured hatch region comprising residues 112–124 were fully mobile during runs, while all remaining residues had fixed backbone atoms and mobile side chains. The standard Xplor potentials BOND, ANGL, and IMPR were used in conjunction with the torsionDB and repel potentials for all elements, and DEER restraints were encoded as square well potentials using the noePot potential term. All potential terms were ensemble averaged across the three input structures. The peak positions used in the modeling were the peak centered at 4.5 nm for 90–188, and the peak at 2.8 nm for 90–237. Full peak widths were used, with the square well stopping at 5% of maximum intensity. Randomization was introduced to the calculations using the randomizeTorsions function on the starting side chain positions of the mobile hatch elements. Following this, 10 rounds of 400 step Powell minimization using all potentials and with the mobile hatch elements were used to satisfy the experimental restraints. Structure calculation was repeated until both restraints were within error.

Data availability

Raw unprocessed DEER data are available in a compressed folder called "SourceData". The Pymol session file used to produce Figure 6b is included as a supplementary file.

References

    1. DeLano WL
    (2002)
    Pymol: an open-source molecular graphics tool
    CCP4 Newsletter on Protein Crystallography 40:82–92.

Decision letter

  1. Janice L Robertson
    Reviewing Editor; Washington University in St Louis, United States
  2. Olga Boudker
    Senior Editor; Weill Cornell Medicine, United States
  3. Reza Dastvan
    Reviewer; Saint Louis University School of Medicine, United States
  4. Phillip Klebba
    Reviewer; Kansas State University, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Acceptance summary:

This is an interesting study that pushes forward the understanding of structural changes associated with the TonB-dependent BtuB transporter in native E. coli membranes. Once of the most intriguing aspects of this investigation, is that it highlights the importance, or necessity in this case, of studying membrane protein dynamics in native settings. The results further the investigation into the mechanism of this protein by linking the structural changes to a salt bridge that is linked to TonB interactions. In addition, the high quality of the EPR spectroscopy data presented, and further development of protein specific labeling in the cell, is expected to advance the ability to study other membrane proteins using similar approaches.

Decision letter after peer review:

Thank you for submitting your article "Structural intermediates observed only in intact Escherichia coli indicate a mechanism for TonB-dependent transport" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Olga Boudker as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Reza Dastvan (Reviewer #2); Phillip Klebba (Reviewer #3).

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

Essential revisions:

This is an interesting study that demonstrates the importance of investigating BtuB in the native E. coli outer membrane environment. Technologically, the DEER data presented were viewed as being of high quality, pushing forward the ability to report conformational changes of membrane proteins in cells. Therefore, this study was regarded as innovative and there was general enthusiasm about the potential to advance the membrane protein field. However, there were several concerns about the interpretation of these conformational changes as directly informing on the transporter mechanism, particularly due to the lack of labeling and functional controls. The complete recommendations by the reviewers have been included below and should be addressed. In particular, the following list describes the essential revisions needed.

1) Please provide quantification of the labeling yields for the various cysteine mutants in this study. In addition, address the question of reproducibility of spectra obtained from different cellular preparations.

2) In order to provide a better framework for interpreting the conformational changes observed, please include a functional assessment of the key mutations examined in this study either by uptake assays or other approach.

3) The proposed mechanism of the breakage of the ionic lock as a transport intermediate is speculative at this point. Please revise the paper to address other possible interpretations that have been raised by the reviewers as detailed below.

Reviewer #1 (Recommendations for the authors):

• The conformational change shown in Figure 6b is not detailed enough. How would such a conformational change affect the other parts of the core domain. Could a more detailed figure be provided?

• The LPS is proposed to be the main factor affecting the differences detected in vivo versus in vitro. Periplasmic components could also be involved, especially the presence of a functional Ton complex. It would be interesting to assess the influence of Ton inhibitors during these experiments. With uptake experiments, the authors could also probe the importance of LPS directly.

• Line 87: there is a third structure of a TBDT bound to TonB, the FoxA-TonB complex (pdb 6I97, PMID 31385808)

• Line 111: define/introduce SB3

• Legend of figure 1: there is a typo in the pdb id, should be 1NQH, not 1NHQ

• Figure 3d, 4th panel from top (90R1-188R1 R14A), what are the 2 marginal peaks at 4 and 5nm for the apo state (blue curve)? And why are these not present in figure 5c 1st panel?

• In the Materials and methods section, lines 563-564, only for the V90C-T188C-R14 samples is indicated a time frame of the experiments. What is the time frame for the other samples?

Reviewer #2 (Recommendations for the authors):

A number of points require clarification.

Details:

1. The introduction would profit from citing Figure 1b when different domains of the transporter are discussed.

2. Concentration of the utilized vitamin B12 is not mentioned in the paper.

3. p. 6, l. 111: Abbreviation SB3 is not introduced.

4. P. 7 (caption of Figure 1): 2Azz is not described in the paper.

5. P. 8 (Figure 2d and corresponding Figure S1): Error analysis or confidence bands are not shown for Figure 2d. In Figure S1-c separating the data and introducing offsets in addition to showing the fits to the data are really helpful.

6. P. 8, l. 173 (caption of Figure 2): "hatch domain" is used instead of "core domain".

7. P. 10 (Figure 3 and corresponding Figure S2): Unless the reviewer is missing a point, the provided distance distributions in Figure 3 for pairs 63-188, 63-188-14A, 65-188, 65-188-14A, 72-188, 72-188-14A, 93-188 does not correspond to the background-corrected DEER data in Figure S2. Even if the dimensionality of the background is presumed to be around 2 because of the two-dimensional distribution of the labeled proteins in the outer membrane or excluded volume is considered, these background corrected data correspond to distance distributions that include broad/long component(s) between 4-8 nm. For instance, in Figure 2b and 2c, these long-distance components are shown. The contribution of such components is higher in Figure 3 and even if the x-axes are between 1.5 to 5 nm, we should see some of those components. A clarification for this review is greatly appreciated.

8. P. 12 (Figure 4): For analyses of pairs 90-188(+B12), 93-188(apo), 93-188-R14A(+B12), please see point 7.

9. P. 12, l. 338: "But significantly, for neither spin pair are the longer substrate-induced shifts that 339 were seen in Figure 3d observed in the reconstituted system." The reviewer agrees with the statement if the distance distributions are correctly presented in Figure 4. Please see points 7 and 8.

10. P. 13. (Figure 5): In the case of DEER distance distribution for the pair 90-188-D316A-R14A(+B12) the error analysis seems a bit underestimated.

11. P. 14, l. 387: "… single broad peak centered around 2.4 nm in the apo state" The corresponding background-corrected DEER spectrum in Figure 5 consists of two components. Please see point 7.

12. P. 16, l. 459: "We do not observe any significant structural changes in the N-terminal side of the core in cells, suggesting that the N-terminus may not move during transport." Please be more specific on the region that is considered the N-terminal side of the core and the supporting experimental data.

Reviewer #3 (Recommendations for the authors):

Biophysical experiments on living cells face more challenges than in vitro studies, like the interactions of probes with other molecules than the target molecule, lower intensity probe responses that may undermine accurate data collection, and interference from cellular background signals. Besides those obstacles, the manuscript by Nilaweera et al., "Structural intermediates observed only in intact Escherichia coli…," attempts to simultaneously label pairs of Cys side chains, in relatively close physical proximity, with extrinsic nitroxide reagents. This framework requires, but the manuscript lacks, controls that confirm the specificity and extent of covalent modification. This deficiency leads to questions about the validity and interpretations of ensuing EPR observations. Secondly, the paper describes structural changes in BtuB when it binds vitamin B12, but whether the conformational changes are pertinent to its transport mechanism is clouded by the impact of the mutations on BtuB structure and physiology. Potential over-interpretation creeps into the title of the paper, "Structural intermediates observed only in intact Escherichia coli…": whether the reorientation of SB3 that occurs in the mutant proteins during B12 binding constitutes a structural intermediate is debatable. Although the authors interpret their data as if they are structural coordinates, the resolution of the ESR observations does not reveal the exact magnitude, overall nature nor direction of probe motion. Protein dynamics are expected during ligand binding, as seen in many proteins, like the closely related TBDT FepA during its binding of ferric enterobactin. Loop motion probably also occurs in the BtuB-B12 binding reaction, but I'm not sure that the motion the authors describe has biological relevance.

1. In both this manuscript, and its predecessor (Nilaweera et al., 2018) the extent and stoichiometry of the MTSL labeling reactions with Cys mutant BtuB proteins are unknown. Because the authors do not provide these data it's difficult to compare one set of Cys pairs to another. To study BtuB in living bacteria the observed EPR spectra must originate from the target protein, in this case BtuB-MTSL. Confirmation requires evaluation of specific (BtuB) and non-specific (other proteins or cell components) labeling, as by western blots with anti-MTSL sera. The manuscript lacks these data, and refers to Nilaweera et al., 2018, that also lacks them. The referenced paper provides CW X-band spectra of cells expressing wild type BtuB or Cys-pair mutant derivatives, but it's not the same parameter. The problem is that the Cys mutant pairs spontaneously form disulfide bonds in vivo, unless the authors inactivate the DsbAB system (that normally facilitates disulfide formation). In this genetic background other cell envelope proteins become susceptible to modification by MTSL. It's relevant that other TBDT contain disulfide bonds in their surface loops, including Fiu (1), FepA (1), FhuA (2) and Cir (1), that are all expressed at high levels when E. coli is grown in minimal media. Besides this uncertainty about the extent of background labeling, the authors tacitly assume that each Cys side chain in each mutant pair is quantitatively labeled, but this may not be the case. One site in a pair may be 90% labeled, and the other 10% labeled. Without knowledge of the labeling distribution, it's hard to interpret the in vivo DEER data. It may be that all of the Cys sulfhydryls in all of the Cys pairs are equivalently quantitatively labeled, but my experience with Cys mutants suggest that this is not the case, and should be evaluated by experiments. I suggest that the authors quantify the MTSL attached to each Cys sulfhydryl of the 5 BtuB Cys pair mutants (e.g., T188C-V90C), and show the background labeling of other cell envelope proteins in each case. This is the only way to impart confidence about the origin of the EPR spectra. These additional data may resolve an inconsistency in the manuscript: CW data on V90C indicate its environment does not change when B12 binds (Figure 1), whereas DEER data on S188C – V90C pair suggest that MTSL attached to V90C relocates 25 Å when B12 binds (Figures 2 and 3). The authors do not address this discrepancy, but misunderstanding of the DEER data is a potential explanation. The authors also encountered a lot of labeling variability. About X-band spectra they stated: "…EPR spectra on live E. coli that did not produce a signal-to-noise ratio of 7 to 10 or greater with a single 20 second field sweep failed to produce pulse echoes at Q-band pulse of adequate amplitude and were discarded." For pulse EPR they encountered similar labeling problems: "… modulation depths obtained from whole cell samples were generally highly variable." Despite the rationalization "… this is a result of the cells actively metabolizing during the labeling and washing steps in their preparation," all the cell samples were presumably identically grown and processed, so what is the basis for the discrepancies? The problem likely originates from irreproducibility in the MTSL labeling reactions. Statistical analyses of the different trials are needed to resolve the variability. It's also necessary to know the exact number of cells that were loaded for CW, and frozen and analyzed by pulsed EPR. Were equivalent amounts of all spin labeled samples used for each CW or pulsed EPR experiments? Because the authors do not mention nor experimentally address these issues, conclusions about the mobility or proximity of the extrinsic probes may be unreliable.

2. The effects of the R14A mutation are interpreted to support the contention of mechanistic relevance, but they may be engendered by the mutation itself. The R14A substitution excludes ionic interactions with D316, but it also substitutes one of the smallest amino acid side chains in place of one of the largest. Besides that, X-ray data show that cobalamin directly contacts both V90 and S93 in SB3 when it binds to BtuB. The attachment of MTSL to these sites adds non-relevant bulk to the surface of the binding pocket. The combination of removing bulk from the BtuB interior, and adding bulk to the SB3 loop directly above it, may impact the tertiary structure of SB3 during the BtuB-B12 binding interaction. Just eliminating the proposed association between R14 and D316 (as in D316A) may similarly destabilize the protein, ultimately altering both affinity for B12 and the structural properties of the OM protein. It seems possible that R14A changes aspects of the binding complex at equilibrium, resulting in unnatural movement of V90 and S93 in unorthodox directions. The proposed movement of the loop toward the periplasm (the actual direction of the motion is not known from the reported data) is consistent with this alternative explanation. The authors previously showed that mutational alterations at R14 or D316 disrupt the structure of the BtuB core, especially with regard to the TonB box region, directly beneath SB3 (Lukasik et al., 2007). Those data originated from purified BtuB Cys mutants, labeled with MTSL. In the current manuscript they observed that the same mutations also affected the configuration of SB3 when B12 bound. In this case the conformational changes were only observed in native outer membranes, for unknown reasons that the authors attribute to the effects of LPS. As a result of these numerous possibilities, it's not certain that R14A or D316A create mechanistically relevant intermediates. It's also conceivable that the distortion of the core caused by the mutations has a domino effect on BtuB structure that ultimately results in aberrant, or novel localization of SB3 during B12 binding. So, the results do not necessarily define a transport reaction intermediate. If not, then neither do they provide mechanistic insight into TonB-dependent transport.

The lack of information about the phenotypes of the mutants themselves, whether it's R14A, D316A, or those that introduce modifiable Cys residues in 6 different positions, is a deficiency of this manuscript. I don't see how to make conclusions about the TBDT transport process without knowing more about the functionality of these constructs relative to native BtuB. What are the affinities of the mutant proteins for B12, and how does modification with MTSL of single and double Cys sulfhydryls affect binding and transport? The authors do not provide information about these key biochemical parameters; any or all of the mutants may be physiologically defective, and hence provide little or no insight into the transport reaction.

3. The authors sometimes misinterpret, misquote or neglect the literature. For example, they ignore the work that involved Jim Feix's lab, on SDSL of FepA 25 years ago, as well as other data on FepA, that demonstrated conformational motion akin to what they now describe for BtuB, both in vitro (Liu et al., 1995) and in vivo (Jiang et al., 1997). Their statement "…Computational work suggests.… that environment, specifically LPS, may be important in controlling the configuration of SB3" neglects other work on TBDT that demonstrated their completely different properties in vitro and in vivo. FepA has >100-fold lower affinity for ferric enterobactin when purified and reconstituted into detergent or liposomes (Payne et al., 1997), than when measured in bacterial cells (Newton et al., 1999). It's more difficult to rationalize an effect of LPS on SB3, that is ensconced with the 22-stranded β-barrel of BtuB, and thereby isolated from interactions with LPS.

https://doi.org/10.7554/eLife.68548.sa1

Author response

Essential revisions:

This is an interesting study that demonstrates the importance of investigating BtuB in the native E. coli outer membrane environment. Technologically, the DEER data presented were viewed as being of high quality, pushing forward the ability to report conformational changes of membrane proteins in cells. Therefore, this study was regarded as innovative and there was general enthusiasm about the potential to advance the membrane protein field. However, there were several concerns about the interpretation of these conformational changes as directly informing on the transporter mechanism, particularly due to the lack of labeling and functional controls. The complete recommendations by the reviewers have been included below and should be addressed. In particular, the following list describes the essential revisions needed.

1) Please provide quantification of the labeling yields for the various cysteine mutants in this study. In addition, address the question of reproducibility of spectra obtained from different cellular preparations.

Quantitation of labeling yields. From the modulation depths in the DEER data, we provide a rough estimate of the efficiency of double labeling relative to the total spin count. Unlike a continuous wave EPR spectrum where a poorly labeled sample can yield usable data, the modulation depth in the DEER signal is dependent upon the fraction of spin-pairs excited. We now include an estimate of labeling efficiency, which typically ranges from 40 to 60% in our samples. Making a more precise measurement of double labeling efficiency in cells relative to the total BtuB count may not be possible; moreover, it is not clear why an exact labeling efficiency matters for the interpretation of these data? Lower or higher labeling efficiencies will alter the quality (signal-to-noise) of the data and affect the error in the distance distributions, but they will not change the result.

Specificity in labeling. The method used here produces labeling that is specific. Previous work using this method on DsbA- cells demonstrated that there was little labeling of the cells when expressing wild-type BtuB (which lacks cysteine) and that no DEER signal can be detected (see: Nilaweera et al. (2019) Biophysical journal 117, 1476-1484). Two previous publications also show DEER signals that arise from specifically labeled BtuB in DsbA- cells (Nyenhuis et al. (2020) JACS 142, 10715-10722; Nyenhuis et al. (2020) Biophys. J. 119, 1550-1557). And in the latter Biophys. J. publication, a wide range of distances is obtained that match well with the predictions from crystal structures.

Reproducibility of spectra. The modulation depths in the DEER data and signal levels from our preparations do vary, despite our best efforts to control for it. However, the EPR spectra are consistent, and DEER data are highly reproducible even when the labeling efficiencies vary.

2) In order to provide a better framework for interpreting the conformational changes observed, please include a functional assessment of the key mutations examined in this study either by uptake assays or other approach.

We performed a growth assay on the ionic mutant used it this study, and it shows that the protein is functional and not damaged in a significant way.

3) The proposed mechanism of the breakage of the ionic lock as a transport intermediate is speculative at this point. Please revise the paper to address other possible interpretations that have been raised by the reviewers as detailed below.

We revised the paper to focus just on what we believe the data tell us, and these changes are detailed below. The data suggest that substrate binding loop 3 undergoes a motion towards the periplasm during the transport cycle. We agree with the reviewers that the exact role of this structural transition, or whether it is necessarily directly involved in transport, is not known at this time.

Reviewer #1 (Recommendations for the authors):

• The conformational change shown in Figure 6b is not detailed enough. How would such a conformational change affect the other parts of the core domain. Could a more detailed figure be provided?

We wanted to see what the DEER data implied regarding the movement of SB3, and Figure 6b is based upon Xplor-NIH modeling using the restraints obtained for the third substrate binding loop SB3. We found that this loop could move and satisfy the EPR restraints without significant rearrangements of the remainder of the core. This was not an MD simulation and we do not have enough data on other regions of the core to know how it might rearrange.

We have provided a Pymol session file showing 300 structures in SB3 that are consistent with the distance data. This should provide the reader with as much detail of this figure as is needed.

• The LPS is proposed to be the main factor affecting the differences detected in vivo versus in vitro. Periplasmic components could also be involved, especially the presence of a functional Ton complex. It would be interesting to assess the influence of Ton inhibitors during these experiments. With uptake experiments, the authors could also probe the importance of LPS directly.

The reviewer is correct, periplasmic components might also be important. However, there is not enough TonB present in these experiments relative to BtuB to account for the altered distributions seen in cell system. Nonetheless looking at the role of TonB directly is something we are attempting to carry out. Because other components such as peptidoglycan might be playing a role, we have amended our comments on the third paragraph in the Discussion to reflect this fact. As mentioned in the Discussion, we focused on LPS because there is already support from computational work for the role of LPS on the energetics of the BtuB core region (see: Balusek and Gumbart, Biophys. J. (2016) 111, 1409-1417).

• Line 87: there is a third structure of a TBDT bound to TonB, the FoxA-TonB complex (pdb 6I97, PMID 31385808)

We added a reference to this structure in the introduction.

• Line 111: define/introduce SB3.

We have now defined “SB3” in the introduction.

• Legend of figure 1: there is a typo in the pdb id, should be 1NQH, not 1NHQ.

The typo has been corrected.

• Figure 3d, 4th panel from top (90R1-188R1 R14A), what are the 2 marginal peaks at 4 and 5nm for the apo state (blue curve)? And why are these not present in figure 5c 1st panel?

These small peaks in the apo state for 90R1-188R1 R14A may indicate a very small incidence of the shifted state of SB3 even in the absence of cobalamin, which is not seen for reconstituted protein (as in Figure 5c). However, these peaks are small enough to preclude reliable interpretation, and so we were not comfortable drawing conclusions from them in the manuscript. Getting true error estimates on distance distributions obtained from DEER is a difficult problem due to the mathematical deconvolution of the dipolar evolution into specific frequency and distance components. Developing methods for error analysis in the processing of DEER data is currently a topic of some interest. The ranges provided in most of the current software packages give a range of solutions (distance distributions) relative to the best fits.

• In the Materials and methods section, lines 563-564, only for the V90C-T188C-R14 samples is indicated a time frame of the experiments. What is the time frame for the other samples?

The timeframe used for the other samples (from label addition to freezing for the DEER experiment) was typically 20 minutes or less, and we have now included this information in the methods.

Reviewer #2 (Recommendations for the authors):

A number of points require clarification.

Details:

1. The introduction would profit from citing Figure 1b when different domains of the transporter are discussed.

We added a reference to Figure 1b in the introduction.

2. Concentration of the utilized vitamin B12 is not mentioned in the paper.

Unless otherwise mentioned, the vitamin B12 concentration was 100 μM. This is mentioned in the Methods section.

. p. 6, l. 111: Abbreviation SB3 is not introduced.

We have now defined this abbreviation in the introduction where it is first used.

4. P. 7 (caption of Figure 1): 2Azz is not described in the paper.

We re-wrote this caption to better indicate the points being measured and to make the reference to 2Azz clearer.

5. P. 8 (Figure 2d and corresponding Figure S1): Error analysis or confidence bands are not shown for Figure 2d. In Figure S1-c separating the data and introducing offsets in addition to showing the fits to the data are really helpful.

The program LongDistances treats error differently for a model-based fit than for a model-free fit. It allows for negative probabilities in the distributions, which generates unrealistic errors for data that have smaller modulation depths. However, the model-based fit does generate an error for each component, and we added a table (Supplementary File 1) which lists the proportions of the two distance components as a function of vitamin B12 as well as the error in the fits.

We modified Figure 2—figure supplement 1, to separate the data with offsets and show fits to the data.

6. P. 8, l. 173 (caption of Figure 2): "hatch domain" is used instead of "core domain".

We made the correction.

7. P. 10 (Figure 3 and corresponding Figure S2): Unless the reviewer is missing a point, the provided distance distributions in Figure 3 for pairs 63-188, 63-188-14A, 65-188, 65-188-14A, 72-188, 72-188-14A, 93-188 does not correspond to the background-corrected DEER data in Figure S2. Even if the dimensionality of the background is presumed to be around 2 because of the two-dimensional distribution of the labeled proteins in the outer membrane or excluded volume is considered, these background corrected data correspond to distance distributions that include broad/long component(s) between 4-8 nm. For instance, in Figure 2b and 2c, these long-distance components are shown. The contribution of such components is higher in Figure 3 and even if the x-axes are between 1.5 to 5 nm, we should see some of those components. A clarification for this review is greatly appreciated.

As the reviewer correctly notes, the difference between the background corrected DEER data and the distance distribution is due to a long-distance contribution. This contribution is in the 6 nm range, and it results from intermolecular BtuB-BtuB interactions due to the formation of what are termed “OMP islands.” The size of this interaction is variable, and it “pollutes” many of our DEER traces in the native system. As a result, we have truncated the distributions at 5 nm. We published a paper last year describing this intermolecular signal and used it to define the interfaces of BtuB that interact in the outer membrane (Nyenhuis et al. (2020) JACS, 142, 10715-10722).

To avoid confusing the reader who carefully looks at the data, we added a sentence to the paragraph that introduces Figure 3 to make the reader aware of this feature.

8. P. 12 (Figure 4): For analyses of pairs 90-188(+B12), 93-188(apo), 93-188-R14A(+B12), please see point 7.

See response to point 7.

9. P. 12, l. 338: "But significantly, for neither spin pair are the longer substrate-induced shifts that 339 were seen in Figure 3d observed in the reconstituted system." The reviewer agrees with the statement if the distance distributions are correctly presented in Figure 4. Please see points 7 and 8.

See response to point 7. Intermolecular BtuB-BtuB interactions generally do not appear in the reconstituted system, and unlike the intermolecular measurements made to SB3, they are not modulated by substrate.

10. P. 13. (Figure 5): In the case of DEER distance distribution for the pair 90-188-D316A-R14A(+B12) the error analysis seems a bit underestimated.

Yes, we agree with the reviewer that the error analysis seems to be underestimated for Figure 5. Nonetheless, this was the output of Christian Altenbach’s program LongDistances. The raw data is available for readers who wish to re-analyze the data or process the data in a different manner. It should be noted that with an echo time of 4 μsec, distances and distributions below 5 nm are expected to be reliable.

11. P. 14, l. 387: "… single broad peak centered around 2.4 nm in the apo state" The corresponding background-corrected DEER spectrum in Figure 5 consists of two components. Please see point 7.

As described above (response to point 7), the data reflects the presence of BtuB-BtuB interactions due to the formation of OMP islands. The effect of intermolecular BtuB-BtuB interactions does not appear past 5 nm. By truncating the distribution at this point, the remaining distribution represents the 90R1-188R1 interaction.

12. P. 16, l. 459: "We do not observe any significant structural changes in the N-terminal side of the core in cells, suggesting that the N-terminus may not move during transport." Please be more specific on the region that is considered the N-terminal side of the core and the supporting experimental data.

We are referring to measurements made to sites 63, 65 and 72 in SB1 and SB2. We modified the discussion to refer to these specific sites so that this is clear to the reader.

Reviewer #3 (Recommendations for the authors):

Biophysical experiments on living cells face more challenges than in vitro studies, like the interactions of probes with other molecules than the target molecule, lower intensity probe responses that may undermine accurate data collection, and interference from cellular background signals. Besides those obstacles, the manuscript by Nilaweera et al., "Structural intermediates observed only in intact Escherichia coli…," attempts to simultaneously label pairs of Cys side chains, in relatively close physical proximity, with extrinsic nitroxide reagents. This framework requires, but the manuscript lacks, controls that confirm the specificity and extent of covalent modification. This deficiency leads to questions about the validity and interpretations of ensuing EPR observations. Secondly, the paper describes structural changes in BtuB when it binds vitamin B12, but whether the conformational changes are pertinent to its transport mechanism is clouded by the impact of the mutations on BtuB structure and physiology. Potential over-interpretation creeps into the title of the paper, "Structural intermediates observed only in intact Escherichia coli…": whether the reorientation of SB3 that occurs in the mutant proteins during B12 binding constitutes a structural intermediate is debatable. Although the authors interpret their data as if they are structural coordinates, the resolution of the ESR observations does not reveal the exact magnitude, overall nature nor direction of probe motion. Protein dynamics are expected during ligand binding, as seen in many proteins, like the closely related TBDT FepA during its binding of ferric enterobactin. Loop motion probably also occurs in the BtuB-B12 binding reaction, but I'm not sure that the motion the authors describe has biological relevance.

There are three points we would like to make regarding these comments. First, as detailed below, previously published work has demonstrated specificity in our labeling – it is the first control experiment we perform. We have also now provided relative estimates of labeling efficiency from the modulation depths in the DEER signal, which is now mentioned in the Methods section where the DEER experiment is described.

Second, we agree with the reviewer that we do not know that the structural transition that we observe in substrate binding loop 3 is related to transport. We have not fully characterized the BtuB core, and other motions may occur that we have not yet observed. All we can say is that the transition we see may occur during the transport process. To address this concern (as well similar concerns raised by the other reviewers), we have modified portions of the text and two paragraphs in the Discussion. Other than the partial unfolding of the very N-terminal segment of BtuB (the Ton box), this movement of SB3 is the only site in the core of BtuB that has been observed to undergo an extensive substrate-induced structural change.

Finally, we do not understand the comment that these EPR measurements do not reveal the exact nature and overall motion of the probe. This is precisely what these measurements do. They provide a measure of the interspin distances in the labeled protein. We demonstrated in an earlier work that the loops in BtuB are relatively static in the intact cell, as a result the movements seen for substrate binding loop 3 (SB3) can be entirely attributed the movement of this loop.

The reviewer may be interested to know that the loops in BtuB do not gate in the native cell environment as they do in a purified phospholipid environment. This may not be true for iron transporters like FepA, but at least for BtuB the loops are relatively static (Nyenhuis et al., Biophys. J. (2020) 119, 1550-1557).

1. In both this manuscript, and its predecessor (Nilaweera et al., 2018) the extent and stoichiometry of the MTSL labeling reactions with Cys mutant BtuB proteins are unknown. Because the authors do not provide these data it's difficult to compare one set of Cys pairs to another. To study BtuB in living bacteria the observed EPR spectra must originate from the target protein, in this case BtuB-MTSL. Confirmation requires evaluation of specific (BtuB) and non-specific (other proteins or cell components) labeling, as by western blots with anti-MTSL sera. The manuscript lacks these data, and refers to Nilaweera et al., 2018, that also lacks them. The referenced paper provides CW X-band spectra of cells expressing wild type BtuB or Cys-pair mutant derivatives, but it's not the same parameter. The problem is that the Cys mutant pairs spontaneously form disulfide bonds in vivo, unless the authors inactivate the DsbAB system (that normally facilitates disulfide formation). In this genetic background other cell envelope proteins become susceptible to modification by MTSL. It's relevant that other TBDT contain disulfide bonds in their surface loops, including Fiu (1), FepA (1), FhuA (2) and Cir (1), that are all expressed at high levels when E. coli is grown in minimal media.

We understand the reviewer’s concern, but this was one of our first control experiments and several published observations indicate that we are not getting significant labeling of proteins other than BtuB in the DsbA- strain. The reviewer mentions the Nilaweera et al. 2018 paper – we presume he is referring to our 2019 paper (Nilaweera et al. (2019) Biophysical J. 117, 1476-1484). First, the DsbA- strain without BtuB expression does not label to any significant extent (see Figure S4 – Nilaweera et al., 2019), and we cannot obtain a DEER signal because there is no echo upon which to set up the pulse experiment. The same is true with WT BtuB is expressed in the DsbA- or DsbB- strains (Figure 2, Nilaweera et al., 2019). Second, the CW EPR spectra we obtain from the DsbA- strain are consistent with our expectations based upon spectra measured from single labeled sites in the RK5016 strain or from purified reconstituted protein. Finally, the interspin distances, which we only observe when BtuB having double cysteines is overexpressed, matches that expected from the crystal structures (see for example Nyenhuis et al. (2020) Biophys. J. 119, 1550-1557).

To ensure that other readers to not miss this important point, we added a note addressing the specificity of labeling in the Methods section under the section “Whole cell sample preparation.”

Besides this uncertainty about the extent of background labeling, the authors tacitly assume that each Cys side chain in each mutant pair is quantitatively labeled, but this may not be the case. One site in a pair may be 90% labeled, and the other 10% labeled. Without knowledge of the labeling distribution, it's hard to interpret the in vivo DEER data. It may be that all of the Cys sulfhydryls in all of the Cys pairs are equivalently quantitatively labeled, but my experience with Cys mutants suggest that this is not the case, and should be evaluated by experiments. I suggest that the authors quantify the MTSL attached to each Cys sulfhydryl of the 5 BtuB Cys pair mutants (e.g., T188C-V90C), and show the background labeling of other cell envelope proteins in each case. This is the only way to impart confidence about the origin of the EPR spectra. These additional data may resolve an inconsistency in the manuscript: CW data on V90C indicate its environment does not change when B12 binds (Figure 1), whereas DEER data on S188C – V90C pair suggest that MTSL attached to V90C relocates 25 Å when B12 binds (Figures 2 and 3). The authors do not address this discrepancy, but misunderstanding of the DEER data is a potential explanation.

The reviewer is in error here. We do not assume or even need quantitative labeling of our cysteine sites. It is in fact not necessary to know the exact extent of labeling to correctly interpret the DEER data – much in the same way that it is not necessary to know labeling efficiency or even have matched donor/acceptor pairs on all proteins in a single molecule FRET measurement. All that is necessary is that sufficient sites be labeled so that an adequate number of spin pairs are excited in the 4-pulse DEER experiment, resulting in a measurable dipolar evolution with good signal-to-noise. Our maximum modulation depths (with our current probe and instrument configuration) are typically around 20% for well labeled protein (both sites near 100%). Taking this value, we can say that our double sites are typically labeled to an efficiency that range from about 40 to 60%. This is adequate but not as good as we would obtain for an in-vitro sample. Unlike a CW EPR spectrum where you might get good signals over quite a wide range of labeling efficiencies with sufficient signal averaging, the DEER signal is very sensitive to labeling efficiency.

We have included an estimate of labeling efficiency in the Methods. Information on the exact level of labeling efficiency in the intact cell is both unnecessary and very difficult to obtain.

Regarding the discrepancy where the label at site 90 shows no change in the EPR spectrum upon ligand binding, we do not agree that there is an inconsistency here. The environment around the label may change, but this does not necessarily mean that the EPR spectrum must change. In the present case, the label is at or near its rigid limit and only an increase in motion is likely to produce a change in EPR lineshape. A change in structure or environment that does not change the motion of the probe on the ns timescale is unlikely to significantly change the CW spectrum. Two labels buried in a protein interior will generally yield very similar if not identical EPR spectra. At the same time, the DEER data provide unequivocal evidence that the label position is changing.

The authors also encountered a lot of labeling variability. About X-band spectra they stated: '…EPR spectra on live E. coli that did not produce a signal-to-noise ratio of 7 to 10 or greater with a single 20 second field sweep failed to produce pulse echoes at Q-band pulse of adequate amplitude and were discarded." For pulse EPR they encountered similar labeling problems: "… modulation depths obtained from whole cell samples were generally highly variable." Despite the rationalization "… this is a result of the cells actively metabolizing during the labeling and washing steps in their preparation," all the cell samples were presumably identically grown and processed, so what is the basis for the discrepancies? The problem likely originates from irreproducibility in the MTSL labeling reactions. Statistical analyses of the different trials are needed to resolve the variability. It's also necessary to know the exact number of cells that were loaded for CW, and frozen and analyzed by pulsed EPR. Were equivalent amounts of all spin labeled samples used for each CW or pulsed EPR experiments? Because the authors do not mention nor experimentally address these issues, conclusions about the mobility or proximity of the extrinsic probes may be unreliable.

We were being quite transparent and honest in stating that not all our attempts at labeling yielded adequate signal-to-noise to produce reasonable DEER data. We were as reproducible as we could be, and we do not know all the reasons for the variability (perhaps variability in the riboswitch expression system intrinsic to BtuB). However, we fail to see why this is an issue. Unlike a CW spectrum, which might be quite easy to acquire over a range of concentrations given sufficient signal averaging, the pulse experiment is more sensitive to relative levels of labeled material and much more time consuming – particularly if labeled protein levels are not adequate. It is not clear what a “statistical analyses of different trials” would tell us or even how it would be carried out.

We do not understand the reviewer’s comment that our conclusions about probe mobility or proximity are unreliable. In every case where we have made a measurement under conditions where a known structure is expected, we get the expected result. We refer the reviewer to our most recent Biophysical Journal paper (Nyenhuis et al. (2020) Biophys. J. 119, 1550-1557) using the current approach where we make a wide range of inter-loop distance measurements. All these measurements yield distances that fall completely within the expectations of the in-surfo crystal structures. There is no evidence that distances we obtain are unreliable.

2. The effects of the R14A mutation are interpreted to support the contention of mechanistic relevance, but they may be engendered by the mutation itself. The R14A substitution excludes ionic interactions with D316, but it also substitutes one of the smallest amino acid side chains in place of one of the largest. Besides that, X-ray data show that cobalamin directly contacts both V90 and S93 in SB3 when it binds to BtuB. The attachment of MTSL to these sites adds non-relevant bulk to the surface of the binding pocket. The combination of removing bulk from the BtuB interior, and adding bulk to the SB3 loop directly above it, may impact the tertiary structure of SB3 during the BtuB-B12 binding interaction. Just eliminating the proposed association between R14 and D316 (as in D316A) may similarly destabilize the protein, ultimately altering both affinity for B12 and the structural properties of the OM protein. It seems possible that R14A changes aspects of the binding complex at equilibrium, resulting in unnatural movement of V90 and S93 in unorthodox directions. The proposed movement of the loop toward the periplasm (the actual direction of the motion is not known from the reported data) is consistent with this alternative explanation. The authors previously showed that mutational alterations at R14 or D316 disrupt the structure of the BtuB core, especially with regard to the TonB box region, directly beneath SB3 (Lukasik et al., 2007). Those data originated from purified BtuB Cys mutants, labeled with MTSL. In the current manuscript they observed that the same mutations also affected the configuration of SB3 when B12 bound. In this case the conformational changes were only observed in native outer membranes, for unknown reasons that the authors attribute to the effects of LPS. As a result of these numerous possibilities, it's not certain that R14A or D316A create mechanistically relevant intermediates. It's also conceivable that the distortion of the core caused by the mutations has a domino effect on BtuB structure that ultimately results in aberrant, or novel localization of SB3 during B12 binding. So, the results do not necessarily define a transport reaction intermediate. If not, then neither do they provide mechanistic insight into TonB-dependent transport.

The lack of information about the phenotypes of the mutants themselves, whether it's R14A, D316A, or those that introduce modifiable Cys residues in 6 different positions, is a deficiency of this manuscript. I don't see how to make conclusions about the TBDT transport process without knowing more about the functionality of these constructs relative to native BtuB. What are the affinities of the mutant proteins for B12, and how does modification with MTSL of single and double Cys sulfhydryls affect binding and transport? The authors do not provide information about these key biochemical parameters; any or all of the mutants may be physiologically defective, and hence provide little or no insight into the transport reaction.

The reviewer raises important points regarding the consequences of the mutations and the incorporation of spin labels into BtuB. First, regarding the ionic interactions surrounding R14 and D316, the reviewer is concerned that the R14A mutation might destabilize the protein and induce unnatural structural rearrangements. An inspection of the TonB-BtuB crystal structure shows that upon TonB binding, R14 is removed from the protein interior so that the ionic interaction to D316 is broken. There is nothing in the structure to indicate that the R14A mutation alone would do anything unusual to BtuB that would not be accomplished by the binding of TonB. To address the concern about function of this mutant, we have performed a growth assay (Figure 3 —figure supplement 3) which shows that the BtuB R14A mutant functions in transport. This is also consistent with earlier work (Lathrop et al., (1995) J. Bacteriol. 177, 6810-6819) showing that a dipeptide insertion that would break the D316-R14A ionic bond does not destroy transport. We have included this information at the Results section. Some of the earlier work from Kadner’s laboratory shows that BtuB is highly tolerant to substitution and insertion mutations.

Regarding the incorporation of the spin label. We would like to point out that two spin labels – at site 90 and 93 – show the same effect. There is the possibility that both are modifying the protein in the same way, but this seems unlikely. Second, spin labels buried into proteins can alter folding energies, but they are generally well-tolerated in proteins and in general they do not dramatically alter structures (Altenbach et al., (2015) Methods in Enzymology, 564, 60). We have published structures with the R1 side chain at position 10 in the protein interior of the protein (Freed et al., (2010), Biophys. J. 99, 1604-1610). Except for some differences in B-factors, the structure is similar to the wild-type structure. Finally, regarding the incorporation of the spin label at site 90, site 90 on BtuB has been derivatized with a wide range of side chains (Mills et al., 2016, BBA 1858, 3105-3112). While these modifications lower B12 affinity by increasing the off rate, the substrate is still able to bind.

We fully admit that the structural changes we observe may or may not be directly related to transport and that more work is needed. But there has been relatively little progress on understanding the molecular details of transport in this system and that the movements that are seen here in SB3 on the C-terminal side of the core suggest that alternate mechanisms should be examined (other than an unfolding of the N-terminus of the core).

The reviewer appears to be uncomfortable with the fact that we attribute changes in the native environment to LPS. We admit that there may be other sources of the differences between the native and reconstituted system, but we do not have a problem proposing that LPS may make the difference. First, BtuB is known to be a highly alloteric protein, with conformational shifts coupled through various regions of the core. Second the computation work of Balusek and Gumbart (Biophys. J. 111, 1409-1417), provides evidence for an energetic coupling of LPS-loop interactions, the ionic interactions between the core and the barrel, and the Ton box. These interactions are consistent – at least qualitatively – with the observations that we have made.

Lastly, like any effort where we see something new, there is always the possibility that we have incorrectly interpreted the experiment. And there is always more to do. But we do not understand why the reviewer insists on raising the possibility of a complicated solution when a relatively simple explanation is available.

3. The authors sometimes misinterpret, misquote or neglect the literature. For example, they ignore the work that involved Jim Feix's lab, on SDSL of FepA 25 years ago, as well as other data on FepA, that demonstrated conformational motion akin to what they now describe for BtuB, both in vitro (Liu et al., 1995) and in vivo (Jiang et al., 1997). Their statement "…Computational work suggests.… that environment, specifically LPS, may be important in controlling the configuration of SB3" neglects other work on TBDT that demonstrated their completely different properties in vitro and in vivo. FepA has >100-fold lower affinity for ferric enterobactin when purified and reconstituted into detergent or liposomes (Payne et al., 1997), than when measured in bacterial cells (Newton et al., 1999). It's more difficult to rationalize an effect of LPS on SB3, that is ensconced with the 22-stranded β-barrel of BtuB, and thereby isolated from interactions with LPS.

We are sorry that the reviewer feels this way, and perhaps we have neglected a study that should have been sited. We are aware of this earlier work, but we did not site this work here because we do not believe it is relevant to the present study. Indeed, we have left out a number of references to earlier work (including our own) because they are not particularly relevant to the present study.

The reviewer mentions Liu et al. (1994), which provides evidence for a change in protein conformation in the ligand binding domain. The exact nature or size of this change was not possible to determine from the CW spectra taken and no distance measurements were made. Moreover, the results were obtained prior to the crystal structure of FepA so that the exact placement of the labeled sites was not known. It is curious that the CW spectra are not consistent with what is expected from a loop region, and inspection of the crystal structure indicates that the sites examined are within the barrel or facing outward near the edge of the barrel at the membrane interface. While this was a groundbreaking paper at the time, we do not see how it is relevant to our current work.

The Jiang et al. 1997 paper describes the spin-labeling of a single site on FepA in vivo. This was an important study that demonstrated that ligand uptake could be followed in vivo by examining the EPR spectrum of a site sensitive to ligand binding. Significantly this was the first case of an in vivo EPR study on an outer membrane protein. However, the exact nature of the structural change observed was not known, and the label (as seen in the subsequent crystal structure) is directed outward from the protein barrel near the extracellular interface. in vivo, FepA can transport, while in vitro it cannot (TBDT has never been reconstituted). As a result, it is perhaps not surprising that an in vivo measurement shows evidence for transport whereas an in-vitro experiment does not. We did not think that referencing this work added to the understanding of our data since it does not reveal significant details of the transport mechanism.

The other two papers Payne, 1997 and Newton, 1999, address ligand affinity, the kinetics of FepA uptake, and the effect of loop deletions on binding and ligand uptake. In the Newton 1999 paper, the difference in ligand affinity that is observed between the in vitro and in vivo cases is attributed to ligand depletion, which presumably results from the different methods used to make the binding measurements. The difference may be due to the difference between the in vivo and in vitro environments, but this was not the reason emphasized in the paper.

https://doi.org/10.7554/eLife.68548.sa2

Article and author information

Author details

  1. Thushani D Nilaweera

    Department of Chemistry and Center for Membrane Biology, University of Virginia, Charlottesville, United States
    Present address
    Genetics and Biochemistry Branch, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, United States
    Contribution
    Conceptualization, Resources, Formal analysis, Validation, Investigation, Methodology, Writing - review and editing
    Contributed equally with
    David A Nyenhuis
    Competing interests
    No competing interests declared

  2. David A Nyenhuis

    Department of Chemistry and Center for Membrane Biology, University of Virginia, Charlottesville, United States
    Present address
    Biochemistry and Biophysics Center, National Heart Lung and Blood Institute, National Institutes of Health, Bethesda, United States
    Contribution
    Conceptualization, Resources, Data curation, Software, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Thushani D Nilaweera
    Competing interests
    No competing interests declared

  3. David S Cafiso

    Department of Chemistry and Center for Membrane Biology, University of Virginia, Charlottesville, United States
    Contribution
    Conceptualization, Resources, Data curation, Supervision, Funding acquisition, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    dsc0b@virginia.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3813-8721

Funding

Office of Extramural Research, National Institutes of Health (NIGMS GM035215)

  • David S Cafiso

Office of Extramural Research, National Institutes of Health (NIGMS S10OD025149)

  • David S Cafiso

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We would like to thank Viranga Wimalasiri (University of Virginia) and Dr Harris Bernstein (NIH, NIDDK) for technical support with the bacterial growth assays.

Senior Editor

  1. Olga Boudker, Weill Cornell Medicine, United States

Reviewing Editor

  1. Janice L Robertson, Washington University in St Louis, United States

Reviewers

  1. Reza Dastvan, Saint Louis University School of Medicine, United States
  2. Phillip Klebba, Kansas State University, United States

Version history

  1. Preprint posted: March 18, 2021 (view preprint)
  2. Received: March 18, 2021
  3. Accepted: July 11, 2021
  4. Accepted Manuscript published: July 12, 2021 (version 1)
  5. Accepted Manuscript updated: July 13, 2021 (version 2)
  6. Accepted Manuscript updated: July 30, 2021 (version 3)
  7. Version of Record published: August 5, 2021 (version 4)

Copyright

© 2021, Nilaweera et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Thushani D Nilaweera
  2. David A Nyenhuis
  3. David S Cafiso
(2021)
Structural intermediates observed only in intact Escherichia coli indicate a mechanism for TonB-dependent transport
eLife 10:e68548.
https://doi.org/10.7554/eLife.68548

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