Cells are surrounded by a plasma membrane that acts like a barrier to help to keep the cell intact. Proteins are embedded in this plasma membrane; and some of these membrane proteins act as channels that allow molecules to enter and leave the cell, while others allow the cell to communicate with its surroundings.

Like all proteins, membrane proteins are chains of amino acids that are joined together by a molecular machine called a ribosome. Most membrane proteins are inserted into the membrane as they are being built. All bacteria contain a protein called YidC that inserts proteins into the plasma membrane of bacterial cells. However, the mechanism behind this activity and the parts of the YidC protein that interact with the ribosome and plasma membrane are unknown.

Wickles et al. have now used data from a range of sources to predict the three-dimensional structure of the YidC protein taken from a bacterium called E. coli. The model shows how the YidC protein is threaded back-and-forth through the membrane, a total of five times. Some of the protein also extends into the inside of the bacterial cell. Wickles et al. then used a technique called cyro-electron microscopy to look at the structure of a YidC protein bound to a ribosome that is building a new protein. Fitting the more detailed model of YidC into this overall structure of the whole complex revealed how a single YidC protein might interact with the ribosome to insert a newly built protein into a membrane.

Wickles et al. then used a combination of theoretical modeling and other experiments to identify the amino acids in the YidC protein that bind to the ribosome: as expected, the binding takes place where the newly formed protein chain exits the ribosome. Further experiments also identified the amino acids in the YidC protein that interact with the newly built membrane protein, thus revealing where it might leave the YidC protein and be inserted into the membrane. The next challenge will be to investigate how the YidC protein assists the folding of new membrane proteins into their own highly specific three-dimensional structure.

At present, a mechanistic understanding of the function of YidC, as well as its mitochondrial and chloroplast counterparts Oxa1 and Alb3, respectively, is limited by a lack of structural information (Kol et al., 2008; Dalbey et al., 2011). High resolution structures are available only for the first periplasmic domain (P1) of Escherichia coli YidC (Figure 1A; Oliver and Paetzel, 2008; Ravaud et al., 2008), however, this domain is poorly conserved, only present in Gram-negative bacteria and not essential for functionality (Jiang et al., 2003). Furthermore, the region(s) of YidC mediating the interaction with the ribosome have not been identified, and the oligomeric state of YidC during co-translational translocation remains controversial (Kohler et al., 2009; Herrmann, 2013; Kedrov et al., 2013). Hence, we set out to determine a molecular model of ribosome-bound YidC during co-translational translocation of the substrate F_Oc (van der Laan et al., 2004), an integral membrane subunit of the ATP synthase complex.

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In order to build an initial structural model of YidC, we predicted contacts between pairs of residues based on covariation analysis (Marks et al., 2011; Hopf et al., 2012). For that purpose, we constructed a multiple sequence alignment of E. coli YidC excluding the nonconserved first transmembrane helix (TM1) and the P1 domain (Figure 1A) and computed direct evolutionary couplings between pairs of YidC residues (Kamisetty et al., 2013). The resulting matrix of coupling strengths (Figure 1B) contains several diagonal and anti-diagonal patterns of stronger coupling coefficients, which are indicative of parallel or anti-parallel helix–helix pairs, respectively. We computed probabilities for each possible helix–helix contact by aggregating the evidence of stronger coupling coefficients over the expected interaction patterns and calibrating the resulting raw scores on an independent dataset of helix–helix interactions to obtain accurate interaction probabilities. Seven helix–helix contacts attained probabilities above 57% (Figure 1B–D) while all other possible contacts scored below 15%, demonstrating the specificity of the method (Figure 1—figure supplement 1B).

We roughly positioned the five TM helices of E. coli YidC relative to each other using the predicted helix–helix contacts as constraints, and rotated them according to their predicted lipid or protein exposure (Lai et al., 2013; Figure 1C). Next, we used MODELLER (Eswar et al., 2008) to create full length models based on the TM core, secondary structure prediction and the 50 residue–residue contacts with the highest coupling coefficients (39 excluding intrahelical contacts, indels and topology violations). In the resulting model (Figure 1E,F), the conserved membrane integrated core of YidC forms a helical bundle arranged like the vertices of a pentagon, in the order 4-5-3-2-6 (clockwise) when viewed from the cytoplasm (Figure 1F). Notably, all the predicted interactions between TM domains can be explained by monomeric YidC suggesting that dimer or oligomer formation may not be strictly required for YidC activity (see also below).

Outside the membrane region, strong helix–helix contacts were predicted within the cytoplasmic loop between TM2 and TM3, which can be explained the by formation of a helical hairpin (Figure 1F). The base of this ‘helical paddle domain’ (HPD) is structurally constrained by predicted contacts with TM3, its tip on the other hand is more mobile and appears to interact with lipid headgroups (see below).

While this manuscript was under review, two crystal structures were published of Bacillus halodurans YidC2 (BhYidC2, 34% sequence identity with E. coli YidC) (Kumazaki et al., 2014), providing us with a unique opportunity to directly assess the accuracy of our model. Overall, the root mean square deviation (RMSD) between the TM helices of our model and those of BhYidC2 is 7.5 Å (3WO6) and 7.3 Å (3WO7) (Table 1), which is within the resolution limits of our method. The global arrangement of TM helices is the same as in BhYidC2, yet, their tilt angle relative to the plane of the membrane is slightly different (Figure 2). The tilt angle of the HPD also differs, as well as its side that faces the membrane (Video 1), which may be indicative of a high degree of flexibility of this domain, consistent with its high crystallographic B-factors (Kumazaki et al., 2014). Notably, the HPD is not essential for YidC function in E. coli since the deletion of the entire domain is possible without compromising cell viability (Jiang et al., 2003).

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A qualitative difference between our model and BhYidC2 that may have more mechanistic importance is the relative position of TM3. In the structure of BhYidC2 a hydrophilic groove is formed on the cytoplasmic side of the TM bundle that has been proposed to form a binding site for YidC substrates (Kumazaki et al., 2014). Interestingly, the opening state of this groove differs between the two crystal forms, that is it is more open in 3WO6 than in 3WO7 (Video 1), largely due to movement of the N-terminal half of TM3 (Figure 2—figure supplement 1). In our model on the other hand, this hydrophilic groove is even more closed than in 3WO7 because we imposed covariation-based constraints between TM3 and TM5 (Pro⁴²⁵-Pro⁴⁹⁹) and between TM3 and TM6 (Cys⁴²³-Gln⁵²⁸ & Phe⁴³³-Thr⁵²⁴) (Figure 2; Video 1). Strikingly, in BhYidC2 the distances between the Cβ atoms of these three pairs are outliers compared to other residue–residue pairs (20.5 Å/20.9 Å/14.9 Å vs an average of 8.2 Å, Table 2). Thus, given that (i) the position of TM3 differs in the two crystal forms, and (ii) that covariation analysis predicts with high accuracy a closer interaction of TM3 with TM6 and one contact with TM5, we conclude that movement of TM3 is a genuine feature of YidC. This movement and the accompanying dynamics of the hydrophilic groove may represent a crucial step in the functional cycle of the YidC insertase.

In summary, the overall structure of our YidC model agrees well with the BhYidC2 crystal structure, and a comparison of both structures reveals dynamic regions in YidC that may be of mechanistic importance. This further illustrates the power of covariation analysis not merely for structure prediction but also for obtaining dynamic insights (Hopf et al., 2012).

Next, in order to further characterize and validate our obtained YidC model, we assessed its stability and biochemical properties in the bacterial membrane by employing traditional molecular dynamics (MD) simulations. Overall, the model was found to be very stable during the simulation. While the five TM helices enable a rigid protein core, the polar loop regions tend to swim on the membrane surface (Figure 3A). An analysis of inter-residue interactions within the TM region (Figure 3B) provides a firm basis to the observed stability of YidC: hydrophobic residues on the exterior of the TM bundle stabilize interactions with the apolar lipid tails. The YidC core, in turn, is stabilized both via short and long-range interactions between the five helices. Residues towards the cytoplasmic side of the core are primarily polar or charged and, therefore, engaged in strong electrostatic or charge–dipole interactions. In contrast, residues on the periplasmic side are primarily aromatic and involved in stacking and other nonpolar dispersion interactions.

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In order to verify the functional relevance of residues suggested by the MD simulations, we created alanine mutants and subjected them to an in vivo complementation assay. Some of the most stabilizing residues, T362 in TM2 and Y517 in TM6, both of which are located at the same height in the membrane, completely inactivated YidC when mutated to alanine (Figure 3D, Figure 3—figure supplement 1). Both mutants were stably expressed, indicating that the lack of complementation was not caused by instability of YidC (Figure 3—figure supplement 2). Several residues close to this pair show intermediate activity levels (F433, M471 and F505), whereas residues further away do not show an effect (Figure 3—figure supplement 1). Taken together, we provide a model for the overall arrangement of the conserved domains of YidC that is in good agreement with our covariation analysis, lipid exposure prediction, MD simulation, in vivo complementation analysis as well as the recent crystal structures.

Interestingly, we observed that YidC induces thinning of the lipid bilayer during the MD simulation. A significant thinning of 7–10 Å results from the hydrophobic mismatch between the TM helices and the membrane (Figure 3E). The thinning is similar in the upper and lower leaflet, and the thinnest region is in proximity of TM3 and TM5. Since membrane inserting YidC substrates have been chemically cross-linked to both these helices (Klenner et al., 2008; Yu et al., 2008; Klenner and Kuhn, 2012), we argue that thinning of this region in particular may be relevant for the molecular mechanism of YidC-dependent membrane insertion. In addition, the distribution of hydrophilic and hydrophobic residues within YidC revealed the presence of a hydrophilic environment on the cytoplasmic side of the YidC TM bundle (Figure 3F), which continues into the mentioned hydrophobic cluster of aromatic residues towards the periplasmic side. It is tempting to speculate that this hydrophilic environment may receive the polar termini and loops of YidC substrates during the initiation of translocation, thus facilitating their transfer across the hydrophobic core of the (thinned) lipid bilayer (see below). Notably, essentially the same conclusions have been drawn on the basis of the BhYidC2 crystal structures and accompanying cross-linking studies (Kumazaki et al., 2014).

In order to provide a molecular model of YidC in its active state, we reconstituted purified full length YidC (extended with the C-terminus of R. baltica YidC [Seitl et al., 2014]) with ribosome nascent chains (RNCs) exposing the first TM helix of F_Oc, and subjected the complex to cryo-EM and single particle analysis to a resolution of ∼8 Å (Figure 4A,B). In agreement with previous structural studies (Kohler et al., 2009; Seitl et al., 2014), YidC binds to the ribosomal exit site, however, the improved resolution now allows for a more detailed interpretation. Firstly, we were able to separate the weaker electron density of the detergent micelle from that of YidC (Figure 4A). Secondly, the presence of elongated structural features (Figure 4D–F) allowed us to dock our molecular model in a distinct orientation (cross correlation coefficient 0.865). Following placement of the YidC-core model, two prominent densities in the membrane region, one next to TM3 and one next to TM5, remained unaccounted for. These could be attributed to either TM1 of YidC or to the TM helix of the nascent chain (NC) F_Oc. Given that (i) YidC substrates are known to crosslink to TM3 (Klenner et al., 2008; Yu et al., 2008; Klenner and Kuhn, 2012), and (ii) that the density neighboring TM3 is aligned with the ribosomal exit tunnel and (iii) that at the same relative position nascent chains have been observed inside the SecY channel (Frauenfeld et al., 2011) (Figure 4—figure supplement 1), the most plausible assignment to the density near TM3 appeared to be the TM helix of F_Oc. To verify this, and to exclude that the density neighboring TM5 corresponds to the nascent chain, we reconstituted single cysteine mutants of YidC either in TM3 (M430C and P431C) or in TM5 (V500C and T503C) with RNCs of a single cysteine mutant of F_Oc^(G23C), and exposed them to disulphide crosslinking. Upon exposure to the oxidator 5,5′-dithiobis-(2-nitrobenzoicacid) (DTNB), only in the TM3 mutants a DTT-sensitive ∼90 kDa product appeared that reacted with antibodies against the nascent chain (NC-tRNA∼30 kDa, Figure 4C) as well as YidC (∼60 kDa, Figure 4C). Thus, the adduct represented indeed the inserting F_Oc TM domain crosslinked to TM3 of YidC. RNCs lacking a cysteine in the nascent chain (Figure 4—figure supplement 2) or YidC mutants with cysteines in TM5 did not yield any crosslinks (Figure 4C). Hence, we conclude that the unaccounted electron density next to TM3 represents the TM of the nascent chain, and that the density neighboring TM5 represents TM1 of YidC (Figure 4D–F).

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We attribute the remaining unaccounted electron density in the periplasmic region to the P1 domain; however, because it is substantially smaller than the crystal structure of P1, we did not include it in our molecular model. Flexibility relative to the conserved membrane region of YidC is the most likely explanation for this finding. We did not observe density for the HPD, in agreement with its flexibility observed in both, the crystal structures of BhYidC2 and the MD simulations (Figure 3C).

In order to validate our molecular model of co-translationally active YidC, we mutated residues that would be in direct contact with the ribosome (Figure 5A,B) and analyzed their effect on functionality in the in vivo complementation test. Indeed, mutation of residues Y370A and Y377A (contacting ribosomal RNA helix 59) and D488K (contacting ribosomal protein uL23) severely interfere with YidC activity (Figure 5C, Figure 5—figure supplement 1) thereby emphasizing their functional significance. All these mutants were stably expressed, indicating that the lack of complementation was not caused by instability of YidC (Figure 5—figure supplement 2). Given that YidC in general is known to be very tolerant to point mutations (Jiang et al., 2003), this provides further support for the overall correctness of our model of ribosome-bound YidC during membrane protein insertion.

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Finally, it is notable that we observe only a single monomer of YidC bound to the active ribosome. This is in agreement with recent literature showing clearly that both YidC (Herrmann, 2013; Kedrov et al., 2013; Seitl et al., 2014) and the SecY complex (Frauenfeld et al., 2011; Park and Rapoport, 2012; Taufik et al., 2013; Park et al., 2014) can be fully active as monomers. However, the comparison of models for active YidC and active SecY (Figure 5E, Figure 4—figure supplement 1) reveals an important difference between the two proteins that has mechanistic implications. While SecY is known to translocate hydrophilic nascent chains through its central aqueous channel (Cannon et al., 2005; Rapoport, 2007; Driessen and Nouwen, 2008) and insert TM domains through a lateral gate (Van den Berg et al., 2004; Gogala et al., 2014), our model suggests that the YidC substrates are inserted at the protein-lipid interface. Two principal findings of our work suggest how YidC may facilitate this process: (i) it provides a hydrophilic environment within the membrane core for receiving the hydrophilic moieties (termini or loops) of a substrate, and (ii) it reduces the thickness of the lipid bilayer: initial interaction of the hydrophilic moieties of YidC substrates with the hydrophilic environment of YidC would allow for a partial insertion into the membrane, while facilitating exposure of the hydrophobic TM domains to the hydrophobic core of the bilayer. The latter in turn may compensate for the energetic penalty of driving the hydrophilic moieties across the (already thinned) bilayer. Further biochemical and structural studies that capture the earlier stages of this translocation process are eagerly awaited to fully elucidate this mechanism.

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Author details

1. Stephan Wickles

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   SW, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

2. Abhishek Singharoy

   Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana-Champaign, Urbana, United States

   Contribution

   AS, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

3. Jessica Andreani

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   JA, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

4. Stefan Seemayer

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   SS, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

5. Lukas Bischoff

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   LB, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

6. Otto Berninghausen

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   OB, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

7. Johannes Soeding

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   JS, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

8. Klaus Schulten

   Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana-Champaign, Urbana, United States

   Contribution

   KS, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

9. Eli O van der Sluis

   1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
   2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

   Contribution

   EOS, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   vandersluis@lmb.uni-muenchen.de

   Competing interests

   The authors declare that no competing interests exist.

10. Roland Beckmann

    1. Gene Center Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany
    2. Center for Integrated Protein Science Munich, Department of Biochemistry, Ludwig-Maximilians-Universität München, Munich, Germany

    Contribution

    RB, Conception and design, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    beckmann@lmb.uni-muenchen.de

    Competing interests

    The authors declare that no competing interests exist.

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