Proteins perform many important jobs inside cells. Each protein is made of a chain of building blocks called amino acids that fold together to form precise shapes. Old, damaged or poorly constructed proteins tend to be harmful to cells so it is important that they are recycled. Ring-shaped enzymes called AAA+ unfoldases help to recycle these proteins by unfolding their amino acid chains. However, since these enzymes only interact with their target proteins for a short time it has been difficult to find out what happens when the target protein binds to the enzyme.

AAA+ unfoldases use chemical energy provided by a molecule called ATP to drive protein unfolding. Ripstein et al. used a technique called electron cryomicroscopy to study a AAA+ unfoldase from a microbe called Thermoplasma acidophilum. The enzymes were supplied with a molecule called ATPγS, which is similar to ATP but releases its energy more slowly. Ripstein et al. used this slowed down reaction, as well as new computer algorithms to analyse the microscopy images, to examine the structure of the target bound to the enzyme.

The experiments show that proteins move through the centre of the enzyme’s ring as they unfold. At any time the enzyme is attached to the amino acid chain of the target protein in several places, which allows it to pull the protein through the ring in a “hand-over-hand” motion. Energy from each ATP molecule drives the enzyme to release one contact with the amino acid chain and form a new contact slightly further along the chain.

It remains unclear how AAA+ unfoldases identify and bind to the proteins they unfold and how they work alongside other recycling proteins inside cells. AAA+ unfoldases have an essential role in the biology of cells, and contribute to cancer and several other human diseases. Therefore, understanding how these proteins work may aid the development of new drugs to treat these diseases.

The 83 kDa VAT protein, which is composed of an N-terminal regulatory domain (NTD) and two tandem nucleotide-binding AAA+ domains (NBD1 and NBD2), assembles into a 500 kDa homohexamer (Figure 1—figure supplement 1). The physiological function of the VAT NTDs remains unclear. Truncation of the first 182 residues of VAT removes the regulatory NTD, producing a construct (ΔN-VAT) that has both higher ATPase and unfoldase activity than the intact protein (Gerega et al., 2005; Rothballer et al., 2007; Barthelme and Sauer, 2012). In solutions of ΔN-VAT, the enzyme itself is degraded in an ATP- and proteasome-dependent manner (Figure 1a–c). The 20S proteasome does not hydrolyze ATP during proteolysis (Barthelme et al., 2014) and therefore this observation demonstrates that ΔN-VAT complexes must unfold other copies of ΔN-VAT via a process that requires ATP hydrolysis prior to degradation by the proteasome. VAT’s ability to unfold pyruvate kinase (Figure 1b) and other VAT particles is consistent with the finding that it can recognize substrates with an unstructured N or C terminus (Gerega et al., 2005) and does not require a specific marker or target sequence for initiating protein unfolding.

Cryo-EM of full-length VAT previously showed that the enzyme adopts a helical split-ring conformation in the presence of ADP (Huang et al., 2016). ΔN-VAT particles with bound ADP assemble into long fibrils (Figure 1d). Power spectra from these fibrils (e.g. Figure 1d, inset), as well as their real space appearance, suggest that they are composed of helical split-ring structures interacting with each other top-to-bottom in a poorly-ordered helical filament. The ~1/55 Å⁻¹ distance from the origin to the first layer line is consistent with the pitch of each NBD helix in the split-ring structure of VAT. These fibrils were not observed in the ADP-bound state of full-length VAT (Huang et al., 2016), possibly because the NTDs block their formation due to steric hindrance. With the slowly-hydrolysable ATP-analogue ATPγS bound, full-length VAT exists in both the split-ring conformation and a six-fold symmetric stacked-ring conformation (Huang et al., 2016). Similarly, ΔN-VAT with bound ATPγS forms a mixture of fibrils (Figure 1e) as well as single particles, the majority of which appeared to be six-fold symmetric (Figure 1e, red circles). Single particle cryo-EM analysis was performed, excluding the fibrils by inspecting micrographs after automatic particle selection and manually deselecting particle images that were part of a filament. This process allowed calculation of a cryo-EM map of the six-fold symmetric stacked-ring conformation of VAT at 3.9 Å resolution. This substantial improvement over the 7.8 Å resolution obtained previously for the six-fold symmetric conformation of full length VAT (Huang et al., 2016) is likely due to a number of factors. First, removal of the NTDs in ΔN-VAT makes the complex more structurally homogeneous and allowed specimens to be prepared with thinner ice than could be used with full length VAT. Specimen preparation with gold grids likely reduced beam-induced movement and improved image quality. Finally, preparation of crowded specimens with ~700 particles per micrograph improved power spectra and allowed more accurate fitting of contrast transfer function parameters at high resolution. The map resolution enabled construction of an atomic model for the complex (Figure 2, Figure 2—figure supplement 1, Supplementary file 1). In the structure, a central pore runs along the six-fold symmetry axis of the complex through the center of the stacked NBD1 and NBD2 rings (Figure 2a, left, hexagon). While it is possible that removal of the NTDs affects the conformations of the NBDs, Fourier shell correlation (FSC) of the map with the earlier map of full-length VAT (Huang et al., 2016) with the NTDs excluded by masking reached 0.5 at 8 Å resolution, consistent with the 7.8 Å resolution of the full-length VAT map (Figure 2—figure supplement 1e). The stacked-ring structure of VAT resembles recent structures of the VAT homologue p97 in the fully ATP-loaded state (Zhang et al., 2000; DeLaBarre and Brunger, 2003; Banerjee et al., 2016), except for the orientations between the NBD1 and NBD2 domains in VAT protomers and the relative rotation of the NBD1 and NBD2 rings. All AAA+ unfoldases have conserved sequences that extend from the NBDs towards the center of the ring, forming ‘pore loops’ that are essential for unfolding substrate (Gerega et al., 2005) (Figure 2—figure supplement 2). In the six-fold symmetric VAT structure, pore loop 1 of NBD1, containing conserved and essential tyrosine residues (Tyr264 and Tyr265) (Gerega et al., 2005), forms an opening that is ~28 Å across while pore loop 2 of NBD1 (residues 299 to 305) forms a slightly tighter aperture that is ~18 Å across (Figure 2—figure supplement 2a, left). Similar apertures are produced by the corresponding pore loop 1 of NBD2 (containing conserved residues Trp 541 and Val 542) and pore loop 2 of NBD2 (residues 575–579) (Figure 2—figure supplement 2b, left). An atomic model could be built for a significant fraction of the map (Figure 2b), including the linker region between NBD1 and NBD2 (Figure 2c, left), and both NBD1 and NBD2 have strong density for ATPγS in their nucleotide-binding pockets (Figure 2c, right) indicating that VAT adopts a rigid and symmetric conformation when fully loaded with nucleotide.

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Unsupervised classification of ATPγS-loaded ΔN-VAT particle images using a new ab initio 3D classification algorithm (Punjani et al., 2017) allowed identification of a second conformation of the protein that was significantly different from known structures. The class corresponding to this conformation, containing ~15% of particle images, showed two ΔN-VAT complexes in close apposition to one another (Figure 3, Figure 3—figure supplement 1). Refinement of this structure to ~4.8 Å resolution revealed it to consist of a ΔN-VAT complex in the process of unfolding a neighboring complex as substrate (Figure 3a). Formation of this state may be dependent on the slow hydrolysis thought to occur with ATPγS. The map shows clear density for the substrate protein in an extended conformation as it runs almost directly through the central pore of the active ΔN-VAT (Figure 3a and b, red density). This substrate-bound conformation displayed a strong preference for particles to lie with their long-axes parallel to the plane of the specimen support, possibly because of the elongated shape of the complex (Figure 2—figure supplement 1d).

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Density for the substrate above NBD1 of the active ΔN-VAT complex is weak and fragmented, and is most clearly visualized without sharpening of the map (Figure 3a, above dashed line) (Zhao et al., 2015a), suggesting that outside of the active enzyme the substrate has a flexible orientation relative to the pore of ΔN-VAT. The length of unfolded protein gripped by the pore of the active ΔN-VAT is ~80 Å, which corresponds to 12 to 14 extended amino acid residues. In both NBD1 and NBD2 five of the six VAT subunits form close contacts with the substrate through their pore loops (Figure 3b and c, and Figure 2—figure supplement 2a and b, middle). The hydrophobic residues of these pore loops create a tight helical sheath around the substrate (Figure 3a, green). The top and bottom of the helix formed from the five subunits that grip the substrate are joined by a ‘seam’ subunit, which does not interact with the substrate and has protomer-protomer contacts that differ dramatically from the other subunits (Figure 3c, gold, Figure 2—figure supplement 2a and b, middle). This helical arrangement of the pore loop residues maximizes the contact area between the unfolded substrate and the pore loops of the five subunits that form the helix (Figure 3b, Video 1). Unfolded substrate that spans the gap between the pore loops of NBD1 and NBD2 has a low density in the cryo-EM map, likely because of flexibility of the substrate in this region. This flexibility is likely due to a lack of tension on the substrate in the observed state. The transition from the stacked-ring to the substrate-engaged conformation of ΔN-VAT occurs with rearrangement of NBD1 relative to NBD2 in each protomer and a rotation of the NBD1 ring against the NBD2 ring (Figure 3—figure supplement 2, Videos 1 and 2). In the resulting asymmetric structure, the NDB1 and NBD2 of each protomer form similar interactions with their neighboring protomers. The two rings are also displaced horizontally relative to each other in the substrate-bound state and the central pore is tilted so that it passes between the rings at an angle of ~15° (Figure 3—figure supplement 2c). This rotation and displacement brings the NBD1 and NBD2 rings closer together than in the stacked-ring conformation, decreasing the distance between their pore loops.

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In this study, only the newly identified conformation described above is found with substrate bound. Consequently, the six-fold symmetric stacked-ring state with ATPγS bound and the split-ring state with ADP bound (Huang et al., 2016) likely correspond to pre- and post-unfolding conformations of the enzyme. In the substrate-bound conformation, the seam protomer is positioned such that it does not interact with the substrate. Notably, recent structures of the 26S proteasome (Chen et al., 2016; Wehmer et al., 2017) also showed that either the Rpt5 or Rpt6 subunit in the hexameric Rpt ring of the 19S regulatory particle base is displaced from the pore, similar to the seam protomer in VAT (Figure 3). Furthermore, the translocation mechanisms of AAA+ DNA helicases such as E1, Rho, and MCM, also involve a helical gripping state in which only five of six protomers interact with a single DNA strand (Lyubimov et al., 2011; Itsathitphaisarn et al., 2012; Enemark and Joshua-Tor, 2006). The structure presented here suggests an analogous substrate translocation mechanism for VAT where the enzyme operates processively with a ‘hand-over-hand’ mechanism, shown schematically in Figure 4a, focusing on protomer 1 outlined in green. In the first step of this cycle (Figure 4a, top) protomer 1 is bound to unfolded substrate at the bottom of the helix of VAT protomers, protomer 2 in the seam position does not contact the substrate (Figure 4a, gold), and protomer 3 binds substrate at the top of the helix near the remaining folded protein. The nucleotide binding sites shared by the seam protomer and protomer 1 are empty (see below). As the seam protomer reengages with substrate at the top of the helix (step 2: Figure 4a, middle), protomer 1 disengages from the substrate and becomes the seam protomer, with protomer 6 now furthest along the unfolded substrate. The process repeats in a cyclical manner for additional protomers (e.g. step 3: Figure 4a, bottom, Video 3). The power stroke for substrate unfolding is provided by ATP hydrolysis, which occurs in sequence around the VAT ring, with ADP released by the protomer adjacent to the seam (counter clockwise from the seam when viewed from NBD1 toward NBD2, Figure 4b). Although the precise position where ATP hydrolysis and ADP release occur in the cycle is not apparent from our data, it is clear that ATP must rebind to the seam protomer as it engages with substrate at the top of the protomer helix. As the cycle continues, each protomer pulls a single section of the substrate, corresponding to ~13 Å or approximately two extended residues, through the central pore until it becomes the lowest protomer in the helix and releases the substrate (Video 3). The cyclic nature of the ATP hydrolysis gives directionality to the substrate translocation, allowing unfolded protein to be passed to the proteasome for degradation.

Figure 4

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The cyclic ‘hand-over-hand’ model requires that each protomer hydrolyze ATP in turn (Figure 4b). A single inactivated subunit would consequently have a severe effect on the rate of ATP hydrolysis, potentially stopping unfolding. To test the model, we measured the ATPase activity of VAT complexes that had been doped with different fractions (r) of catalytically-inactive protomers bearing mutations in the Walker B motifs of both NBD1 and NBD2 (E291Q, E568Q) (Werbeck et al., 2008) relative to wild type VAT ATPase activity (A). These experiments were carried out in the presence of GFP with an 11-residue ssRA tag as substrate (Barthelme et al., 2014). For multimeric enzymes where each subunit functions independently, ATPase activity is expected to decrease linearly as increasing amounts of catalytically dead mutant are incorporated randomly into the oligomers (Figure 4c, blue line). In contrast, the decrease in ATPase activity of the mixed complexes as a function of increasing concentration of catalytically dead mutant closely follows the $A= {(1-r)}^6$ curve expected for the proposed hand-over-hand model (Figure 4c, gray line). The activity of VAT with increasing r decreases faster than the ${\displaystyle A={\left(1-r\right)}^6+5r{\left(1-r\right)}^5}$ curve (Figure 4c, dashed gray line) that is expected if ATP hydrolysis only stops when two of the six VAT protomers are inactivated. A similar curve is obtained when measuring the unfolding rate of the ssRA tagged GFP substrate rather than the ATPase rate (Figure 4d).

The slight elevation relative to the $A= {(1-r)}^6$ profile in Figure 4c and d may reflect an ‘escape’ mechanism that allows stalled VAT complexes to disengage and then reengage with substrate, as seen with other molecular machines (Werbeck et al., 2008). In the absence of an escape mechanism ATP hydrolysis would proceed from one protomer to the next in a cyclical manner (Figure 4b) until a defective subunit is reached, preventing further hydrolysis and, thus, subsequent substrate unfolding. In contrast, if the machine is able to disengage at this point it would be possible for other functional subunits to hydrolyze ATP, although at reduced rates. The existence of an escape mechanism therefore has important implications for substrate unfolding, even in the context of wild type VAT, by allowing the release of stalled substrate. This release minimizes both the loss of active enzyme and increases catalytic efficiency. Our ATPase and unfolding assay data in Figure 4c and d cannot distinguish between a model where each protomer hydrolyzes ATP simultaneously or a sequential binding/hydrolysis mechanism as described in Figure 4a and b. However, previous ATPase measurements with VAT followed Michaelis-Menten kinetics (Gerega et al., 2005), with little cooperativity for ATP hydrolysis. This low cooperativity is inconsistent with simultaneous hydrolysis at all twelve nucleotide binding sites but is consistent with the proposed hand-over-hand mechanism, where only one NBD needs to bind and hydrolyze nucleotide at a time. The sequential model for ATP hydrolysis is also consistent with the asymmetry of the complex and nucleotide binding sites observed here, and the homology between VAT and other AAA+ enzymes known to hydrolyze ATP in a sequential fashion (Enemark and Joshua-Tor, 2006; Kim et al., 2015). ATP hydrolysis in the two NBDs has been shown to be coupled, with mutations in one domain also affecting ATP hydrolysis by the other (Gerega et al., 2005). The mechanism proposed here suggests that the two NBDs of each protomer move along the substrate in unison with each other (Figure 4a, Video 3). Both NBD1 and NBD2 of each protomer form similar interactions with their neighboring protomers. Preserving these similar interactions would require that they function in concert with each other. These conformations may be perturbed by the presence of the NTDs in full-length VAT. The conformations could also differ when the enzyme is in a physiological mixture of ATP, ADP, and inorganic phosphate, which could also affect the appearance of the bound substrate.

The similar conformations of both NDB rings suggests that the mechanism of substrate unfolding proposed here may well operate in single-ring AAA+ unfoldases as well, including Rpt_1-6 of the 19S proteasome, and the hexameric PAN and ClpX unfoldases. Thus, a picture emerges whereby AAA+ unfoldases adopt asymmetric substrate-bound conformations, creating narrow channels that sequester unfolded stretches of target proteins. Notably, even in molecular machines comprised of homo-oligomers, asymmetry can play an important functional role, in the case of VAT by priming the enzyme for processive translocation of its substrate.

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Author details

1. Zev A Ripstein

   1. The Hospital for Sick Children Research Institute, Toronto, Canada
   2. Department of Biochemistry, University of Toronto, Toronto, Canada

   Contribution

   ZAR, Performed protein purification, electron microscopy, image analysis, atomic model building, and enzyme assays, Wrote the manuscript with input from the other authors

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0003-3601-0596

2. Rui Huang

   1. Department of Biochemistry, University of Toronto, Toronto, Canada
   2. Department of Molecular Genetics, University of Toronto, Toronto, Canada
   3. Department of Chemistry, University of Toronto, Toronto, Canada

   Contribution

   RH, Created the ΔN-VAT construct , Performed protein degradation assays

   Competing interests

   No competing interests declared.

3. Rafal Augustyniak

   1. Department of Biochemistry, University of Toronto, Toronto, Canada
   2. Department of Molecular Genetics, University of Toronto, Toronto, Canada
   3. Department of Chemistry, University of Toronto, Toronto, Canada

   Contribution

   RA, Created the E291Q, E568Q VAT mutant and assisted with the purification of this protein

   Competing interests

   No competing interests declared.

4. Lewis E Kay

   1. The Hospital for Sick Children Research Institute, Toronto, Canada
   2. Department of Biochemistry, University of Toronto, Toronto, Canada
   3. Department of Molecular Genetics, University of Toronto, Toronto, Canada
   4. Department of Chemistry, University of Toronto, Toronto, Canada

   Contribution

   LEK, Supervised research

   For correspondence

   kay@pound.med.utoronto.ca

   Competing interests

   LEK: Reviewing editor, eLife

   "This ORCID iD identifies the author of this article:" 0000-0002-4054-4083

5. John L Rubinstein

   1. The Hospital for Sick Children Research Institute, Toronto, Canada
   2. Department of Biochemistry, University of Toronto, Toronto, Canada
   3. Department of Medical Biophysics, University of Toronto, Toronto, Canada

   Contribution

   JLR, Supervised research, Wrote the manuscript with input from the other authors

   For correspondence

   john.rubinstein@utoronto.ca

   Competing interests

   No competing interests declared.

   "This ORCID iD identifies the author of this article:" 0000-0003-0566-2209

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