Cells are filled with activity—proteins must be folded into intricate shapes and unfolded, complex molecules must be built and taken apart—and all this activity requires energy. Cells use a molecule called ATP as a source of energy, with the energy being released when enzymes called ATPases remove a phosphate group from the ATP molecule.

There are many different ATPases, and when one of them does not work properly, the consequences can be severe. A single mutation in the gene for an ATPase called TorsinA, for example, can lead to a painful and severely disabling disorder called primary dystonia. However, scientists have yet to discover what the TorsinA enzyme does in cells and how mutated TorsinA causes primary dystonia.

It is known that the TorsinA enzyme is found in the endoplasmic reticulum—a region of the cell where, among other things, proteins are folded and unfolded—and that it interacts with two membrane proteins, LAP1 and LULL1. Also, TorsinA is a member of the AAA+ family of ATPases (AAA+ is short for ATPases Associated with diverse cellular Activities), and these enzymes tend to combine with each other to form functional ring-shaped arrays.

Now Sosa et al. have used a combination of X-ray crystallography, electron microscopy, computer modeling, and biochemistry to study the interactions between TorsinA and the two membrane proteins. This work revealed that, somewhat surprisingly, certain parts of the membrane proteins have structures that are similar to those of AAA+ ATPases. This allows the TorsinA enzymes and the membrane proteins to combine to form rings that contain three enzymes and three LAP1 proteins or three LULL1 proteins.

Within these rings, the LAP1 and LULL1 proteins activate the TorsinA enzyme by supplying an amino acid called arginine to the neighboring TorsinA molecule. The amino acid is supplied to a site on the enzyme that can bind nucleotides (which are the building blocks of DNA and RNA); the LAP1 and LULL1 proteins themselves cannot bind nucleotides.

The work of Sosa et al. may help scientists to better understand how TorsinA causes primary dystonia. One of the important next steps is to work out what molecules TorsinA acts on.

Lamina-Associated Polypeptide 1 (LAP1) and Luminal domain-Like LAP1 (LULL1) are type II integral membrane proteins with ∼30 kDa luminal domains that share 62% identity. LAP1 localizes to the nuclear envelope (NE) via its lamin-interacting domain, whereas LULL1 is found throughout the endoplasmic reticulum (ER) (Senior and Gerace, 1988; Goodchild and Dauer, 2005). LAP1 and LULL1 associate with Torsins (Goodchild and Dauer, 2005; Naismith et al., 2009; Zhao et al., 2013), which are ER-resident members of the ATPases Associated with a variety of cellular Activities (AAA+ ATPases) superfamily (Iyer et al., 2004; Hanson and Whiteheart, 2005; Erzberger and Berger, 2006). AAA+ ATPases use ATP hydrolysis to undergo conformational changes and to exert mechanical force on a substrate. AAA+ ATPases are involved in many processes, including vesicle fusion and scission, protein folding/unfolding, complex assembly/disassembly, protein transport, and nucleic acid remodeling. Detailed mechanistic knowledge exists for a number of AAA+ ATPases, but not for Torsins. Torsins are found in all animals, but not in single-cell organisms and plants. Deletion of a single glutamate in TorsinA (humans have four Torsin orthologs: TorsinA, TorsinB, Torsin2A, and Torsin3A) at position 302/303 causes the movement disorder DYT1 primary dystonia (Ozelius et al., 1997; Breakefield et al., 2008). Mammalian TorsinA has been assigned to a variety of possible functions including nuclear envelope (NE) organization, synaptic vesicle transport and turnover, operation of the secretory pathway, protein degradation, cytoskeletal organization and transport via NE budding (Vander Heyden et al., 2009; Granata and Warner, 2010; Jokhi et al., 2013). While LAP1 and LULL1 were first considered as Torsin substrates (Naismith et al., 2009), they are now being proposed as possible activators of Torsin (Zhao et al., 2013).

To better understand the function of LAP1 in relation to Torsin, we set out to determine its structure. Our data suggest that LAP1 and LULL1 can both form heterohexameric ring assemblies with Torsin, whereby Torsin is activated through an arginine finger at amino acid (aa) R563 of LAP1 (R449 of LULL1). Thus LAP1 and LULL1 each have dual roles, namely the targeting and activation of Torsins.

The conserved luminal domain of LAP1 (aa356–583) was recombinantly expressed, purified, and set up for crystallization. Because the protein failed to yield crystals alone, we produced a camelid single domain (VHH) antibody fragment, VHH-BS1, against LAP1 as a crystallization chaperone. The complex of VHH-BS1 and LAP1 yielded well-diffracting crystals (Table 1). The structure was solved using molecular replacement with VHH as the search model (for details, see ‘Materials and methods’). In the asymmetric unit we find two LAP1 molecules related by a non-crystallographic symmetry axis, each identically bound by one VHH. The final structure is completely built, except for four N-terminal residues in LAP1, which seem disordered in the crystal (six N-terminal residues in the second LAP1 copy).

The LAP1 structure clearly has an AAA+ like fold (Figure 1). AAA+ proteins are a functionally diverse group within the vast family of ‘P-loop’-type NTP-binding proteins (Iyer et al., 2004). Typically, AAA+ proteins are characterized by strongly conserved sequence motifs involved in nucleotide recognition. Subcategories within AAA+ proteins are based on distinct secondary structure topologies. The D2 domain of the two-ring AAA+ ATPase ClpB superimposes on LAP1 with an rmsd of 3.28 Å over 111 Cα positions (Figure 1; Zeymer et al., 2014). ClpB belongs to the additional strand conserved E (ASCE) family (Iyer et al., 2004). Like ASCE-type AAA+ proteins, such as ClpA, ClpB, and p97, LAP1 also has a central, five-stranded parallel β-sheet, surrounded on both sides by 10 helices in total (Figure 1A,B). The secondary structure topology is α(−1)α(0)β1α(1)β2-α(2a)α(2b)β3α(3a)α(3b)β4α(4)β5α(5)α(6), using the AAA+ nomenclature (Erzberger and Berger, 2006). In contrast to canonical ASCE-type ATPases (Figure 1C), LAP1 lacks a C-terminal domain. Instead the C terminus is attached to helix α1 via a disulfide bond between the highly conserved residues C424 and C582 (Figure 1—figure supplement 1). In a AAA+ ATPase, the nucleotide binding pocket is the most sequence-conserved region, defined by the essential Walker A and B motifs. LAP1 lacks a recognizable Walker A region, typically located between β1 and α1 (Figure 2A,B, Figure 1—figure supplement 1). Instead, helix α1 is N-terminally extended compared to AAA+ ATPases, thereby sterically blocking the nucleotide-binding pocket. The Walker B motif with the canonical sequence hhhhDE (h, hydrophobic), is instead hhhhHR in human LAP1, and therefore cannot interact with nucleotide. Finally, the conserved disulfide bridge would also interfere with nucleotide binding (Figure 2B, Figure 1—figure supplement 1).

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Having established that LAP1 represents a nucleotide-free AAA+ domain, we asked how this might inform us about LAP1's interaction with Torsins, which can be modeled with great confidence as AAA+ ATPases (Zhu et al., 2010). The four canonical nucleotide-sensing elements, Walker A, Walker B, Sensor 1, and Sensor 2 (Erzberger and Berger, 2006), are well conserved and immediately recognizable in TorsinA when compared to ClpB-D2 (Figure 2C,D, Figure 2—figure supplement 1; Kock et al., 2006; Zhu et al., 2010). AAA+ ATPases are often activated by an arginine residue (‘arginine finger’) in the neighboring protomer in the hexameric ring assembly, such that this arginine is positioned to reach the phosphate binding site (Wendler et al., 2012). While we do not find a conserved arginine in TorsinA (Figure 2—figure supplement 1) at the expected position at the end of helix α5, in LAP1 this residue is a strictly conserved arginine (R563). Consequently, modeling and phylogenetic analysis strongly suggest that LAP1 and TorsinA might form a heterohexameric ring (Figure 3A,B) with three active sites in which TorsinA and its partner subunits alternate to form the ring. This hypothesis is further strengthened by the recent observation that LAP1 and LULL1 activate TorsinA in vitro (Zhao et al., 2013).

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To experimentally confirm assembly of LAP1 and TorsinA into a heterohexameric ring, we co-expressed and co-purified TorsinA(E171Q) in complex with either LAP1 and LULL1 via nickel-affinity and size-exclusion chromatography. We obtained complexes with TorsinA(E171Q):LAP1 and TorsinA(E171Q):LULL1 1:1 stoichiometry (Figure 3—figure supplement 1). The E171Q mutation traps TorsinA in the ATP-bound form, which helps to stabilize the interaction with LAP1 and LULL1, respectively (Goodchild and Dauer, 2005). For negative-stain electron microscopic analysis, the nickel eluate was used directly. On individual micrographs, rings of expected size were observed for the two-complex preparation, but not for individually purified LAP1(356–583) or LULL1(322–570) (Figure 3—figure supplement 2). TorsinA(E171Q) is insoluble without binding partner, therefore the observed rings should not be homomeric (Figure 1—figure supplement 3). The best micrographs, obtained with the TorsinA(E171Q):LULL1 preparation, were further processed. Class averaging of 808 TorsinA(E171Q):LULL1 particles yielded multiple classes showing ring assembly (Figure 3C). The rings have a diameter of ∼120 Å, in good agreement with typical hexameric AAA+ ATPase assemblies.

We then tested the ATPase activity of TorsinA in the context of LAP1 and LULL1. In comparison to wild-type TorsinA:LAP1, the mutants TorsinA:LAP1(R563A) and TorsinA:LULL1(R449A) showed substantial reduction in ATPase activity, while both TorsinA(E171Q):LAP1 and TorsinA(E171Q):LULL1 are essentially inactive (Figure 4). We note that the ATP hydrolysis rates are very slow and that the effect of the arginine finger mutation is not as drastic as seen with other AAA+ ATPases. We speculate that the ATP hydrolysis rate in the presence of substrate is likely higher, and that the effect of the arginine finger is likely more pronounced in a more physiological context.

Figure 4

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In summary, we conclude that LAP1 and LULL1 both activate Torsins by providing the arginine finger in a heterohexameric ring assembly in which LAP1 and LULL1 alternate with Torsin. Since LAP1 and LULL1 contain transmembrane helices immediately N-terminal to their luminal, nucleotide-free AAA+ Activator domain, they bring Torsin into close proximity to the membrane (Figure 5).

Figure 5

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Here we provide structural evidence to explain how LAP1 and LULL1 function as activating proteins for Torsin. This confirms recent work which biochemically showed that LAP1 and LULL1 activate TorsinA (Zhao et al., 2013). A heterohexameric ring assembly for AAA+ ATPases is unusual, but not unprecedented (Gribun et al., 2008; Saffian et al., 2012). However, to our knowledge this is the first description of a heterohexameric assembly where a canonical AAA+ domain alternates with a nucleotide-free AAA+ like domain, which we now call AAA+ Activator domain. It is striking that LAP1 and LULL1 lack all canonical nucleotide-binding elements. This feature has undoubtedly complicated their detection by sequence-based methods. Using the LAP1 structure, we performed a reverse hidden Markov model search to possibly identify structurally similar proteins in various organisms using Backphyre (Kelley and Sternberg, 2009). Interestingly, the top hits that we find besides LAP1 homologs are the Torsins. This indicates that both proteins are possibly derived from a common ancestor. In Drosophila as well as in Caenorhabditis, we find LAP1 homologs that have so far escaped detection (Figure 1—figure supplement 1). Gratifyingly, for all species where so far an orphaned Torsin has been found, we now also find a LAP1/LULL1 homolog (for many species, it is difficult to distinguish between LAP1 and LULL1 based on sequence alone). This lends further support to our model suggesting that a heterohexameric LAP1-Torsin ring is the functionally relevant protein assembly.

Our structure was obtained using a VHH raised against LAP1. We can only speculate as to why VHH-BS1 helped in crystallizing the protein. Perhaps VHH-BS1 fortuitously generates packing contacts important for crystal formation. We note that VHH-BS1 binds to the area immediately surrounding the arginine finger of LAP1, thereby interfering with TorsinA binding (Figure 1—figure supplement 3). Inclusion of VHH-BS1 might prevent cryptic and poorly ordered hexamerization of LAP1 at high protein concentrations and therefore favor a more ordered conformation conducive to crystallization. Interestingly, VHH-BS1 can compete with LAP1 for TorsinA binding in vitro (Figure 1—figure supplement 3). When expressed intracellularly, it might be a useful tool to further characterize TorsinA-LAP1 function in vivo.

What are the substrate(s) of Torsin? Torsin acts in close proximity to the NE or the ER membrane, depending on whether it interacts with membrane-bound LAP1 or LULL1, respectively. Membrane proximity is further ensured by the N-terminal hydrophobic region within TorsinA (Vander Heyden et al., 2011). Using an ATP-trapped mutant, it was shown that TorsinA is involved in the exit of large ribonucleoprotein (mRNP) granules from the nucleus through a pathway akin to the nuclear egress of herpes-type viruses (Rose and Schlieker, 2012; Speese et al., 2012; Jokhi et al., 2013). A substrate-trap mutant of TorsinA localized to the neck of mRNP-filled vesicles that bud from the inner nuclear membrane (INM) into the perinuclear space (Jokhi et al., 2013), consistent with our model of LAP1-mediated activation of TorsinA close to the INM. We therefore speculate that the so far elusive Torsin substrates include proteins involved in membrane scission. This is akin to the action of the AAA+ ATPase Vps4 involved in the scission of narrow membrane necks mediated by ESCRT-III (endosomal sorting complexes required for transport) components (Hill and Babst, 2012; McCullough et al., 2013), but with an important difference: while Vps4 acts on the inside of the necks, Torsins presumably act on the outside. We further speculate that under more physiological conditions, that is, properly complexed with LAP1 or LULL1 and engaged with substrate, the ATPase activity of Torsin might be substantially higher than that reported under the in vitro conditions tested so far.

Apart from the identification of Torsin substrate(s), understanding the catalytic mechanism of the Torsin-LAP1/LULL1 machinery is equally important. Because of the unusual heterohexameric architecture with only three active sites, we expect substantial differences between this system and those that are better understood. One important aspect will be to unravel the role of the highly conserved cysteines, both in LAP1/LULL1 and in Torsin. While both proteins have conserved disulfides, these do not account for all cysteines and suggest a possible redox mechanism, as has been discussed previously (Zhu et al., 2010, 2008). We note that in our model the most conserved cysteine C496 in LAP1 cannot engage in an intramolecular disulfide bridge, but is close to the nucleotide-binding site of the neighboring TorsinA. This remarkable conservation might indicate a role in a redox mechanism that is coupled to the catalytic cycle of Torsin-LAP1/LULL1. The lumen of the endoplasmic reticulum is a site rich in oxidoreductases that assist in protein folding and assembly. The catalytic function(s) of the Torsin-LAP1/LULL1 assemblies may have co-opted part of this machinery.

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Author details

1. Brian A Sosa

   Department of Biology, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

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

   Competing interests

   No competing interests declared.

2. F Esra Demircioglu

   Department of Biology, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

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

   Competing interests

   No competing interests declared.

3. James Z Chen

   Department of Biology, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

   JZC, Acquisition of data, Analysis and interpretation of data

   Competing interests

   JZC: Reviewing editor, eLife.

4. Jessica Ingram

   1. Department of Biology, Massachusetts Institute of Technology, Cambridge, United States
   2. Whitehead Institute for Biomedical Research, Cambridge, United States

   Contribution

   JI, Acquisition of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

5. Hidde L Ploegh

   1. Department of Biology, Massachusetts Institute of Technology, Cambridge, United States
   2. Whitehead Institute for Biomedical Research, Cambridge, United States

   Contribution

   HLP, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

6. Thomas U Schwartz

   Department of Biology, Massachusetts Institute of Technology, Cambridge, United States

   Contribution

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

   For correspondence

   tus@mit.edu

   Competing interests

   No competing interests declared.

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