Truncation constructs coding for the RPV1 TIR domain were prepared by amplification of the corresponding fragments from a plasmid template of MrRPV1 full-length cDNA (Feechan et al., 2013) with primers containing Gateway attB sites, using polymerase chain reaction (PCR). The PCR products were cloned into the donor vector pDONR223 (Invitrogen) using BP clonase. To create protein fusions with yellow fluorescent protein (YFP) tags, the desired entry clone and destination vector (pEarlyGate100-L-YFP) containing attR1 and attR2 sites were recombined by LR reaction. The vector pEarlygate100-L-YFP was derived from pEarlyGate100 (Earley et al., 2006) by the introduction of an AvrII-SpeI GA linker-vYFP fragment from ER082 (Tucker et al., 2012). All MrRPV1 TIR domain mutants were created from the plasmid of the entry clone with the QuikChange Site-Directed Mutagenesis Kit (Agilent Technologies) according to the manufacturer’s instructions. The desired mutations were recombined into pEarlygate100-L-YFP by LR reaction. For yeast-two-hybrid (Y2H) studies, RPV1^TIR1-193 was recombined into the Gateway-compatible Y2H vectors based on pGADT7 and pGBKT7 (Clontech) kindly provided by Dr Maud Bernoux (CSIRO Agriculture, Canberra). All constructs were sequenced for verification. All primers used to generate the above constructs are listed in Supplementary Table S1.

Tobacco (N. tabacum cv. White Burley) plants were grown in a greenhouse at the Commonwealth Scientific and Industrial Research Organisation (CSIRO), Adelaide, Australia. Agrobacterium (strain EHA105) cells were in grown in yeast extract peptone (YEP) media supplemented with 25 μg/mL rifampicin and 50 μg/mL kanamycin at 28°C for 2 days. Approximately 0.5 ml culture was inoculated into 50 ml of YM+MES (pH 5.6) media supplemented with 20 μM acetosyringone and 50 μg/mL kanamycin and grown at 28°C until the OD_{600 nm} was >0.5. Cells were pelleted and resuspended in infiltration medium (10 mM MgCl₂, 10 mM MES pH 5.6, 200 μM acetosyringone) to give a final OD_{600 nm} of 0.5 and incubated at room temperature for 2–3 h. Infiltration of rapidly expanding tobacco leaves was carried out with a blunt needleless syringe. Plants were maintained at a constant temperate of 23°C for 48 h before being transferred to the glasshouse.

Yeast transformation and growth assays were performed essentially as described in the Yeast Protocols Handbook (Clontech). Yeast cells (AH109) were co-transformed with the prey and bait vectors using a lithium acetate-based protocol. Transformants were first spread on CSM-Trp-Leu plates, and then co-transformed positive colonies were grown on CSM-Trp-Leu-His + 5 mM 3-AT plates to detect activation of the HIS3 reporter gene. Prey and bait vectors containing the L6^TIR29-233 (Bernoux et al., 2011) were used as a positive control. Empty prey and bait vectors were used as a negative control.

Leaf tissue (∼1 cm diameter disk) was collected from the middle of an infiltrated area 48 h after agroinfiltration. Tissue was ground up in 100 μl of 2X Laemmli extraction buffer, centrifuged at 23,000 x g for 10 min at 4°C and the supernatant fraction collected. Proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes (Pall). Membranes were blocked in 5% skim milk and probed with anti-GFP (Roche) followed by goat anti-mouse antibodies conjugated with horseradish peroxidase (Thermo Scientific). Labeling was detected with a SuperSignal West Pico chemiluminescence kit (Thermo Scientific) following the manufacturer’s instructions. Membranes were stained with Ponceau S staining solution for protein loading.

RPV1^TIR20-193 and RPV1^{TIR20-193 H42A} were amplified from plasmid templates described above using the primer combinations RPV1^TIR20-F and RPV1^TIR193-R (Supplementary Table S1). The PCR product was cloned into the Escherichia coli expression vector pMCSG7 (Stols et al., 2002) by ligation-independent cloning and verified by sequencing. The proteins were expressed in E. coli BL21 (DE3) cells using the autoinduction method (Studier, 2005). All media was supplemented with 100 μg/mL ampicillin for plasmid selection. An overnight culture was used to inoculate large-scale cultures. Cells were grown by continuous shaking at 37°C in 2 L flasks containing 500 mL of media until the OD_{600 nm} reached 0.6–0.8. At this point, the temperature was reduced to 20°C and the cells were grown for an additional 18 h before harvesting by centrifugation.

Cells expressing the protein of interest were resuspended in a lysis buffer containing 50 mM HEPES (pH 8.0), 300 mM NaCl and 1 mM DTT. The cells were lysed using sonication, clarified by centrifugation and the resulting supernatant was applied to a 5 mL HisTrap FF column (GE Healthcare). The column was washed with the lysis buffer supplemented with 30 mM imidazole to remove non-specifically bound proteins. The bound protein was eluted using a linear gradient of imidazole from 30 to 250 mM. Fractions containing the protein of interest were pooled, concentrated and buffer-exchanged (to remove imidazole) into 50 mM Tris pH 8.0, 250 mM NaCl, 1 mM DTT and 0.1 mM EDTA for overnight treatment with His-tagged TEV protease at 4°C. The cleaved protein was reapplied to the HisTrap FF column to remove the histidine tag, TEV protease and other contaminants and the flow-through was concentrated and separated further on a Superdex 75 HiLoad 26/60 gel-filtration column (GE Healthcare) pre-equilibrated with the gel filtration buffer containing 10 mM HEPES (pH 7.5), 150 mM NaCl and 1 mM DTT. The peak fractions were pooled and concentrated to 10 mg/mL. Protein was stored in aliquots at -80°C for future biophysical and crystallization studies.

Initial screening was prepared using a Mosquito robot (TTP LabTech, UK) in a 96-well format. The hanging drop vapor-diffusion method of crystallization was used, and drops consisting of 100 nl protein solution and 100 nl reservoir solution were equilibrated against 80 μl reservoir solution. Eight commercial screens were utilized: Index, PEG/Ion and PEGRx (Hampton Research), Morpheus, ProPlex, JCSG Plus, PACT Premier (Molecular Dimensions) and Precipitant Synergy (Jena Biosciences). A number of single crystals were observed in the screening plates and these were harvested directly from the screens and after cryo-protection with 20% glycerol in well solution the crystals were vitrified in liquid nitrogen. The crystals were subjected to X-ray radiation at the Australian Synchrotron MX1 beamline. X-ray diffraction data to approximately 2.3 Å resolution was obtained from a crystal (approximate dimensions 200 μm × 30 μm × 30 μm) harvested from the PEG/Ion screen, grown in condition 16% PEG3350, 0.1 M Tris pH 8.5 and 2% Tacsimate. Data was collected using a wavelength of 0.9537 Å with a detector distance of 200 mm. The resulting dataset was indexed and integrated with XDS (Kabsch, 2010) and scaled with Aimless (Evans and Murshudov, 2013). Molecular replacement was performed using the program phaser (McCoy et al., 2007) using the RPS4^TIR structure (PDB ID 4c6r) as a template. Automated model building was performed in the Phenix package using autobuild, while phenix refine combined with manual inspection and corrections using coot (Emsley et al., 2010) were combined to produce the final atomic model.

Multi-angle laser light scattering (MALS) and small-angle x-ray scattering (SAXS) were used in conjunction with size-exclusion chromatography (SEC) to assess the molecular mass of RPV1^TIR20-193 in solution. SEC-SAXS was performed at the SAXS/WAXS beamline of the Australian Synchrotron. A Pilatus 1M detector at a sample-to-detector distance of 1.6 m and an energy of 12 keV yielded data over a q-range of 0.007–0.361 Å^-1, where q=4π.sin (θ)/λ. 1 mg of both RPV1^TIR and RPV1^TIR20-193H34A were separated over an inline 3 mL Superdex S200 5/150 GL Increase column (GE Healthcare) at 16°C, at a flow rate of 0.2 mL/min in 10 mM HEPES (pH 7.5), 150 mM NaCl buffer with 1 mM DTT. Frames were collected in 2 s exposures. Data reduction and subtraction was performed using scatterBrain¹. 100 frames immediately preceding each peak were summed and normalized to obtain buffer blanks, which were subtracted from each individual frame across the peak. Values of R_g and I(0) were calculated from each frame via Autorg in the PRIMUS suite (Petoukhov et al., 2012), and molecular masses were calculated from the volume of correlation for points where 0 < q < 0.3 Å^-1 (Rambo and Tainer, 2013).

To gain insight into whether the AE interface is conserved across a wider sample of plant species, TIR domain-annotated sequences from Pfam version 29.0 (Finn et al., 2016) were aligned and used to construct profile Hidden Markov Models (profile HMMs). These profile HMMs were then used to search plant genomes from the more extensive Phytozome database (Supplementary Figure S1). Pfam sequences were mapped to four clades and four individual sequences within Phytozome in order to include representative sequences from a greater number of plant species than present in Pfam (Supplementary Figure S2). We were able to uncover a larger set of TIR domains by expanding the search from Pfam to Phytozome, except in the cases of Arabidopsis thaliana and V. vinifera, in which mapping to the Phytozome database resulted in fewer proteins than originally identified within Pfam (Supplementary Table S2 and Supplementary Data Sheet 1).

The Pfam description of TIR domains (PF01582) does not always account for the entire sequence of the domain, judging from the available structures, with sequences often truncated at the C-terminus of the domain (i.e., missing the αE helix). To address these errors, the full-length protein sequences for each Pfam entry were retrieved from UniProt (UniProt, 2015).

When an individual sequence had multiple annotated TIR domains, it was divided into regions that maximized the length of each region. If an individual region’s new length was below 100 amino-acids, it was deemed unlikely to represent a true TIR domain and was re-joined with the neighboring region in the sequence. If two or more neighboring region’s new lengths were greater than 100 amino-acids the original sequence was split at the boundary and the regions were included as separate sequences within the set.

The previously reported AE interface within RPS4 (Williams et al., 2014) was used as a reference to exclude sequences that did not contain the αA and αE regions. As RPS4 was only present within the Malvids clade and Pentapetalae clade, for all other sets RPS4 was manually added and aligned to identify a new sequence that aligned to RPS4 in the AE interface regions and could serve as a reference sequence for its set. After identification of this reference sequence, RPS4 was removed from the alignments for subsequent analysis.

The sequences were clustered using CD-HIT (Li and Godzik, 2006) at 100% identity to identify any duplicate sequences and remove them. Sequences were aligned using MAFFT (Katoh and Standley, 2013) with the default FFT-NS-2 strategy. Any sequence in the alignment that had less than 50% of positions containing an amino-acid in either the αA or αE regions defined by the reference sequence were tentatively excluded. The remaining sequences were realigned and this exclusion process was repeated until no sequences were present in the alignment that had less than 50% of positions containing an amino-acid in either of these regions.

Excluded sequences were checked for the possibility of truncation of the UniProt record. A BLAST search was conducted that only accepted hits longer than the query sequence but that otherwise exactly matched the query sequence. The results from this BLAST search were realigned to the set of sequences without 50% or higher gaps in the αA or αE regions, and all sequences with 50% or higher gaps were definitively excluded.

The remaining sequences within these clades were aligned and used to build four profile HMMs specific to their containing sequences. To identify TIR domains in species not annotated by Pfam, the characteristic profile HMMs were used to query sets of predicted proteins from primary transcripts of genomes found in the Phytozome database. The primary transcripts were from clades containing the original sets of species used to derive each profile.

A profile HMM was constructed using HMMER 3.1b2 (Eddy, 2011) and used to search the relevant subset of the Phytozome database. Hits from the profile HMM search of the Phytozome databases with an E-value cut-off of 1e-5 were used to construct a multiple sequence alignment following the previous protocol of clustering at 100% identity with CD-HIT, identification of a reference sequence, alignment with MAFFT FFT-NS-2 strategy, and exclusion of sequences missing characters at over 50% of the reference sequence’s designated αA or αE regions. The resulting multiple sequence alignments were trimmed of columns that had gaps in 75% or more of the sequences, and visualized as sequence logos. Note that residues at positions 18–20 representing the start of the αE region are presented here in the sequence logos for completeness; however, there is a higher presence of gaps in the alignments at positions 18–20 and alignment error could account for the differences of amino-acids at these positions.

In order to highlight patterns occurring at the species level, individual species from the four clades were chosen—A. thaliana, Glycine max, P. persica, Populus trichocarpa, and V. vinifera. The same search procedure as for the larger sets was repeated for each of these species. Alignments and profile HMMs were constructed specific to each individual species and the subsets of the Phytozome database used for the more extensive search were restricted to sequences from each individual species.

The RPV1 gene from M. rotundifolia encodes a TIR-NB-LRR protein that confers resistance to the oomycte pathogen P. viticola (Feechan et al., 2013). Based on the protein structures of RPS4^TIR and L6^TIR (Bernoux et al., 2011; Williams et al., 2014), the TIR domain is predicted to be located between residues 20 and 188 (Figure 1A). As TIR-domain autoactivity has been demonstrated for a number of plant NLR proteins (Frost et al., 2004; Swiderski et al., 2009; Krasileva et al., 2010; Bernoux et al., 2011), we tested whether the RPV1 TIR domain was also autoactive. Agroinfiltration experiments confirmed that RPV1^TIR1-193 and RPV1^TIR20-193 were capable of causing rapid cell death in N. tabacum (Figures 1B,D). To determine the minimal functional region of RPV1 required for autoactivity, we tested a series of RPV1^TIR truncated fragments fused to YFP for autoactivity in tobacco. The addition of the YFP tag had no effect on RPV1^TIR1-193 autoactivity (Figure 1B). Truncation of the αE helix led to a significant decrease in RPV1^TIR autoactivity (Figure 1B), in agreement with similar results obtained with L6^TIR (Bernoux et al., 2011). However, whereas corresponding truncations in the αE helix of L6^TIR were found to affect protein stability, this was not the case for the truncated RPV1^TIR proteins (Figure 1C). These results strongly support the secondary-structure predictions, suggesting that RPV1^TIR is located between residues 20–188.

On the basis of these results, we expressed the N-terminal residues 20–193 of RPV1 (designated RPV1^TIR20-193) in E. coli and purified it to homogeneity. Crystals of RPV1^TIR20-193 diffracted x-rays to 2.3 Å resolution and the structure was solved by molecular replacement (Table 1, Figure 2). The RPV1^TIR20-193 structure resembles closely the AtTIR (PDB ID 3jrn), L6^TIR (3ozi) and RPS4^TIR (4c6r; Chan et al., 2010; Bernoux et al., 2011; Williams et al., 2014) structures with an overall Cα RMSD (root-mean-square-distance) value of ∼1.2, ∼1.3, ∼1.6 Å (for 141, 147, 148 superimposed residues), respectively. Within the asymmetric unit, we observe a dimer (Figure 2A) that resembles the heterodimer of the TIR domains from Arabidopsis RRS1 and RPS4 (Williams et al., 2014). This interaction involves the αA and αE helices and the loop regions that precede both helices (Figure 2A); we consequently define this protein-protein interface as the AE interface. Analysis of the RPV1^TIR structure using the program PISA (Krissinel and Henrick, 2007) identified that the AE interface was the largest crystal lattice contact, contributing a combined buried surface area of ∼1340 Ų (∼670 Ų from each molecule). This is similar to the combined ∼1300 Ų buried surface in the RRS1^TIR:RPS4^TIR heterodimer (Williams et al., 2014). Within the AE interface, 17 surface-exposed amino-acids from each RPV1^TIR20-193 monomer are buried within the dimer and contribute to the interaction (Figure 2B). At the core of the interface, His42 forms an important stacking interaction between the two monomers and hydrogen bonds with Glu170 from the opposing monomer (Figure 2C). Hydrogen bonding between Asp41 and Ser171 is also prominent and these equivalent residues were also important in the RPS4^TIR:RRS1^TIR interaction (Williams et al., 2014).

Based on the analysis of the crystal structure of RPV1^TIR20-193, we were interested to understand further the potential role of the AE interface in RPV1 function. To do this, we generated mutations to specific residues in the core and peripheral regions of the AE interface of RPV1^TIR1-193 (fused to YFP) and assessed the impact of these mutations on autoactivity. Mutations of residues His42 and Asp41 to alanine within the core of the AE interface, were found to abolish and markedly reduce, RPV1^TIR1-193-YFP mediated cell death in tobacco. Mutation of R49 to alanine at the periphery of the interface had no effect on RPV1^TIR1-193 autoactivity (Figure 2D), while mutation of R36 to alanine, which is also on the periphery of the interface, abolished cell death.

Importantly, these results could not be explained by differences in protein stability. Indeed, levels of RPV1^TIR1-193-YFP protein recovered from tobacco-leaf tissue agroinfiltrated for the loss-of-function mutants (R36A, D41A and H42A) appeared higher, 48 h post-infiltration, than from the constructs that displayed strong necrosis at day 5 (Figure 2E), potentially due to the impact of cell death on protein yield from agroinfiltrated sectors. These results demonstrate that residues R36, D41, and H42 within the AE interface play a key role in the RPV1^TIR signaling.

The crystal structure of L6^TIR revealed an interface that is spatially distinct to the AE interface, involving the αD and αE helices and the βE strand. In L6^TIR, this region was shown to be important in mediating L6^TIR self-association, which is a requirement for the cell death signaling function of the L6^TIR (Bernoux et al., 2011). To investigate if residues in this region affect RPV1^TIR signaling, we generated mutations P121Y, R125A and G161R (Figure 3A); equivalent to P160Y, R164A and G201R in L6^TIR, and found that these mutants were compromised in their ability to mediate cell death in tobacco (Figure 3B). In the case of L6^TIR, cell-death signaling was shown to be compromised when residues outside the L6^TIR self-association interface were mutated (Bernoux et al., 2011). We also observed disruption of cell-death signaling in W94A and C95S RPV1^TIR1-193-YFP constructs (equivalent to W131A and C132S in L6^TIR), consistent with findings for L6. Importantly not all mutations to surface-exposed residues disrupted cell-death signaling, as demonstrated by the fact that mutation of the non-conserved L108 to a valine did not affect the protein (Figures 3A,B). All mutated RPV1^TIR1-193YFP proteins were detectable by western-blot analysis (Figure 3C), suggesting that these results are not influenced by in planta protein stability.

Homo- and hetero-meric TIR:TIR domain interactions are responsible for the biological functions of TIR domains (Ve et al., 2014). Interactions have been observed in the TIR domains from RRS1, RPS4, and L6 by both Y2H assays and by studies in solution with recombinant proteins (Bernoux et al., 2011; Williams et al., 2014). For RPV1^TIR1-193, we did not observe an interaction in Y2H assays (Supplementary Figure S3) under the conditions tested, despite observing self-association of L6^TIR29-233, which was consistent with previous observations (Bernoux et al., 2011). Solution studies using SEC-coupled MALS and SAXS suggested that only limited self-interaction of RPV1^TIR20-193 may be present (Figure 4). The average SEC-MALS-derived molecular mass of RPV1^TIR20-193 was 22.3 kDa, while the theoretical molecular mass of RPV1^TIR20-193 is 20.7 kDa. We also tested, by SEC-MALS, the recombinant RPV1^TIR20-193 protein carrying an alanine mutation at position H42 (RPV1^{TIR20-193H42A}). The equivalent mutation in RPS4^TIR had previously been shown to disrupt RPS4^TIR:RRS1^TIR dimerisation and inhibited RPS4^TIR self-association. This analysis produced a molecular mass of 20.8 kDa, slightly lower than that of RPV1^TIR20-193 (Figure 4A). Using SEC-SAXS, the averaged molecular masses observed for both proteins were lower than masses determined by SEC-MALS, 19.6 kDa for RPV1^TIR20-193 and 18.7 kDa for RPV1^{TIR20-193H42A}, and small shifts in elution time between RPV1^TIR20-193 and RPV1^{TIR20-193H42A} were also observed (Figure 4B). Molecular masses higher than monomer were previously observed in L6, RPS4, and RRS1 by SEC-MALS (Bernoux et al., 2011; Williams et al., 2014) and in these cases, this behavior could be convincingly linked to self-association. However, in the case of RPV1^TIR, the absolute difference in molecular mass is small and may be beyond the sensitivity of these scattering techniques. Therefore, while these data do not exclude the possibility that the RPV1^TIR domain can self-associate, a link to this function cannot be made based on the Y2H and in-solution experiments presented here.

We previously highlighted the potential conservation of surface-exposed residues in the AE interface after defining its role in RPS4 and RRS1 defense signaling (Williams et al., 2014). In light of our findings presented here for RPV1, we embarked on a more extensive analysis to characterize the conservation of the AE interface across a broader range of plant species. We created species-specific profiles to capture >2000 TIR domain-containing sequences from across 29 plant species and used profile HMMs to create and inspect TIR-domain sequence logos across this wide species range with a focus on the AE interface (See Supplementary Table S2, Supplementary Data Sheet 1, and Materials and Methods). Sequences identified from Pfam were collated into four clades for comparison based on the species tree from Phytozome (Supplementary Figure S2): Malvids, containing A. thaliana, A. lyrata, Brassica rapa, and Eutrema salsugineum; Fabales, containing G. max and P. persica; Malpighiales, containing P. trichocarpa and Ricinus communis; and Pentapetalae, that contained all of the three previous sets, plus Solanum lycopersicum, S. tuberosum, and V. vinifera—representing all classified TIR domain sequences from Pfam within the Pentapetalae clade (Figure 5B). We also undertook a more specific analysis into five individual species to highlight residue differences occurring at the species level (Figure 5C).

Conservation of residues across a wide range of species in the αA and αE regions showed that the αA region (positions 1–17) was more conserved than the αE region (positions 18–30). A number of hydrophobic residues are highly conserved, including positions 7, 11, 15, 26, 29, and 30 (Figures 5A–C). These residues are mostly buried and appear to assist in stabilizing the positioning of the αA and αE helices. The well-conserved position 26 (RPV1 Ile174) is surface-exposed and contributes to a pocket that in the RPV1^TIR20-193 structure is occupied by His42 from the interacting RPV1^TIR20-193 protomer. The most important surface-exposed residues in the AE interface are at positions 9, 10, and 22, which form the core of the AE interface (Figures 5A and 2C). This analysis reveals that at these positions, there is conservation of histidine and glutamate at position 10 and 22, respectively (RPV1 His42 and Glu170). The amino-acid distribution at position 9 (RPV1 Asp41) is more varied across the clades (Figure 5B), with evidence of species-specific conservation, for example the conservation of serine in A. thaliana (Figure 5C). There is also a general trend for position 23 to contain a small amino-acid such as alanine or serine. While histidine at position 10 is well conserved, there is minor representation of aromatic residues including phenylalanine and tyrosine. Interestingly, in the case of AtTIR, a phenylalanine occupies this position, indicating that phenylalanine is compatible with dimerisation through the AE interface, at least within the context of AtTIR crystals. At the species level, G. max (soybean) appeared to deviate most significantly of the species compared across the important AE interface residues. G. max appears to have a similar preference for histidine and asparagine at position 10 (Figure 5C) and at position 22, the negatively charged residue glutamate is replaced by aspartate and glutamine in some instances. In general, there is significant variation in other interface interacting residues at both the clade and species levels, with the exception of position 4, which is essentially an invariant arginine.