1. Molecular cloning

Hts and Mud sequences were amplified from a cDNA library prepared from Drosophila S2 cells. Additional aldolase enzyme and domain genes were amplified from plasmids constructed through complete gene synthesis by Genewiz (Azenta Life Sciences, South Plainfield, NJ, USA). All aldolase enzyme and domain sequences were cloned into the pMAL-C5x plasmid using 5’-NdeI and 3’-SalI restriction sites. The Mud^CC domain was cloned into pBH with an N-terminal TEV-cleavable hexahistidine tag using 5’-BamHI and 3’-XhoI restriction sites.

2. Protein purification

All proteins were expressed in BL21(DE3) E. coli under induction of isopropyl β-D-1-thiogalactopyranoside (IPTG) and grown in standard Luria–Bertani broth supplemented with 100 μg/ml ampicillin. Transformed cells were grown at 37°C to an OD₆₀₀ ∼0.6 and induced with 0.2 mM IPTG overnight at 18°C. Cells were harvested by centrifugation (5000 × g for 10 min), and bacterial pellets were resuspended in lysis buffer and flash-frozen in liquid nitrogen. Cells were lysed using a Branson digital sonifier and clarified by centrifugation (12,000 × g for 30 min).

For 6 × His-tagged Mud^CC, cells were lysed in N1 buffer (50 mM Tris pH8, 300 mM NaCl, 10 mM imidazole) and coupled to Ni-NTA resin (Thermo Fisher Scientific, catalog #88222) for 3 h at 4°C. Following extensive washing, protein was eluted with N2 buffer (50 mM Tris pH8, 300 mM NaCl, 300 mM imidazole). The 6 × His tag was removed using TEV protease during overnight dialysis into N1 buffer. Cleaved product was reverse affinity purified by a second incubation with Ni-NTA resin and collection of the unbound fraction. Final purification was carried out using an S200-sephadex size exclusion column equilibrated in storage buffer (20 mM Tris pH8, 200 mM NaCl, 2 mM DTT).

For all MBP-tagged aldolase proteins, cells were lysed in Phosphate Buffered Saline (PBS), and lysate was then clarified by centrifugation (12,000 × g for 30 min) and immediately flash frozen in liquid nitrogen for storage at −80°C. Proteins were isolated by coupling to amylose resin followed by extensive washing prior to their direct use in pulldown assays (see below).

3. Pulldown assays

MBP-fused aldolase proteins were absorbed to amylose agarose for 30 min at 4°C and washed three times in wash buffer (20 mM Tris, pH 8, 100 mM NaCl, 1 mM DTT, and 0.2% Triton-X100) to remove unbound protein. Subsequently, soluble Mud^CC prey protein was added at varying concentrations for 2 h at 4°C with constant rocking in wash buffer. Incubation for different times (e.g., 1 or 3 h at 4°C, or 1 h at room temperature) produced similar results, indicating that this experimental framework had established equilibrium binding conditions. Reactions were then washed four times in wash buffer, and resolved samples were analyzed by Coomassie blue staining of SDS-PAGE gels. All gels shown in figures are representative of 3–5 independent experiments. Each independent experiment measured a single data point for indicated conditions. Data shown in binding isotherms represent the mean standard ± deviation for measurements from the 3–5 replicates examining the specific indicated concentration and binding conditions.

All interactions were quantified using ImageJ software. Briefly, gel images were converted to greyscale and individual band intensities were measured using the boxed ‘Measure’ analysis tool. The size of measurement box was kept the same across all concentrations. To ensure consistent measurements of bound proteins across experimental conditions, the measurement box was set by the size of the MBP-fusion protein bait. Binding curves shown in figures plot measured band intensities (expressed as arbitrary units, ‘AU’) for bound Mud protein calculated by subtracting background signal quantified from the zero added Mud control in each experiment. Dissociation binding constants were calculated in GraphPad Prism using a one-site binding isotherm regression analysis. All plots and statistics were also performed in Prism.

4. Structural modeling

Protein structural modeling was primarily performed using the AlphaFold server (www.alphafoldserver.com) utilizing the current AlphaFold 3 code [12]. Primary protein sequences were input along with the desired copy number (e.g., ‘4’ for prediction and modeling of tetrameric assemblies). Confidence metrics were interpreted for model accuracy. Monomeric structures were assessed using predicted template modeling (pTM) scores; tetrameric structures were assessed using this along with the interface predicted template modeling (ipTM) scores, both of which assess the accuracy of the entire structure [13,14]. All models presented scored above the 0.6 threshold for likelihood, with most all scoring above the 0.8 threshold for high-quality predictions. Models were further interpreted and shown images were rendered in PyMOL (www.pymol.org).

Two additional models of the Hts ALDO^DOM tetramer were constructed for comparison and validation of the AF3 model. First, Rosetta Comparative Modeling (RosettaCM) was used to generate an Hts homology model based on the porcine Adducin heterotetramer [8,15]. Modeling was performed using the Robetta server (http://robetta.bakerlab.org), where Hts amino acids 110–401 were input for sequence alignment and subsequent modeling against the PBDid 8IAH template. The SWISS-MODEL homology modeling server (https://swissmodel.expasy.org) was used to generate the second Hts homology model, here using the Pseudomonas syringae aldolase enzyme homotetramer (PDBid 3OCR) as a template. Hts residues 110–401 were input, and a template search was performed. This search identified 8IAH as the top hit, followed by the 3OCR structure with a GMQE score of 0.64, which was selected as the input template.

5. Molecular Dynamics (MD) simulations

All MD simulations were performed by Creative Proteomics (Shirley, NY, USA) using the GROMACS MD simulation engine [16]. Simulations were performed in triplicate using a Linux GPU cluster. The AF3-generated Hts tetramer model was prepared for analysis using the PDB2PQR tool by adjusting pH to 8 to match buffer conditions of the biochemical pulldown experiments. Similarly, counter ions were added, and the system was solvated with 100 mM NaCl. Additional parameters were set for 1 bar pressure and 4°C temperature. MD simulations were performed for 200 ns using Leap-Frog integrator and results were analyzed using GROMACS analysis tools.

6. Sequence database analysis

Database searches were performed using BLAST (www.blast.ncbi.nlm.nih.gov). Information related to the sequences used as inputs for respective aldolase class searches can be found in Supplementary S1 Table. Class I and II enzymes are highly divergent in sequence, easily allowing distinctions of the Class I subgroup [4]. Class II enzymes were confirmed by AlphaFold modeling and structural alignment to that of the query sequence. For all sequences inferred as class IIa enzymes, identified proteins were not part of larger sequences typical of Adducin genes and were subsequently modeled using AlphaFold with a Zn²⁺ ion to demonstrate predicted binding at the conserved histidine triad conserved within the active site of bona fide class II aldolase enzyme [17]. Class IIa/adducin domains were identified as those located at the N-terminal region of proteins also containing sequence C-terminal to the aldolase sequence that most likely constitutes additional Adducin-specific function [7]. Phyla searched were selected as those representing the three kingdoms forming the tree of life along with additional ones at significant positions in animal evolution [18]. All multiple sequence alignments were performed using ClustalW (https://www.genome.jp/tools-bin/clustalw), and images shown were constructed in SnapGene software (https://www.snapgene.com).

Structural modeling of Hts^ALDO domain reveals a distinct loop insertion in its CThelix

The recently determined cryo-EM structure of Adducin bound to a spectrin-actin complex found a heterotetrameric α/β-Adducin ALDO^DOM associated with the barbed actin end (PDBid: 8IAH; [8]). This tetrameric assembly, as well as folding of individual monomers, shares high homology with those of a subgroup of bacterial aldolase enzymes (e.g. PBDid: 3OCR; RMSD = 0.735 for tetramer alignment; Fig 1). To examine how the ALDO^DOM of Hts compares, we constructed a tetrameric model using AlphaFold 3 (AF3; [12]), with an input sequence of Hts amino acids (residues 110–401) that align with those that constitute the complete aldolase fold in α-Adducin [8]. Drosophila contain only a single Adducin gene that corresponds to α-Adducin in mammals, and thus the AF3 model generated constitutes an Hts homotetramer. This AF3 model showed a high overall confidence (ipTM = 0.87; pTM = 0.88), with a majority of positions within the ‘Very high’ (pIDDT > 90) confidence range apart from an extended loop within the final, ‘C-terminal helix’ (CThelix) of each aldolase domain (Fig 1A). This model showed high homology to both the mammalian Adducin domain and bacterial enzyme tetramers (Fig 1B,1C). Results of AF3 modeling thus demonstrate that the Hts aldolase domain is structurally conserved with that of mammalian Adducin as well as the subclass of bacterial aldolase enzymes, both in its globular fold and tetrameric assembly. This structural model will serve as the basis to further explore how the Hts aldolase domain binds its partner protein Mud and for comparison with the Adducin/actin complex.

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Fig 1. AlphaFold modeling identifies an extended CThelix loop in a tetrameric Hts aldolase domain assembly.

(A): Left: AF3 modeling output results for the Hts tetramer color coded by per-atom confidence estimate (pLDDT) on 0-100 scale (Blue – Very high: pLDDT > 90, Cyan – Confident: 90 > pLDDT > 70, Yellow – Low: 70 > pLDDT > 50, Orange – Very low: pLDDT < 50). The core aldolase fold in each of the four Hts monomers contains a majority of ‘Very high’ scoring positions, with a minor fraction of ‘Confident’. In contrast, the extended CThelix loop showed scores in the ‘Low’ and ‘Very low’ categories. Right: Predicted alignment error plot shows the expected positional error for each residue, also indicating a high confidence level (dark green) in model accuracy apart from the CThelix loop positions (light green). (B): Multiple sequence alignment of CThelix sequences from bacterial aldolase enzymes (P. syringae and Pseudomonas) and aldolase domains from pig (Sus scrofa, S.scrofa), human (Homo sapiens, H.sapiens), and fruit fly (Drosophila melanogaster, D.mel) demonstrates the extended insert sequence in fly Hts domain that occurs between the N- and C-terminal regions of the CThelix. Also indicated are positions conserved among enzymes and domains (dashed green boxes) and those only conserved among domains (dashed red boxes). (C): Overlay of fly Hts (bluegreen) and porcine Adducin (black) tetramers shows a high degree of similarities, with an RMSD = 0.979, calculated from a superimposition alignment in PyMOL. (D): Overlay of fly Hts (bluegreen) and bacterial aldolase enzyme (white) tetramers shows a high degree of similarities, with an RMSD = 0.882. (E): Overlay of Hts, Adducin, and aldolase enzyme monomers indicating conserved YLE motif positions (numbered according to fly Hts) that are positioned adjacent to the bacterial enzyme active site containing the histidine triad (‘His triad’; white sticks) and catalytic Zn²⁺ (magenta). (F): Overlay of Hts, Adducin, and aldolase enzyme monomers demonstrating the conserved structure of the N- and C-terminal segments of the CThelix (‘CThelix_N’ and ‘CThelix_C’, respectively) as well as the divergent loop insert segment (‘CThelix_loop’). Fly Hts contains a large CThelix loop, Adducin shows a smaller insert, and the bacterial enzyme lacks this segment entirely.

https://doi.org/10.1371/journal.pone.0316787.g001

To corroborate the AF3 structure, we generated additional Hts tetramer assemblies using homology modeling approaches. Rosetta Comparative Modeling (RosettaCM) was used to generate a model from the cryo-EM structure of porcine Adducin heterotetramer as a template [8,15]. An additional homology model was generated using SWISS-MODEL from the crystal structure of Pseudomonas syringae Aldolase enzyme homotetramer as a template [PDBid 3OCR; [19]]. Both homology models showed strong overall similarity to the AF3-generated structure (RMSD = 1.179 and 1.507, respectively; S1A, S1C Fig). However, the models showed divergent conformations of the extended CThelix loop, consistent with the lower confidence predictions within this region and its potentially flexible nature (see further discussion below).

We also performed molecular dynamics (MD) simulation analysis to evaluate the stability to the AF3 structure. Triplicate simulations each demonstrated that the tetramer is stable, with all four subunits remaining intact throughout the simulation (S2 Fig). RMSD for the Ca backbone equilibrated quickly and stabilized at an average 3–3.5 Å over the 200 ns simulation (S2D Fig). Radius of gyration (Rg) values remained stable between 3.24–3.26 nm, as did solvent accessible surface area (SASA) values (S2E-S2F Fig). Comparison of the three simulated structures with the AF3 model revealed overall alignment RMSDs of 1.646, 1.486, and 1.769 Å (S2A-S2C Fig). Alterations in the CThelix conformation, particularly the extended loop insert, were observable in each simulated structure suggesting a higher intrinsic flexibility of this region (S2A-S2C Fig). Overall, from these combined modeling and simulation approaches, we conclude that the Hts ALDO^DOM forms a tetrameric assembly with high structural homology to related aldolase fold proteins.

As mentioned, each Hts domain monomer has a large, conspicuous loop within its CThelix (Fig 1). Alignment of single monomers demonstrated that Adducin has a smaller loop insertion, and the bacterial enzyme has no significant loop but instead a short break in an otherwise contiguous CThelix (Fig 1B-1D). AF3 modeling of the bacterial enzyme was able to accurately place the active site catalytic Zn²⁺ ion (Fig 1D). In contrast, neither Adducin nor Hts modeling showed such placement, consistent with divergence in the catalytic site and loss of enzymatic function in transition to aldolase domains [7]. Overall, these results demonstrate the conserved nature of the core aldolase fold, highlight a marked difference in the CThelix, and substantiate the accuracy of AF3 modeling for both enzymes and domains.

The CThelix loop regulates Mud^CC direct interaction with the Hts^ALDO domain

Having identified the CThelix as the most structurally divergent element in aldolase proteins, we next inspected the role for this Hts sequence in regulating Mud binding. To do so, we immobilized Maltose-binding protein (MBP)-fused Hts proteins on agarose resin and co-incubated with purified Mud^CC domain, which is responsible for direct Hts interaction (Fig 2A). The full-length Hts ALDO^DOM strongly bound Mud with low-micromolar affinity, similar to our previous studies (Fig 2B; [11]). Deletion of the large CThelix loop insertion resulted in a significant loss of Mud binding, as quantified by a reduction in the maximal binding (B_max), with minimal effects on the affinity of this diminished interaction efficacy (Fig 2C). Note that for a complete list of mutants tested in this study, along with their evolutionary significance and functional impact, see Table 1. This result suggests that Mud binding could occur directly within this loop insertion. However, truncation of the entire CThelix, including the loop insertion, surprisingly restored complete Mud binding comparable to the full-length ALDO^DOM (Fig 2D), indicating that the binding site occurs within the minimal aldolase core fold. Taken together (Fig 2E), these data suggest a possible regulatory role for the CThelix in Mud binding, with the loop insertion playing an important but indirect part of the interaction.

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Table 1. List of key mutants examined together with their evolutionary significance and functional impact.

https://doi.org/10.1371/journal.pone.0316787.t001

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Fig 2. The CThelix regulates direct interaction between Hts aldolase domain and Mud^CC.

(A): Domain diagrams for Mud and Hts proteins are shown (not to scale). Mud (top) contains extended coiled coils at the N-terminus (grey), followed by a shorter, C-terminal coiled coil (CC, purple) that is the focus of the current study. This Mud^CC domain is followed by the Pins-binding domain (PBD, red). Hts (bottom) contains the N-terminal Aldolase domain (bluegreen) that is also the focus of this study, followed by the Neck (dark grey) and disordered Tail (light orange) regions. (B): Representative gel image depicting interaction between the full-length Hts aldolase domain and Mud^CC. MBP-fused Hts [MBP:Hts (110-401)] or MBP alone control were incubated in the absence or presence of increasing concentrations of Mud^CC. Gel shown is representative of 4 independent experiments. (C): Representative gel image depicting interaction between a CThelix loop deletion in the Hts aldolase domain and Mud^CC. MBP-fused Hts [MBP:Hts (110-401)-·CTloop] or MBP alone control were incubated in the absence or presence of increasing concentrations of Mud^CC. Gel shown is representative of 4 independent experiments. (D): Representative gel image depicting interaction between a CThelix truncation in Hts aldolase domain and Mud^CC. MBP-fused Hts [MBP:Hts (110-310)] or MBP alone control were incubated in the absence or presence of increasing concentrations of Mud^CC. Gel shown is representative of 4 independent experiments. (E): Saturation binding curves for all the above experiments demonstrate the effects of CTloop deletion and CThelix truncation mutants on Mud^CC binding to the Hts aldolase domain. Full-length Hts domain (green) shows a robust, dose-dependent binding. Deletion of the CTloop (red) significantly impairs Mud binding, whereas truncation of the entire CThelix (blue) restores strong binding indistinguishable from the full-length domain.

https://doi.org/10.1371/journal.pone.0316787.g002

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Fig 3. Conserved surface exposed Hts residues do not contribute to Mud binding.

(A): Left: Lateral view of Hts tetramer highlighting three conserved surfaces residue pairs targeted for mutational analysis. Right: Rotation depicting the tetramer face with the conserved actin-contacting D310 residue also targeted for mutational analysis. (B): Representative gel images depicting interaction between in the Hts aldolase domain mutants and Mud^CC. Each indicated MBP-fused Hts mutant or MBP alone control were incubated in the absence or presence of increasing concentrations of Mud^CC. Gels shown are each representative of 3-5 independent experiments. (C): Saturation binding curves for Hts aldolase domain wild-type and mutants binding Mud^CC. All mutants showed similar binding dynamics compared to control Hts.

https://doi.org/10.1371/journal.pone.0316787.g003

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Fig 4. CThelix deletion confers Mud binding function to a bacterial aldolase enzyme.

(A): Overlay of the full-length P. syringae aldolase enzyme (PDBid: 3OCR; white with CThelix colored black) tetramer with that of an AF3-generated model following deletion of the CThelix (ΔCThelix; yellow). The ·CThelix enzyme retains a confident prediction for tetrameric assembly (pTM score = 0.7; ipTM score = 0.62) and aligns well to the full-length enzyme structure (RMSD = 1.1). Cyan box and asterisks depict interaction of the CThelix (*) with loops in the aldolase core of its adjacent protomer. (B): Representative gel images depicting Mud^CC interaction with full-length (left) and ΔCThelix (right) aldolase enzyme from P. syringae. Each indicated MBP-fused aldolase enzyme or MBP alone control were incubated in the absence or presence of increasing concentrations of Mud^CC. Gels shown are representative of 3-5 independent experiments. (C): Saturation binding curves for conditions described in (B). Whereas the full-length aldolase enzyme shows only minimal Mud-binding capacity, truncation of the CThelix results in a robust Mud interaction. (D): Similar experiments to those described in (C) were performed using the class II aldolase enzyme from an alternative Pseudomonas species. No apparent Mud binding was observed with the wild-type, full length enzyme; however, deletion of the CThelix also conferred strong Mud binding to this bacterial enzyme.

https://doi.org/10.1371/journal.pone.0316787.g004

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Fig 5. Chimeric aldolase proteins support a key regulatory role for the CThelix in Mud binding function.

(A): Schematic representation of chimeric aldolase proteins shows strategy to swap CThelices (‘CT’) between the Pseudomonas syringae aldolase enzyme (light grey; ALDO^ENZ) and the Hts aldolase domain (black; ALDO^DOM). (B): Saturation binding curves for the full-length aldolase enzyme (ENZ/ENZ^CT; grey circles) and its chimeric version with the Hts CThelix (ENZ/Hts^CT; black squares). Whereas the aldolase enzyme shows only minimal Mud-binding capacity, the chimeric aldolase shows a robust Mud interaction. (C): Saturation binding curves for the Hts aldolase domain (Hts/Hts^CT; black circles) and its chimeric version with the aldolase enzyme CThelix (Hts/ENZ^CT; grey squares). The chimeric aldolase shows a moderately impaired Mud binding compared to the full-length Hts domain.

https://doi.org/10.1371/journal.pone.0316787.g005

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Fig 6. Hts shares a conserved YLE motif with primitive aldolase enzymes that contacts the adjacent protomer CThelix in the tetrameric assembly.

(A): Multiple sequence alignment among several bacterial aldolase enzymes and aldolase domains. Green boxes indicate conserved residues in the YLE motif, whereas red boxes indicate additional conserved residues that neighbor the YLE motif in the core aldolase structure. These residues were tested for contributions to Mud binding (see Fig 7). (B): Left: face view tetrameric assembly overlay for Hts aldolase domain (black; chain A is red, chain B CThelix is green) and the Pseudomonas syringae aldolase enzyme (white; chain A is yellow, chain B CThelix is purple). Right: lateral view of contacts between chain A and B protomers with the conserved, CThelix-contacting Y/F-LE motif residues shown as sticks. (C): Zoomed views of Y/F-LE/CThelix contacts in the aldolase enzyme (left; ALDO enzyme) and aldolase domain (right; ALDO domain) demonstrate the conserved nature of these interactions. Specific contacting residues are indicated.

https://doi.org/10.1371/journal.pone.0316787.g006

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Fig 7. Mutation of the conserved YLE motif impairs Mud binding.

(A): Left: YLE residues were mutated as a triple-alanine mutant (YLEaaa) in either the full-length Hts aldolase domain (Hts401) or the CThelix truncation (Hts310) and tested for Mud binding compared to wild-type (WT). In both cases, the YLEaaa mutant significantly impaired Mud binding. Right: a single alanine mutation at a conserved L139 in the core aldolase domain (Hts310_L139A) also negatively impacted Mud binding dynamics. (B): Left: A double alanine mutation at two conserved positions adjacent to the YLE motif was made in the core domain (Hts310_HTaa). This mutant retained full Mud binding function. Right: Similarly, a single R128A mutation did not impair Mud binding. (C): Triple-alanine mutation of the FLE motif residues in the Pseudomonas syringae aldolase enzyme (3OCR) significantly reduced the gain-of-function Mud binding seen following truncation of the enzyme CThelix. The full-length (FL) enzyme again did not bind Mud. (D): AF3-generated tetrameric assembly of the full-length Hts aldolase domain (white with CThelices colored black). YLE motif and L139 residues, those whose mutation impaired Mud binding, are shown in green, whereas the adjacent residues that were not required for Mud binding are shown in red.

https://doi.org/10.1371/journal.pone.0316787.g007

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Fig 8. Divergent aldolase enzymes do not bind Mud despite lack of a complete CThelix.

(A): Multiple sequence alignment among several bacterial aldolase enzymes (with ‘aldolase’ label and respective PDBid codes where applicable) and aldolase domains (black text). Enzyme sequences shown include fuculose aldolases lacking a CThelix (blue text), those with a partial CThelix (green text), and those with a full CThelix (red text). Also shown are the fructose aldolases with full CThelix sequences tested in previous figures (grey text). Dots indicate positions corresponding to the YLE motif and L139 in the Hts domain, with those species showing conservation in these positions boxed in red. Note that these residues are divergent in fuculose enzymes, particularly those lacking a full CThelix. (B): Images depicting structural superimpositions between the Pseudomonas syringae fructose aldolase enzyme (3OCR; grey in each case) with indicated fuculose aldolases (colors corresponding to that described in panel A). RMSD values are shown for each. Each fuculose aldolase enzyme shares structural homology with the fructose aldolase despite significant primary sequence divergence. (C): Saturation binding curves for indicated fuculose aldolase enzyme interaction with Mud compared to the Hts aldolase domain. Left: the ‘no CT’ enzyme does not bind Mud despite lack of an intact regulatory CThelix. Mutation of its divergent amino acids to YLE does not confer Mud binding. Middle: the ‘half CT’ enzyme also does not bind Mud, and truncation of its partial CThelix (ΔCT) does not confer binding. Right: the ‘full CT’ enzyme shows modest Mud binding relative to Hts and other fuculose enzymes. This binding is moderately improved upon truncation of the CThelix (ΔCT).

https://doi.org/10.1371/journal.pone.0316787.g008

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Fig 9. Sequence database analysis suggest a Placozoan origin for aldolase domain proteins.

(A): Structural representations of each subgroup of aldolase proteins are shown. (B): Dots represent the positive identification of an annotated gene for the indicated aldolase protein in BLAST search analysis. For class IIa enzymes, the open dots represent ‘no CT’, grey dots represent ‘half CT’, and black dots represent ‘full CT’ sequences. Higher eukaryotes generally lack class II enzymes but gain expression of aldolase domain proteins (demarked by dashed horizontal line), whereas prokaryotes and lower eukaryotes lack domains and generally show a more diverse enzyme expression profile. Emergence of the aldolase domain-containing Adducin gene family appears similarly in Placozoa and Porifera (dotted line), with each also retaining class IIa enzyme genes.

https://doi.org/10.1371/journal.pone.0316787.g009

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Fig 10. The Placozoan aldolase enzyme has full Mud binding activity and a CThelix with aldolase domain-like regulatory function.

(A): Overlay of AF3-generated tetramer models for the T. adhearens (Ta) aldolase enzyme (white) and Hts-like domain (green) shows similarities to other aldolase proteins. AF3 accurately models Zn²⁺ ions (magenta) into the conserved histidine triad within the enzyme catalytic site. Multiple sequence alignment shows conservation in the YLE motif (*) in the Ta enzyme and domain compared to fly Hts and Pseudomonas syringae enzyme (3OCR). Sequence for a ‘no CT’ fuculose enzyme was also identified in Ta (enz**), which shows high sequence divergence in the YLE positions. (B): Zoom views of active sites from the Pseudomonas syringae aldolase enzyme (3OCR; top), Ta aldolase enzyme (middle), and Ta aldolase domain (bottom) with the AF3-modeled Zn²⁺ in the Ta enzyme coordinated by the histidine triad nearly identical to the bacterial enzyme. In contrast, the Ta domain has diverged in two of these histidine residues and does not contain a catalytic metal. (C): Saturation binding curves for the Ta aldolase proteins. Notably, the full-length (FL) enzyme containing a full CThelix binds Mud equally to the aldolase domain. Truncation of the enzyme CThelix does not improve or impair binding, similar to Hts (see Fig 2). In contrast, the divergent ‘no CT’ enzyme (**) does not bind Mud. (D): Saturation binding curves for chimeric Trichoplax and Pseudomonas syringae aldolase enzymes are shown. Fusion of the Trichoplax CThelix onto the core Pseudomonas enzyme (3OCR/Ta_enz-CT) shows robust Mud binding compared to the lack of interaction in the full-length bacterial enzyme (see Fig 4). In contrast, fusion of the Pseudomonas CThelix onto the core Trichoplax enzyme shows a moderate suppression of binding compared to the strong Mud binding seen in full-length Ta enzyme (compare to panel C).

https://doi.org/10.1371/journal.pone.0316787.g010

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Fig 11. A single Arginine-to-Glycine substitution permits Mud binding in a bacterial aldolase enzyme.

(A): Multiple sequence alignment of aldolase enzyme and domain sequences demonstrates two distinct subgroups based on the highlighted position. A group of prokaryotic enzymes have a conserved arginine (R) or lysine (K) (blue box). Additional prokaryotic enzymes, along with primitive eukaryotic enzymes, have substituted this position with a glycine (G) (green box). Aldolase domains also have a strictly conserved glycine in this position. (B): Tetrameric assembly of the P. syringae aldolase enzyme (RCSBid: 3OCR) is shown in surface rendering with the core fold in white and CThelix in black. Arginine at position 47 (R47) in each protomer is colored blue, the side chains of which point toward a central core of the tetramer to create a cavity of ~8.6Å diameter, denoted by the dashed red circle. (C): AF3 model for the tetrameric assembly of D. melanogaster Hts aldolase domain is shown in surface rendering with the core fold in white and CThelix in black. Glycine at position 134 (G134) in each protomer is colored green. The absence of a non-hydrogen side chain creates a cavity of ~19Å diameter, denoted by the dashed red circle. (D): Saturation binding curves for Wild-type (blue) and R47G (green) P. syringae aldolase enzymes are shown. The R47G mutant shows significant Mud binding compared to the wild-type enzyme. (E): Saturation binding curves for Wild-type (green) and G134R (blue) Hts aldolase domains are shown. The G134R mutant shows reduced Mud binding affinity compared to the wild-type domain.

https://doi.org/10.1371/journal.pone.0316787.g011

Mutations at conserved surface residues on the Hts tetramer do not impair Mud binding

We next sought to identify a specific Mud binding site within the Hts aldolase domain. The ability of Mud to bind following removal of the Hts CThelix narrows the binding site to a minimal aldolase fold (residues 111–310), eliminating the small number of conserved contacts between the Adducin CThelix and actin as likely contributors. The CThelix loop insertion is notably disordered in the Adducin/Actin complex [8], consistent with it not being directly involved similar to our results above with Hts/Mud binding. Furthermore, a majority of Adducin contacts with actin involve residues in extended sequences N- and C-terminal to the minimal aldolase core [8], with D339 (D310 in Hts) representing one exception. These findings together suggest that Mud binding occurs at a unique site within Hts. To examine this directly, we constructed a single alanine mutant at the conserved D310, along with three additional double mutants that were identified as conserved residues on surface-exposed regions of the Hts core tetramer (Q151A/E152A, Q177A/V181A, R205A/D207A; Fig 3A). Saturation binding experiments found that none of these mutants impaired Mud binding relative to a wild-type control Hts (Fig 3B-3C). These data suggest that Mud interacts with a distinct site within the Hts ALDO^DOM that lies outside of conserved regions of the tetrameric complex surface.

CThelix deletion confers Mud binding to an ancient class II aldolase enzyme

We next explored Mud binding to a primitive aldolase enzyme from Pseudomonas syringae, whose CThelix lacks the loop insertion found in Hts and α-Adducin domains but nevertheless makes similar contacts to its adjacent protomer within an otherwise conserved tetrameric assembly (Fig 1D and 4A). This ALDO^ENZ lacked significant Mud binding, a result not unexpected considering its suspected role as a metabolic enzyme (Fig 4B-4C). Remarkably, deletion of the CThelix from this ALDO^ENZ conferred robust Mud binding (Fig 4B-4C), which was nearly equivalent to that seen with the ALDO^DOM of Hts, both ‘full length’ and ΔCThelix. AF3 modeling indicated that CThelix removal from the Pseudomonas syringae enzyme does not prevent tetramer assembly (Fig 4A), consistent with the known tetramerization of aldolase enzymes naturally lacking a CThelix (see below). To corroborate this finding, we performed similar experiments with the homologous aldolase enzyme from an alternative Pseudomonas species (labeled simply ‘Pseudomonas’ throughout). We found this enzyme also lacked Mud binding in its full-length, wild-type form, but showed a robust interaction following deletion of its CThelix similar to the Pseudomonas syringae aldolase (Fig 4D).

Chimeric CThelix aldolase proteins display differential Mud binding dynamics

We next constructed two chimeric aldolase proteins, swapping the CThelices between the Hts ALDO^DOM and the Pseudomonas syringae ALDO^ENZ, to further investigate the impact of this divergent structural element in Mud binding (Fig 5A). Compared to the wild-type enzyme that lacks significant Mud binding, chimeric fusion with the Hts CThelix resulted in a strong interaction similar to that seen following deletion of the enzyme CThelix (Fig 5B). In contrast, fusion of the Hts domain with the CThelix from the bacterial enzyme reduced Mud binding affinity compared to the wild-type domain (Fig 5C). However, this inhibitory effect of the enzyme CThelix on Hts function was weaker compared to the strong gain-of-function seen in the reciprocal chimera. Taken together, these results further highlight a key regulatory role for the CThelix in controlling the Mud binding activity within the aldolase fold.

Identification of a conserved, CThelix-binding ‘YLE motif’ within the core aldolase fold

Considering the gain-of-function Mud binding following truncation or chimerism of the aldolase enzyme CThelix, we next considered that the binding site may occur in a location repressed by this sequence in the full-length enzyme. To explore this idea, we constructed a multiple sequence alignment of Hts with several aldolase enzymes and Adducin domains to identify conserved residues near protomer interfaces that contact the CThelix in tetrameric structures (Fig 6). This analysis revealed several such positions within interconnecting loops of the core aldolase fold that directly contact the CThelix of the neighboring protomer (Fig 6A). Close inspection of this interface shows a particularly remarkable conservation within a triad of residues (F88/L91/E94 in the Pseudomonas syringae enzyme and Y159/L162/E165 in the Hts domain; termed the ‘YLE motif’ herein) that contact a series of CThelix residues, some of which display similar conservation along with additional residues that constitute unique contacts in the aldolase domain (Fig 6C). The RosettaCM and SWISS-MODEL homology models both showed a similar disposition of the YLE motif residues and their critical contacts in the CThelix compared to the AF3-generated structure (Fig S1B,D).

The ‘YLE motif’ defines a putative Mud binding site

To determine the role of these conserved positions in the Hts ALDO^DOM, we constructed a series of alanine mutants and measured their Mud binding capacity relative to wild-type. Triple mutation of the YLE motif resulted in a complete loss of Mud binding when examined within the context of either the full-length domain or the minimal core (e.g., ΔCThelix; Fig 7A). A single L139A also significantly impaired binding (Fig 7A). In contrast, double mutation of H164/T167 bound Mud similar to wild-type, as did a single R128A mutant (Fig 7B). These data highlight a critical role specifically for the YLE motif in Mud binding, while discounting a role for the adjacent H164/T167 and R128 residues. To further examine the importance of the YLE motif, we tested the effects of mutations to the equivalent FLE motif in the Pseudomonas syringae aldolase enzyme. When examined within the context of the gain-of-function ΔCThelix enzyme, this mutation resulted in significantly impaired Mud interaction, similar to the YLE mutation in Hts (Fig 7C). Overall, these data implicate an important role for the conserved YLE motif in Mud binding (Fig 7D).

Mud binding varies widely across aldolase isozymes

Having identified a putative Mud interaction site conserved between the Hts aldolase domain and Pseudomonas syringae enzyme, we next widened our analysis to additional aldolase enzymes. Searching structural and sequence databases, we noted three subgroups within the Adducin-like class II aldolase enzymes (those with structural homology to the aldolase domain, referred to as ‘class IIa’ herein), which could be distinguished by the length of their CThelix (Fig 8A-8B). These enzymes mostly belong to a group of tetrameric L-fuculose-1-phosphate aldolases found in many pathogenic bacteria [20]. The first subgroup completely lacks a CThelix (termed ‘no CT’ herein); nevertheless, they share a core structure and tetrameric assembly with those aldolases already discussed (e.g. PBDid: 2IRP; Fig 8B). Ostensibly, the 2IRP enzyme mimics the Pseudomonas syringae enzyme following truncation of its CThelix. In contrast to this enzyme, however, we found that the ‘no CT’ enzyme was completely incapable of binding Mud (Fig 8C). Primary sequence comparison revealed that the ‘no CT’ enzyme is highly divergent in the core loop sequences, including the YLE motif (Fig 8A). Additional ‘no CT’ aldolase structures also demonstrate that, unlike ‘full CT’ enzymes, the divergent residues corresponding to the YLE motif in these ‘no CT’ enzymes do not contact neighboring protomers in what are otherwise similar tetrameric assemblies [21]. Mutation of these corresponding residues to YLE in the 2IRP enzyme did not lead to a gain of Mud binding function (Fig 8C), indicating this motif is necessary for Mud binding in relevant aldolase proteins but not sufficient for those naturally lacking this activity.

The second enzyme subgroup identified contains a short CThelix sequence (termed ‘half CT’ herein), which is structurally analogous to the sequence found prior to the loop insertion in Hts (Fig 8B). Structural studies have determined a similar tetrameric assembly in this enzyme subgroup as well [e.g. PDBid: 2FUA; [22]]. As with the ‘no CT’ enzyme, we found that the ‘half CT’ enzyme was completely devoid of Mud binding. Furthermore, truncation of the shortened CThelix did not permit Mud binding (Fig 8C). Notably, as with ‘no CT’ enzymes, the ‘half CT’ aldolases also lacked conservation of the YLE motif (Fig 8A).

Lastly, database searches identified a third subgroup of fuculose aldolase enzymes, distributed across diverse fungi, that contain a CThelix of similar length to that of Pseudomonas syringae enzyme and Hts/Adducin domain (e.g. PDBid: 4XXF; termed ‘full CT’ herein; Fig 8B). This ‘full CT’ fuculose aldolase showed a moderate Mud binding activity that was improved upon truncation of its CThelix, although both responses were weaker than that measured with Hts (Fig 8C). Examination of its primary sequence found a partially conserved YLE motif (Fig 8A). Overall, these analyses further substantiate the CThelix and YLE motif as key molecular determinants for Mud binding in the aldolase protein fold.

Aldolase gene expression suggests a potential Placozoan emergence of the Adducin family

Aldolases represent a large family of surprisingly diverse enzymes with divergent sequences, structures, and reaction mechanisms that nonetheless contribute similar metabolic functions through convergent evolution [4]. The starkest of these contrasts distinguish the class I and II subfamilies. Class I aldolases adopt a classic (and eponymous) TIM barrel fold and use a lysine-dependent Schiff base reaction mechanism [23,24]. Class II enzymes, in contrast, use a metal (most often Zn²⁺) catalyzed reaction scheme, with a conserved histidine triad functioning to coordinate this catalytic ion [17,25]. Class II enzymes can be further categorized structurally, with one subset also adopting a TIM barrel fold and a second showing high homology to the aldolase domain of Adducin proteins, along with other subgroups of sugar lyases (e.g., ‘class IIa’ enzymes; [26–28]). As presented above, class IIa enzymes have a similar core structural fold but diverge in the length of their CThelices.

Considering these sequence and structural distinctions across the aldolase enzyme family, as well as the dramatic functional divergence between class IIa enzymes and the Adducin aldolase domain, we next extended our search of the protein sequence database (https://blast.ncbi.nlm.nih.gov/Blast.cgi) to identify expression patterns of these unique aldolase proteins across major phyla in an effort to further explore a potential pathway of functional evolution within this protein fold (Fig 9). We searched selected phyla for class I enzyme, class II enzyme, class IIa enzyme (‘no CT’, ‘half CT’, and ‘full CT’ versions), and Adducin-like aldolase domain sequences as depicted in Fig 9A. We found class I enzyme sequences present across all phyla apart from the Choanoflagellates (Fig 9B). Our searches also indicated that class I enzymes represents the lone subtype found across higher eukaryotes (with the exception of a class II sequence also identified in the protostome Lophotrochozoa), consistent with previous studies finding a loss of class II genes in these phyla [29–31]. Class II enzymes were found in bacteria and most of lower eukaryotic phyla, typically together with class I sequences (Fig 9B). Class IIa enzymes were identified among similar phyla, albeit slightly more restricted than the class II sequences (Fig 9B). Searches for the aldolase domain found a restricted pattern of expression to metazoan phyla (Fig 9B). Based on these search results, the aldolase domain appears to have emerged in Placozoa (or possibly Porifera), which were also found to be the last phylum with apparent class IIa enzyme sequences, with both a ‘no CT’ and ‘full CT’ sequence identified (Fig 9B). Notably, searches of several Choanoflagellate species (an outgroup of metazoa), which have been shown to be a critical origin for other multi-domain scaffold proteins similar to Adducin [32–36], revealed neither class IIa enzymes nor aldolase domain-containing Adducin sequences (Fig 9B). Overall, our sequence analysis supports existing theories of the aldolase enzyme family evolution and highlight a potentially important role for Placozoa, a phylum of basal animals, in the class IIa and Adducin gene subgroups. This phylum of marine invertebrates forms aggregate ‘blobs’ of cells and are thought to represent one of the simplest forms of animal life. As such, they represent a critical position in the move to complex, multicellular life that could reveal essential molecular insights into this evolutionary transition [37].

A Class IIa aldolase enzyme from Trichoplax functions as a Mud binding protein

To empirically evaluate the Placozoan class IIa enzyme and domain genes identified, we began by modeling tetrameric structures of those from a prototype member, Trichoplax adhaerens [38]. Trichoplax form a ciliated bilayer of cells that perform fundamental coordinated functions such as locomotion and signal transduction [38,39]. As shown in Fig 10A, AF3 predicted assemblies similar to those found for bacterial enzyme and Hts domain aldolase tetramers. The lower ipTM and pTM confidence scores for the domain assembly likely result from ‘Low’ confidence per-residue plDDT scores found in each protomer CThelix (not depicted). Notably, AF3 modeled Zn²⁺ ions within the conserved histidine triad retained in the Trichoplax enzyme sequence and structure, supporting the notion that these retain its aldolase activity (Fig 10A-10B). Similar metal placement was not predicted in the aldolase domain, which has diverged significantly within this catalytic triad (Fig 10B).

We next examined the ability of the Trichoplax aldolase proteins to directly bind Mud in pulldown experiments. The Trichoplax ALDO^DOM bound Mud similarly to Hts, indicating that protein binding is a function present in Adducin proteins in the most basal metazoan (Fig 10C). Surprisingly, we found that the ‘full CT’ ALDO^ENZ from Trichoplax bound Mud to an equal extent, and truncation of its CThelix did not improve or impair binding (Fig 10C). Notably, this enzyme, as well as the identified aldolase domain, both contain a partially conserved YLE motif (Fig 10A). We also identified a ‘no CT’ enzyme in Trichoplax, which showed a divergent sequence at the YLE positions and was completely devoid of Mud binding function (Fig 10A, 10C).

Having found that the Trichoplax ‘full CT’ enzyme binds Mud at a level similar to the Hts aldolase domain, we next constructed enzyme chimeras with the Pseudomonas syringae aldolase, which is devoid of Mud binding (Fig 4). Specifically, we swapped the CThelix between these enzymes and examined Mud binding ability. Chimeric Pseudomonas syringae aldolase that has its CThelix replaced with that from Trichoplax was found to have a gain-of-function, showing Mud binding similar to that of the full-length Trichoplax enzyme (Fig 10D). In contrast, chimeric Trichoplax aldolase with its CThelix replaced with that from Pseudomonas syringae was found to have impaired Mud binding relative to the wild-type enzyme (Fig 10D). These data further highlight a regulatory role for the CThelix in protein binding function of the aldolase fold.

Identification of a conserved Glycine substitution across aldolase domains

Truncation and chimerism of the CThelix, both of which lead to Mud binding in a primitive aldolase enzyme (Figs 4 and 5), represent relatively nontrivial sequence alterations. As such, we sought to identify changes that would have a minimal impact on aldolase enzyme sequence but a similarly large impact on Mud binding function. We considered two criteria to identify possible amino acid substitutions: [1] those located at or near protomer interfaces within the aldolase tetramer, and [2] those with significant distinctions between enzymes and domains conserved across evolution. This analysis revealed one residue in particular, which shows a strong conservation for arginine (or lysine; R/K) across several bacterial aldolase enzymes but that is substituted for glycine (G) in Adducin aldolase domains (Fig 11A). Although glycine was also identified in a subset of aldolase enzymes, it was unanimously present across domain sequences analyzed. Substitution of a cationic arginine for glycine represents a significant chemical change, and the strong conservation of these distinct amino acids further suggests a potential functional consequence. In both aldolase enzyme and domain structures, this residue is positioned immediately after the first α-helix of the aldolase fold, which packs against the CThelix (Fig 11B-11C). In the aldolase enzyme, the arginine side chain points towards a central cavity formed in the tetrameric assembly. This results in the guanidinium R-groups of all four protomer arginine residues in the tetramer facing one another and packing closely together to form a narrow central cavity (8.6Å diameter; Fig 11B). In contrast, the lack of a non-hydrogen side chain in the aldolase domain glycine results in a loss of R-group packing and the formation of a significantly wider central cavity in the AF3 structural model of the Hts tetramer (19Å diameter; Fig 11C).

Single arginine-to-glycine change in an aldolase enzyme confers Mud binding

To directly examine the role of this R→G change across aldolase proteins, we constructed an R47G mutation in the Pseudomonas syringae enzyme and the reciprocal G134R substitution in the Hts domain and measured Mud binding compared to their respective wild-type controls. Mud binding was significantly enhanced in the single R47G aldolase enzyme substitution (Fig 11D). Binding to the R47G enzyme was not as strong as that seen following truncation of the enzyme CThelix, but nevertheless represents a significant gain-of-function. In contrast, Mud binding was reduced in the G134R substituted Hts aldolase domain, although this effect was more modest compared to the gain-of-function seen in the R47G enzyme (Fig 11E). These results agree with those from alterations in the CThelix, with domain-mimicking changes in the aldolase enzyme producing significantly stronger effects than enzyme-mimicking changes in the Hts aldolase domain. Overall, our results spotlight a single amino acid substitution that may represent a significant step in the evolution of aldolase protein function. It should be noted again that although many bacterial enzymes were found to contain an R/K, others were identified with a G at this position. The consequences of this substitution to the normal enzyme function, for example with respect to their catalytic activity, will require further investigation. Deviation from either R/K or G amino acids in this position was rarely seen across aldolase sequences, underscoring a unique functional role.