Mechanism of APTX nicked DNA sensing and pleiotropic inactivation in neurodegenerative disease
2018; Springer Nature; Volume: 37; Issue: 14 Linguagem: Inglês
10.15252/embj.201798875
ISSN1460-2075
AutoresPercy Tumbale, Matthew J. Schellenberg, Geoffrey A. Mueller, Emma Fairweather, Mandy Watson, Jessica Neville Little, J.M. Krahn, Ian D. Waddell, Robert E. London, R. Scott Williams,
Tópico(s)Prion Diseases and Protein Misfolding
ResumoArticle22 June 2018Open Access Transparent process Mechanism of APTX nicked DNA sensing and pleiotropic inactivation in neurodegenerative disease Percy Tumbale Percy Tumbale Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Matthew J Schellenberg Matthew J Schellenberg Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Geoffrey A Mueller Geoffrey A Mueller Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Emma Fairweather Emma Fairweather Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK Search for more papers by this author Mandy Watson Mandy Watson Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK Search for more papers by this author Jessica N Little Jessica N Little Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Juno Krahn Juno Krahn Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Ian Waddell Ian Waddell Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK Search for more papers by this author Robert E London Robert E London Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author R Scott Williams Corresponding Author R Scott Williams [email protected] orcid.org/0000-0002-4610-8397 Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Percy Tumbale Percy Tumbale Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Matthew J Schellenberg Matthew J Schellenberg Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Geoffrey A Mueller Geoffrey A Mueller Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Emma Fairweather Emma Fairweather Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK Search for more papers by this author Mandy Watson Mandy Watson Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK Search for more papers by this author Jessica N Little Jessica N Little Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Juno Krahn Juno Krahn Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Ian Waddell Ian Waddell Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK Search for more papers by this author Robert E London Robert E London Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author R Scott Williams Corresponding Author R Scott Williams [email protected] orcid.org/0000-0002-4610-8397 Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA Search for more papers by this author Author Information Percy Tumbale1,‡,‡, Matthew J Schellenberg1,‡,‡, Geoffrey A Mueller1,‡,‡, Emma Fairweather2, Mandy Watson2, Jessica N Little1,‡, Juno Krahn1,‡, Ian Waddell2, Robert E London1,‡ and R Scott Williams *,1,‡ 1Genome Integrity and Structural Biology Laboratory, Department of Health and Human Services, National Institute of Environmental Health Sciences, US National Institutes of Health, Research Triangle Park, NC, USA 2Drug Discovery Group Cancer Research UK Manchester Institute, Manchester, UK ‡These authors contributed equally to this work ‡This article has been contributed to by US Government employees and their work is in the public domain in the USA *Corresponding author. Tel: +1 984 287 3542; E-mail: [email protected] The EMBO Journal (2018)37:e98875https://doi.org/10.15252/embj.201798875 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract The failure of DNA ligases to complete their catalytic reactions generates cytotoxic adenylated DNA strand breaks. The APTX RNA-DNA deadenylase protects genome integrity and corrects abortive DNA ligation arising during ribonucleotide excision repair and base excision DNA repair, and APTX human mutations cause the neurodegenerative disorder ataxia with oculomotor ataxia 1 (AOA1). How APTX senses cognate DNA nicks and is inactivated in AOA1 remains incompletely defined. Here, we report X-ray structures of APTX engaging nicked RNA-DNA substrates that provide direct evidence for a wedge-pivot-cut strategy for 5′-AMP resolution shared with the alternate 5′-AMP processing enzymes POLβ and FEN1. Our results uncover a DNA-induced fit mechanism regulating APTX active site loop conformations and assembly of a catalytically competent active center. Further, based on comprehensive biochemical, X-ray and solution NMR results, we define a complex hierarchy for the differential impacts of the AOA1 mutational spectrum on APTX structure and activity. Sixteen AOA1 variants impact APTX protein stability, one mutation directly alters deadenylation reaction chemistry, and a dominant AOA1 variant unexpectedly allosterically modulates APTX active site conformations. Synopsis Aprataxin (APTX) deadenylase protects genomic integrity by reversing DNA adenylation arising during ribonucleotide excision and base excision repair. New structures of APTX-substrate complexes reveal the mechanism of nicked DNA sensing and the basis for differential impact of APTX mutations in neurodegenerative disorder. Molecular snapshots of APTX in complex with nicked RNA-DNA substrates show that APTX bends DNA. X-ray and NMR data define APTX conformations throughout its reaction cycle, supporting a substrate-induced fit active site assembly mechanism. A wedge-pivot-cut strategy for 5′-AMP resolution is shared with the alternate 5′-AMP-processing enzymes POLβ and FEN1. Comprehensive survey of APTX mutations in Ataxia with Oculomotor Apraxia 1 (AOA1) shows their differential impact on APTX protein stability, deadenylation reaction chemistry, and APTX active site conformational changes. Introduction DNA ligation is a central process in biology that finalizes genome maintenance metabolic processes including DNA replication, recombination, and DNA repair. Eukaryotic DNA ligases catalyze ligation via a three-step, ATP-dependent reaction. First, the DNA ligase active site lysine is adenylated. Second, the adenylate is transferred to a DNA 5′ phosphate to facilitate the third nick-sealing step. Third, nucleophilic attack of a 3′-OH on the activated 5′-adenylate facilitates phosphodiester bond formation, and sealing of the DNA break (Pascal et al, 2004; Tomkinson et al, 2006). Environmental and metabolic sources of DNA damage prompt "abortive ligation", the failure of ligase to complete step 3, and generation of 5′-adenylated (5′-AMP) DNA strand breaks (Ahel et al, 2006; Andres et al, 2015; Schellenberg et al, 2015) (Fig 1A). Triggers of abortive ligation include RNA-DNA junction intermediates in ribonucleotide excision repair (RER) (Tumbale et al, 2014), single-strand breaks with oxidative DNA base damage (e.g. 3′-8-oxo-dG) (Parsons et al, 2005; Ahel et al, 2006; Harris et al, 2009), and DNA nicks bearing 5′ deoxyribose phosphate groups, such as those generated by base excision repair (BER) (Rass et al, 2007). In these contexts, it is hypothesized that DNA ligase exacerbates genome instability with the creation of complex 5′-adenylated damage comprised of the instigating lesion, compounded by adenylation of the DNA 5′-terminus. Figure 1. X-ray structure of hAPTX-nicked-RNA-DNA complex Schematic presentation of APTX 2-step deadenylation mechanism. Concatemerized nicked RNA-DNA duplex substrate bound in the structure of hAPTX-nicked-RNA-DNA complex. The 6 bp upstream region is shown in blue, 8 bp downstream region, and the complementary template strand in black, 5′ ribonucleotide positioned at the nick junction in yellow, and N-terminal α1-helix in orange. Transparent surface representation of two APTX protomers (Pro1, cyan and Pro2, gray) bound to a single RNA-DNA molecule containing two nick sub-sites in each asymmetric unit. Simulated annealing omit 2Fo-Fc electron density for the bound nicked RNA-DNA (magenta) illustrates APTX engagement to DNA nick induces large-scale DNA distortions at the nick. X-ray structure of hAPTX-nicked-RNA-DNA complex. N-terminal α1-helix is colored orange, HIT domain in light blue, Znf domain in light green, DNA in pink, RNA in yellow, AMP in teal, and HIT-loop in dark green. Download figure Download PowerPoint The aprataxin (APTX) RNA/DNA polynucleotide deadenylase directly reverses 5′-AMP damage (Fig 1A) (Ahel et al, 2006; Harris et al, 2009; Reynolds et al, 2009; Tumbale et al, 2011, 2014) and associates with DNA repair scaffolds XRCC1 (Cherry et al, 2015) and XRCC4 (Breslin & Caldecott, 2009). Mutational inactivation of APTX is associated with elevated levels of oxidative DNA damage (Hirano et al, 2007; Harris et al, 2009), and increased DNA damage following treatment with the anticancer topoisomerase inhibitor CPT (Mosesso et al, 2005), and S. pombe Aptx mutants are sensitive to 4NQO (Deshpande et al, 2009). Genetic evidence implicates the budding yeast APTX homolog Hnt3p in repair of RNA-triggered abortive ligation during ribonucleotide excision repair (RER) (Tumbale et al, 2014), as well as alkylation and oxidative DNA damage repair pathways (Daley et al, 2010). Aptx-deficient mice expressing mutant superoxide dismutase (SOD1) show cellular survival defects in cultured cells (Carroll et al, 2015), and the genetic combination of Aptx and Tdp1 (tyrosyl-DNA phosphodiesterase 1) knockout results in global defects in repair of oxidative and alkylation-induced DNA damage (El-Khamisy et al, 2009). The consequences of APTX dysfunction in humans are severe. APTX mutations are linked to the progressive neurodegenerative diseases ataxia with Oculomotor Apraxia 1 (AOA1) (Date et al, 2001; Moreira et al, 2001), ataxia with coenzyme Q10 (coQ10) deficiency (Quinzii et al, 2005), and a multiple system atrophy resembling Parkinson's disease (Baba et al, 2007). AOA1 mutations map to the APTX catalytic domain (Barbot et al, 2001; Moreira et al, 2001; Shimazaki et al, 2002; Le Ber et al, 2003; Tranchant et al, 2003; Criscuolo et al, 2004, 2005; Ito et al, 2005; Mosesso et al, 2005; Baba et al, 2007; Yokoseki et al, 2011) and cause variable age of disease onset. Yet, how these mutations impact the APTX structure and its polynucleotide deadenylase functions remains largely unknown. Previous molecular structural interrogations of S. pombe (Tumbale et al, 2011; Chauleau et al, 2015) and human (Tumbale et al, 2014) APTX homologs by us and others have resolved how APTX engages blunt DNA duplex structures, and provided a basis for understanding the APTX direct-reversal DNA deadenylation reaction. However, the presumed cognate substrates for APTX that arise during ribonucleotide excision repair (RER) and base excision repair are nicked DNA and RNA-DNA junctions. How APTX senses DNA nicks and processes adenylation damage in these contexts remains undefined. It is also unknown whether the APTX structure and activity is regulated, and if so, how disease states might impact such regulation. To help resolve these questions, we have secured molecular snapshots of APTX in complex with nicked RNA-DNA substrates and investigated its conformations throughout its reaction cycle using nuclear magnetic resonance (NMR) spectroscopy. We uncovered a DNA nick-induced fit active site assembly mechanism that modulates APTX active site loop conformations. Furthermore, results from a comprehensive structural and functional characterization of AOA1 mutants provide a framework for understanding APTX inactivation in neurodegenerative disease. We find AOA1 mutations impair APTX function by impacting protein folding, altering active site chemistry, or by perturbing ligand-dependent APTX conformational changes. Results Molecular architecture of APTX bound to nicked RNA-DNA To visualize APTX bound to a cognate reaction product harboring nicked RNA-DNA and a cleaved AMP lesion, we surveyed 80 combinations of protein/RNA-DNA complexes for co-crystallization. Our combinatorial crystallization approach involved varying upstream and downstream duplex lengths surrounding a DNA nick, varying position of the nick, as well as concatemerization of DNA nicks within a single target nicked RNA/DNA substrate (Fig 1B). We obtained crystals (Appendix Fig S1A) diffracting to 2.4 Å resolution that contain two APTX catalytic domain (APTXcat, residues 165–342) protomers engaging a doubly nicked target structure (Fig 1B and C). Successful crystallization strategies ultimately utilized a concatemerized nick substrate with a 6 bp upstream and 8 bp downstream duplexes (Fig 1B). In addition, a 5′-ribonucleotide is positioned at the nick junction. In this configuration, the structural snapshots captured in our crystal structures are representative of APTX bound to a nicked RNA-5′-3′-DNA junction reaction product complex formed by APTX during repair of abortive ligation products created during ribonucleotide excision repair (RER) (Tumbale et al, 2014; Schellenberg et al, 2015). The structure reveals how APTX induces large-scale DNA duplex substrate distortions in order to access the 5′-terminal adenylated lesion in the context of a nicked duplex. In each of the complexes captured in our crystals, abrupt DNA bending is governed by the APTX amino-terminal helix, as previously hypothesized (Tumbale et al, 2011). This structure illuminates how APTX senses DNA damage through two discontinuous nucleic acid binding sub-sites, dictated by both the HIT and Znf domains which together collaborate to sense the nick. The electron density for the bound nick target is well defined in the crystals (Fig 1C). Although the doubly nicked RNA-DNA sequence is symmetrical, about an 8 bp double-stranded palindromic region, in three-dimensional space this twofold symmetry, is broken. Two nick-bound APTX molecules bind the upstream region of the nick with distinct conformations (Appendix Fig S1B, and Materials and Methods). In both binding modes captured, APTX-directed DNA wedging and penetration into the base stack results in the redirection of the base stack approximately orthogonal to the helical axis of the bound downstream region. The major molecular interface at the DNA nick is mediated by the amino-terminal HIT domain α-helix (α1) that infiltrates the DNA base stack. Here, α1 serves as a doubly barbed "wedge" that splays the base stack apart (Figs 1D and 2A, orange helix). The planar rings of His166 and Trp167 together redirect the DNA duplex, imparting a ~90° bend to the substrate RNA-DNA. The protein displaces the upstream half of the nick and unwinds both the 3′ and 5′ sides of the nick. This has two effects: First, extraction of the 5′-terminus facilitates positioning of the adenylated lesion into the active site; and second, disruption of the 3′-terminal side of the nick exposes the 3′ end. Figure 2. DNA-induced conformational ordering of N-terminal α1-helix Molecular details of APTX-DNA nick interface illustrate the N-terminal α1-helix wedges into the DNA base stack. His166 and Trp167 stack against the bases at the nick, bending the DNA duplex at a ˜90° and unwinding both the 3′ and 5′ ends of the damaged strand. Limited chymotryptic proteolysis of APTX. The unliganded full-length APTX was proteolysed to C1 fragment (blue) mapped to the N-terminus of the HIT domain. The C1 fragment was further degraded to C2 (magenta) and C3 (green), and mapped to the N-term α1-helix (orange box). Addition of blunt-ended DNA, nicked DNA, and transition mimic DNA (adenosine (Ade) and orthovanadate ()) substrates resulted in proteolytic protection of C1. APTX was not protected from proteolysis when incubated with relaxed circular plasmid DNA. AOA1 K197Q active site mutant that is severely defective in DNA binding displays impaired DNA-end-dependent proteolytic protection. Locations of [methyl-13C]-labeled methionine residues (cyan) mapped in the catalytic core of APTX. Overlays of assigned 1H-13C HSQC spectra of [methyl-13C]methionine-labeled wild-type APTX in unliganded state (black), blunt-ended RNA-DNA substrate bound state (cyan), and nicked RNA-DNA substrate bound state (red). The arrows indicate resonance shifts in response to DNA substrates. Overlays of assigned 1H-13C HSQC spectra of [methyl-13C]methionine-labeled wild-type APTX in unliganded state (black), adenosine-VO3-RNA-DNA transition mimic state (blue), and nicked AMP and RNA-DNA product bound state (purple). The arrows indicate resonance shifts in response to RNA-DNA substrates. Kinetic parameters of APTX processing of 5′-adenylated RNA-DNA substrates, nicked (red) and blunt-ended (cyan). Mean ± 1 SD (n = 3 technical replicates) is displayed. Download figure Download PowerPoint DNA-induced fit APTX conformational changes Previous X-ray structures of APTX bound to blunt-ended DNA displayed two major APTX conformations differing by a conformational change of α1, a series of linked rearrangements of the histidine triad (HIT, HxHxH) substrate engagement loop, and additional compensating movements throughout the fused HIT-Znf catalytic core (Tumbale et al, 2014). In a "closed", and catalytically competent conformer, α1 packs against the HIT-loop and aligns the active site nucleophile His260 for attack on the 5′-AMP phosphorous atom. In comparison, a catalytically incompetent "open" conformer is typified by partial disengagement of the α1 helix from the HIT-loop, and a misaligned active site (Appendix Fig S2A). Structural superpositions indicate the present nick-bound APTX protomers are both found in the closed conformation, with the active site poised for catalysis (Appendix Fig S2B). Furthermore, in the DNA nick-bound forms, His166 (α1) stacks against the upstream duplex (Fig 2A). Modeling the opened state with this DNA shows it would clash in the open conformation, suggesting the interaction of His166 with the upstream duplex reinforces a closed and catalytically competent active site. Collectively, these observations suggest that DNA substrate engagement regulates conformations of the HIT wedge helix and, in turn, active site assembly. Difference distance matrix plot (DDMP) analysis (Appendix Fig S3) and a structural interpolation comparing the open and closed APTX conformational states (Movie EV1) reveals significant overall rearrangements of the HIT-Znf scaffold. We thus hypothesized that substrate binding and catalysis are intimately associated with α1 conformational change. To test this, we probed the structural response of full-length human APTX (hAPTXFL) to substrate and AMP lesion binding using limited proteolysis coupled to mass spectrometry. In the absence of DNA and the AMP lesion, chymotrypsin protease efficiently degrades hAPTXFL, yielding a metastable fragment (fragment "C1" aa 152-342, blue, Fig 2B, and Appendix Fig S4). C1 is digested further by cleavages at Trp167 (C2) and Leu171 (C3) in the N-terminal helix α1, revealing that in the absence of DNA, relative to the HIT-Znf core, helix α1 is flexible, accessible to protease, and therefore partially unfolded in solution. Addition of blunt-ended DNA or a nicked DNA oligonucleotide in the presence of AMP resulted in proteolytic protection of α1 (sites C2 and C3), consistent with a DNA-induced conformational ordering of α1 (Fig 2B). Similar proteolytic protection was observed with incubation with nicked DNA, adenosine (Ade), and orthovanadate () under solution conditions that have been previously demonstrated to covalently trap an APTX transition-like state (Tumbale et al, 2014; Fig 2B, lanes 11–15). That APTX was not protected from proteolysis when incubated with relaxed circular plasmid DNA indicates that helix α1 is specifically responsive to DNA damage binding at DNA ends and nicks (Fig 2B, lane 21–25 Appendix Fig S4). Moreover, an AOA1 active site mutant K197Q which impacts the active site DNA and AMP binding pocket displays impaired DNA-dependent proteolytic protection (Fig 2B, lanes 27–45). Thus, DNA-binding linked α1 conformational change is altered by AOA1 mutations linked to neurological disease. Global conformational responses of the APTX catalytic domain during catalysis To better define the response of the APTX catalytic domain (APTXcat) to substrate binding in solution, we investigated the behavior of 13CH3-Met-labeled APTXcat by NMR. The 13CH3-Met labels provide seven probes distributed throughout APTXcat located proximal to α1 (Met164, Met175, and Met180), the HIT-loop (Met256), and at three added positions remote from α1 and the active center (Met227, Met309, and Met296) (Fig 2C and Appendix Table S1). These probes are also coincident with regions shown to undergo motion in X-ray structures (Appendix Fig S3). Distinct 13CH3-Met resonance shifts are observed for apo (unliganded structure) compared with blunt-ended or nicked DNA substrates (DNA in twofold molar excess; Fig 2D). The 13CH3 shifts for Met164 are close to those expected for a methionine residue in a random coil (Butterfoss et al, 2010), consistent with the origin of this residue as a residual fragment from an N-terminal His-tag. Nevertheless, the M164 1H shift is sensitive to the presence of nicked DNA, most probably as a result of the stacking of nearby Trp167 against the rG1 residue at the nick. As a result of its position near the N-terminus of helix α1, this observation underscores a key role for α1 in DNA nick recognition. Met175/Met180 (α1) and Met256 (HIT-loop) also display substrate-specific chemical shift perturbations. The large, downfield 1H shift of ~0.4 ppm for M256 is consistent with its position at the edge of the Trp167 indole side chain in the nicked DNA complex (Appendix Table S1, Fig 2D). Consistent with these substrate-dependent conformational responses, kinetic analysis of APTX deadenylation reaction (Fig 2F) yielded distinct parameters for APTX deadenylase activity on blunt (Km = 37.1, kcat = 0.51/s) versus nicked (Km = 17.1 nM, kcat = 0.38/s) substrates, with APTX displaying superior catalytic efficiency on nicked DNA (kcat/Km = 0.022 nicked, kcat/Km = 0.014 blunt DNA). To obtain further insight into catalysis-related conformational changes, we measured the 13CH3-Met resonance shifts to compare (i) Apo (Black, Fig 2E), (ii) reaction transition state (blue, Fig 2E), and (iii) product states (purple, Fig 2E). Comparison of 13CH3-Met spectra shows that major changes in the molecular environment of Met256 (HIT-loop), Met175, and Met180 (α1) occur during the APTX reaction cycle. Strikingly, the methyl resonances of Met227, Met309, and to a lesser extent, Met296, corresponding to residues located removed from the active site (> 10–15 Å, Appendix Table S1) and substrate binding regions of the enzyme, also exhibit significant substrate (Fig 2D) and reaction-state (Fig 2E)-dependent chemical shift behaviors. The chemical shift sensitivity of residues remote from the active site may be amplified by proximity to shift-inducing aromatic residues. For instance, the M227 shift may be responsive to small positional variations relative to Y250. Overall, these NMR solution results are consistent with our X-ray structures and proteolysis indicating conformational change in α1 and the HIT-loop is coincident with a global APTX conformational response to DNA engagement. These observations further implicate a DNA regulated active site assembly/disassembly cycle in catalysis. APTX mutations variably impact protein folding and activity The crystallographically defined APTX intermediate states and NMR characterized ligand-dependent conformational changes provide a framework for understanding the molecular consequences of the many known AOA1-linked missense mutations. A subset of APTX mutants have been demonstrated to harbor defects in a weak APTX AMP-lysine hydrolase activity (Seidle et al, 2005). However, for the majority of AOA1 variants, the molecular and functional impacts on APTX RNA-DNA deadenylase activity have not been assessed. Several AOA1 mutations are single amino acid substitutions within the protein core or lead to premature protein truncations (Fig 3A and Appendix Fig S5). We thus evaluated the impact of AOA1 mutations on protein folding. First, we tested the solubility of 17 AOA1-linked missense and nonsense mutants expressed in E. coli. With the exception of two nonsense variants (R247X and W279X), all proteins expressed at comparable levels as assessed by Western blotting to a 6x-His-tag in the recombinant protein (Fig 3B). In contrast, analysis of the soluble protein fractions revealed marked variability in solubility. Six mutants (A198V, H201R, P206L, G231E, R247X, W279X) were entirely insoluble, revealing these substitutions are poorly tolerated in the APTX structure. Examination of the molecular environments of these amino acid positions (Appendix Fig S5) shows these mutations are likely to disrupt folding of the HIT domain. The remaining 11 AOA1 variants (D185E, K197Q, R199H, H201Q, L223P, S242N, L248M, V263G, D267G, W279R, R306X) (Fig 3B) display wide ranging solubility, suggesting these mutations differentially impact protein stability and activity. Figure 3. AOA1 mutagenic effects on APTX solubility, stability, and catalytic activity APTX mutations in AOA1 map to the catalytic core of APTX comprising the HIT (blue) and Znf (green) domains. Solubility of 17 AOA1-linked APTX missense and nonsense mutants expressed in E. coli was assessed by Western blotting. With the exception of two nonsense variants (R247X and W279X), all proteins expressed at comparable levels. The soluble protein fractions revealed marked variability in solubility. Thermal melting profile of APTX mutants. Unfolding of APTX proteins was monitored using SYPRO Orange fluorescent dye. The midpoint temperature (Tm) for APTX wild-type and mutants in unliganded (bottom) and AMP bound (top) conditions is displayed. X = unable to determine Tm, *P-value < 0.05, **P-value < 0.01, N.S. = not-significant, two-tailed t-test. Mean ± 1 SD (three technical replicates) is shown. Deadenylation activity of APTX mutants. Tenfold dilutions of APTX mutant proteins were tested for deadenylation on a TAMRA labeled 5′-adenylated nicked RNA-DNA substrate at 25 and 37°C. Reaction products were resolved on 15% TBE-urea denaturing gels and visualized by fluorescence scan. Fold increase in protein to reach 50% activity relative to wild-type APTX is displayed. Mean ± 1 SD (3 technical replicates) is displayed. Download figure Download PowerPoint To further define the molecular basis for AOA1 defects, we purified soluble variants (Fig 3B, Appendix Fig S6) and measured protein thermal stabilities and RNA-DNA deadenylase activities (Fig 3C, Appendix Figs S6 and S7A, and Dataset EV1). Thermal shift assays derive a thermal melting point transition from fluorescence of Sypro Orange dye binding to the protein hydrophobic core as the protein undergoes heat-induced unfolding, in the presence or absence of bound ligands (Huynh & Partch, 2015) (Fig 3C). With the exception of L248M, all mutants displayed statistically significant decreases in protein thermal stability compared to WT. In the absence of ligand, accurate Tm values for D185E, H201Q, V263G, and L223P could not be determined due to their intrinsic instabil
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