A structural role for the PHP domain in E. coli DNA polymerase III
- Tiago Barros†1,
- Joel Guenther†1,
- Brian Kelch1,
- Jordan Anaya1,
- Arjun Prabhakar1,
- Mike O’Donnell2,
- John Kuriyan1, 3Email author and
- Meindert H Lamers1, 4Email author
© Barros et al.; licensee BioMed Central Ltd. 2013
Received: 5 March 2013
Accepted: 7 May 2013
Published: 14 May 2013
In addition to the core catalytic machinery, bacterial replicative DNA polymerases contain a Polymerase and Histidinol Phosphatase (PHP) domain whose function is not entirely understood. The PHP domains of some bacterial replicases are active metal-dependent nucleases that may play a role in proofreading. In E. coli DNA polymerase III, however, the PHP domain has lost several metal-coordinating residues and is likely to be catalytically inactive.
Genomic searches show that the loss of metal-coordinating residues in polymerase PHP domains is likely to have coevolved with the presence of a separate proofreading exonuclease that works with the polymerase. Although the E. coli Pol III PHP domain has lost metal-coordinating residues, the structure of the domain has been conserved to a remarkable degree when compared to that of metal-binding PHP domains. This is demonstrated by our ability to restore metal binding with only three point mutations, as confirmed by the metal-bound crystal structure of this mutant determined at 2.9 Å resolution. We also show that Pol III, a large multi-domain protein, unfolds cooperatively and that mutations in the degenerate metal-binding site of the PHP domain decrease the overall stability of Pol III and reduce its activity.
While the presence of a PHP domain in replicative bacterial polymerases is strictly conserved, its ability to coordinate metals and to perform proofreading exonuclease activity is not, suggesting additional non-enzymatic roles for the domain. Our results show that the PHP domain is a major structural element in Pol III and its integrity modulates both the stability and activity of the polymerase.
The DNA polymerases at the core of every bacterial replisome belong to the C-family of DNA polymerases . All members of this family contain a set of four domains that are organised within a single polypeptide in the following order: Polymerase and Histidinol Phosphatase (PHP), Palm, Thumb and Fingers. While these four domains always appear in the same order, the DNA polymerase III (Pol III) and DNA polymerase C (Pol C) subfamilies can be distinguished within the C-family of DNA polymerases, depending on the arrangement of additional accessory domains, such as the OB-fold domain that binds single-stranded DNA [2–4]. Neither Pol III nor Pol C share detectable sequence homology with other DNA polymerases, including bacterial DNA polymerases such as Pol I and Pol II and the eukaryotic replicative DNA polymerases ϵ and δ.
The crystal structures of Escherichia coli (E. coli) Pol III , Thermus aquaticus (T. aquaticus) Pol III  and Geobacillus kaustophilus (G. kaustophilus) Pol C  have shown that the active sites of these polymerases are structurally related to that of human DNA polymerase β, an atypical DNA polymerase that is involved in base excision repair and belongs to the X-family of DNA polymerases. This is surprising, as X-family polymerases are typically slow and exhibit low fidelity and processivity, in contrast to the high-fidelity replicative C-family polymerases, which are amongst the fastest polymerases known.
Pol III and Pol C polymerases are also unique in that they contain a Polymerase and Histidinol Phosphatase (PHP) domain that is not found in other polymerases, except for some bacterial Pol X family members . The PHP domain in Pol III and Pol C is a barrel-shaped domain located at the side of the polymerase, near the Thumb domain [5, 6]. The active site of typical PHP domains is a shallow cavity located at the top of the barrel-shaped domain, usually consisting of seven β-strands that provide most of the residues that coordinate the catalytic metals. Several crystal structures of PHP domains have been determined. These include PHP domains that are not part of a larger protein: E. coli YcdX , Thermus thermophilus (T. thermophilus) histidinol phosphate phosphatase (ppPHP)  and tm0559 from Thermatoga maritima (T. maritima) (pdb code 2ANU); or those that are part of polymerases: E. coli Pol III , T. aquaticus Pol III , G. kaustophilus Pol C  and Deinococcus radiodurans (D. radiodurans) Pol X .
The role of the PHP domain in Pol III and Pol C polymerases remains unclear. When first identified in sequence alignments, the PHP domain in Pol III/Pol C was hypothesized to act as a pyrophosphatase, removing the by-product of DNA synthesis in order to drive the polymerization reaction in the direction of DNA synthesis . However, no such activity has been detected as yet for a polymerase PHP domain. Instead, the PHP domains of T. thermophilus Pol III and T. aquaticus Pol III, which have a complete set of metal-coordinating residues and have been shown to bind metals, have exonuclease activity [12, 13], which presumably serves to proofread newly synthesized DNA. Likewise, the PHP domains of Pol X from both Bacillus subtilis (B. subtilis)  and T. thermophilus  have exonuclease activity. In contrast, no exonuclease activity could be detected for the PHP domain of G. kaustophilus Pol C .
The invariable presence of the PHP domain in all C-family polymerases, even those lacking exonuclease activity, suggests that this domain must play an essential, yet non-enzymatic, role in maintaining the activities of these polymerases. We show, using sequence analysis, that the loss of metal-coordinating residues in the Pol III PHP domain is correlated with the presence in bacterial genomes of a protein homologous to the E. coli Pol III proofreading exonuclease ϵ subunit. Despite the apparent loss of catalytic function, the structural scaffold of the PHP domain has been conserved to a remarkable degree. This observation is strongly supported by our ability to restore metal binding to the E. coli Pol III PHP domain by introducing only three point mutations. We further show that the structural integrity of the PHP domain is important for the stability and activity of the E. coli Pol III.
Results and discussion
A complete set of metal-coordinating residues is not universally conserved in DNA polymerase PHP domains
Conservation of the nine residues required for metal binding in PHP domains
Metal binding residue
Eco Pol III
Taq Pol III
The PHP active site in E. coli Pol III has 5 replacements compared to the consensus sequence (Figure 1). In addition to the three histidines and the cysteine at position 7 mentioned previously, a glutamate to aspartate replacement at position 5 (Asp 69) is present. Indeed, in our hands the E. coli Pol III subunit does not show any nuclease activity (see below). The lack of activity by the E. coli PHP domain has been predicted . It is surprising, however, that the G. kaustophilus PHP domain is also inactive  as it presents an almost intact active site that has been shown to bind metals. The only replacement in this active site is an aspartate to asparagine substitution at position 8. It is likely that additional residues not directly involved in metal coordination are also necessary for robust exonuclease activity.
The presence of variant PHP domains in bacterial replicative polymerases is correlated with the presence of separate proofreading exonucleases
In E. coli Pol III, the proofreading exonuclease function is provided by the ϵ subunit of the DNA polymerase III holoenzyme . We wondered if the presence of an ϵ subunit was a general feature of holoenzymes containing a Pol III protein with a variant PHP domain. The ϵ subunit of the DNA polymerase III holoenzyme, also known as DnaQ, is a member of the DEDD superfamily of DNAses and RNAses, which have in common a set of four strictly conserved acidic residues (DEDD) that are responsible for binding two catalytic metal ions . Within the DEDD superfamily, a distinction is made between the DEDDh and DEDDy subfamilies, based on whether a fifth conserved residue is a tyrosine (DEDDy) or a histidine (DEDDh), as in the E. coli DNA polymerase III ϵ subunit.
From the genomes of the 47 bacterial species represented in our C-family DNA polymerase sequence alignment, which include 12 from α-, β- or γ-proteobacteria, we extracted the sequences of 72 proteins containing a DEDD exonuclease domain (sequence alignment in Additional file 2). Visualizing the relatedness of the sequences as a phylogenetic tree (Figure 2B), we observed that the E. coli DNA polymerase III ϵ subunit is part of a clade of 12 sequences. The sequences within this clade belong exclusively to genomes of α-, β- or γ-proteobacteria. Each species is represented by one DEDDh sequence. No other class of bacteria was represented in the clade. We also observed that all the exonucleases in the clade contain a C-terminal tail homologous to the one that the E. coli ϵ subunit uses to bind to its Pol III α subunit partner, and these contain multiple highly conserved residue, two of which have been demonstrated to be important for binding to the PHP domain of the polymerase (in E. coli: His 225, Trp 241) [22, 23]. From these observations, we hypothesize that a bona fide ϵ subunit, which we define as being both a member of the DEDDh family of exonucleases and containing a C-terminal tail that binds to the α subunit, is only found in α-, β- and γ- proteobacteria. We stress that, while members of the DEDDh family of exonucleases are very common, few have been functionally characterized. Indeed, E. coli contains five such DNA exonuclease paralogs, but only its DNA polymerase III ϵ subunit is essential for viability . Although our sequence analysis cannot exclude the possibility that a different DEDDh subtype may be essential for some bacteria and involved in DNA replication proofreading, we find no support for the existence of a canonical DNA polymerase III ϵ subunit outside of α-, β- or γ- proteobacteria, whose DNA polymerase III PHP domain seems to have lost metal-binding capability. Therefore, we hypothesize that the proofreading activity for Pol III is supplied by either a metal-binding PHP domain or, in α-, β- or γ-proteobacteria, by a separate protein equivalent to the DNA polymerase III holoenzyme ϵ subunit in E. coli.
Re-introduction of the catalytic residues in the PHP domain of E. coli Pol III restores metal binding
Superposition of the PHP domains of E. coli Pol III (Eco), T. aquaticus Pol III (Taq) and G. kaustophilus Pol C (Gka) reveals a striking structural conservation of the domain. There is relatively low sequence identity between the domains of the three species: 38% over 270 residues between Eco and Taq, 26% over 270 residues between Eco and Gka and 24% over 280 residues between Taq and Gka; however the root mean square deviation of the Cα positions is only 1.20 Å (over 220 aligned atoms), 1.34 Å (over 182 aligned atoms) and 1.10 Å (over 174 aligned atoms) for the Eco/Taq, Eco/Gka and Taq/Gka superpositions, respectively.
Given that the structural similarity among the PHP domain is greater than that expected based on the level of sequence identity , we endeavoured to restore metal binding by reverting the variant residues to those found in canonical PHP domains. For this, we made three variants of E. coli Pol III with three, four or five mutations in the PHP domain, termed 3mPHP, 4mPHP, and 5mPHP, respectively (see Table 1). The first of these mutants, 3mPHP, has three histidine residues restored at positions 1, 4 and 9 (i.e. R10H, F44H, R203H). 4mPHP has an additional glycine to cysteine mutation introduced at position 7 (i.e. R10H, F44H, G134C, R203H), while 5mPHP has a fifth and final introduced mutation of Asp69 to glutamate at position 5 (i.e. R10H, F44H, D69E, G134C, R203H) to complete the canonical PHP metal-binding motif. All mutations were made in the truncated version of the polymerase (truncated after residue 917) that was used to determine the crystal structure .
X-ray diffraction data processing and model refinement statistics
E. coli 3mPHP
Unit cell (a,b, c in Å)
82.43, 98.94, 139.88
46.94 - 2.90
Rwork / Rfree
19.4 / 24.5
RMS(bonds) / RMS (angles)
0.003 / 0.733
Ramachandran favored (%)
Ramachandran outliers (%)
To identify the metals observed in the 3mPHP crystal structure, we analysed a protein sample treated in an identical manner to the solution used for crystallisation using X-ray fluorescence. We found that the X-ray fluorescence spectrum of our sample very closely resembles that of a zinc standard (Figure 4B). The two metal ions in the 3mPHP structure were, therefore, modelled as zinc.
Re-introduction of the catalytic residues in the PHP domain does not result in exonuclease activity
In vivo, the exonuclease activity of the ϵ subunit is dependent on the binding of Mg2+, while in vitro this activity is enhanced by replacing the Mg2+ with Mn2+ . As shown in Figure 5B and 5C, we observed that the exonuclease activity present in the 5mPHP sample is stimulated by both metals and that the activity with Mn2+ is higher than with Mg2+, as with the ϵ subunit. On the other hand, we found that Zn2+ has an inhibitory effect (Figure 5D). This is in contrast to the T. thermophilus Pol IIII exonuclease activity which was found to be Zn2+ dependent . It is therefore likely that the exonuclease activity measured in E. coli Pol III preparations, and especially in 5mPHP preparations, is caused by an impurity in the protein sample, most likely endogenous E. coli ϵ subunit.
The PHP domain provides stability to the polymerase
Interestingly, the unfolding of the protein appears to be a cooperative event as indicated by the single sigmoidal curve obtained by both melting temperature experiments monitored by circular dichroism and tryptophan fluorescence. Taken together, the two curves reveal that the secondary structure (as reported by circular dichroism) and tertiary structure (as reported by tryptophan fluorescence) of Pol III break down simultaneously in a two-state transition. The same single step unfolding was observed when the unfolding of the protein with the chemical denaturant guanidinium chloride as measured by circular dichroism (Figure 6B). Here too, we find that the protein apparently unfolds in a single step, with the 3mPHP mutant being less stable than the wild-type protein.
It is rather surprising that a protein as large as this Pol III catalytic construct (100 kDa) apparently unfolds in single step. Large multi-domain proteins often exhibit intermediate states of unfolding due to the sequential unfolding of the individual domains [27, 28]. The single step unfolding of E. coli Pol III indicates that the different domains unfold cooperatively. The correlation between the number of experimental mutations at the PHP domain and decrease in the Tm values of Pol III highlights how the PHP domain is structurally integrated with the rest of the polymerase. Our data therefore suggest that the PHP domain plays a crucial non-enzymatic role in stabilising the entire structure of Pol III.
Mutations at the PHP domain modulate polymerase activity
Our results emphasize the fact that bacterial replicative polymerases have maintained the structure of PHP domains that have lost metal-binding residues. The extent of this conservation is remarkable, as it has survived the billions of years of evolution subsequent to the split between those bacterial species that retained metal-binding residues (with presumed retention of the enzymatic activity) and those that have lost it, of which E. coli is the primary example. Biochemical studies have indicated that the association between the ϵ and α subunits involves the polymerase PHP domain and the C-terminal tail of ϵ [29, 30]. The correlation between loss of PHP domain activity and the presence of an ϵ homologue in the corresponding genome suggests that the strict structural conservation of the PHP domain might arise from the necessity to precisely position the active site of the trans exonuclease near the PHP cleft. Furthermore, the substantial decrease in polymerization activity and global stability of our PHP mutants clearly indicates that the scaffolding role of the PHP domain goes beyond the positioning of trans exonucleases. It suggests that the PHP domain is a major structural element for the stabilization of Pol III and plays a key role in the provision of optimal Pol III activity.
Protein sequence analysis
Replicative C-family polymerase sequences were retrieved using Protein BLAST at the Bioinformatics Toolkit hosted by the Max-Planck Institute for Developmental Biology (http://toolkit.tuebingen.mpg.de/prot_blast). The input sequence was E. coli Pol III, the database was nr_bac70 (the bacterial sequences in NCBI protein database filtered to a maximum of 70% identity), and all other parameters were defaults. The resulting alignment was manually edited to include only sequences identified as Pol III with high confidence. For easier visualization, this alignment was reduced to its 47 most diverse sequences using AlignmentViewer (http://toolkit.tuebingen.mpg.de/alnviz). To broaden the scientific interest of the polymerase sample set, some sequences in the reduced alignment were replaced with homologs from organisms of medical, industrial and/or historical interest. Because the alignment from Protein BLAST included only those sequence regions scored as homologous to E. coli Pol III, the complete sequence for each polymerase chosen for the sample set was retrieved using its GID number and RetrieveSeq (http://toolkit.tuebingen.mpg.de/gi2seq). These sequences were aligned using MAFFT L-INS-i (http://mafft.cbrc.jp/alignment/server/index.html), and a neighbor-joining phylogenetic tree was generated using the MAFFT server (settings: all gap-free sites, WAG substitution model, estimate heterogeneity among sites, bootstrap resampling = 100).
Proteins homologous to the DNA polymerase ϵ exonuclease subunit were gathered by (1) retrieving from the Conserved Domains Database  the sequence alignment of all 232 proteins used to define the conserved domain PRK05711 (DNA polymerase III subunit epsilon; Provisional) and (2) performing a blastp search (http://blast.ncbi.nlm.nih.gov/Blast.cgi?PROGRAM=blastp&BLAST_PROGRAMS=blastp&PAGE_TYPE=BlastSearch) limited to the organisms included in the Pol III sequence analysis. All sequences with expect scores equal to or better than that of the worst scoring sequence annotated as an ϵ exonuclease were kept and aligned using MAFFT L-INS-i, and a phylogenetic tree was generated using FastTree 2.1.7 . All sequence alignments were visualized using Jalview 2.8  and all trees with FigTree 1.3.1 and 1.4 (http://tree.bio.ed.ac.uk/software/figtree/).
Mutagenesis and protein purification
Mutations were introduced in a truncated version of E. coli DNA polymerase III α subunit (residues 1-917)  according to Table 1 using the Quikchange kit from Stratagene. All E. coli Pol III constructs included a N-terminal His6 tag, followed by a Prescission protease cleavage site and were expressed and purified using a protocol based on the method described in  , with the addition of a Ni-resin chromatography purification step after cell lysis.
Crystallisation and structure determination
Crystals of E. coli 3mPHP were grown using the hanging drop vapour diffusion method under conditions similar to the WT version  albeit with slightly lower precipitant concentration. Concentrated protein at 10-15 mg/ml was mixed with 15%-20% PEG3350, 0.2-0.4 M NaH2PO4, 100 mM HEPES pH 7.5. Crystals were frozen in mother liquor including 20% glycerol. The structure was solved by molecular replacement using the WT structure  as search model in PHASER . The model was further improved by multiple rounds of manual rebuilding in COOT  and refinement in REFMAC  and phenix.refine . Coordinates and structure factors for 3mPHP were deposited in the Protein Data Bank data with the accession code 4JOM.
Polymerase preparation for biochemical assays
Polymerase stock solutions were thawed from storage at -80°C, diluted with a concentrated monomerization buffer (supplemental concentrations after addition: 15% glycerol, 20 mM HEPES pH 7.4, 100 mM NaCl, 0.1 mM TCEP), and monomerized by incubation overnight at 15°C. Polymerases were confirmed to behave as monomers by size-exclusion chromatography on an S200 SMART column. For storage over several days at -20°C, a high-glycerol buffer was added as a cryoprotectant (supplemental concentrations after addition: 50% glycerol, 20 mM TAPS pH 8.5, 100 mM NaCl, 2.25 mM TCEP).
Exonuclease activity assays
Activity was detected using a novel, real-time fluorescence anisotropy assay. Single-stranded DNA labeled on its 3′-end was purchased from IDT (TAGGACAGTTCACGCTTCTTGG-TAMRA). Exonuclease activity at the 3′-end of the DNA first cleaves the TAMRA label from the DNA in a reaction with apparent first-order kinetics. Reactions (150 μL) were initiated by adding 2.8 μM protein to aliquots of 50 nM labeled DNA (buffer: 15% glycerol, 0.2 mg/mL BSA, 20 mM TAPS pH 8.5, 100 mM K glutamate, 10 mM MgCl2, 10 mM βME) in opaque black 96-well plates and monitored using a Perkin-Elmer Victor3 fluorescence plate reader with a 535/30 nm excitation filter, 595/60 nm emission filter, and an averaging time of 1 sec. The metal-dependence of the exonuclease activity was tested using an assay based on . The activity was tested in a buffer containing 40% glycerol, 1 mg/ml lysozyme, 20 mM HEPES at pH 7.5, 100 mM K glutamate and 0.5 mM TCEP. The Zn2+ titration was performed in the presence of 0.3 mM MnCl2.
Determination of melting temperature
Samples (1.5 mL) were individually subjected to temperature titrations with 1°C intervals separated by 1 minute equilibration periods. Data collection occurred in a sealed 4-mL Hellma quartz cuvette using a FluoroMax-3 (Jorbin Yvon Horiba) fluorometer with a Wavelength Electronics Model LFI-3751 temperature controller. Excitation occurred at 280 nm (slit width 3.5 nm), and emission scans were collected from 295 to 397 nm (slit width 7 nm) in 2 nm increments with 0.5 s of integration time and then converted into scan centres of mass. KaleidaGraph (Synergy Software) was used for curve fitting to a standard temperature melt equation. Details on data reduction and curve fitting are provided in Additional file 3.
An individual sample (1 mL) was prepared for each data point, and all samples were allowed to equilibrate at 25°C overnight (roughly 18 h). Samples were held in a 4-mL Hellma quartz cuvette during analysis. Circular dichroism was measured in kinetic mode on a Circular Dichroism Spec 410 (AVIV Biomedical) at 226 nm using 60 separate 1-sec reads. The reads for each sample were averaged and normalized by conversion into units of mean residue ellipticity. After circular dichroism analysis, a FluoroMax-3 (Jorbin Yvon Horiba) was used to assay tryptophan fluorescence. Excitation occurred at 280 nm (slit width 2 nm), and emission scans were collected from 295 to 397 nm (slit width 4 nm) in 2 nm increments with 0.5 s of integration time. Each scan was reduced to its centre of mass. Increasing Gdn•HCl caused a non-linear shift of the scan centres of mass toward longer wavelengths. To simplify curve fitting, the points corresponding to the highest Gdn•HCl concentration (3 M) samples were omitted during data analysis. KaleidaGraph (Synergy Software) was used for curve fitting to a standard denaturant melt equation. Details on data reduction and curve fitting are provided in Additional file 3.
DNA polymerization assay
DNA polymerization was monitored by fluorescence intensity using a slightly modified version of the standard PicoGreen-based quench assay [38–40] in 96-well format. The substrate was a DNA primer-template complex generated by annealing two oligomers: Template 5′-TTGTGGGTAGATAAATACAGACCTAAGTCCTTGAATGCCGCGTGCGTCCC and Primer 5′-GGGACGCACGCGGCATTCAAGGA. The assays were performed at 250 nM polymerase, 7.5 nM DNA and 50 μM of each dNTP. The reaction buffer included 20 mM HEPES at pH 7.5, 100 mM NaCl, 0.2 mg/ml BSA, 3 mM MgCl2 and 0.5 mM TCEP. Time point samples were taken until the reactions reached completion. After quenching the last sample, 90 minutes were allowed for fluorescence development before data acquisition on a Perkin-Elmer Victor3 fluorescence plate reader using a 535/30 nm excitation filter, 595/60 nm emission filter, and an averaging time of 1 sec. To achieve stable readings, samples were scanned 9 times, and the last 5 reads were used for curve fitting. Rate constants were fit in KaleidaGraph using a standard exponential function.
The authors thank members of the Kuriyan laboratory, in particular Markus Seeliger, Jeff Iwig and Margaret Stratton for helpful discussions. In addition, we thank Tiffany Chou for help with construction of the PHP domain mutants. This work was supported in part by a grant from the National Institutes of Health to JK (GM45547) and by a EMBO Long-term fellowship to TB (ALTF 576-2009).
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