Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Biomol NMR Assign. Author manuscript; available in PMC 2017 April 1.
Published in final edited form as:
PMCID: PMC4789116

Backbone chemical shift assignments for Xanthomonas campestris peroxiredoxin Q in the reduced and oxidized states: a dramatic change in backbone dynamics


Peroxiredoxins (Prx) are ubiquitous enzymes that reduce peroxides as part of antioxidant defenses and redox signaling. While Prx catalytic activity and sensitivity to hyperoxidative inactivation depend on their dynamic properties, there are few examples where their dynamics has been characterized by NMR spectroscopy. Here, we provide a foundation for studies of the solution properties of peroxiredoxin Q from the plant pathogen Xanthomonas campestris (XcPrxQ) by assigning the observable 1HN, 15N, 13Cα, 13Cβ, and 13C′ chemical shifts for both the reduced (dithiol) and oxidized (disulfide) states. In the reduced state, most of the backbone amide resonances (149/152, 98%) can be assigned in the XcPrxQ 1H-15N HSQC spectrum. In contrast, a remarkable 51% (77) of these amide resonances are not visible in the 1H-15N HSQC spectrum of the disulfide state of the enzyme, indicating a substantial change in backbone dynamics associated with the formation of an intramolecular C48-C84 disulfide bond.

Keywords: plant diseases, peroxidase, drug design, conformational change

Biological context

Under aerobic conditions cells are constantly exposed to reactive oxygen species, such as H2O2, organic peroxides, and hydroxyl radicals, which can damage cellular DNA, proteins, and lipids (Winterbourn, 2008). Aerobic organisms counter with an arsenal of antioxidant proteins including peroxiredoxins (Prxs), a family of ubiquitous, cysteine-based peroxidases that play influential roles in the turnover of peroxides including peroxynitrite (recently reviewed in Perkins et al. 2014). In eukaryotes Prxs also appear to regulate “non-oxidative-stress-related” signaling events (Rhee et al. 2012). All six known Prx subgroups – Prx1, Prx5, Prx6, Tpx, AhpE, and PrxQ (also called BCP) – require for catalysis a “peroxidatic” Cys (CP) in a “fully folded” (FF) conformation that binds and reacts with the peroxide substrate to form Cys-sulfenic acid (CP-SOH) and H2O. For most Prxs, the protein then undergoes substantial rearrangement to some type of a “locally unfolded” (LU) conformation allowing the CP-SOH to form a disulfide bond with a “resolving” Cys (CR) that can be located in various parts of the protein. Despite the importance of this FF/LU conformational change, little is still known about its thermodynamics or dynamics. Among Prxs, only the PrxQ subgroup includes members that, as monomers of ~20 kD, are amenable to facile study by NMR spectroscopy; and Arabidopsis thaliana PrxQ (AtPrxQ), with its CR in helix-2, is the only PrxQ so far characterized by NMR. This latter study revealed a substantial dynamical disorder on the μs-ms timescale especially for the reduced (dithiol) form (Ådén et al. 2011).

We are initiating NMR studies of another monomeric PrxQ that undergoes a different conformational change, because it has its CR residue in helix-3. Xanthomonas campestris PrxQ (XcPrxQ), from an economically important plant pathogen (Leyns et al. 1984), is 160-residues long and has relevance for drug design (Myler et al 2009) as it is representative of PrxQs from various genera of human pathogens such as Burkholderia (63% sequence identity), Helicobacter (46%), Cryptosporidium (42%), Toxoplasma (41%), Mycobacterium (40%), and Acanthamoeba (38%). Also, high resolution crystal structures for the disulfide (LU) conformation and the pseudo-dithiol (FF) conformation (Cys→Ser double variant) show that the transition from FF to LU involves a substantial unfolding of helix-3 (Liao et al. 2009). With plans to directly measure how redox state, mutations, and post-translational modifications modulate the enzyme’s structure and dynamics, and to enable chemical shift perturbation studies of ligand binding (e.g. Buchko et al. 1999), we have fully assigned the observable 1HN, 15N, 13Cα, 13Cβ, and 13C′ chemical shifts of native XcPrxQ in both the disulfide (LU) and dithiol (FF) states.

Methods and experiments

Cloning, expression, and purification

The full length X. campestris strain 17 prxQ gene (GI:499347373) was synthesized by GenScript (Cambridge, MA) and cloned into the NcoI and HindIII restriction sites of the expression vector pTHCm (Nelson et al. 2008) such that the expressed protein used in our studies contained no non-native residues. The expression vector was then transformed into Escherichia coli BL21(DE3) cells. Uniformly 15N-, 13C-labeled XcPrxQ was obtained by growing the transformed cells (310 K) in 750 mL of minimal medium containing 15NH4Cl (1 mg/mL) and D-[13C6]glucose (2.0 mg/mL), NaCl (50 μg/mL), MgSO4 (120 μg/mL), CaCl2 (11 μg/mL), FeCl3 (10 ng/mL) and the antibiotic ampicillin (100 μg/mL). When the cell culture reached an OD600 reading of ~0.8, it was transferred to a 298 K incubator and protein expression induced with isopropyl β-D-1-thiogalactopyranoside (0.026 μg/mL). Cells were harvested approximately four hours later by mild centrifugation, and frozen at 193 K. After thawing the frozen pellet, the cells were resuspended in 40 mL of 20 mM TrisHCl, pH 7.5 and lysed by three passes through a French Press (SLM Instruments, Valencia, CA) followed by sonication for 1 min. Insoluble cell debris was removed by centrifugation at 25,000g for 1 h with the supernatant dialyzed overnight (278 K) in 5 L of 20 mM TrisHCl, pH 7.5. XcPrxQ was purified from this solution using two chromatography steps: High Q ion exchange (5 mL Bio-Rad Econo-Pac cartridge (Bio-Rad, Hercules, CA)) and Superdex75 gel-filtration (HiLoad 16/60 column). In the first chromatography step, approximately 5 – 10 mg of total protein was applied to the High Q column pre-equilibrated with 25 mM TrisHCl, pH 7.5 (flow rate = 5 mL/min). Oxidized XcPrxQ did not bind to the column and eluted in the flow-through, which was concentrated to ~10 mg/mL before application to a size exclusion column pre-equilibrated with 100 mM NaCl, 20 mM Tris-HCl, pH 7.1 (flow rate = 1 mL/min). To obtain XcPrxQ in the reduced state, the size exclusion buffer was made 1.0 mM in dithiothreitiol (DTT) prior to the addition of the High Q fraction (which was also made 1.0 mM in DTT before loading onto the column). The major fractions contained XcPrxQ and these were pooled and concentrated to approximately 25 mg/mL for the NMR studies.

Nuclear magnetic resonance spectroscopy

The NMR data were all collected at 293 K on a double-labeled (13C, 15N) sample (~1.1 mM) of XcPrxQ in the oxidized and reduced states using four-channel Varian Inova-600, -750, and -800 spectrometers equipped with triple resonance probes and pulse field gradients. For the reduced state, assignment of the 1HN, 13C, and 15N backbone and some of the 13C side chain resonances were made from a standard two-dimensional 1H-15N HSQC experiment and three-dimensional HNCACB, HNCA, HNCACO, HNCO, CC-TOCSY-NNH and 15N-edited NOESY experiments using the Varian Biopack suite of pulse programs. A mixing time of 80 ms was used in the 15N-edited NOESY experiment and the HNCACB experiment was optimized for the detection of β-carbons using a tauCC value of 7 ms. For the oxidized state, assignment of the 1HN, 13C, and 15N backbone and some of the 13C side chain resonances were made from a standard two-dimensional 1H-15N HSQC experiment and three-dimensional HNCACB (13C β-optimized), HNCO, and HNN experiments. An overall rotational correlation time (τc) for XcPrxQ in both states was estimated from backbone amide 15N T1/T ratios. Felix2007 (MSI, San Diego, CA) and Sparky (v3.115) were used to process and analyze, respectively, all the NMR data. Indirect methods (DSS = 0 ppm) were used to reference the 1H, 13C, and 15N chemical shifts that were deposited into the Biological Magnetic Resonance Bank database ( under the BMRB accession numbers 25557 (reduced) and 25566 (oxidized).

Assignments and data deposition

The expectation that XcPrxQ is monomeric in solution (Liao et al. 2009) is corroborated by the elution of both the disulfide and dithiol forms of the enzyme from the gel filtration column with an identical retention time of ~89 min characteristic of a ~17 kDa protein under our conditions (data not shown). Also, the isotropic overall rotational correlation times (τc) inferred from 15N spin relaxation times were the same for XcPrxQ in the oxidized and reduced states, 10.6 ± 0.1 ns at 293 K, a value consistent with an ~17 kDa protein. Finally, the reasonably efficient magnetization transfer through the side chain carbons in the three-dimensional NMR experiments used to make chemical shift assignments in both the reduced and oxidized states would be unlikely with a protonated protein if XcPrxQ formed ~35 kDa dimers (Yee et al., 2002).

The features in the 1H-15N HSQC spectrum for reduced (dithiol) XcPrxQ (Fig. 1A) are characteristic of a folded protein (Yee et al. 2002): cross peaks of near uniform intensity that are dispersed in both the proton and nitrogen dimensions. Out of the 152 expected backbone amide resonances (160 – (7 Pro + Met1), 149 were assigned (98%) to cross peaks in the 1H-15N HSQC spectrum. The three unassigned residues, Y40, S44, and G47, could not be unambiguously assigned to the three remaining unassigned amide cross peaks due to poor magnetization transfer to carbon resonances in the three-dimensional backbone assignment experiments. The chemical shifts for the β-carbons of the two catalytic cysteine residues CP (C48) and CR (C84) are 32.2 and 26.9 ppm, respectively. Such chemical shifts are consistent with cysteine residues in the Cys-thiol/thiolate state (Sharma and Rajarsthnam, 2000) expected in the presence of the reducing agent DTT.

Fig. 1Fig. 1
1H-15N HSQC spectra of dithiol and disulfide forms of XcPrxQ. (A) 1H-15N HSQC spectrum for dithiol XcPrxQ (~1.5 mM) collected at a proton resonance frequency of 750 MHz, 293 K, in 100 mM NaCl, 20 mM Tris, 1 mM DTT, pH 7.0, with labels for assigned amide ...

For oxidized XcPrxQ, the wide chemical shift dispersion of the cross peaks in both the proton and nitrogen dimensions in the 1H-15N HSQC spectrum (Fig. 1B), is similarly characteristic of a folded protein (Yee et al. 2002). However, relative to the 1H-15N HSQC spectrum for reduced XcPrxQ (Fig. 1A), the intensity of the cross peaks are not as uniform and approximately half (51%, 77/152) of the expected amide resonances are missing under conditions that are identical except for the absence of the reducing agent DTT. Collecting the reduced 1H-15N HSQC spectrum at 283 K instead of 293 K did not increase the number of observable amide resonances. Of the remaining amide cross peaks observed for disulfide XcPrxQ, many overlay with amide cross peaks for reduced XcPrxQ, allowing a rapid tentative assignment based on the assignments made for the dithiol form. The analysis of the HNCACB (β-optimized) and HNN data collected on oxidized XcPrxQ confirmed the tentative assignments and also allowed the assignment of the remaining amide cross peaks (Fig. 1B). In the disulfide XcPrxQ spectrum, a second set of less intense cross peaks were observed for residues A4 – L6 (blue in Fig. 1B) reflecting a second minor orientation at the N-terminus, a region that is disordered in both crystal structures (Laio et al. 2009). The amides of C48 and C84 are not observed in the 1H-15N HSQC spectrum for oxidized XcPrxQ, and therefore, the presence of the disulfide bond cannot be confirmed from cysteine β-carbon chemical shifts.

While there are some intensity differences in corresponding amide resonances in the reduced and oxidized forms of XcPrxQ, most of the chemical shift differences are small, less than 0.1 ppm (Fig. 2A). This suggests that in the regions of the protein where amide resonances are observed in both states, the structure is rather similar. To see which parts of the protein correspond to the amide resonances missing in the 1H-15N HSQC spectrum of oxidized XcPrxQ, we have mapped them onto the primary amino acid sequence of XcPrxQ (Fig. 2B). Interestingly, all five amide resonances with average chemical shift perturbations greater than 0.1 ppm – S20, G22, A57, F91, and L119 (Fig. 2A) – are adjacent to or just one residue away from a segment with missing amide resonances suggesting that a structural change is associated with at least some of the missing amide resonances. Such a structural difference is indeed seen in comparing the XcPrxQ LU disulfide (3GKK) and FF pseudo-dithiol (3GKM) crystal structures (Fig. 2C). Though much of the structure changes minimally, substantial rearrangements occur in the length and orientation of helix-3 (circled in Fig. 2C), which contains the resolving cysteine, CR, and extends from V77 – Q87 in the pseudo-dithiol structure (blue), but only from C84 – Q87 in the disulfide structure (gold), with electron density absent from K78 – D81 consistent with that region being disordered (Laio et al. 2009).

Fig. 2
Mapping dithiol versus disulfide XcPrxQ chemical shift changes onto the sequence and structure. (A) Plot of the root-mean-square combined chemical shift perturbation in the 1H and 15N resonances (Δrms = [((Δ1HN)2 + (Δ15N/5)2)/2)] ...

Missing amide cross peaks in 1H-15N HSQC spectra of proteins typically identify regions undergoing motion or chemical exchange in the intermediate timescale (ms to μs) (e.g. Buchko et al. 1999; Ådén et al. 2011) and are often associated with regions involved in binding and catalysis (e.g. Müller et al. 1992). A mapping of the amide resonances missing in the 1H-15N HSQC spectrum of oxidized XcPrxQ onto the crystal structure of pseudo-dithiol XcPrxQ (Fig. 2D) reveals that the extensive set of missing resonances (red) are all surrounding the region (helix-3) that is the focal point of the FF/LU conformation change. The disappearance of ~50% of the amide resonances in the 1H-15N HSQC spectrum of reduced XcPrxQ upon the formation of a single intramolecular CP-CR disulfide bond indicates the protein has undergone a drastic change in backbone dynamics even for the nearby residues that do not apparently change much in their average conformation. This is fascinatingly reminiscent of, but opposite to, the large changes in dynamics observed for AtPrxQ. For AtPrxQ nearly 50% of the backbone amide peaks in the 1H-15N HSQC spectrum were absent when in the reduced state (Ådén et al. 2011). As noted in the introduction, the AtPrxQ CR is located in helix-2 rather than helix-3, and so it is plausible that these enzymes truly do experience very different structural and dynamics changes in their FF/LU transitions. Although further analysis is beyond the scope of this assignments report, these results make clear that different Prxs can behave very differently and that the chemical shift assignments presented here for XcPrxQ in the dithiol and disulfide states provide a basis for what will be very informative studies that provide residue-level insights into the interplay of Prx function, structure, and dynamics.


This research was supported by the National Institute of Health R01 Grant number GM050389 (LBP and PAK). A large part of this research was performed at the W.R. Wiley Environmental Molecular Sciences Laboratory (EMSL), a national scientific user facility located at Pacific Northwest National Laboratory (PNNL) and sponsored by U.S. Department of Energy’s Office of Biological and Environmental Research (BER) program. Battelle operates PNNL for the U.S. Department of Energy.


  • Ådén J, Wallgren M, Storm P, Weise CF, Christiansen A, Schroder WP, Funk C, Wolf-Watz M. Extraordinary μs-ms backbone dynamics in Arabidopsis thaliana peroxiredoxin Q. Biochem Biophys Acta. 2011;1814:1880–1890. [PubMed]
  • Buchko GW, Daughdrill GW, de Lorimier R, Rao BK, Isern NG, Lingbeck JM, Taylor JS, Wold MS, Gochin M, Spicer LD, Lowry DF, Kennedy MA. Interactions of human nucleotide excision repair protein XPA with DNA and RPA70ΔC327: Chemical shift mapping and 15N NMR relaxation studies. Biochemistry. 1999;38:15116–15128. [PMC free article] [PubMed]
  • Leyns F, De Cleene M, Swings J, De Ley J. The host range of the genus Xanthomonas. Bot Rev. 1984;50:305–355.
  • Liao S-J, Yang C-Y, Chin K-H, Wang A, H-J, Chou S-H. Insights into the alkyl peroxide reduction pathway of Xanthomonas campestris bacterioferritin comigratory protein from the trapped intermediate–ligand complex structures. J Mol Biol. 2009;390:951–966. [PubMed]
  • Maiti R, Van Domselaar GH, Zhang H, Wishart DS. Superpose: a simple server for sophisticated structural superposition. Nucl Acids Res. 2004;32:W590–W594. [PMC free article] [PubMed]
  • Müller CW, Schlauderer GJ, Reinstein J, Schulz GE. Adenylate kinase motions during catalysis: an energetic counterweight balancing substrate binding. Structure. 1992;4:147–156. [PubMed]
  • Myler PJ, Stacy R, Stewart LJ, Staker BL, Van Voorhis WC, Varani G, Buchko GW. The Seattle Structural Genomics Center for Infectious Disease (SSGCID) Infect Diseases Drug Targets. 2009;9:493–506. [PMC free article] [PubMed]
  • Nelson KJ, Parsonage D, Hall A, Karplus PA, Poole LB. Cysteine pKa values for the bacterial peroxiredoxin AhpC. Biochemistry. 2008;47:12860–12868. [PMC free article] [PubMed]
  • Perkins A, Poole LB, Karplus PA. Tuning of peroxiredoxin catalysis for various physiological roles. Biochemistry. 2014;53:7693–7705. [PMC free article] [PubMed]
  • Rhee SG, Woo HA, Kil IS, Bae SH. Peroxiredoxin functions as a peroxidase and a regulator and sensor of local peroxides. J Biol Chem. 2012;287:4403–4410. [PMC free article] [PubMed]
  • Sharma D, Rajarathnam K. 13C NMR chemical shifts can predict disulfide bond formation. J Biomol NMR. 2000;18:165–171. [PubMed]
  • Winterbourn CC. Reconciling the chemistry and biology of reactive oxygen species. Nat Chem Biol. 2008;4:278–286. [PubMed]
  • Yee A, Chang X, Pineda-Lucena A, Wu B, Semesi A, Le B, Ramelot T, Lee G, Bhattacharyya S, Gutierrez P, Denisov A, Lee C-H, Cort JR, Kozlov G, Liao J, Finak G, Chen L, Wishart D, Lee W, McIntosh LP, Gehring K, Kennedy MA, Edwards AM, Arrowsmith CH. An NMR approach to structural proteomics. Proc Natl Acad Sci USA. 2002;99:1825–1930. [PubMed]