Determination of Redox Sensitivity in Structurally Similar Biological

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Determination of Redox Sensitivity in Structurally Similar Biological Redox Sensors Anil Kumar Jamithireddy, Rudra Narayan Samajdar, Balasubramanian Gopal, and Aninda Jiban Bhattacharyya J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.7b02081 • Publication Date (Web): 28 Jun 2017 Downloaded from http://pubs.acs.org on July 3, 2017

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The Journal of Physical Chemistry

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Determination of Redox Sensitivity in Structurally

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Similar Biological Redox Sensors

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Anil K. Jamithireddy a, ‡, Rudra N. Samajdar b, ‡, B. Gopal a and Aninda J. Bhattacharyya b, *

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a

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Molecular Biophysics Unit, Indian Institute of Science, Bangalore 560012, India.

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Solid State and Structural Chemistry Unit, Indian Institute of Science, Bangalore 560012,

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India.

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ABSTRACT: Redox stimuli govern a variety of biological processes. The relative sensitivity of

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redox sensors plays an important role in providing a calibrated response to environmental stimuli

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and cellular homeostasis. This cellular machinery plays a crucial role in the human pathogen

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Mycobacterium tuberculosis as it encounters diverse microenvironments in the host. The redox

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sensory mechanism in Mycobacterium tuberculosis is governed by two component and one-

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component systems, alongside a class of transcription factors called the extra cytoplasmic

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function (ECF) σ factors. ECF σ factors that govern the cellular response to redox stimuli are

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negatively regulated by forming a complex with proteins called zinc associated anti – σ factors

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(ZAS). ZAS proteins release their cognate σ factor in response to oxidative stress. The relative

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sensitivity of the ZAS sensors to redox processes dictate the concentration of free ECF σ –

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factors in the cell. However, factors governing the redox threshold of these sensors remain

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unclear. We describe here, the molecular characterization of three σ – factor/ZAS pairs -

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σL/RslA, σE/RseA and σH/RshA using a combination of biophysical and electrochemical

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techniques. This study reveals conclusively, the differences in redox sensitivity in these proteins

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despite apparent structural similarity, and rationalizes the hierarchy in the activation of the

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cognate ECF σ factors. Put together, the study provides a basis for examining sequence and

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conformational features that modulate redox sensitivity within the confines of a conserved

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structural scaffold. The findings provide the guiding principles for the design of intracellular

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redox sensors with tailored sensitivity and predictable redox thresholds providing a much needed

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biochemical tool for understanding host-pathogen interaction.

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Mechanisms sensitive to redox events govern diverse biological processes. In prokaryotes, redox

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sensors regulate the cellular response to changing microenvironments and are often coupled with

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other sensing mechanisms to provide a nuanced cellular response to environmental stimuli.

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Mycobacterium tuberculosis is an important respiratory pathogen in humans. Once the bacterium

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infects the host, as a part of the immune response against the invading pathogen, macrophages

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phagocytize the bacterium. The growth of the bacterium inside the host is predominantly

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determined by these activated macrophages. Inside the macrophages, reactive oxygen

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intermediates (ROI) and reactive nitrogen intermediates (RNI) are produced by the host

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enzymes. These ROIs and RNIs are critical for controlling growth of Mycobacterium

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tuberculosis in macrophages1. Despite the toxic effects of ROIs and RNIs, Mycobacterium

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tuberculosis persists within macrophages, with its defense system involving cell wall-associated

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lipids, secretory redox buffers and secretory antioxidant enzymes as anatomical barriers2.

INTRODUCTION


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Redox enzymes carry out the detoxification of oxidative damage to proteins and DNA in

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Mycobacterium tuberculosis. Thioredoxins and ribonucleotide reductases are some of the

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important redox proteins in Mycobacterium tuberculosis. These proteins with either CXXC or

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TXXC motifs sense and maintain the redox balance3. The CXXC motif is a prevalent motif in

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these proteins that helps in maintaining thiol-disulphide homeostasis4. Disulphide bonds play an

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important role in the maintenance of protein structure and function5.

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Mycobacterium tuberculosis explores multiple mechanisms in order to survive the severe

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oxidative stress conditions in the host5. Regulation of transcription is one such mechanism that

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helps the bacterium to adapt to or overcome adverse conditions. Among the several factors

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responsible for global transcriptional regulation, sigma (σ) factors have a prominent role. σ

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factors are small interchangeable subunits of RNA-polymerase that are required for specific

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transcription initiation. Mycobacterium tuberculosis has thirteen σ factors of which σH, σE, σL, σF

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and the stress response σ factor σB have been reported to be involved in the redox stress

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response6. These σ factors rely on anti-σ factors or other receptors (as in the case of σB), as they

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are unable to sense and respond to external stress signals alone.

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The disulfide based oxidative response is diverse in bacteria. It mainly involves negatively

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regulating receptor proteins such as anti-σ factors7, 8. The anti-σ domain in an anti – σ factor was

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first identified based on a compact arrangement of an α helical bundle that interacts with two σ

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domains9. Anti – σ factors that negatively regulate redox σ factors possess a conserved

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HXXXCXXC motif that can coordinate with zinc (II)6. The zinc ions participate in the thiol-

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disulfide redox switch. The anti – σ factors that associate with zinc (II) are termed as zinc –

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associated anti – σ factors10. These ZAS proteins detect oxidative stress and release the bound σ

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factor as shown in Figure 1A. There are three anti – σ factors regulating the ECF σ factors σH,


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σL, and σE, that have the HXXXCXXC motif8. Structural zinc (II) ion coordinates with the ZAS

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proteins and stabilizes them; thus increasing the reactivity of the cysteine thiol groups in the

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CXXC motif. The zinc (II) acts as a Lewis acid and lowers the pKA of cysteine-thiols, increasing

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their nucleophilicity and ability to interact with host induced electrophilic peroxide and

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superoxide radicals. The metal coordination also prevents the two conserved cysteines from

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forming a disulfide bond in the absence of redox stress. These three proteins contain a

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structurally well-conserved N-terminal anti-σ domain (ASD) within which the HXXXCXXC

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motif is located.

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Figure 1A. Mechanism by which redox sensitive anti σ factors regulate their corresponding σ

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factor. Figure 1B. Sequence analysis of well – characterized zinc associated anti σ factors

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reveals a conserved HXXXCXXC motif that coordinates with zinc (II). The residues that

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coordinate zinc (II) are marked by red asterix and blue inverted triangles. The interactions of His,

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Cys and Cys from the conserved motif with the zinc moiety are depicted in solid blue lines. The

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fourth coordination with Zinc (II), formed with His / Cys residue, located 21-26 residues towards

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the N-terminal from the HXXXCXXC motif, is depicted by a dotted red line. The conserved anti

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sigma domain is marked by green colored helices. RspChrR = Rhodomonas spheroides ChrR,

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Mtb-RseA/RshA/RslA = Mycobacterium tuberculosis RseA/RshA/RslA respectively, and

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ScoRsrA = Streptomyces coelicolor RsrA.

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Comparison between the structures of diverse ASDs reveal that local sequence and

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conformational variations guide the sensitivity of the zinc sensor. Zinc (II) is bound by two

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cysteine residues in the CXXC motif as shown in Figure 1B and Figure 2. The nature of the

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residues between the two cysteines is reported to govern the functional role of this motif in

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different proteins10. In an effort to understand if conformational features specific to the ASD lead

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to differences in the observed redox-sensitivity, we examine the location of the two zinc (II)

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coordinating residues in the Ramachandran plot (Figure 3). Other anti-σ domains, which are not

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redox sensitive, are also included in this analysis. A sequence propensity profile of different

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ASDs is shown in Figure 1B. We note that these features can contribute to local changes in the

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pKA of the zinc (II) coordinating cysteines thereby resulting in altered redox sensitivity.


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Figure 2. Comparison of the ASDs of Rhodobacter sphaeroides ChrR (Lime green), Escherichia

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coli RseA (Cyan), Mycobacterium tuberculosis RslA(Magenta), RskA(Sandy brown) and

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RsdA(Deep blue). The ASDs represented as backbone traces are superimposed over the first

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three α helices (H1–H3). Despite the poor sequence similarity, the structures of ASDs are well

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conserved. Shown in-set is the Rsp ChrR HXXXCXXC motif coordinated to zinc (II).

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Figure 3. Ramachandran plot for the HXXXCXXC cysteine residues for 496 structurally

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characterized proteins. Shown in red are the cysteine residues of the two structurally

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characterized ZAS proteins (Mtb RslA and Rsp ChrR).

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In this work, we examine the redox response of the σL/RslA, σH/RshA, and σE/RseA

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complexes. All three complexes are known to bind zinc (II) ion through the CXXC motif in their

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native state. Upon oxidative stress, the zinc (II) ion is released freeing the two proximal cysteine


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thiols thereby enabling the formation of a disulfide bond between the proximal cysteines and

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conformational changes in the anti – σ factors. The change in conformation, coupled with the

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release of zinc (II) ion, and formation of disulfide linkage have been studied using spectroscopic

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and electrochemical techniques.

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We attempt to establish the redox sensitivities of these complexes from the complementary

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electrochemical and spectroscopic data and use a structural argument for rationalizing the

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observed redox behavior. This work provides a new perspective by virtue of the diverse

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techniques used to understand, both qualitatively and quantitatively, the response of different

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proteins towards oxidative stress. It also gives a comparative picture of the responses of the three

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different ZAS anti – σ factors in Mycobacterium tuberculosis, to oxidative stress. We believe

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that these studies could have potential implications for engineering genetic tools to understand

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host – pathogen interactions, eventually leading to therapeutic intervention.

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EXPERIMENTAL SECTION

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Expression and purification of proteins. The σE/RseA, σL/RslA, and σH/RshA complexes are

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purified with a co-expression and co-purification strategy using the pETDuet-1 (Novagen, Inc.)

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expression vector. All protein samples are purified using the same protocol. Plasmids with genes

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encoding σE/RseA, σL/RslA, and σH/RshA are transformed into E. coli Lemo21 (DE3), Rosetta

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(DE3) pLysS, and BL21 (DE3)* expression cell strains (Novagen, Inc.) respectively. The cells

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are grown in Luria broth with antibiotic (100 µg/ml ampicillin) to an A600 of 0.5–0.6. The cells

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are induced with 0.5 mM IPTG. Subsequently, growth temperature is lowered to 290 K, and the

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cells are grown for 12–18 h before they are spun down. Following this, cells are lysed by

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sonication in lysis buffer (60 mM Tris (pH 7.5) and 250 mM NaCl). Cell-free lysate is incubated


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with Ni+2-NTA beads and eluted by an imidazole gradient in the elution buffer (60 mM

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phosphate, pH 7.5, 250 mM NaCl, and 200 mM imidazole). Recombinant proteins are further

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purified by size-exclusion chromatography using a Sephacryl S-200 column (GE Healthcare Life

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Sciences) after the affinity chromatography step. For analytical gel-filtration experiments, ∼ 0.2-

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mg to 0.5-mg protein samples are passed through a Superdex S-200 10/300 GL column

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equilibrated in 25 mM phosphate (pH 7.5) and 128 mM NaCl at 6 °C. The proteins are then

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concentrated using a membrane-based centrifugal ultrafiltration system (Amicon-Ultra).

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Spectroscopic assay for the kinetics of zinc (II) release. Metal chelator 4-(2-

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pyridylazo)resorcinol (PAR), when complexed with free zinc (II), absorbs light intensely at 500

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nm11. Protein samples are diluted to 10 µM in a buffer containing 0.1 mM PAR for the

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experiment. The buffer is purged with nitrogen gas prior to the experiment. Nonspecifically

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bound zinc in the protein solution is trapped by PAR in the absence of oxidation, and seen by a

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slight increase in absorbance at 500 nm after addition of PAR. The solution is allowed to

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stabilize for five to ten minutes after addition of PAR. After the stabilization of absorbance, 10

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mM H2O2 is added and the absorbance at 500 nm is monitored every 10 seconds for 40 minutes

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at 25 °C. To avoid generation of air bubbles, the solution in the cuvette is continuously stirred

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via a magnetic fly. The absorbance measurements are done on a V 630Bio Jasco UV – vis

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spectrophotometer. The absorbance kinetics is fit using a standard first order kinetic model12 as

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shown in Equation 1.

150 = − (1) 151


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Here A refers to the absorbance at 500 nm at time t sec, Amax refers to the maximum absorbance,

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while k refers to the pseudo first order rate constant in s-1. The half-lives are calculated from the

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pseudo first order rate constant k obtained from the fitting using the transformation valid for first

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order kinetics12 (Equation 2):

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=

. (2)

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Circular Dichroism. Circular dichroism spectra are recorded on a JASCO-J715 instrument.

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All spectra are recorded using 5 mM Phosphate buffer adjusted to pH 7.4, with 1 mM DTT,

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using a 1 mm path length cuvette. Protein concentrations are kept in the range of 5–10 µM. Each

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reported spectrum is an average of three scans. The spectra of each protein in the absence and

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presence of 10 mM H2O2 are compared to understand the secondary structural changes induced

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by oxidative stress.

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Spectroscopic assay for determining exposed cysteines in the CXXC motif. The ready

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availability of cysteine thiols on the surface of the protein molecules determines the sensitivity of

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the proteins to redox stress. Ellman’s reagent (5,5'–dithiobis-2-nitrobenzoic acid) (DTNB) is

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known to react with exposed cysteine thiols in proteins leading to the formation of a yellow

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colored compound (TNB2-) with an absorption maximum at 412 nm13. This reaction is employed

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to determine the surface exposure of cysteines in the proteins. The reaction is carried out in 25

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mM tris – HCl, pH 7.4, and 150 mM NaCl buffer. To a final concentration of 10 µM protein in

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100 µL reaction mixture, 10 µL of 4mM DTNB solution is added at room temperature and

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absorbance at 412 nm is recorded at 10 second intervals up to 40 minutes at 25 °C on a V 630Bio


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Jasco UV–vis spectrophotometer. The number of reactive cysteine residues in these molecules is

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calculated using the standard value of molecular coefficient (ε) for TNB2- in dilute buffer as

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14150 M-1 cm-1 following literature14.

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Cyclic Voltammetry. The cyclic voltammetry measurements on the proteins are performed on

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a screen printed carbon electrode (SPCE) using a DropSens µStat 400 electrochemical

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workstation. The SPCEs contain the working electrode (carbon), reference electrode (silver), and

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the counter electrode (carbon) integrated within a single ceramic chip approximately 3 cm X 1cm

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X 0.5 cm in dimensions. Pre-treatment of the SPCEs include running a cyclic voltammogram

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with 1 M sulphuric acid within the potential window 0 to 1.4 V over several cycles till a stable

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response is seen, followed by cyclic voltammogram with 1 mM K3[Fe(CN)6] – K4[Fe(CN)6]

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mixture within the potential window -1.0 to 1.0 V (Supporting Figure S1). Following this, the

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SPCEs are thoroughly washed in deionized water and incubated in buffer overnight before

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measurement with the proteins.

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25 mM tris – HCl buffer, pH 7.4 containing 150 mM NaCl is used as the supporting electrolyte

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for the cyclic voltammetry measurements. The buffer is incubated with 1 mM dithiothreitol

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(DTT) and thoroughly purged with nitrogen prior to the measurement to prevent oxidation of the

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protein complexes in situ within the buffer. For σL/RslA, σH/RshA and σE/RseA, 10 µM

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solutions of the protein are used for the measurement. Cyclic voltammogram of the blank buffer

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are recorded to distinguish the redox activity of the buffer and dissolved salts from that of the

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protein.

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To study the response to oxidative stress, voltammograms of the protein solution in presence

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of different concentrations of H2O2 (10 mM to 90 mM) are also recorded. Thus any changes in


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these voltammograms can be ascribed to the redox response of the protein against the oxidative

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stress induced by the peroxide. The scan rates for the measurements are optimized to 50 mV/s at

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which distinct redox peaks can be obtained.

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Spectro-electrochemical measurements. About 100 µL of the analyte solution (10 µM

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protein with 0.1 mM PAR in buffer) is placed on the SPCE for electrochemical measurements

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through DropSens µStat 400. 50 µL aliquots of this solution are collected for the UV

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measurements using a cuvette of appropriate path – length, on an Avantes Avalight DHS

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spectrophotometer. The solution after UV measurements is transferred back to the SPCE for

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resumption of electrochemical cycling. The measurements are done in ambient conditions (i.e.

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exposed to air, at room temperature).

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Analytical size exclusion chromatographic studies. Freshly purified σ/anti-σ complexes are

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used for analytical size exclusion chromatography experiments. For the oxidized samples, the

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σ/anti σ complexes are incubated with 10mM H2O2 for 45 minutes and passed through Superdex

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200, 10/300 analytical column (GE healthcare life science, Inc.) in a buffer containing 25 mM

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Tris (pH 7.5) and 100 mM NaCl to monitor the oxidative stress induced release of σ from the

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complex.

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Raman Spectroscopy. Raman spectra of solid samples are collected on a LabRam HR system

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with 532 nm diode pump solid state LASER. Each spectrum is averaged over three scans. 0.2

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mW of LASER power and 50X long working distance objective is used. The measurements are

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done at room temperature.

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Covalent Chromatography and Mass Spectrometry. For mass spectrometry, both oxidized

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and reduced forms of proteins are digested with trypsin. Tryptic peptides with free thiols under

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both the conditions are enriched using thiopropyl sepharose 6B resin15 (GE Healthcare Life


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Sciences). The bound peptides are eluted with 50 mM 2-mercapto ethanol in water. The eluted

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peptides are desalted in water using a HiTrap desalting column (GE Healthcare life sciences),

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and examined by positive ionization MS-MS analysis (Bruker Daltonics, Inc).

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RESULTS AND DISCUSSION

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Spectroscopic assay for the kinetics of zinc (II) release. 4-(2-pyridylazo)-resorcinol (PAR),

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chelates with free Zinc (II) in solution to give an intense absorption band centered around 500

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nm11. Upon addition of 10 mM peroxide to the protein complexes, σL/RslA, σH/RshA and

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σE/RseA, the change in absorption at 500 nm is studied as a function of time (shown in Figure

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4). A first order kinetic model is used to fit the absorbance versus time data and calculate the

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half-lives of zinc (II) release from the complexes (Table 1). The data indicate that zinc (II)

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release under oxidative stress is fastest for σE/RseA, followed by σH/RshA and σL/RslA.


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Figure 4. Kinetics of zinc (II) release from the proteins, following addition of peroxide. Open

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circles represent the experimental data point while solid lines represent the first order kinetic fits.

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σE/RseA is represented in orange, σH/RshA is represented in blue, while σL/RslA is represented

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in dark green. The discontinuity in the graph, observed for σL/RslA, is due to generation of

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oxygen bubbles in situ within the cuvette due to addition of peroxide.

238 239 240


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Table 1. Half-lives calculated from PAR assay data. Protein Complex

Half – life / s

σE/RseA

111

σH/RshA

144

σL/RslA

220

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Circular Dichroism. Circular Dichroism in the far – UV region (180 – 240 nm) reflects the

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secondary structure of proteins involving the amino acid backbone16. Thus, changes in secondary

245

structure can be probed by circular dichroism spectroscopy. The circular dichroism spectra of the

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protein before and after addition of peroxide are compared for σE/RseA, σH/RshA, and σL/RslA

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in Figure 5. The data indicate changes in secondary structure in all three proteins, albeit to

248

different extents, with the addition of H2O2. Prior to addition of peroxide, there is no change in

249

the secondary structure of the protein even after 45 minutes from the start of the experiment

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(Supporting Figure S2). The change, after peroxide addition, is more pronounced for σE/RseA

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and σH/RshA, compared to σL/RslA. We note that the mean residue ellipticities become more

252

negative upon addition of peroxide to σH/RshA and σL/RslA, while the reverse is observed for

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σE/RseA. Keeping in mind the standard CD spectra for well – known protein secondary

254

structures16, this may be due to opposing trends in secondary structure change (increase in α

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helicity for σH/RshA and σL/RslA, and decrease for σE/RseA upon application of oxidative

256

stress). However, we do not have enough information about the structures to comment on why

257

two opposing trends are observed in the same family of protein.


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Figure 5. Secondary structural changes in the protein complexes under oxidative stress as seen

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in circular dichroism. Spectra in black represent native protein, spectra in red represent protein

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after peroxide addition. Shown from top to bottom are the spectra for σE/RseA, σH/RshA, and

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σL/RslA.

263

Cyclic voltammetry and Raman Spectroscopy. The effect of oxidative stress on the proteins

264

is also studied using voltammetry. The cyclic voltammograms of the proteins are recorded in

265

their native state. No changes are observed even after several electrochemical cycles

266

(Supporting Figure S3). This shows the electrochemical stability of these proteins during the

267

course of the experiment. Upon addition of H2O2, cathodic and anodic peaks are observed, as

268

shown in Figure 6. We ascribe these peaks to redox phenomena arising due to changes in the

269

secondary structure of the protein, accompanied by release of zinc (II), due to the oxidative

270

stress. Further, we observe the evolution of these redox peaks with increasing concentration of

271

oxidizing agent (Supporting Figure S4). Beyond 90 mM H2O2, the voltammograms are noisy

272

presumably due to generation of a high concentration of reactive oxygen species (ROS) in

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solution.


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Figure 6. Cyclic voltammograms of the three proteins (σE/RseA, σH/RshA, and σL/RslA) in the

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native state (black), and in the presence of peroxide (30 mM – red, and 90 mM – blue).

278 279

From our circular dichroism and PAR-Zn (II) reaction assay data, we know that on application of

280

oxidative stress to these proteins, the following redox active phenomena occur – release of zinc

281

(II) in the buffer and exposure of cysteines from the HXXXCXXC motif and subsequent

282

oxidation to disulfide. Voltammograms of zinc (II) in aqueous media are rare in literature17. The

283

zinc (II/0) couple is irreversible and shows metalation. This prevents the observation of well-

284

defined reversible redox peaks18. Cysteine residues, however, are easily oxidizable to disulfides

285

if exposed to the electrode surface under oxidizing conditions19,20. To deconvolute the

286

contribution of the cysteine – cysteine redox couple to the protein voltammograms, we examined

287

the electrochemistry of pure L – cysteine (Figure 7). The amino acid shows an anodic

288

(oxidation) peak around 0.8 V, corresponding to the oxidation of cysteine to cystine. This peak is

289

found to rapidly disappear with repeated electrochemical cycling indicating depletion of

290

electroactive layer at the electrode surface due to an irreversible phase change21,22 (cysteine is

291

water – soluble, cystine is not). The formation of cystine is seen in the form of a fine white

292

precipitate that increases on addition of an external oxidizing agent like peroxide. Raman spectra

293

with this precipitate (Figure 7) confirms the formation of cystine as S – S stretching bands are

294

seen, which match with standard values reported in literature23. On widening the potential

295

window, the cathodic (reduction) peak is observed. Comparing the voltammograms of the

296

proteins and the amino acid leads us to infer that the primary signatures observed in the protein

297

voltammograms after addition of oxidative stress (cathodic peak around -1 V, anodic peak


20

Page 21 of 38

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298

around 1 V) arise due to the cysteine – cysteine redox couple. The reduced forms of the proteins

299

are stable and unaffected by repeated electrochemical cycling; but in the presence of an oxidizing

300

agent like H2O2, disulfide formation becomes facile and the cysteine – cystine redox couple

301

becomes visible in the voltammograms. A possible reason for this may be the exposure of the

302

cysteines in the HXXXHCXXC motif to the electrode surface following zinc (II) release after

303

application of oxidative stress. We note that addition of peroxide also causes the broadening of

304

the peaks observed in the voltammograms, as peroxide is also electrochemically active within

305

this potential window24. Dissolved reducing agent DTT inside the buffer also contributes to the

306

peaks observed in the native protein solutions, primarily the cathodic peak observed around 1.0

307

V (Voltammogram of DTT shown in Supporting Figure S5). Ideally, the released zinc (II)

308

should be redox active under electrochemical cycling. However, this occurs at potentials too low

309

to be accessible under the stability window of aqueous electrolytes25-28. Additionally, the

310

concentration of zinc is also too low (≈ 1 ppm) in the protein solutions. Hence, we cannot

311

identify zinc redox signatures directly in the protein voltammograms. To indirectly correlate the

312

redox signals observed in voltammograms, to zinc (II) release, we perform voltammetry coupled

313

with

UV

–

visible

absorption

spectroscopy

as

described

in

the

next

section.


21


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Page 22 of 38

314 315

Figure 7. Voltammetry of the pure amino acid cysteine to understand the cysteine – cystine

316

redox couple. Clockwise from top (A). Electrochemical oxidation of cysteine to cystine (the

317

cathodic oxidation peak is marked by an arrow) (B) Raman spectrum of cystine formed from (A)

318

with the characteristic S – S stretch signals marked in red. (C) Cysteine to cystine oxidation and

319

corresponding reduction as seen in the voltammograms on widening the potential window. (D)

320

Chemical representation of the oxidation – reduction process.

321 322

Spectroelectrochemical measurements. UV - vis absorption spectra are collected for the

323

proteins in the presence of PAR in tandem with electrochemical measurements. Pure PAR has


22

Page 23 of 38

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324

absorption peak centered at 410 nm (P1) while careful analysis of the Zn – PAR complex

325

absorption data shows that the maxima can be deconvoluted into two peaks centered around 493

326

nm (P2) and 520 nm (P3) respectively29. The evolution of P2 and P3 are studied as a function of

327

electrochemical cycling. The ratio of areas of the Zn – PAR peaks (P2 + P3) to that of the parent

328

PAR peak (P1) is calculated and found to increase with the addition of hydrogen peroxide,

329

corresponding to an increase in the cathodic peak current in the cyclic voltammogram (Figure

330

8). This is seen during the time course of zinc release after addition of peroxide. Once the entire

331

zinc is released (45 minutes post peroxide addition), the current stabilizes and repeated cycling

332

leads to a slight decrease in peak current, possibly due to depletion of electroactive layer21, 22 at

333

the electrode surface. This set of data leads to the conclusion that zinc (II) release under

334

oxidative stress also contributes to the cathodic (reduction) peak current in the protein

335

voltammograms, along with the cysteine – cystine redox couple. This also supports the model of

336

disulfide mediated zinc (II) release from the protein under oxidative stress30-32. However, we also

337

note that no metalation due to zinc reduction is observed at the electrode surface, and believe that

338

this may be due to very low concentration of zinc (≈ 1 ppm) in the protein samples.


23


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 38

339 340

Figure 8. Spectroelectrochemistry data for the protein (σH / RshA) in buffer. Initial value of the

341

peak area ratio is due to free divalent ions including zinc (II) in solution. Growth of the peak area

342

ratio is seen immediately after addition of peroxide corresponding to increase in peak current for

343

the cathodic peak. This current increases along with electrochemical cycling, due to continuous

344

release of zinc (II). Beyond 45 minutes, there is no further release of zinc and the peak area ratio

345

stabilizes along with reduction peak current. There is a decrease in peak current with repeated

346

cycling in this stage, presumably due to irreversible depletion of active layer at the electrode

347

surface.

348

Spectroscopic assay for determining exposed cysteines in the CXXC motif. To understand

349

the structural environment of the cysteine residues in the protein and rationalize their


24

Page 25 of 38

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350

participation in the redox response, we employ a standard assay with Elman’s reagent (DTNB).

351

Absorption maxima at 412 nm arising due to binding of surface exposed thiols in the protein to

352

the reagent is studied as a function of time. From the absorption kinetics shown in Figure 9, it is

353

evident that the cysteine residues in σH/RshA have maximum solvent exposure followed by

354

σE/RseA and σL/RslA. The cysteine residues exposed to the solvent are also exposed to the

355

electrode surface. These cysteines are responsible for disulfide formation and hence the redox

356

response of the protein under oxidizing conditions. This rationalizes the greater sensitivity of

357

σE/RseA and σH/RshA to redox stress compared to σL/RslA. We note that the absorbance value

358

for σH/RshA is greater compared to σE/RseA. This is in spite of the fact that the other

359

experiments suggest σE/RseA being more sensitive to redox stress. We believe that the greater

360

DTNB absorbance in σH/RshA is because it has a larger number of cysteine residues in the anti-

361

sigma domain8, 33, which gradually unfold and bind to the chromophore as the reaction is allowed

362

to proceed for a very long time (> 1000 sec). The number of reactive cysteines in these proteins

363

is calculated using the extinction coefficient (for TNB2-)14 tabulated in the supporting

364

information (Supporting Method SM1, Supporting Table S3). This experiment suggests that

365

the number of reacted cysteines is maximum for σH/RshA, followed by σE/RseA, and σL/RslA.


25


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Page 26 of 38

366 367

Figure 9. Kinetics of DTNB binding showing the relative solvent accessibility of the cysteines in

368

the three proteins: σH/RshA is represented in black, σE/RseA is represented in green, and σL/RslA

369

is represented in red.

370

The separation of a σ factor from an inactive σ/anti-σ complex in oxidative environments is

371

evaluated using analytical size exclusion chromatography. The chromatograms clearly reveal the

372

disassociation of the σ/anti-σ complex in oxidizing conditions.

373

oxidative stress levels could be different from those examined here, the in vitro data

374

demonstrates the redox sensitivity of these complexes leading to an increase in the cellular

375

concentration of free, active σ factor in response to oxidative stimuli. This proves the biological

While the physiological


26

Page 27 of 38

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376

consequence of oxidative stress response in these protein complexes, leading to the release of σ.

377

A representative chromatogram is shown in Figure 10 (Additional related data in

378

SupportingFigures S6-S10).

379 380

Figure 10. The native σL/RslA protein complex (shown in black) eluted as a single peak on an

381

analytical size exclusion column, while the complex subjected to oxidative stress eluted as two

382

peaks (shown in red) denoting the separation of constituent proteins of the complex under non -

383

reducing conditions.

384

To further confirm the disulfide bond between proximal cysteines in the HXXXCXXC motif

385

under oxidizing conditions, we studied the thiol containing peptides by MS – MS analysis. The


27


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Page 28 of 38

386

formation of a disulfide bond between proximal cysteines makes that peptide incapable of

387

binding to the thiopropyl sepharose resin. The HXXXCXXC motif containing peptides could be

388

enriched and detected only in reduced sample but not in oxidized sample. These results are

389

consistent with the suggestion that the proximal cysteines of the CXXC motif form a disulfide

390

bond upon the removal of the bound Zn2+. A representative mass spectra (σL/RslA) is shown in

391

Supporting Figure S8.

392 393

Calculation of theoretical pKA s of the cysteines in RslA, RshA, and RseA34-36 (tabulated in

394

Supporting Table S1) shows that the cysteines in RslA tends to remain in the conjugate base

395

form, thus forming strong coordination with zinc (II). This also supports the low half-life of zinc

396

(II) release and lesser changes in secondary structure upon application of oxidative stress seen in

397

σL/RslA, and thus the overall sluggish response to oxidative stress.

398

To validate the hypothesis that disulfide bond formation requires the exposure of cysteines to

399

solvent, we compare the cysteines in the HXXXCXXC motif from the crystal structures of

400

σL/RslA37 and ChrR anti – sigma domain9 (Supporting Figures S11 and S12). We chose this

401

pair as ChrR is not known to form disulfides, whereas σL/RslA does. As expected, we find that

402

the HXXXCXXC cysteines in σL/RslA are more solvent exposed compared to those in ChrR.

403

This shows the importance of solvent exposure for facile disulfide formation between cysteines,

404

which in turn controls the redox response.

405 406

CONCLUSIONS


28

Page 29 of 38

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407

In this study, which is a first of its kind, the redox sensitivities of three structurally conserved

408

Zinc associated anti – σ factor systems are evaluated. Oxidative stress on these proteins results in

409

distinct chemical changes. We note a perturbation in the secondary structure of the anti-σ factors

410

due of zinc release and disulfide bond formation between the proximal cysteines of

411

HXXXCXXC motif. We use circular dichroism to monitor the change in secondary structure,

412

and UV absorption to follow the zinc (II) release. Cyclic voltammetry of the proteins under

413

oxidative stress shows the combined effects of zinc (II) release and disulfide formation.

414

Spectroscopic assay with Ellman’s reagent establishes the relative solvent accessibility of the

415

thiol groups across the three different anti-σ factors, determining the ready availability of these

416

cysteine side chains to sense oxidative stress.

417

The voltammetry data provide a basis to rationalize structural changes that coincide with zinc

418

(II) release. Release of zinc (II) from the HXXXCXXC motif is the primary trigger for the

419

disulfide bond formation between the proximal cysteines. The propensity to form this disulfide is

420

not the same for all three proteins. In an effort to understand if constrained disulfides led to

421

apparent changes within an identical scaffold, the conformational propensity of the cysteines for

422

disulfide bond formation is evaluated. We examine known conformational parameters that

423

contribute to favorable disulfide formation with those observed in the ZAS domain.

424

While RseA, RshA, and RslA adopt similar anti σ domain structures, they have dissimilar

425

redox responses. We can qualitatively understand the response from secondary structure

426

perturbation; circular dichroism shows this to be maximum for σE/RseA, followed by σH/RshA

427

and σL/RslA. The cyclic voltammograms show the redox properties that arise because of the

428

response to oxidative stress. The kinetics of zinc (II) release from the HXXXCXXC motif gives

429

us an insight into the temporal features of the stress response. The half-life of zinc (II) release is


29


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 30 of 38

430

fastest for σE/RseA, followed by σH/RshA and σL/RslA. This indicates that σE/RseA is likely the

431

fastest to respond to oxidative stress.

432

To summarize, these studies provide an experimental validation for the sensitivities of

433

structurally similar σ factor/ZAS complexes from both kinetic and thermodynamic points of

434

view. Oxidative stress results in changes in secondary structure induced by disulfide formation

435

and release of zinc (II) ion. We note that these two phenomena occur either simultaneously, or in

436

a time frame that is impossible to resolve using the experimental techniques examined. Half-lives

437

of zinc release indicate kinetic sensitivity while the secondary structural changes (as observed

438

directly from circular dichroism, and indirectly through availability of cysteine residues for

439

redox cycling in the voltammograms) indicate the thermodynamic sensitivity. In the family of σ

440

– ZAS pairs examined, σE/RseA is found to be both kinetically and thermodynamically the most

441

sensitive to oxidative stress. We rationalize this in terms of the local chemical environment and

442

solvent accessibility around the HXXXCXXC motif coordinating with the zinc ion.

443

Put together, this study provides an alternative explanation for the hierarchy in σ factor

444

activation in Mycobacterium tuberculosis in response to a redox stress. While this study is

445

confined to Mycobacterium tuberculosis σ/anti σ complexes that respond to redox stimuli, it also

446

provides a sequence – structure – redox sensitivity dataset that can be utilized for protein

447

engineering applications.

448

449

Supporting Information. The following files are available free of charge.

450

Cyclic voltammogram of [Fe(CN)6]3- / [Fe(CN)6]4- at SPCE; circular dichroism of protein as a

451

function of time; cyclic voltammogram of protein as functions of time, peroxide concentrations

ASSOCIATED CONTENT


30

Page 31 of 38

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452

and scan rates; cyclic voltammogram of DTT; purification pattern and SDS Poly – acrylamide

453

gel; mass spectrometry data of separated protein components; size exclusion chromatography

454

data; SDS – Page of peak fractions from analytical size exclusion chromatography; estimation of

455

solvent accessibility from reported crystal structures; calculation of pKA s of cysteine residues;

456

structure based sequence alignment of anti – σ domains; thiol concentration calculation from

457

DTNB assay. (PDF)

458 459

460

Corresponding Author

461

*[email protected]

462

Author Contributions

463 464

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

465 466 467 468

ASSOCIATED CONTENT

‡A. K. J. and R. N. S. contributed equally to this work.

ACKNOWLEDGEMENT The authors thank Mr. Prahlada BL, Center for Nano Science and Engineering (CeNSE),

Indian Institute of Science, Bangalore for the Raman Spectroscopy measurements.

469


31


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Page 32 of 38

470 471

Funding Sources

472

This work is supported in part by a grant from the Department of Biotechnology, Government

473

of India; and from Project No. EMR/2014/000029 implemented by the Department of Science

474

and Technology, Science and Engineering Research Board, Government of India

475 476

477

ZAS, Zinc Associated Anti σ factor; ECF, Etra Cytoplasmic Function; PAR, 4-(2-pyridylazo)-

478

resorcinol ; DTNB, 5,5'–dithiobis-2-nitrobenzoic acid; ASD, Anti sigma domain; DTT,

479

dithiothreitol; CD, Circular dichroism; CV, Cyclic voltammetry; SPCE, Screen printed carbon

480

electrode; Mtb, Mycobacterium tuberculosis; Rsp, Rhodomonas spheroids; Sco, Streptomyces

481

coelicolor.

482

ABBREVIATIONS

REFERENCES

483

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Resistance to Pulmonary Tuberculosis in p47phox -/- Mice. Infect. Immun. 2000, 68, 1231 –

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1234.

486

(2) Kumar, A.; Farhana, A.; Guidry, L.; Saini, V.; Hondalus, M.; Steyn, A. J. Redox

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Homeostasis in Mycobacteria: The Key to Tuberculosis Control?

488

Med. 2011, 13, e39

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(5) Tyagi, P.; Dharmaraja, A. T.; Bhaskar, A.; Chakrapani, H.; Singh, A. Mycobacterium

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tuberculosis has Diminished Capacity to Counteract Redox Stress Induced by Elevated

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Determination of Redox Sensitivity in Structurally Similar Biological

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