Free Radical Biology and Medicine 65 (2013) 541–551

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Free Radical Biology and Medicine

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n CorrE-m

journal homepage: www.elsevier.com/locate/freeradbiomed

Original Contribution

Cytochrome c peroxidase is a mitochondrial heme-based H2O2 sensorthat modulates antioxidant defense

Dorival Martins, Meena Kathiresan, Ann M. English n

PROTEO and Department of Chemistry and Biochemistry, Concordia University, Montreal, QC, Canada, H4B 1R6

a r t i c l e i n f o

Article history:Received 23 April 2013Received in revised form17 June 2013Accepted 20 June 2013Available online 2 July 2013

Keywords:H2O2 sensingH2O2 signalingHeme-based sensorsMitochondrial ROS regulationAntioxidant enzymesFree radicals

49/$ - see front matter & 2013 Elsevier Inc. Ax.doi.org/10.1016/j.freeradbiomed.2013.06.037

esponding author. Fax: +514 848 2868.ail address: ann.english@concordia.ca (A.M. En

a b s t r a c t

Hydrogen peroxide (H2O2) is a key signaling molecule that also induces apoptosis. Thus, cells mustrapidly sense and tightly control H2O2 levels. Well-characterized cellular responses to exogenous H2O2

involve oxidation of specific cytosolic protein-based thiols but sensing of H2O2 generated by mitochon-drial respiration is less well described. Here we provide substantial biochemical evidence that the hemeenzyme Ccp1 (cytochrome c peroxidase), which is targeted to the intermembrane space, functionsprimarily as a mitochondrial H2O2 sensing and signaling protein in Saccharomyces cerevisiae. Keyevidence for a sensing role for Ccp1 is the significantly higher H2O2 accumulation in ccp1-null cells(ccp1Δ) vs ccp1W191F cells producing the catalytically inactive Ccp1W191F variant. In fact, intracellularH2O2 levels (ccp1Δ4wildtype 4ccp1W191F) correlate inversely with the activity of the mitochondrial(and peroxisomal) heme catalase, Cta1 (ccp1Δowildtype occp1W191F). Mitochondrial Sod2 activity alsovaries in the three strains (ccp1Δ4wildtype 4ccp1W191F) and ccp1Δ cells exhibit low superoxide levels.Notably, Ccp1W191F is a more persistent H2O2 signaling protein than wild-type Ccp1, and this enhancedmitochondrial H2O2 signaling decreases the mitochondrial fitness of ccp1W191F cells. However, these cellsare fully protected from a bolus (0.4 mM) of exogenous H2O2 added after 12 h of growth, whereas theviability of ccp1Δ cells drops below 20%, which additionally associates Ccp1 with Yap1-dependent H2O2

signaling. Combined, our results strongly implicate Ccp1, independent of its peroxidase activity, inmitochondrial H2O2 sensing and signaling to maintain reactive oxygen species homeostasis.

& 2013 Elsevier Inc. All rights reserved.

Introduction

It is now well established that hydrogen peroxide (H2O2) is notjust a harmful oxidant but also a key signaling molecule thatmodulates diverse biological processes from cell differentiation toapoptosis [1]. Consistent with its potentially toxic nature, H2O2

signals are transmitted over short distances and cellular responsesto H2O2 are both site and concentration dependent [1]. The focuson cellular defenses to exogenous H2O2 has provided significantinsight into thiol-based cytosolic H2O2 sensors [2,3]. For example,glutathione peroxidase 3 (Gpx3), a cytosolic thiol peroxidase,senses excess H2O2 through oxidation of its active-site cysteineand conveys this redox signal via disulfide exchange to thetranscription factor Yap1 on exposure of Saccharomyces cerevisiaeto nonlethal doses of H2O2 [3,4]. Oxidized Yap1 is translocated tothe nucleus to regulate the transcription of several antioxidantgenes [5]. However, oxidative environments exist within eukar-yotic cells but sensing of endogenously generated H2O2 or otherreactive oxygen species (ROS) within these regions is poorly

ll rights reserved.

glish).

documented. For example, mitochondria are the main sites ofH2O2 production in respiring yeast [6] but to date only a singlecase of mitochondrial H2O2 sensing has been reported [7],although ROS sensing by mitochondria has been extensivelydiscussed in the context of oxygen availability [8,9].

Yeast switch from anaerobic to aerobic metabolism when afermentable carbon source such as glucose becomes limiting.Because mitochondrial ROS spike during this diauxic shift [10],yeast cells must be prepared for an imminent spike in mitochon-drial ROS. Given its chemical properties [11] and known signalingfunctions [1], H2O2 is probably a crucial messenger of changes inmitochondrial ROS. We demonstrated previously that cytochromec peroxidase (Ccp1), a heme peroxidase targeted to the mitochon-drial intermembrane space [12,13], protects yeast from challengewith a bolus of exogenous H2O2 [14]. This was not surprising giventhat in vitro Ccp1 serves as a highly efficient H2O2 scavenger in thepresence of excess donor ferrocytochrome c (Cyc1II; Scheme 1)[15]. Unexpectedly, we also found that ccp1W191F cells producingthe Ccp1W191F variant with negligible cytochrome c peroxidaseactivity are less sensitive to H2O2 than wild-type cells. The addedprotective effect of Ccp1W191F cannot be attributed to H2O2

scavenging because, unlike the oxidized form of wild-type Ccp1(CmpdI, Scheme 1), CmpdI bearing the W191F mutation [16]is not

Scheme 1. Cytochrome c peroxidase catalysis. Resting ferric (FeIII) Ccp1IIIis oxi-dized by H2O2 to compound I (CmpdI), which has a FeIV heme and a cation radicalon residue W191. CmpdI reacts with ferrous (FeII) Cyc1II to form compound II with aFeIV heme but no W191 radical [16]. Reaction with a second Cyc1II reduces the FeIV

heme to yield resting Ccp1III. The Ccp1W191F variant rapidly reacts with H2O2 (stepa) but is very slowly reduced by Cyc1II (steps b and c) such that it exhibits negligibleCyc1II-oxidizing activity [16].

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551542

reduced by Cyc1II back to the resting ferric enzyme. Consistentwith our results, it has been demonstrated that both Ccp1 andits inactive variant Ccp1W191F convey an oxidative stress signal tothe nuclear transcription factor Skn7 [17], which regulates theexpression of many antioxidant enzymes, including Tsa1, a thiolperoxidase that protects cytosolic proteins against oxidativedamage [18].

In this work we report on the function of Ccp1 and itsCcp1W191F variant in responding to endogenously producedH2O2. In support of independent H2O2 scavenging and H2O2

signaling roles for Ccp1, we find that respiring ccp1W191F yeastcells accumulate less H2O2 than wild-type cells. To establish howccp1W191F cells suppress H2O2, we performed a detailed biochem-ical analysis of the enzyme activities involved in H2O2 breakdown(peroxidases and catalases) in the wild-type, ccp1Δ, and ccp1W191F

strains. We additionally monitored accumulation of the super-oxide anion (O2

d�), the direct chemical precursor to mitochondrialH2O2, as well as superoxide dismutase (SOD) activity, whichcatalyzes O2

d� dismutation to H2O2 and O2.Our findings clearly establish a role for Ccp1 as a heme-based

mitochondrial H2O2 sensor and signaling protein. Like cytosolicGpx3, Ccp1 reacts rapidly with H2O2 [19], which is a hallmark ofsensor specificity in signaling pathways [11]. Another commonfeature of these peroxidases is their utilization of small electron-donor proteins as reducing substrates. Cyc1II reduces Ccp1 CmpdI(Scheme 1) back to the resting ferric enzyme, and thioredoxinreduces the sulfenic acid formed in Gpx3 back to the sulfhydryl.Thus, as is the case for Gpx3–thioredoxin [3,4], H2O2 signaling byCcp1 CmpdI is probably controlled by the availability of Cyc1II.Furthermore, we provide evidence that Ccp1 signals via distinctpathways in response to a steady flux of H2O2 generated endo-genously by the mitochondria vs challenge with a bolus ofexogenous H2O2. We re-examined the effects of exogenous H2O2

on yeast in the BY4741 genetic background because the W303-1Bstrains we studied previously [14] exhibit impaired Yap1 signaling[20]. Yap1 is a master regulator of the response to H2O2 challenge[21] so the analysis of strains with functional Yap1 signaling iscritical in understanding Ccp1′s role in sensing both exogenousand endogenous H2O2.

Materials and methods

Yeast strains, media, and growth conditions

The BY4741 S. cerevisiae strains used in this study are listed inTable 1. Wild-type and ccp1Δ cells were purchased from theEuropean Saccharomyces cerevisiae Archive for Functional Analysis

(EUROSCARF, Frankfurt, Germany). The DNA template for catalyti-cally inactive mature Ccp1 (mccp1W191F) was kindly provided byProfessor Yi Lu (Department of Chemistry, University of Illinois,Urbana, IL, USA) and chromosomally integrated under control ofthe CCP1 promoter by homologous recombination [22]. Briefly, theURA3 gene was amplified by PCR from the plasmid pRS306-URA3using primers 1f and 1r (Table 1), which also contain the homo-logous recombination sequences for genomic integration of theURA3 gene in place of CCP1 (underlined in Table 1). The ampliconwas used to transform wild-type BY4741 using the lithium acetatemethod [23]. Recombinants were selected in synthetic complete(SC) medium lacking uracil and the resulting strain was labeled asDC1-4 (Table 1). Next, mccp1W191F was amplified by PCR usingprimers 2f and 2r and this product was used to replace URA3 instrain DC1-4. After transformation, cells were incubated for 48 h inYPD (1% yeast extract, 2% peptone, and 2% glucose) liquid mediumand spread on SC-agar medium containing 5 mM 5-fluoro-oroticacid for selection of recombinants [24]. The sequence ofmccp1W191F integrated in the chromosome of the recombinantswas confirmed by sequencing the genomic DNA with primers 3fand 3r (Table 1).

Yeast cells were grown under high aeration in YPD liquidmedium at a flask-to-medium volume ratio of ≥5. The culturesat an initial OD600 of 0.01 were incubated at 30 1C with shaking at225 rpm. YPD mediumwas chosen for these studies because its pHof 7.2 is close to physiological pH. We previously used syntheticcomplete (SCD) medium [14], which has a pH ∼3 [25], and asnoted below the choice of medium strongly influences the results.Additionally, the spent YPD mediumwas replaced with 0.85% (w/v)NaCl solution after 72 h to extend culture viability [26] and avoidthe accumulation of acetic acid, which may create a toxic environ-ment [25].

Detection of intracellular ROS

The profluorescent dyes dihydrorhodamine 123 (DHR) anddihydroethidine (DHE) (Molecular Probes) which become fluores-cent on oxidation (Supplementary Fig. S1), were used to probe therelative H2O2 and superoxide levels, respectively, in live cells. DHRis also oxidized to rhodamine 123 (Rhod123) by peroxynitrite [27]but because we detected negligible 3-nitrotyrosine, a peroxynitritemarker [28], in mitochondria isolated from 2-day wild-type orccp1Δ cells (D. Martins et al., unpublished results), we concludethat DHR is oxidized mainly by H2O2 (Supplementary Fig. S1). DHEoxidation to 2-hydroxyethidine (2-HE) correlates with aconitaseinactivation (D. Martins et al., manuscript in preparation), whichvalidates 2-HE fluorescence as a readout of superoxide levels(Supplementary Fig. S1) [29,30]. Cells were stained as describedin [31] with slight modifications. Briefly, 107 cells were harvestedby centrifugation at 2000 g for 10 min, incubated at 30 1C with1 ml of 5 mM dye in PBS (10 mM sodium phosphate and 150 mMNaCl, pH 7.0) for 60 min (2-HE and DHE) or 120 min (Rhod123 andDHR), centrifuged, washed twice, and diluted to 106 cells/ml inPBS. To compare dye uptake by the strains, cells were stained withRhod123 (Molecular Probes) and 2-HE (2-hydroxyethidine pre-pared by air oxidation of DHE) [32], the fluorescent oxidized formsof DHR and DHE, respectively (Supplementary Fig. S1). Dye stocks(5 mM) dissolved in ethanol (Rhod123 and DHR) or dimethylsulfoxide (2-HE and DHE) were stored at �20 1C in the darkuntil use.

Flow cytometry (FACS) analysis was performed with a BDFACScan flow cytometer equipped with a 15-mW argon laseremitting at 488 nm (Becton–Dickinson). The fluorescence of10,000 stained cells and the autofluorescence of 10,000 unstainedcontrol cells were recorded at a flow rate of 1000 cells/s using 530/30 nm (Rhod123 and DHR) and 580/45 nm (2-HE and DHE)

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551 543

bandpass filters. A minimum fluorescence-to-autofluorescence(signal-to-noise) ratio of 10 was obtained under these conditions,and signals were processed using FlowJo software version 7.5.5(Tree Star, Ashland, OR, USA). The fluorescence of the mutantstrains plotted in Figs. 1B and 4B and Supplementary Figs. S2C andS5C is normalized to wild-type cells to account for variation in theFACS instrumentation over 15 days.

DHR- and DHE-stained cells were also visualized by fluores-cence microscopy using a wide-field fluorescence microscope(Model BX 51, Olympus). An equal volume of 90% aqueous glycerol

Table 1S. cerevisiae strains, plasmids, and primers.

Strain, plasmid, or primer Description

Wild-type BY4741 strain MAT a his3Δ1 leu2Δ0 met15Δ0 ura3Δ0ccp1Δ strain ccp1::KAN4MXpRS306-URA3 pRS306 plasmid encoding URA3, which replaces CCP1DC1-4 strain BY4741 cells with ccp1::URA3ccp1W191F gene Produces the CcpW191F protein variantpET17b-mccp1W191F pET17b plasmid encoding mccp1W191F

ccp1W191F strain BY4741 cells with ccp1::ccp1W191F

Primer 1 f 5′-AAGCTGAGGGAAGATGACGAATATGACAACTATATAGGPrimer 1r 5′-TAAATGGACTGGGCGCGTCTTTAGGGAAAGTGATACCGPrimer 2 f 5′-CTACACCGCTCGTTCATGTC-3′Primer 2r 5′-CCTTGTTCCTCTAAAGTCTTGA-3′Primer 3 f 5′-CATGGTCAGGCCCAGATTTG-3′Primer 3r 5′-CATAGACGTACCGTACAAACG-3′

Fig. 1. H2O2 accumulates above wild-type levels in ccp1Δ but not in ccp1W191F cells. (A)(DHR). Data points are the median fluorescence/cell (RFU, relative fluorescence unitsfluorescence/cell normalized to wild-type values in (A) for ccp1W191F and ccp1Δ. Yeast ceNaCl at the vertical dotted line. Cells (107) were stained with 1 ml of 5 mM DHR for 2 h arecorded using 488 nm laser excitation and a 530/30 nm emission filter. The statistical sStudent′s t-test and the lines between the data points in (A) and (B) are for visualization(green), and ccp1Δ (blue) cells (counts¼number of cells with a given RFU). (D) Photom(bottom) and the corresponding bright-field images (top). Rhod123-positive cells were vexposure) and emission at 520 nm (long-pass filter). (E) Normalized rates of H2O2 releaseccp1Δ cells measured by Amplex red peroxidation in a fluorescence microplate readernormalized to those of wild-type mitochondria, and the average rates of H2O2 releaseprotein for the 2- and 7-day cells, respectively. Data points are the averages of three seprotein, 50 μM Amplex red, 1 U/ml horseradish peroxidase in 50 mM KPi buffer (pH 7.4

was added to 104 cells/ml in PBS, 10-ml aliquots were spotted perslide, and samples were analyzed using a 50� objective withnumerical aperture of 0.5. Cells were excited at 490 nm using atungsten–halogen lamp and emission was collected using a520 nm longpass filter.

H2O2 release assay from intact mitochondria

Intact mitochondria were isolated as described elsewhere [33].Cells grown as outlined under Yeast strains, media, and growth

Reference

EUROSCARFEUROSCARF

(ORF YKR066C) in the first step of homologous recombination [64]This work[65][65]This work

CTATGGGCCCTGTGCGTTTCACACCG-3′ This workTTTTCCAACAGTAGATTGTACTGAGAGTGCAC-3′ This work

This workThis workThis workThis work

FACS measurements of H2O2 in wild-type cells stained with dihydrorhodamine 123) measured in 10,000 cells/sample for three separate cultures7SD. (B) Medianlls were grown at 30 1C with 225 rpm stirring in YPD medium and changed to 0.85%t 30 1C and diluted to 106 cells/ml with PBS (pH 7.0), and Rhod123 fluorescence wasignificance in (B) between ccp1W191Fand ccp1Δ cells (*Po0.05) was determined byonly. (C) Histograms of the FACS measurements of 3-day wild-type (red), ccp1W191F

icrographs of 3-day cultures (diluted to 104 cells/ml) showing intracellular H2O2

isualized by wide-field fluorescence microscopy with excitation at 490 nm (220msfrom intact mitochondria of 2-day (gray bars) and 7-day (black bars) ccp1W191F andwith 550/20 nm excitation and 590/30 nm emission band-pass filters. Values areby wild-type mitochondria were 51.9775.84 and 16.0071.12 pmol H2O2/min/mgparate cultures7SD. The final mitochondrial suspensions contained 0.1–0.3 mg/ml). See Materials and methods for additional information.

Fig. 2. Catalase activity is depressed in ccp1Δ cells and Prx activity is depressed in ccp1W191Fcells. (A) Total Prx (solid squares) and total catalase activities (open circles) in thesoluble protein extracts from wild-type cells vs cell age. (B) Prx and (C) catalase activities normalized to the corresponding wild-type values in (A) of ccp1W191F andccp1Δ cells. Data points are the averages of three separate cultures7 SD. The statistical significance in (B) and (C) between ccp1W191Fand ccp1Δ cells (*Po0.05) wasdetermined by Student′s t-test and the lines between the data points were added for visualization only. (D) In-gel assay of Cta1 and Ctt1 catalase activities in the solubleprotein extracts (2.5 μg protein/lane) from 7-day cells. See Materials and methods for additional information.

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conditions were harvested and washed twice with 100 mM KPibuffer (pH 7.0). The cell pellets were resuspended in 10 ml ofprewarmed 100 mM Tris–H2SO4 (pH 9.4) with 10 mM dithiothrei-tol (DTT), incubated for 10 min at 30 1C and 80 rpm, harvested,washed twice with 10 ml of 10 mM potassium phosphate buffer(pH 7.4) containing 1.2 M sorbitol, and treated with 3 mg ofZymolyase 20T per gram of wet cells for 40 min at 30 1C. Theresulting spheroplasts were washed twice with 10 ml of the samebuffer and resuspended in 5 ml of 10 mM Tris–HCl (pH 7.4)containing 0.6 M sorbitol, 1 mM EDTA, and 1 mM phenylmetha-nesulfonyl fluoride. Spheroplasts at 4 1C were disrupted by 15strokes of a glass–Teflon homogenizer, the homogenates werespun at 2000 g, and the supernatants (i.e., denucleated lysates orS2 fractions) were collected. The S2 fractions were further cen-trifuged at 10,000 g for 15 min at 4 1C, and the mitochondria-depleted supernatants (S10 fractions) were separated from themitochondria-enriched pellets (P10 fractions). P10 fractions werecarefully resuspended in 250 mM sucrose, 1 mM EDTA, and 10 mMMops–KOH (pH 7.2) and applied to a step gradient of 60/32/23/15%sucrose (w/v) in 1 mM EDTA and 10 mM Mops–KOH (pH 7.2). Thegradient was ultracentrifuged at 134,000 g for 1 h at 2 1C and thehighly purified mitochondrial fraction was collected between the60/32% layer of the gradient.

H2O2 release from intact mitochondrial suspensions was fol-lowed using the Amplex Red Hydrogen Peroxide/Peroxidase AssayKit from Molecular Probes according to the manufacturer′sinstructions. An assay solution containing 100 μM Amplex redand 2 U/ml horseradish peroxidase (HRP) in 100 mM KPi buffer

(pH 7.4) was prepared using the kit components. Mitochondriawere resuspended at a protein concentration of 0.2–0.6 mg/ml in10 mM Mops–KOH buffer (pH 7.2) with 250 mM sucrose and1 mM EDTA. After mixing of 50 μl of mitochondrial suspensionwith 50 μl of assay solution, the HRP-catalyzed oxidation ofAmplex red by mitochondria-derived H2O2 was monitored byfluorescence at 22 1C in a plate reader (SpectraFluor Plus, Tecan)with 550/20 nm excitation and 590/30 nm emission bandpassfilters. Rates of H2O2 released from mitochondria (pmol/min/mgprotein) were calculated using a linear calibration plot of fluores-cence intensity vs 20–600 pmol of H2O2.

Preparation of soluble protein extracts and enzyme activity assays

The proteins soluble in aqueous buffer were extracted asdescribed previously [14]. Briefly, cells were centrifuged at 2000 gand washed twice with 100 mM KPi buffer (pH 7.0); the pellets wereresuspended in this buffer containing Complete Protease InhibitorCocktail (Roche) and mixed with an equal volume of acid-washedglass beads. Suspensions were subjected to four cycles of vortexing,15 s each, and the cell debris was removed by centrifugation at13,000 g for 10 min at 4 1C. The total protein in the supernatants wasdetermined by the Bradford assay with bovine serum albumin as astandard [34], and enzyme activities were assayed at 22 1C using adiode-array UV–Vis spectrophotometer (Agilent) or the SpectraFluorPlus plate reader.

To determine catalase activity, 5.0–20-μl aliquots of solubleprotein extract were added to 1.0 ml of 20 mM H2O2 in 50 mM KPi

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551 545

buffer (pH 7.0). H2O2 decomposition was monitored at 240 nm(ε240¼43.6 M�1 cm�1) [35], and 1 U of catalase activity catalyzedthe degradation of 1 mmol of H2O2/min. Cta1 and Ctt1 activities wereassayed separately after extract fractionation by native PAGE asdescribed in [36] with slight modification. Briefly, extracts containing1.0–5.0 μg total protein were electrophoresed on 4% stacking(10�4�0.1 cm) and 8% resolving (10�6�0.1 cm) gels at 50 mAfor 3 h at 4 1C. Importantly, we found that a long stacking gel wasrequired to separate Cta1 and Ctt1. The gels were incubated for10 min at 20 1C in the dark with 10 mM H2O2 in 100 mM KPi buffer(pH 7.0), rinsed once with distilled water, and stained with 60 mMK3[Fe(CN)6] and 75 mM FeCl3 for 5 min at 20 1C. Active catalasecreates a white band in the gel by degrading H2O2 and preventingK3[Fe(CN)6] reduction to K2[Fe(CN)6], which reacts with FeCl3 to forma Prussian blue precipitate [36].

SOD (Sod1 and Sod2) in-gel activities also were assayedseparately after native PAGE [36]. Extracts containing 0.75–2.0 μgtotal protein were electrophoresed on 12% resolving gels at 50 mAfor 2 h at 4 1C. The gels were incubated for 30 min at 20 1C in thedark with the staining solution (0.17 mM nitroblue tetrazolium(NBT), 6.7 mM TEMED, and 0.3 mM riboflavin in 100 mM KPibuffer, pH 7.2), rinsed twice with distilled water, and exposed towhite light from a 60-W mercury lamp for 60 min. Active enzymeinhibits reduction by O2

d� (generated by excited-state riboflavinautoxidation) of NBT to blue formazan and gives white bands [36].These were quantified on an AlphaImager (ProteinSimple) andconverted to units of SOD activity based on the calibration plot inSupplementary Fig. S3 prepared using purified human Sod1 (seebelow). Note that negative images of the gels (light areas appeardark and vice versa) are shown in Figs. 2D, 5A, and 7A andSupplementary Fig. S3A. To discriminate between Sod1 andSod2, the resolved gels were incubated for 1 h before stainingwith 5 mM KCN to specifically inhibit Sod1.

Human recombinant thioredoxin used in the thioredoxin perox-idase (Prx) activity assays was kindly provided by Professor WilliamMontfort (University of Arizona, Tucson, AZ, USA). Prx activity wasdetermined by measuring NADPH oxidation at 340 nm (ε340¼6.22 mM�1 cm�1) in a coupled thioredoxin/thioredoxin reductaseassay [37]. Aliquots (10–50 μl) of soluble protein extract were addedto 1.0 ml of Prx assay solution (90 mMH2O2, 6 mg/ml thioredoxin, 1 mg/ml thioredoxin reductase, and 250 μMNADPH in 50mMHepes–NaOHbuffer, pH 7.0), and 1 U of Prx activity catalyzed the oxidation by H2O2

of 1 mmol of NADPH/min. Gpx activity was assayed by monitoringNADPH oxidation in a coupled glutathione (GSH)/glutathione reduc-tase assay [38]. Aliquots (5.0–10 μl) of soluble protein extract wereadded to 100 μl of Gpx assay solution (2 mM GSH, 100 μM tert-butylhydroperoxide, 120 μM NADPH, and 1 U/ml glutathione reductase in50 mM Tris–HCl/0.1 mM EDTA buffer, pH 7.6), and 1 U of Gpx activitycatalyzed the oxidation of 1 mmol of NADPH by tBHP/min.

Expression and purification of recombinant human Sod1(rhCuZnSOD)

The DNA template for rhCuZnSOD was kindly provided byProfessor P. John Hart (University of Texas Health Science Center,San Antonio, TX, USA). The cDNA for human Sod1 was subclonedinto the NcoI and HindIII (Fermentas Life Sciences) sites of theNovagen pET22b(+) vector (EMD Millipore), and Escherichia coliBL21(DE3) cells were transformed with pET22b-hSod1. Cells weregrown to an OD600 of 0.6 in LB medium with 100 mg/ ml ampicillinat 37 1C and 250 rpm. After this time, 1 mM isopropyl-β-D-1-thiogalactopyranoside was added to induce Sod1 overexpressionand the culture was further incubated at 23 1C and 250 rpm for16 h. Cells were lysed at 75 1C in 50 mM Tris–HCl (pH 8.0) bufferwith 100 mM NaCl, 0.1 mM EDTA, and 5 mM DTT and centrifugedat 30,000 g for 15 min. Sod1 in the supernatant was purified on a

2.6�10-cm Q-Sepharose anion-exchange column (GE HealthcareLife Sciences) at 20 1C using a linear 0.1–1 M NaCl gradient at aflow rate of 2 ml/min. The purified protein (49 mg/L culture) wasincubated with 0.5 mM CuCl2 and 0.5 mM ZnCl2 in 0.5 M sodiumacetate buffer (pH 5.5) at 4 1C for 20 h, desalted on a NAP-5column (Sephadex G-25; GE Healthcare Life Sciences), and ana-lyzed for metal ion content on an Agilent 7500ce ICP-MS [39].Sod1 was found to be 25% Cu- and Zn-loaded, and the ng/well inSupplementary Fig. S3A are corrected for metal loading.

Ccp1 and Ccp1W191F protein levels

Rabbit anti-Ccp1 serumwas kindly provided by Professor DavidGoodin (University of California, Davis, CA, USA). Immunoblotswere used to determine the levels of the Ccp1 and Ccp1W191F

proteins in wild-type and ccp1W191F cells, respectively. Solubleprotein extracts were electrophoresed under reducing conditionsin 6% stacking and 12% resolving SDS–PAGE for 1 h at 150 V. Wettransfer of electrophoresed proteins to a polyvinylidene difluoride(PVDF; Bio-Rad) membrane was carried out at 100 mA for 3 h atroom temperature. After blocking for 1 h at room temperaturewith 5% (w/v) skim milk in TBST (50 mM Tris, 150 mM NaCl, and0.05% v/v Tween 20, pH 7.6), membranes were incubated withrabbit anti-Ccp1 serum (1:10,000 dilution) for 2 h, washed threetimes with TBST, and incubated with goat anti-rabbit HRP-con-jugated secondary antibody (1:20,000; Bio-Rad) for 1.5 h at roomtemperature. Blots were visualized using the SuperSignal WestPico enhanced chemiluminescence kit from Thermo Fisher in theAlphaImager. To generate loading controls, membranes werestained with Coomassie blue (ICN Biomedicals) after immunode-tection, and the integrated intensity of each anti-Ccp1 reactiveband was normalized by the sum of the integrated intensity of allCoomassie bands in the same lane [40]. Each lane contained 3 μgof total protein because the Ccp1 signal (2-min exposure) waslinear over 0.5–4 μg total protein in 5-day cells, which exhibitedthe highest Ccp1 production.

Challenge of exponentially growing cells with a bolus of exogenousH2O2

Cells grown for 12 h (OD600 0.5) in YPD medium were exposedto 0.4 mM H2O2. After a further 1.5-h incubation at 30 1C and225 rpm, the cultures were diluted 10-fold and plated on YPD-agarmedium. Colony forming units were counted after incubation at30 1C for 2 days [10]. Aqueous 0.85% NaCl was added to the controlcells. Catalase, Prx, and SOD activities in the soluble proteinextracts were determined as described above.

Statistical analyses

Three biological replicates (i.e., three independent cultures)were analyzed in each experiment. Statistical significance betweenthe mutant strains (Po0.05) was determined by Student′s t-testcalculated using Origin 9.1 software (OriginLab).

Results

Independently of its peroxidase activity, Ccp1 senses H2O2 andregulates catalase activity

DHR-derived fluorescence, and hence H2O2 levels, spike in wild-type cells at day 2 (Fig. 1A). This corresponds to the diauxic shift [10]when most cells are respiring and generating H2O2 in their mitochon-dria owing to electron leakage from complex III of the respiratorychain to O2 [41]. Wild-type cells quickly increase their antioxidant

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551546

defenses since on day 3 their H2O2 level is dramatically lower (Fig. 1A).Because they exhibit higher DHR-derived fluorescence (see DHRactivation, Supplementary Fig. S1), ccp1Δ cells appear to respondmore weakly to the spike in mitochondrial H2O2 relative to wild-typecells at day 3 (Fig. 1B–D, Supplementary Fig. S2). An additional H2O2

spike is seen in the null mutant around day 7 when cells are instationary phase, which is associated with increased oxidative stress[10]. This reflects again their lower than wild-type H2O2 scavengingcapacity. In contrast, ccp1W191F

fluorescence drops below that of wild-type cells at day 3 (Fig. 1B–D; Supplementary Fig. S2), revealing thatthese cells mount a stronger defense than wild-type to elevated H2O2.Consistent with the intracellular results, mitochondria from 2- and 7-day ccp1Δ cells release more H2O2 than those from wild-type cells,whereas sub-wild-type levels of H2O2 are released from mitochondriaisolated from ccp1W191F cells (Fig. 1E).

If Ccp1 were to function solely as a mitochondrial H2O2

scavenger [6,14,42], production in cells of the catalytically inactiveCcp1W191F variant should not depress H2O2 levels. Thus, wehypothesized that Ccp1 might modulate the activities of otheryeast H2O2-metabolizing enzymes such as Prx, Gpx, or catalase.

Wild-type and ccp1Δ cells possess comparable Prx activity butthis is clearly depressed in ccp1W191F cells (Fig. 2A and B), whichaccumulate less H2O2 than wild-type cells (Fig. 1B and E). Further-more, Gpx activity is similar in the three strains (Supplementary Fig.S4). Thus, the lack of correlation between Prx or Gpx activity andmitochondrial H2O2 levels, especially in ccp1W191F cells, suggests thatH2O2 accumulation (Fig. 1) is controlled by another enzymaticactivity. Catalases effectively scavenge H2O2 by catalyzing its dis-proportionation with high efficiency [43]. The ccp1Δ cells displaysignificantly lower catalase activity than ccp1W191F cells (Fig. 2C),notably around the diauxic shift (day 2) and between day 5 and day10 of stationary-phase cells, when this activity peaks in wild-typecells (Fig. 2A). Because yeast express Ctt1, a cytosolic catalase, andCta1 [44,45] that is co-targeted to mitochondria and peroxisomes[46], we separately monitored the activity of each catalase isoformusing in-gel assays. Fig. 2D reveals that whereas Ctt1 activity iscomparable in the three strains, Cta1 activity varies dramatically.Importantly, the high and low H2O2 levels in ccp1Δ and ccp1W191F

cells (Fig. 1B and E) can be associated with their low and highCta1 activities, respectively (Fig. 2D). This leads us to proposethat respiration-derived H2O2 is removed principally by mitochon-drial Cta1, which is regulated in a H2O2-dependent manner byCcp1, defining a H2O2 sensing and signaling function for thisperoxidase.

Fig. 3. Ccp1 and Ccp1W19F production is comparable in wild-type and ccp1W191F strain(bottom) cells over 0.5 to 15 days. Extracts of total soluble protein (3 μg/lane) were sepaover 3 h at room temperature; the membranes were blocked for 1 h in 5% milk–TBST aantibody for 1.5 h. The anti-Ccp1-reactive bands were detected by enhanced chemiluminccp1W191Fcells (black bars) corrected for loading. The membranes were stained with Coomserves as a loading control [40]. Data points are the averages of three separate cultures

Ccp1W191F is a more persistent H2O2 sensor protein than wild-typeCcp1

Their elevated Cta1 activity (Fig. 2D) suggests that H2O2 signaling ismore persistent in ccp1W191F cells. Persistent H2O2 signaling byoxidized or activated Ccp1W191F is likely because its reduction ordeactivation by Cyc1II (steps b and c in Scheme 1) is very slow relativeto wild-type Ccp1 deactivation [16]. Notably, increased Ccp1W191F

protein production does not contribute to enhanced H2O2 signalingbecause the Ccp1W191F and Ccp1 levels are comparable in theirrespective strains (Fig. 3).

Ccp1 H2O2 sensing and signaling regulate Sod2 activity to controlsuperoxide levels

H2O2 in respiring yeast cells is derived mainly from thedismutation of superoxide (O2

d�) generated by one-electronreduction of O2 in the electron-transport chain. Although O2

d�

spontaneously dismutates to H2O2 and O2 (k∼105–106 M�1s�1), itslevel in cells is controlled by SODs that catalyze this reaction(k∼109 M�1 s�1) [47]. Yeast possess two SODs, an abundant andlargely cytosolic Sod1 (CuZnSOD), which is additionally present inthe mitochondrial intermembrane space [48], and Sod2 (MnSOD),localized in the mitochondrial matrix [49]. DHE staining revealsthat at all time points, ccp1Δ cells have lower O2

�levels than theCcp1-producing strains (Fig. 4A and B; Supplementary Fig. S5),consistent with the enhanced SOD activity of the null mutant(Fig. 5; Supplementary Fig. S6). However, O2

d�levels in ccp1Δ cellsdrop significantly below wild-type levels (Fig. 4B) only when Sod2activity spikes at day 3 (Fig. 5A and C). For Sod2 activity to controlits accumulation, the O2

d�flux from the electron transport chain

must be directed largely to the matrix [41] and trapped therebecause its charge impedes diffusion across membranes [28].

Sod2 activity is significantly lower in ccp1W191F vs ccp1Δ cells(Fig. 5C). Thus, low Sod2 activity combined with high Cta1 activity(Fig. 2D) prevents H2O2 accumulation in ccp1W191F cells (Fig. 1),whereas the opposite situation permits H2O2 accumulation in ccp1Δcells. Notably, staining with Rhod123 (a probe of mitochondrialmembrane potential, Supplementary Fig. S2C) reveals a postdiauxicdrop at day 3 in mitochondrial function after the spike inO2

d�accumulation in ccp1W191F cells (Fig. 4B). In contrast, elevatedH2O2 in the null mutant (Fig. 1B and E) has little effect on membranepotential (Supplementary Fig. S2C). Also noteworthy is the surge inO2

d�accumulation in wild-type and ccp1W191F cells in stationary phase

s. (A) Western blotting of Ccp1 and Ccp1W191Ffrom wild-type (top) and ccp1W191F

rated under reducing conditions by SDS–PAGE and transferred to PVDF membranesnd probed with rabbit anti-Ccp1 for 2 h followed by a HRP-conjugated anti-rabbitescence. (B) Intensities of the anti-Ccp1 reactive bands in wild-type (gray bars) andassie after probing for Ccp1 and the sum of Coomassie band intensities in each lane7SD.

Fig. 4. Superoxide (O2d�) accumulates above wild-type levels in ccp1W191F but not in ccp1Δ cells. (A) FACS measurements of O2

d�in wild-type cells stained with DHE. Datapoints are the median fluorescence/cell (RFU) measured in 10,000 cells/sample in three separate cultures7SD. 2-HE fluorescence was recorded using 488 nm laser excitationand a 580/45 nm emission filter. (B) Median fluorescence/cell normalized to wild-type values in (A) for ccp1W191F and ccp1Δ. Data points are the averages of three separatecultures7SD. The statistical significance in (B) between ccp1W191F and ccp1Δ cells (*Po0.05) was determined by Student′s t-test and the lines between the data points in(A) and (B) were added for visualization only. Experimental data as in Fig. 1 except that cells were stained with DHE for 1 h. See Materials and methods for additionalinformation.

Fig. 5. Sod2 activity is upregulated in ccp1Δ but not in ccp1W191F cells. (A) In-gel analysis of SOD activity vs cell age in the soluble protein extracts (0.75 μg total protein/lane).The activity of 1 ng of purified rhCuZnSOD is shown in lane S. (B) Sod2 activity in wild-type cells was determined from the calibration curve in Supplementary Fig. S3.(C) Sod2 activity normalized to wild-type values in (B) of ccp1W191F and ccp1Δ. Data points are the averages of three separate cultures7SD. The statistical significance in(C) between ccp1W191F and ccp1Δ cells (*Po0.05) was determined by Student′s t-test and the lines between the data points in (B) and (C) were added for visualization only.Enzyme activities were assayed after native PAGE as described under Materials and methods.

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551 547

(Fig. 4), which is partially offset by elevated Sod2 activity in ccp1Δ cells(Fig. 5C). In sum, H2O2 sensing and signaling via Ccp1 stronglyinfluence Sod2 activity and O2

d�levels in respiring cells.

Ccp1 signaling protects exponentially growing cells against challengewith exogenous H2O2

Previously, we [14] and others [42,50] reported that ccp1Δ cells inW303-1B, JM7490, and BY4741 genetic backgrounds exhibit poor

viability when challenged with H2O2. We confirm here that expo-nentially growing BY4741 ccp1Δ cells are sensitive to 0.4 mMexogenous H2O2, whereas ccp1W191F cells are resistant to thischallenge (Fig. 6A). Because Prx and catalase activities are depressedin ccp1W191F and ccp1Δ cells, respectively (Fig. 2B and C), wequestioned how challenge with exogenous H2O2 might alter theseactivities. Surprisingly, Prx activity increased 10-fold in ccp1W191F vs2-fold in wild-type cells but remained uninduced in ccp1Δ (Fig. 6B).Cytosolic Tsa1 (Tpx1) is the most abundant Prx in S. cerevisiae [51]

Fig. 6. ccp1Δ cells exhibit low viability and do not upregulate Prx or catalase activity after challenge with a bolus of exogenous H2O2. (A) Viability of 0.5-day wild-type,ccp1W191F and ccp1Δ cells after challenge with 0.4 mM H2O2. Total (B) Prx and (C) catalase activities with and without H2O2 challenge. Cultures at OD600 0.5 were challengedwith saline solution (control, gray bars) or 0.4 mM H2O2 (black bars) for 1.5 h at 30 1C and 225 rpm. Cell viability and enzyme activities were assayed as described underMaterials and methods. Data points are the averages for three separate cultures7SD.

Fig. 7. Sod2 activity is more sensitive than Sod1 activity to challenge with a bolus (0.4 mM) of exogenous H2O2. (A) Top: in-gel SOD activity (2.5 μg total protein/lane).Bottom: Sod2 activity (10 μg total protein/lane) after Sod1 inhibition with 5 mM KCN. (B) Sod1 and (C) Sod2 activities were determined from the calibration curve inSupplementary Fig. S3. Cultures at OD600 0.5 were exposed to saline solution (control, gray bars) or 0.4 mM H2O2 (black bars) for 1.5 h at 30 1C and 225 rpm. Sod1 and Sod2activities were assayed after native PAGE as described under Materials and methods. Data points are the averages for three separate cultures7SD.

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551548

and its mRNA was upregulated when exponentially growing wild-type and ccp1W191F cells (in the CEN.PK2-1C genetic background)were challenged with exogenous H2O2 [17]. Furthermore, Tsa1 mRNAremained elevated longer in ccp1W191F vs wild-type cells, consistentwith Ccp1W191F being a more persistent H2O2 signaling moleculethan Ccp1 because the latter is readily deactivated by Cyc1II

(Scheme 1).Exogenous H2O2 induces catalase activity by ∼2.5-fold in wild-

type cells to reach the high constitutive level found in ccp1W191F

\(Fig. 6C). This increases by �20% on H2O2 challenge but remainsat its constitutively low level in the ccp1Δ mutant (Fig. 6C). Aprevious study of the yeast H2O2 stimulon [2] reported upregula-tion of cytosolic Ctt1 and Tsa1 protein expression. The catalaseactivity of Ctt1 will scavenge cytosolic H2O2, whereas Tsa1 func-tions both as a ribosome-associated antioxidant [18] and as achaperone [52]. Thus, the low viability of ccp1Δ cells underexogenous H2O2 challenge can be attributed in part to their failureto increase Ctt1 and Tsa1 production in the absence of Ccp1-mediated H2O2 signaling.

We previously reported that W303-1B ccp1Δ cells grown inSCD medium exhibited higher catalase activity on H2O2 challengethan wild-type cells [14]. Here we note that BY4741 ccp1Δ cellsgrown in SCD medium possess lower catalase activity than wild-type cells in both the presence and the absence of H2O2 challenge(Supplementary Fig. S7), although both cell viability and catalaseinduction are lower in SCD vs YPD (data not shown). Thus, thediscrepancy in our previous [14] and present results may reflectdifferences in the parent strains. W303-1B cells have a nonsensemutation in the YBP1gene [53,54] encoding Ybp1, a criticalactivator of the transcriptional regulatory protein, Yap1 [20,53].

Because Yap1 is a master regulator of the H2O2 stimulon [2,21], thedissimilar response to H2O2 challenge of cells in the W303-1B andBY4741 genetic backgrounds is not surprising. Additionally, in ourprevious study we expressed the CCP1 and ccp1W191F genes on alow-copy-number plasmid, pYCplac33 [14], whereas for the pre-sent study all constructs were expressed chromosomally (Table 1).

Interestingly, we observed that H2O2 challenge significantlydecreases Sod2 activity in the three strains and Sod1 activity to alesser extent (Fig. 7). In contrast, Sod2 and Sod1 protein levelswere induced approximately six- and fourfold, respectively, onH2O2 challenge of cells in the YPH98 genetic background [21]. Thedivergence between Sod1 protein levels and activity may beexplained in part by H2O2-induced inactivation of Sod1, whichhas been observed in vitro [55]. However, Sod2 is reportedlyresistant to H2O2 in vitro [56] but a channel for H2O2-inducedSod2 inactivation may exist in vivo that is absent in vitro. Thus,although both endogenous (Fig. 5 and Supplementary Fig. S6) andexogenous H2O2 [21] induce Sod1 and Sod2 production, wespeculate that SOD activity is decreased by the high H2O2 con-centrations used to challenge cells (Fig. 7).

Discussion

We provide data that identify Ccp1 as a mitochondrial H2O2

sensor and signaling molecule. Its role as a scavenger of H2O2 inrespiring mitochondria seems to be secondary to its signaling rolebecause H2O2 levels are higher in wild-type cells than in ccp1W191F

cells (Fig. 1) with negligible CCP activity (Scheme 1). Ccp1-mediated H2O2 signaling increases Cta1 activity (Fig. 2D) and,

Scheme 2. Summary of Ccp1 sensing and signaling of mitochondrially generated H2O2 or of a bolus of exogenously added H2O2. Solid and dashed arrows represent processesassumed to be stimulated and repressed, respectively, by Ccp1 sensing and signaling. (A) Ccp1 in the mitochondrial intermembrane space (IMS) is oxidized to signaling-competent CmpdI by H2O2 generated during mitochondrial respiration. CmpdI is deactivated by Cyc1II or signals to enhance Cta1 activity and suppress Prx activity by anunknown mechanism. Increased Cta1 activity blocks H2O2 accumulation, thereby downregulating Sod2, which increases O2

d� levels. (B) Exogenously added H2O2 oxidizesCcp1, which upregulates Ctt1 catalase and Tsa1 Prx activities to promote resistance to exogenous H2O2. We speculate that the Ccp1 signal is transmitted to the nucleus via theSkn7/Yap1 pathway, because Ccp1 was shown to activate Skn7 on H2O2 challenge, also by an unknown mechanism [17].

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551 549

although Ccp1W191F and Ccp1 production is similar (Fig. 3), Cta1induction is higher in ccp1W191F vs wild-type cells (Fig. 2D)because of sluggish deactivation of activated Ccp1W191F by Cyc1II

(steps b and c, Scheme 1) [16]. Devoid of Ccp1, ccp1Δ cellsaccumulate the highest amount of H2O2 (Fig. 1) and exhibit thelowest Cta1 activity. However, this is beneficial because H2O2

induces Sod2 [2,10,30] as reflected here in the elevated Sod2activity in the null mutant vs ccp1W191F (Fig. 5C). Downregulationof this key mitochondrial antioxidant activity increases intracel-lular O2

d� levels (Fig. 4B) and impairs mitochondrial function inccp1W191F cells (Supplementary Fig. S2C). Scheme 2A summarizeshow H2O2 sensing and signaling by Ccp1 influences endogenousROS levels and antioxidant activities. Ccp1 is activated on oxida-tion by H2O2, and signaling by the active form, CmpdI (or aderivative), increases Cta1 activity, which scavenges H2O2. Thedrop in H2O2 levels inhibits stimulation of Sod2 activity, leading toincreased O2

d� levels (Fig. 4B) and also depressed Prx activity(Fig. 2B), especially in the presence of persistent signaling byCcp1W191F. Importantly, the opposing control of H2O2 and O2

d�

levels by Ccp1 signaling influences yeast chronological lifespanbecause ccp1Δ and ccp1W191F cells live longer and shorter, respec-tively, than wild-type cells (D. Martins et al., manuscript inpreparation). Chronological life-span extension has been pre-viously reported for ctt1Δ and cta1Δ yeast strains, which, likeccp1Δ cells, accumulate more H2O2 and less O2

d� than wild-typecells [30].

Despite its negative effects on mitochondrial fitness (Supple-mentary Fig. S2) and lifespan (D. Martins et al., manuscript inpreparation), Ccp1 signaling clearly protects against exogenousH2O2 (Fig. 6). After challenge with a bolus (0.4 mM) of H2O2, ccp1Δcells exhibit o20% viability and insignificant upregulation ofcatalase or Prx activities (Fig. 6). In contrast, ccp1W191F cells are495% viable and exhibit high catalase and Prx activities, whereaswild-type cells mount a slightly less effective defense (Fig. 6).Increased catalase and Prx activities are consistent with thereported increase on challenge with exogenous H2O2 of Ctt1 [2]and Tsa1 protein levels [2] as well as Tsa1 mRNA levels [17], asoutlined in Scheme 2B. Thus, Ccp1 signaling gives rise to differentoutcomes in response to endogenous and exogenous H2O2, moststriking being the dramatic upregulation of Prx activity in ccp1W191F

cells in response to exogenous H2O2 (Fig. 6B) vs the depression of

this activity in unchallenged ccp1W191F (Fig. 2B). This serves as aclear example of how H2O2 signaling is site and concentrationdependent [1], and more specifically how Ccp1-mediated H2O2

signaling reduces mitochondrial fitness but protects against H2O2

challenge. In essence, Ccp1 is nonessential in unstressed cells butstrongly defends cells against exogenous H2O2, like other antiox-idant enzymes studied in model organisms [57].

Heme-based H2O2 sensing has not been previously reportedalthough heme proteins are known to sense gases including O2,NO, and CO [58]. Also, heme-activated O2 sensors such as Hap1and Hap2/3/4/5 modulate expression of antioxidant enzymes[59,60] but do not respond to H2O2 [61]. Like thiol-based H2O2

sensors such as Gpx3 [1], Ccp1 is rapidly oxidized by H2O2 (step a,Scheme 1) [19] but it is unclear how H2O2 signals through Ccp1.We observe accumulation of catalytically inactive and probablyheme-free Ccp1 outside mitochondria after the diauxic shift(M. Kathiresan et al., manuscript in preparation) when catalaseactivity starts to increase in wild-type and ccp1W191F strains(Fig. 2C and D). This raises the intriguing possibility of hemetransfer from “activated” Ccp1 to apoCta1, which we are currentlyinvestigating. This hypothesis gains credence from the fact thatCcp1 translation is independent of heme availability and apoCcp1is rapidly converted to the holoprotein [62] once heme is synthe-sized [61], whereas catalase translation requires heme [63].

Because Ccp1 is the first mitochondrial heme-based H2O2sensorand signaling protein identified, unraveling its novel mechanism(s) shouldadvance our understanding of mitochondrial retrogradesignaling. Finally, we note that because Ccp1 shares its reducingsubstrate Cyc1II with cytochrome c oxidase, reducing equivalents inthe electron transport chain can be directed to H2O2 or O2 to controlrespiration.

Acknowledgments

This work was funded by grants from the Natural Sciences andEngineering Research Council of Canada (NSERC) and ConcordiaUniversity to A.M.E., who also holds a Concordia University ResearchChair. D.M. and M.K. acknowledge doctoral scholarships from FRQ-NT(Quebec) and NSERC, respectively, and additional awards fromConcordia University and PROTEO, the FRQ-NT Network for Research

D. Martins et al. / Free Radical Biology and Medicine 65 (2013) 541–551550

on Protein Function, Structure, and Engineering. We thank Dr. PamHanic-Joyce for constructing the ccp1W191F mutant strain and Drs.Mengwei Ye and Michael Osbourne for the expression and purifica-tion of rhCuZnSOD. We are grateful to Professor François Dragon andDenis Flipo (Université du Québec à Montréal) for access to their flowcytometry facility and for helpful discussions. We thank ProfessorPaul Joyce for critically reading the manuscript and ProfessorVladimir Titorenko for many helpful discussions.

Appendix A. Supporting information

Supplementary data associated with this article can be found inthe online version at http://dx.doi.org/10.1016/j.freeradbiomed.2013.06.037.

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