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Journal of Biotechnology 167 (2013) 309– 315

Contents lists available at ScienceDirect

Journal of Biotechnology

jo u r n al homep age: www.elsev ier .com/ locate / jb io tec

ydrogen peroxide stress provokes a metabolic reprogramming inseudomonas fluorescens: Enhanced production of pyruvate

dam Bignucolo, Varun P. Appanna, Sean C. Thomas, Christopher Auger,ungwon Han, Abdelwahab Omri, Vasu D. Appanna ∗

epartment of Chemistry and Biochemistry, Laurentian University, Sudbury, Ontario P3E 2C6, Canada

r t i c l e i n f o

rticle history:eceived 10 April 2013eceived in revised form 28 June 2013ccepted 2 July 2013vailable online 16 July 2013

a b s t r a c t

Pseudomonas fluorescens invoked a metabolic reconfiguration that resulted in enhanced production ofpyruvate under the challenge of hydrogen peroxide (H2O2). Although this stress led to a sharp reductionin the activities of numerous tricarboxylic acid (TCA) cycle enzymes, there was a marked increase in theactivities of catalase and various NADPH-generating enzymes to counter the oxidative burden. The upre-gulation of phosphoenolpyruvate synthase (PEPS) and pyruvate kinase (PK) coupled with the reduction

eywords:yruvate productionetabolic reconfiguration

hosphoenolpyruvate synthaseADPH-generating enzymesioconversion

of pyruvate dehydrogenase (PDH) in the H2O2-challenged cells appear to be important contributors tothe elevated levels of pyruvate found in these bacteria. Increased pyruvate synthesis was evident in thepresence of a variety of carbon sources including d-glucose. Intact cells rapidly consumed d-glucose withthe concomitant formation of this monocarboxylic acid. At least a 12-fold increase in pyruvate produc-tion within 1 h was observed in the stressed cells. These findings may be exploited in the developmentof technologies aimed at the conversion of carbohydrates into pyruvate.

. Introduction

Microorganisms are crucial effectors of numerous biotechnolo-ical processes that are commercially utilized to produce a varietyf value-added products. These include the production of ethanolhrough the use of anaerobic respiration in yeast, and the gen-ration of lactic acid in dairy products, a process mediated byhe bacterium Lactobacillus plantarum (Fu and Mathews, 1999).omplex biomolecules such as surfactants and proteases maylso be manufactured by microbial systems. Indeed, various Pseu-omonas species are known to produce rhamnolipids, while Bacilluspecies commercially produce proteases which are widely uti-ized as detergents (Hadj-Ali et al., 2007). In an effort to renderhese biotechnological processes effective, the microbes often need

o undergo genetic modifications that are tailored for these spe-ific duties. These manipulations may include the alteration of aicrobe’s genetic makeup by deleting genes, introducing new ones

Abbreviations: 6PGDH, 6-phosphogluconate dehydrogenase; BN-PAGE, blueative polyacrylamide gel electrophoresis; BSA, bovine serum albumin; CFE, cell freextract; ETC, electron transport chain; FUM, fumarase; GDH, glutamate dehydroge-ase; G6PDH, glucose-6-phosphate dehydrogenase; HPLC, high performance liquidhromatography; ICDH, isocitrate dehydrogenase; KGDH, alpha-ketoglutarateehydrogenase; PDH, pyruvate dehydrogenase; PEPS, phosphoenolpyruvate syn-hase; PK, pyruvate kinase; TCA, tricarboxylic acid.∗ Corresponding author. Tel.: +1 705 675 1151x2112; fax: +1 705 675 4844.

E-mail address: vappanna@laurentian.ca (V.D. Appanna).

168-1656/$ – see front matter © 2013 Elsevier B.V. All rights reserved.ttp://dx.doi.org/10.1016/j.jbiotec.2013.07.002

© 2013 Elsevier B.V. All rights reserved.

from other organisms, or over-expressing the desired genetic infor-mation (Daniell et al., 2002; Masukawa et al., 2012). Recombinanttechnology is often utilized to create bacteria with unique commer-cial properties. The production of human insulin by Escherichia coli,where the gene for this protein is over-expressed, is an example ofsuch a technique (Goeddel et al., 1979; Min et al., 2011).

The enhancement of a given trait by natural means is also acommon method utilized to tailor microbes to perform certain des-ignated tasks. Modulation of environmental factors can also helpstimulate bacteria to produce specific by-products. These effectorscan either be physical or chemical in nature (Akbari et al., 2012;Huang et al., 2010). A common example of this is phosphate star-vation, which is known to trigger the synthesis of phosphatase(Del Pozo et al., 1999). Adaptation to metal-polluted environmentstends to favor microbial consortia that are very effective in biore-mediation processes. Indeed numerous microbial systems isolatedfrom extreme surroundings are utilized in various industrial pro-cesses. The goal of these modifications is to help facilitate theproduction and secretion of a specific product (Aislabie et al., 2006;FitzPatrick et al., 2010).

In this study, we have utilized oxidative stress in the form ofhydrogen peroxide (H2O2) on the nutrient versatile soil microbePseudomonas fluorescens in an effort to remodel its metabolic

networks, with the aim of generating pyruvate from simple car-bohydrates. Pyruvate is an important ingredient in the synthesisof food additives, pharmaceuticals, nutraceuticals, weight controlsupplements, and amino acids such as tyrosine and tryptophan

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10 A. Bignucolo et al. / Journal of B

Esposito et al., 2007; Kalman et al., 1999; Panke and Wubbolts,005). The chemical production of pyruvate via reactions with tar-arate is not environmentally friendly and necessitates the infusionf excessive amounts of energy (Causey et al., 2004). Currentlyhe fermentation process is mediated primarily by microbes thatequire well-defined media with complex supplements, all factorsssociated with high cost (Hua et al., 2001). Here, we have treated P.uorescens with H2O2 to coerce the microbe into the production ofyruvate. This oxidative stress effectively impedes the tricarboxyliccid cycle (TCA) and oxidative phosphorylation, events conduciveo the accumulation of this monocarboxylic acid. The metaboliceconfiguration mediated by phosphenolpyruvate synthase (PEPS)nd pyruvate kinase (PK) favors the formation of pyruvate. Theignificance of this finding in the manufacture of environmentallyriendly pyruvate is discussed.

. Materials and methods

.1. Cell culturing and fractionation

A bacterial strain of P. fluorescens 13525 obtained from Amer-can Type Culture Collection (ATCC) was grown in a phosphate

ineral medium containing Na2HPO4 (6 g), KH2PO4 (3 g), MgSO40.2 g), NH4Cl (0.8 g) and glucose (1.35 g) solubilized in 1 L of dis-illed and deionized H2O (ddH2O). The pH of the culture mediumas adjusted to 6.8 with 2 M NaOH and trace elements were added

o a final concentration of 1% (v/v) (Han et al., 2012). The stressededium contained 100 or 500 �M H2O2.A 200 mL aliquot of medium was autoclaved and subsequently

noculated with 1 mL of stationary phase cells grown in mediumot containing H2O2 (control medium). The cultures were aer-ted on a gyrotary water bath shaker (Model 76; New Brunswickcientific) at 26 ◦C and 140 rpm. At the same growth phase (20 hor the control and 24 h for the stressed culture), the cells werearvested by centrifugation at 10,000 × g and 4 ◦C for 15 min. Theellets were washed with 0.85% (w/v) NaCl and resuspended in celltorage buffer (CSB; 50 mM Tris–HCl pH 7.3, 5 mM MgCl2, 1 mMhenylmethylsulfonylfluoride, and 1 mM dithiothreitol [pH 7.0]).o obtain the soluble and membrane cell-free extracts, the pelletas disrupted by sonication on ice using a Brunswick sonicator onower level 4 for 15 s 4 times and with 5 min pauses. Centrifugationt 180,000 × g resulted in a supernatant that was the soluble cell-ree extract (sCFE) and the pellet that was the membrane cell-freeraction. The purity of these fractions was ascertained enzymati-ally via the analysis of glucose-6-phosphate dehydrogenase for theoluble component and complex IV for the membrane component.rotein levels were determined by the Bradford assay with bovineerum albumin (BSA) used as a standard (Swamy et al., 2006).

.2. Enzymatic studies

BN-PAGE was executed as per the protocol described (Augert al., 2012; Schagger and von Jagow, 1991). For these assays, a–16% gradient gel was prepared and the protein (4 �g/�L) wasrepared in blue native buffer (400 mM 6-amino hexanoic acid,0 mM Bis–Tris [pH 7.0]). To solubilize membrane bound proteins

n order to ensure optimal protein separation, a final concen-ration of 1% dodecyl-�-maltoside was added to the membraneractions. Protein samples were loaded into each well of the nativeel (10–60 �g) and electrophoresed at 4 ◦C under native conditionst 80 V and 15 mA for proper stacking followed by 150 V and 25 mA

n the resolving gel for the migration of the protein until it trav-led half-way through the gel. At the halfway point, blue cathodeuffer (50 mM Tricine, 15 mM Bis–Tris, 0.02% (w/v) Coomassie G-50, [pH 7.0]) was changed to a colorless cathode buffer (50 mM

nology 167 (2013) 309– 315

Tricine, 15 mM Bis–Tris, [pH 7.0]) to provide improved detectionof the protein bands. After which, electrophoresis was performedat 300 V and 25 mA. For 15 min following the electrophoresis, thegel was incubated in reaction buffer (25 mM Tris–HCl, 5 mM MgCl2[pH 7.4]), after which, the in-gel activity assay was performed byusing a reaction mixture containing equilibrium buffer, substratesand cofactors as described (Table 1). In-gel activity was ascertainedusing iodonitrotetrazolium chloride (INT) which precipitates asformazan when reduced, coupled to phenazine methosulfate (PMS)or dichloroindophenol (DCIP) as described (Han et al., 2013).

Destaining solution (40% methanol and 10% glacial acetic acid)was used to stop the reactions. Coomassie staining was used toensure equal protein loading. Enzyme specificity was confirmedby performing the in-gel reaction in the absence of a substrateor addition of specific inhibitors. To further ascertain the natureof enzymes, activity bands were excised from the gel and incu-bated with their respective substrates. The appearance of productswas then monitored by high performance liquid chromatography(HPLC).

2.3. Metabolite analysis

To evaluate the influence of H2O2 stress on metabolicnetworks, various metabolites were analyzed by HPLC. An AllianceHPLC with C18 reverse-phase column (Synergi Hydro-RP; 4 �m;250 mm × 4.6 mm, Phenomenex) and Waters dual absorbancedetector were utilized. Activity bands were excised from the geland placed in a reaction mixture containing 2 mM substrates.After 30 min of incubation, 100 �L of the sample was collectedand diluted with Milli-Q water for HPLC analysis. Mobile phasecontaining 20 mM KH2PO4 (pH 2.9) was used at a flow rate of0.7 mL/min at ambient temperature to separate the substratesand products, which were measured at 210 nm to detect car-bonyl groups. Metabolites were identified by spiking biologicalsamples using known standards, and peaks were quantified usingthe Empower software (Waters Corporation). HPLC analyses wereperformed immediately after the reactions in order to minimizesubstrate and product degradation.

2.4. Glucose consumption

A modified phenol–sulfuric acid assay was performed to gaugeglucose consumption (Masuko et al., 2005). One hundred micro-liters of spent medium was added to a glass test tube alongwith 500 �L of 4% phenol followed by 2.5 mL of 96% sulfuric acid.Absorbance was measured at 490 nm and concentrations were cal-culated from a standard curve.

2.5. Catalase activity and oxidized protein/lipid assays

The thiobarbituric acid reactive species assay (TBARS) was per-formed to evaluate the amount of oxidized lipids (Aydin et al.,2005). Determination of protein carbonyl content was achievedwith the dinitrophenyl hydrazine assay (Frank et al., 2000). Catalaseactivity was assessed using the method described in Beriault et al.(2007). Briefly, 2 mg/mL equivalent of soluble CFE was incubatedin a reaction buffer (25 mM Tris, 5 mM MgCl2, pH 7.0) containing10 mM p-anisidine. The reaction was stopped after 30 min and theabsorbance was recorded at 458 nm.

2.6. Statistical analysis

All experiments were performed in triplicate and at leastin biological duplicate. Data were expressed as mean ± standard

A. Bignucolo et al. / Journal of Biotechnology 167 (2013) 309– 315 311

Table 1Reaction mixtures for the in-gel detection of enzymes described within.

Enzyme Reaction mixture

ICDH-NAD(P) 5 mM isocitrate, 0.5 mM NAD(P), 0.2 mg/mL PMS and 0.4 mg/mL INTKGDH 0.3 mM KG, 0.5 mM CoA, 0.5 mM NAD, 0.2 mg/mL PMS and 0.4 mg/mL INTFUM 5 mM fumarate, 13 U/mL malate dehydrogenase, 0.5 mM NAD, 0.2 mg/mL PMS and 0.4 mg/mL INTPDH 5 mM pyruvate, 0. 1 mM CoA, 0.5 mM NAD, 0.2 mg/mL PMS and 0.4 mg/mL INTComplex I 5 mM NADH, 5 mM KCN and 0.4 mg/mL INTComplex IV 10 mg/mL diaminobenzidine, 10 mg/mL cyctochrome C and 562.5 mg/mL sucroseGDH 5 mM glutamate, 0.5 mM NADP, 0.2 mg/mL PMS and 0.4 mg/mL INTG6PDH 5 mM glucose-6-phosphate, 0.5 mM NADP, 0.2 mg/mL PMS and 0.4 mg/mL INT6PGDH 5 mM 6-phosphogluconate, 0.5 mM NADP, 0.2 mg/mL PMS and 0.4 mg/mL INTPK 5 mM PEP, 0.5 mM NADH, 10 U lactate dehydrogenase (LDH), 16.7 �g/mL DCIP, 0.5 mg/mL INT and 0.5 mM ADP

7 �g/mL DCIP, 0.5 mg/mL INT, 0.5 mM AMP and 1.0 mM Pi

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Table 2Oxidized lipids, proteins and catalase activity analyses (n = 3 ± SD).

Control 100 �M H2O2

Oxidized lipids (�mol ofmalondialdehyde/mg of biomass)

0.76 ± 0.01 2.3 ± 0.2

Oxidized protein (nmol of carbonyl/mg 0.021 ± 0.003 0.069 ± 0.003

F((

PEPS 5 mM PEP, 0.5 mM NADH, 10 U LDH, 16.

eviation (SD). Statistical correlations of data were checked forignificance using the Student’s t-test.

. Results and discussion

P. fluorescens was found to readily adapt to 100 �M H2O2 withinimal impact on cellular yield (Fig. 1A). d-Glucose, the sole car-

on source, was consumed at a slower rate than in the controlultures (Fig. 1B). However, no significant growth was detected

hen the bacterium was grown in a media with 500 �M H2O2

Fig. 1A). The effects of the oxidative environment were evident ashe H2O2-stressed bacterium was characterized with higher lev-ls of oxidized lipids and proteins, as well as an increase in the

ig. 1. Cellular yield and glucose consumption in P. fluorescens. (A) Biomass was determin�) 500 �M H2O2-treated cultures. (B) Glucose consumption was monitored via the phenon = 3).

of protein)Catalase activity (�mol of H2O2

decomposed mg min−1)0.13 ± 0.04 0.82 ± 0.02

activity of catalase, an important H2O2-detoxifying enzyme(Table 2). The metabolic profiles in the control and stressed cul-tures were significantly different, indicating that H2O2 was havinga marked influence on the metabolic networks. The spent fluid from

ed via the Bradford assay. (�) Control cultures, (�) 100 �M H2O2-treated cultures,l–sulfuric acid method. (�) Control cultures and (�) 100 �M H2O2-treated cultures

312 A. Bignucolo et al. / Journal of Biotechnology 167 (2013) 309– 315

Fig. 2. HPLC analysis of extracellular and intracellular metabolites in H2O2-stressed P. fluorescens. (A) Representative chromatographs (n = 3) displaying metabolite levels inthe spent fluid of control and stressed P. fluorescens. (B) Metabolite levels in the soluble cell free extract as measured by HPLC. n = 3 ± SD. (�) Control cultures, (�) 100 �MH2O2-treated cultures, ( ) cultures treated with 100 �M H2O2 twice (the second dosage was given after 15 h of growth in an effort to confirm the role of H2O2 in thisprocess).

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he H2O2-containing medium had peaks attributable to pyruvatend acetate. There was approximately a 10-fold increase in pyru-ate compared to the control spent fluid (Fig. 2A). The soluble cellree extract (sCFE) had characteristic peaks of pyruvate, acetate anddenosine monophosphate (AMP) that were markedly prominentn the stressed bacteria (Fig. 2B).

As expected, H2O2 was having a very pronounced impact on thericarboxylic acid (TCA) cycle and electron transport chain (ETC).he activities of enzymes such as ICDH-NAD, KGDH, PDH, and FUMere severely perturbed (Fig. 3A). The ability to generate energy in

n O2-dependent manner was also attenuated since the activitiesf Complexes I and IV were barely discernible in the H2O2-stressedells (Fig. 3B). This metabolic disruption in the NADH-producingystem was accompanied by an increase in NADPH formation viahe enhanced activities of G6PDH, 6PGDH, ICDH-NADP, and GDHFig. 3C). These enzymes were observed to have markedly increased

ctivity bands in the CFE from the stressed cells, compared to thosen the control.

Thus, in order to survive this major assault on its aerobicifestyle, the microbe was compelled to exploit two enzymes

involved in pyruvate synthesis with the concomitant productionof ATP. There was an upregulation in the activity of PEPS, which isknown to interact with PEP in the presence of AMP and Pi, in orderto produce ATP and pyruvate (Fig. 4A). There was an approximately10-fold increase in activity when compared to the control. Theenhancement of PEPS activity was dependent on the presence ofH2O2 in the media, as the incubation of H2O2-stressed cells in con-trol medium led to a sharp reduction in the activity band attributedto PEPS (Fig. 4B). This finding indicates a pivotal role played byH2O2 in promoting this metabolic event. The overexpression ofPEPS activity appeared to be responsible for the elevated levels ofpyruvate in the spent fluid.

This fortuitous observation led us to evaluate if indeed pyru-vate, a value-added product, might be produced by the intact cells.Indeed when H2O2-stressed cells were incubated in a non-dividingmedium, glucose was rapidly consumed with the formation of

pyruvate in the spent fluid. In 1 h, there was a 12-fold increasein pyruvate levels (Fig. 4C). Increased pyruvate synthesis wasalso observed when d-xylose, d-glycerol, d-fructose, and d-sucrosewere the sources of carbon (personal observation).

A. Bignucolo et al. / Journal of Biotechnology 167 (2013) 309– 315 313

Fig. 3. In-gel enzyme activity assays. (A) Representative gels (n = 3) displaying the down-regulation of TCA cycle enzymes. ICDH, isocitrate dehydrogenase; �-KGDH, alpha-ketoglutarate dehydrogenase; FUM, fumarase; PDH, pyruvate dehydrogenase. (B) Representative gels (n = 3) displaying the down-regulation of the ETC. Negative controlswere performed without NADH. (C) Representative gels (n = 3) displaying the up-regulation of NADPH-producing enzymes. Densitometry was performed using ImageJ forWindows. GDH, glutamate dehydrogenase; G6PDH, glucose-6-phosphate dehydrogenase; 6PGDH, 6-phosphogluconate dehydrogenase.

Fig. 4. Pyruvate production in H2O2-stressed P. fluorescens. (A) In-gel activity of enzymes mediating the formation of pyruvate. Densitometry was performed using ImageJfor Windows. PK, pyruvate kinase; PEPS, phosphoenolpyruvate synthase. (B) Regulation of PEPS. After growing to the stationary phase, control cells were incubated inH2O2-treated media for 8 h and vice versa. (C) HPLC analysis showing pyruvate production in non-dividing control (�) and 100 �M H2O2-stressed ( ) cells after 1 h. n = 3 ± SD.

314 A. Bignucolo et al. / Journal of Biotechnology 167 (2013) 309– 315

Fig. 5. Metabolic reconfiguration aimed at the production of pyruvate in H2O2-stressed P. fluorescens. ETC, electron transport chain; PEPS, phosphoenolpyruvate synthase;P

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. Conclusion

Pyruvate, a central metabolite, is utilized in a variety of prod-cts including food-additives, nutraceuticals, and pharmaceuticalsEsposito et al., 2007; Kalman et al., 1999; Panke and Wubbolts,005). It is critical to find effective means of generating this ketocid in an environmentally friendly manner (Hua et al., 2001).enetic modification is a common strategy used to evoke pyru-ate production in microbes on a commercial scale (Hua et al.,001). However genetically modified organisms have inherent risksnd are required to undergo strict regulatory verification (Schmidtt al., 2009). The H2O2 stress that led to enhanced pyruvate syn-hesis in P. fluorescens may provide a facile strategy to acclimatize

icrobial systems for this task. It appears that the metabolic repro-ramming provoked by the oxidant forces the bacterium to fulfill itsTP needs via substrate-level phosphorylation with the concomi-

ant secretion of pyruvate (Fig. 5). This by-product of this metabolichift is indeed a value-added commodity with multiple applica-ions. Hence, the H2O2-instigated metabolic remodeling may be anffective and green tool in the production of this keto acid.

cknowledgements

This work was supported by Laurentian University and Industryanada. Christopher Auger is a recipient of the NSERC PGS-D.

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